Department of Pharmacology, University of Minnesota Medical School, Minneapolis, Minnesota, USA
Address correspondence and reprint requests to Stanley A. Thayer, Department of Pharmacology, University of Minnesota, 6-120 Jackson Hall, 321 Church Street SE, Minneapolis, MN 55455, USA. E-mail: email@example.com
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HIV-associated neurocognitive disorders afflict about half of HIV-infected patients. HIV-infected cells shed viral proteins, such as the transactivator of transcription (Tat), which can cause neurotoxicity by over activation of NMDA receptors. Here, we show that Tat causes a time-dependent, biphasic change in NMDA-evoked increases in intracellular Ca2+ concentration ([Ca2+]i). NMDA-evoked responses were potentiated following 2-h exposure to Tat (50 ng/mL). Tat-induced potentiation of NMDA-evoked increases in [Ca2+]i peaked by 8 h and then adapted by gradually reversing to baseline by 24 h and eventually dropping below control by 48 h. Tat-induced potentiation of NMDA-evoked responses was blocked by inhibition of lipoprotein receptor-related protein (LRP) or Src tyrosine kinase. Potentiation was unaffected by inhibition of nitric oxide synthase (NOS). However, NOS activity was required for adaptation. Adaptation was also prevented by inhibition of soluble guanylate cyclase (sGC) and cyclic guanosine monophosphate-dependent protein kinase G (PKG). Together, these findings indicate that Tat potentiates NMDA-evoked increases in [Ca2+]i via LRP-dependent activation of Src and that this potentiation adapts via activation of the NOS/sGC/PKG pathway. Adaptation may protect neurons from excessive Ca2+ influx and could reveal targets for the treatment of HIV-associated neurocognitive disorders.
HIV-associated neurocognitive disorders (HAND) afflict about half of HIV-infected patients. HIV-infected cells shed viral proteins, such as the transactivator of transcription (Tat), which can cause neurotoxicity by over activation of NMDA receptors (NMDARs). We show that HIV-1 Tat evoked biphasic changes in NMDA-evoked [Ca2+]i responses. Initially, Tat potentiated NMDA-evoked responses following LRP-mediated activation of Src kinase. Subsequently, Tat-induced NMDAR potentiation adapted by activation of a NOS/sGC/PKG pathway that attenuated NMDA-evoked increases in [Ca2+]i. Adaptation may be a novel neuroprotective mechanism to prevent excessive Ca2+ influx. Solid and dashed arrows represent direct and potentially indirect connections, respectively.
Cognitive function is impaired in 30–55% of patients infected with human immunodeficiency virus (HIV) (Cysique et al. 2004; Tozzi et al. 2005; Heaton et al. 2011). HIV-associated neurocognitive disorders (HAND) range in severity from a subtle reduction in information processing speed to significant functional impairment (Heaton et al. 2004; Antinori et al. 2007). Despite effectively managing viral load with combined anti-retroviral therapy (cART), the prevalence of HAND remains persistently high (Heaton et al. 2010) and may be increasing because of prolonged patient life spans. Currently, the efficacy of drugs to treat HAND is insufficient and the options are few.
In the brain, HIV infects macrophages and microglia, but not neurons (Watkins et al. 1990). Thus, HIV-induced neurotoxicity is indirect and results from the release of neurotoxic agents such as inflammatory cytokines, nitric oxide (NO), glutamate, and viral proteins (Genis et al. 1992; Jiang et al. 2001; Kaul et al. 2001; Nath 2002; Eugenin et al. 2007). The transactivator of transcription (Tat) is a protein shed from HIV-infected cells and detected in the sera and CNS of HIV-infected patients (Chang et al. 1997; Hudson et al. 2000). The level of anti-Tat antibodies in the CSF of HIV-infected patients without cognitive dysfunction is higher than in patients with HAND, suggesting that antibody responses against Tat may be neuroprotective (Bachani et al. 2013). Despite significant improvement in the efficacy of cART for treating HIV infection, current regimens remain unable to halt the production of Tat (Li et al. 2009).
Cognitive decline in patients with HAND correlates with synaptodendritic damage (Ellis et al. 2007). Expression of Tat in transgenic rodent models causes loss of excitatory synapses resulting in learning and memory impairment (Carey et al. 2012; Fitting et al. 2012). In vitro, Tat-induced synapse loss (Kim et al. 2008) and neuronal death (Eugenin et al. 2007) are initiated by NMDA receptor (NMDAR)-mediated Ca2+ influx. Tat potentiates NMDA-evoked increases in intracellular Ca2+ concentration ([Ca2+]i) in hippocampal neurons (Haughey et al. 2001). Most studies of Tat-induced changes in NMDAR function are acute (min to h) while the neurotoxic effects of Tat occur over a prolonged time scale (h to days). Treating primary hippocampal neurons with Tat for 24 h causes loss of excitatory synapses (Kim et al. 2008) and simultaneous gain of inhibitory synapses (Hargus and Thayer 2013) indicating that Tat evokes adaptive changes in the synaptic composition of neurons. Such neuroadaptations may be a mechanism to cope with excess excitatory input. Notably, these adaptive changes are prevented by pharmacologic inhibition of the NMDAR (Shin et al. 2012) indicating that the NMDAR is essential for synaptic neuroadaptation. How NMDA-evoked [Ca2+]i responses are affected by prolonged exposure to Tat, when adaptive changes in synaptic composition occur, is the focus of this study.
Here, we examined changes in NMDA-evoked increases in [Ca2+]i during 48-h exposure to Tat. We found that Tat evoked a biphasic change in NMDA-evoked [Ca2+]i responses. Tat initially potentiated the NMDA-evoked increase in [Ca2+]i via the low-density lipoprotein receptor-related protein (LRP) and activation of Src kinase. Tat-induced potentiation subsequently adapted by gradually returning to baseline levels. Adaptation resulted from activation of the nitric oxide synthase (NOS)/soluble guanylate cyclase (sGC)/protein kinase G (PKG) signaling pathway. This study suggests a changing role for the NMDAR during the course of HIV neurotoxicity with implications for treatment of HAND.
Drugs and reagents
Materials were obtained from the following sources: HIV-1 Tat (Clade B, full-length recombinant) was from the NIH AIDS Research and Reference Reagent Program. HIV-1 Tat protein (full length, Clade B) from John Brady and DAIDS, NIAID and from Prospec Tany TechnoGene Ltd. (Rehovot, Israel); recombinant rat low-density lipoprotein receptor-related protein-associated protein 1 (RAP) was from Fitzgerald Industries International (Concord, MA, USA); Dulbecco's modified Eagle's medium (DMEM), fetal bovine serum, horse serum, fura-2-acetomethyl ester (Fura-2-AM), and glycine were from Invitrogen (Carlsbad, CA, USA); NMDA, NG-nitro-l-arginine methyl ester (L-NAME), and 1H-(1,2,4)oxadiazolo[4,3-a]quinoxalin-1-one (ODQ) were from Sigma-Aldrich (St. Louis, MO, USA); 4-amino-5-(4-chloro- phenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine (PP2) and 4-amino-7-phenylpyrazol[3,4-d]pyrimidine (PP3), 8R,9S,11S)-(-)-9-methoxy-carbamyl-8-methyl-2,3,9,10-tetrahydro-8,11-epoxy-1H,8H,11H-2,7b,11a-trizadibenzo-(a,g)-cycloocta-(c,d,e)-trinden-1-one (KT5823), and (1R*,2S*)-erythro-2-(4-Benzylpiperidino)-1-(4-hydroxyphenyl)-1-propanol hemitartrate (ifenprodil) were from Tocris (Bristol, UK); 2-(2-amino-3-methyoxyphenyl)-4 H-1-benzopyran-4-one (PD98059) was from Cell Signaling Technology (Danvers, MA, USA); recombinant rat interleukin-1β was from R & D Systems (Minneapolis, MN, USA).
Dominant-negative (DN)-Fyn with the K299M mutation and DN-Src with the K295M mutation in pRK5 were kindly provided by Dr Filippo Giancotti (Memorial Sloan Kettering Cancer Center, New York, NY, USA); pTagRFP-N was from Evrogen (Moscow, Russia). The catalytically inactive mutant of full-length PKG1α containing the amino terminal regulatory region fused to green fluorescent protein (GFP) (G1αR-GFP) in pEGFP-N1 (Clontech) was kindly provided by Darren Browning (Georgia Regents University). The G1αR fragment was removed via digestion with EcoR1 and BamH1 (New England Biolabs, Ipswich, MA, USA), and then inserted into the multiple cloning site of DsRed2-N1 (Clontech). Mammalian expression vectors containing genes for the NMDAR subunits GluN1, GluN2A, and GluN2B were kindly provided by Stephen Traynelis (Emory University).
In accordance with the University of Minnesota's Institutional Animal Care and Use Committee and the NIH guide for the care and use of laboratory animals, maternal rats were killed by CO2 inhalation and fetuses were removed on embryonic day 17. Rat hippocampal neurons were grown in primary culture as described previously (Li et al. 2012). Hippocampi were dissected and placed in Ca2+- and Mg2+-free HEPES-buffered Hanks' salt solution (HHSS), pH 7.45. HHSS contained the following (in mM): HEPES 20, NaCl 137, CaCl2 1.3, MgSO4 0.4, MgCl2 0.5, KCl 5.0, KH2PO4 0.4, Na2HPO4 0.6, NaHCO3 3.0, and glucose 5.6. Cells were dissociated by triturating through a 5-mL pipette and a flame-narrowed Pasteur pipette and then resuspended in DMEM without glutamine, supplemented with 10% fetal bovine serum and penicillin/streptomycin (100 U/mL and 100 mg/mL, respectively). Dissociated cells were then plated at a density of 60 000 to 80 000 cells/dish onto a 25-mm-round cover glass (#1) pre-coated with matrigel (150 μL, 0.2 mg/mL). Neurons were grown in a humidified atmosphere of 10% CO2 at 37°C and fed on days 1 and 7 by exchange of 75% of the media with DMEM supplemented with 10% horse serum and penicillin/streptomycin. Cells used in these experiments were cultured without mitotic inhibitors resulting in a mixed glial-neuronal culture consisting of 18 ± 2% neurons, 70 ± 3% astrocytes, and 9 ± 3% microglia as indicated by immunocytochemistry (Kim et al. 2011). The cultures used for experimentation were grown for 12–15 days in vitro.
Intracellular Ca2+ concentration ([Ca2+]i) was recorded as described previously (Li et al. 2013) with minor modifications. Cells were loaded by incubation with 5 μM fura-2 AM in 0.04% pluronic acid in HHSS for 30 min at 37°C followed by washing in the absence of indicator for 10 min. HIV-1 Tat and respective drugs were present during fura-2-AM loading, but were absent during the wash. Coverslips containing fura-2 loaded cells were transferred to a recording chamber, placed on the stage of an IX71 microscope (Olympus, Melville, NY, USA), and viewed through a 20X objective. Excitation wavelength was selected with a galvanometer-driven monochromator (8-nm slit width) coupled to a 75-W xenon arc lamp (Optoscan; Cairn Research, Faversham, Kent, UK). [Ca2+]i was monitored using sequential excitation of fura-2 at 340 and 380 nm; image pairs were collected every 1 s. For experimental recordings, cells were superfused at a rate of 1–2 mL/min with HHSS for 1 min followed by 30 s perfusion of Mg2+-free HHSS that contained 200 μM glycine and either 10 or 100 μM NMDA. Fluorescence images (510/40 nm) were projected onto a cooled charge-coupled device camera (Cascade 512B; Roper Scientific, Tucson, AZ, USA) controlled by MetaFluor software (Molecular Devices, Palo Alto, CA, USA). After background subtraction, the 340- and 380-nm image pairs were converted to [Ca2+]i using the formula [Ca2+]i = Kdβ(R−Rmin)/(Rmax−R) (Grynkiewicz et al. 1985). The dissociation constant (Kd) for fura-2 was 145 nM. β is the ratio of fluorescence intensity acquired with 380-nm excitation measured in Ca2+-free buffer (1 mM EGTA) and buffer containing saturating Ca2+ (5 mM). R is 340 nm/380 nm fluorescence intensity ratio. Rmin, Rmax, and β were determined in a series of calibration experiments on intact cells. Rmin and Rmax values were generated by applying 10 μM ionomycin in Ca2+-free buffer (1 mM EGTA) and saturating Ca2+ (5 mM), respectively. Values for Rmin, Rmax, and β were 0.37, 9.38, and 6.46, respectively. These calibration constants were applied to all experimental recordings. To generate pseudocolor images, a binary mask was generated by applying an intensity threshold to the 380 nm image and applied to [Ca2+]i images with colors assigned as indicated by the calibration bars in the figures. The neuronal cell body was selected as the region of interest for all recordings. All neurons within the imaging field were included in the analysis and no exclusions were made. For time course experiments, coverslips from the same cell culture plating were treated in parallel and each coverslip imaged only once.
Rat hippocampal neurons were transfected between 11 and 12 days in vitro using a modification of a calcium phosphate protocol as described previously (Li et al. 2012). Briefly, hippocampal cultures were incubated for 30 min in DMEM supplemented with 1 mM kynurenic acid, 10 mM MgCl2, and 5 mM HEPES. A DNA/calcium phosphate precipitate containing 1 μg plasmid DNA per well was prepared, allowed to form for 30 min at 21°C then added to the culture. After 90 min of incubation, cells were washed once with DMEM supplemented with MgCl2 and HEPES and then returned to conditioned media.
Biotinylation and immunoblotting
For biotinylation experiments, 7.5 × 106 hippocampal cells were plated in 100 × 20 mm petri dishes. Cells were treated with Tat alone or with an inhibitor of protein kinase G (KT5823) 1 h prior to the addition of Tat. Forty-eight hours later the media were removed and cells were washed three times with ice-cold phosphate-buffered saline (PBS) (pH 8.0) then incubated with 2 mM NHS-PEG4 Biotin (Thermo Fisher Scientific Inc., Waltham, MA, USA) in ice-cold PBS with slow shaking at 4°C for 30 min protected from light. Cells processed in parallel but not treated with biotin served as negative controls. Biotin was gently removed and cells were washed four times with ice-cold PBS. Cells were collected in PBS and centrifuged at 10 000 g for 1 min. Following centrifugation, supernatants were aspirated and pellets were resuspended in 250 μL lysis buffer consisting of PBS, 10% Triton, 20% sodium dodecyl sulfate, and a mixture of protease inhibitors (10 μL phenylmethylsulfonyl fluoride, 100 μL Halt protease inhibitor cocktail). Samples were sonicated with 3 × 1 s pulses and incubated with rocking for 30 min at 4°C. Debris was pelleted in a microcentrifuge at 14 000 g at 4°C for 15 min and supernatants retained. Sample of 30 μL of (‘Total protein’) was removed, and the rest of the sample was mixed by rotation for 30 min at 4°C with 60 μL of 50% slurry of NeutrAvidin beads (Thermo Fisher Scientific Inc.). Beads were washed three times with lysis buffer and bound proteins eluted in 75 μL of 2X sodium dodecyl sulfate sample buffer plus 25 μL 1 M dithiothreitol by heating at 75°C for 30 min. For immunoblotting, 3 μL 1M dithiothreitol was added to 15 μL of sample, and the mixture was incubated at 75°C for 10 min. Samples were then loaded onto 8% Bis-Tris gels and run in a Tris–glycine buffer under reducing conditions. Samples were transferred using the iBlot transfer system (Invitrogen) onto nitrocellulose membranes and probed with a rabbit anti-GluN2B polyclonal antibody (1 : 200 dilution, Millipore Corporation, Bedford, MA, USA). An IRDye 800CW donkey anti-rabbit secondary antibody (LiCor Biosciences, Lincoln, NE, USA) was used at a dilution of 1 : 1000. Visualization and quantification of band intensity were performed using the Odyssey Imaging System (LiCor Biosciences). Integrated intensity values were recorded for all lanes and normalized to the intensity of the untreated control.
For [Ca2+]i imaging studies, an individual experiment (n = 1) was defined as the response from a single neuron on a single coverslip. Changes in NMDA-evoked increases in [Ca2+]i are presented as mean ± SEM. Each experiment was replicated using at least four separate coverslips from at least two separate cultures. For biotinylation experiments, an individual experiment (n = 1) was defined as the change in surface-to-total GluN2B from a single 100 × 20 mm petri dish. Changes in GluN2B surface expression are presented mean ± SEM. Each experiment was replicated using at least 10 separate petri dishes from 10 separate cultures. Significance was determined by one-way anova with Tukey's post hoc test for multiple comparisons (OriginPro v8.5, OriginLab Corp., Northampton, MA, USA).
Tat potentiates NMDA-evoked [Ca2+]i responses via LRP and a Src family kinase
Treating primary hippocampal neurons with Tat potentiated NMDA-evoked increases in [Ca2+]i (Haughey et al. 2001). We replicated this finding using fura-2-based digital imaging of rat hippocampal neurons in vitro. As shown in Fig. 1, increases in [Ca2+]i evoked by 100 μM NMDA were potentiated following 40-min treatment with 100 nM Tat, but not by 100 nM heat-inactivated (HI, 85°C × 30 min) Tat.
Tat is internalized via the LRP (Liu et al. 2000) and Tat-induced neurotoxicity requires binding to the LRP with subsequent NMDAR-mediated Ca2+ influx (Eugenin et al. 2007; Kim et al. 2008). We sought to determine whether the LRP was required for Tat-induced NMDAR potentiation by using the LRP antagonist, receptor-associated protein (RAP). RAP binds to the LRP and prevents neuronal uptake of Tat (Bu 2001). Application of RAP (50 nM) 1 h prior to Tat application completely blocked Tat-induced potentiation (Fig. 1g, h), suggesting that Tat potentiates NMDAR function via the LRP.
Tyrosine phosphorylation can potentiate NMDAR currents (Wang and Salter 1994). Tat-induced NMDAR potentiation occurs via tyrosine kinase-dependent phosphorylation of GluN2A and GluN2B (Haughey et al. 2001), although the specific tyrosine kinase involved in NMDAR potentiation remains unknown. Src family kinases (SFKs) enhance NMDAR function (Kohr and Seeburg 1996), therefore we used the SFK inhibitor, PP2, to determine the role of SFKs in Tat-induced potentiation of NMDARs. Pre-treatment for 1 h with PP2 (10 μM), but not its inactive analog PP3 (10 μM), completely blocked the Tat-induced potentiation of NMDA-evoked responses (Fig. 1g, h). Thus, Tat-induced potentiation of NMDA-evoked increases in [Ca2+]i requires the activation of a SFK.
Tat induces a biphasic change in NMDA-evoked increases in [Ca2+]i
Up to 40 ng/mL of Tat has been detected in the sera of HIV-infected patients (Xiao et al. 2000). Previously, we have observed Tat-induced changes in synapse number following treatment with 50 ng/mL (3.6 nM) Tat for 24 h (Kim et al. 2008; Hargus and Thayer 2013). Thus, we hypothesized that 3.6 nM Tat would potentiate the NMDA-evoked increase in [Ca2+]i, but that these changes might require longer exposure to develop compared with 100 nM Tat. To test this hypothesis, we recorded NMDA-evoked [Ca2+]i responses following treatment with 50 ng/mL Tat within a 48-h window (Fig. 2). Control and Tat-treated cultures were imaged in parallel. Exposure to Tat for 2 h potentiated NMDA-evoked increases in [Ca2+]i. Tat-induced potentiation of the NMDA-evoked response peaked by 8 h then adapted, returning to baseline by 24 h and dropping below control by 48 h.
Tat is susceptible to degradation by the Ca2+-activated protease, calpain-1 (Passiatore et al. 2009). To examine the possibility that adaptation of NMDAR potentiation resulted from the degradation of Tat, cultures received either one (0 h) or two (0 h, 12 h) applications of Tat (50 ng/mL). Following 16-h exposure to Tat, NMDAR function was potentiated in neurons treated with Tat once or twice at 39 ± 2% and 22 ± 2% larger than control, respectively. By 24 h, amplitudes of NMDA-evoked Ca2+ influx from single and double Tat-treated groups returned to baseline levels and were comparable to control suggesting that adaptation of the NMDA-evoked response is not because of the degradation of Tat. Taken together, these data indicate that Tat evokes a biphasic change in NMDAR function. In an attempt to determine whether enhanced Ca2+ flux during the Tat-induced potentiation phase was required for subsequent adaptation, we blocked NMDAR-mediated Ca2+ influx with the readily reversible NMDAR antagonist AP5 during Tat exposure. However, chronic NMDAR antagonism in the absence of Tat enhanced NMDA-evoked responses, consistent with a previous report showing that chronic NMDAR blockade produced a compensatory increase in the surface expression of NMDARs (Crump et al. 2001). Thus, we were unable to determine whether potentiation was necessary for adaptation.
Application of 100 μM NMDA-evoked [Ca2+]i increases in Tat-treated cells that peaked near saturation of the fura-2 Ca2+ indicator. A concentration-response experiment was conducted to determine if Tat-induced potentiation was observed when lower NMDA concentrations were used to probe NMDAR function. Tat-induced potentiation of NMDA-evoked responses was independent of NMDA concentration and was 35 ± 2%, 14 ± 3%, and 32 ± 1% larger than control for 10, 30, and 100 μM NMDA, respectively. Thus, to enable more accurate measurement of the [Ca2+]i responses, we conducted the remaining experiments using 10 μM NMDA.
Tat-induced potentiation of the NMDA-evoked increase in [Ca2+]i is reversible
Tat-induced potentiation was prevented by inhibition of LRP or SFKs (Fig. 1). We next determined whether Tat-induced potentiation of NMDARs could be reversed by inhibition of LRP or SFKs after the potentiation was already established. Changes in NMDA-evoked increases in [Ca2+]i were studied after 16-, 24-, 32-, or 48-h exposure to Tat. Either RAP (50 nM) or PP2 (10 μM) was applied 1 h prior to Ca2+ imaging in the continued presence of Tat (50 ng/mL). Exposure to Tat for 16 h potentiated NMDA-evoked increases in [Ca2+]i (Fig. 3a, e). Potentiation was reversed by application of RAP or PP2 1 h prior to evoking the test response with NMDA (Fig. 3a, e). Potentiation of NMDA-evoked responses adapted by 24 h (Fig. 3b, f) and then dropped below control by 32 h (Fig. 3c, g) and 48 h (Fig. 3d, h). RAP and PP2 did not affect control cultures, but further reduced NMDA-evoked [Ca2+]i responses in cultures treated with Tat for 24–48 h (Fig. 3b–d, f–h). These data indicate that Tat-induced potentiation of NMDA-evoked increase in [Ca2+]i requires sustained activation of LRP and SFK. Furthermore, this potentiation pathway remains activated during the adaptation process as indicated by the reduced amplitude of NMDA-evoked [Ca2+]i responses in fully adapted cultures (24–48 h treated with Tat).
Src kinase mediates Tat-induced potentiation of NMDA-evoked increases in [Ca2+]i
The SFKs, Src and Fyn, have been shown to enhance NMDAR function (Kohr and Seeburg 1996). Following exposure to Tat, Src but not Fyn, associates with NMDARs (King et al. 2010). Because of the potential off-target effects of PP2 (Bain et al. 2007) and the lack of drugs that are selective for Src versus Fyn, we next used a genetic approach to determine which SFK is responsible for potentiating the NMDA-evoked increase in [Ca2+]i. Primary hippocampal cultures were cotransfected with plasmids encoding a red-fluorescent protein (pTag-RFP-N) and dominant-negative (DN)-Fyn or DN-Src (Fig. 4a, b). Twenty-four hours after transfection, cells were treated with Tat (50 ng/mL) for 16 h, loaded with fura-2, and then imaged. Non-expressing cells in the same imaging field served as controls. Tat potentiated NMDA-evoked [Ca2+]i responses in non-expressing controls and cells expressing DN-Fyn (Fig. 4c, d). However, expression of DN-Src completely blocked Tat-induced potentiation (Fig. 4c, d). These data suggest that Tat-induced activation of Src kinase potentiates NMDA-evoked [Ca2+]i responses.
Tat-induced potentiation of NMDA-evoked increases in [Ca2+]i adapts via the NOS/sGC/PKG pathway
We next wanted to better understand the mechanism of adaptation following Tat-induced NMDAR potentiation. Because adaptation of Tat-induced NMDAR potentiation occurred in the presence of sustained Src activation, we hypothesized that adaptation resulted from a sustained increase in NMDAR-mediated Ca2+ influx. NMDAR-mediated Ca2+ influx following exposure to Tat leads to sustained production of NO (Eugenin et al. 2007). To determine whether NO production was mediating the Tat-induced adaptation in NMDAR function, we used a pharmacological approach. Cultures were pre-treated with L-NAME (100 μM), an inhibitor of NOS, 1 h prior to and during exposure to Tat for 16, 24, 32, and 48 h. L-NAME did not affect Tat-induced NMDAR potentiation (Fig. 5a, e) but did prevent adaptation of the NMDA-evoked increase in [Ca2+]i for 48 h (Fig. 5b–d, f–h).
NO can activate soluble guanylate cyclase (sGC) resulting in the synthesis of cyclic guanosine monophosphate (cGMP) (Arnold et al. 1977). To determine whether Tat-induced NO production was mediating the adaptation of NMDA-evoked responses by elevating cGMP, we treated cultures with ODQ (1 μM), an inhibitor of sGC, 1 h prior to and during exposure to Tat for 16, 24, 32, and 48 h. ODQ did not affect Tat-induced NMDAR potentiation (Fig. 5a, e) but did prevent adaption of NMDA-evoked [Ca2+]i responses for 48 h (Fig. 5b–d, f–h).
Elevated cGMP activates cGMP-dependent protein kinase (Kuo and Greengard 1972). To determine whether Tat-induced NO production was mediating the adaptation of NMDA-evoked [Ca2+]i responses by activating PKG, we treated cultures with the PKG inhibitor, KT5823 (10 μM), 1 h prior to and during exposure to Tat for 16, 24, 32, and 48 h. KT5823 did not affect Tat-induced NMDAR potentiation (Fig 5a, e), but did prevent adaptation of NMDA-evoked increases in [Ca2+]i for 48 h (Fig. 5b–d, f–h).
To confirm that adaptation was mediated by PKG we complimented these pharmacological studies using a genetic approach. The catalytically inactive mutant of PKG1α, G1αR, acts in a dominant-negative manner to inhibit PKG activity (Browning et al. 2001). This dominant-negative PKG construct (DN-PKG) was expressed in neurons; cells were then treated with Tat for 16 or 24 h, loaded with Fura-2, and then imaged. Exposure to Tat for 16 h potentiated the NMDA-evoked increase in [Ca2+]i in non-expressing controls and neurons expressing DN-PKG. By 24 h, NMDA-evoked responses adapted in non-expressing cells, but expression of DN-PKG prevented adaptation (Fig. 6). Taken together, these data suggest that adaptation of NMDA-evoked responses following Tat-induced potentiation occurs via a NOS/cGMP/PKG pathway.
PKG activates extracellular signal-regulated kinase (ERK) leading to the expression of neuroplasticity-associated proteins such as c-Fos, Egr-1, Arc, and brain-derived neurotrophic factor (Gallo and Iadecola 2011). Therefore, we tested the hypothesis that Tat-induced activation of PKG stimulates ERK signaling to affect NMDA-evoked [Ca2+]i responses. The MAP kinase 1 inhibitor PD98059 (50 μM) was applied 1 h prior to and during treatment with Tat for 16 or 24 h. PD98059 inhibits ERK1/2 phosphorylation and prevents expression of the aforementioned neuroplasticity-associated proteins (Gallo and Iadecola 2011). Tat-induced NMDAR potentiation was comparable in the absence and presence of PD98059 at 49 ± 3% and 62 ± 8% larger than control, respectively. By 24 h, amplitudes of NMDA-evoked increases in [Ca2+]i from both treatment groups returned to baseline and were comparable to control, suggesting that activation of ERK signaling is not required for Tat-induced potentiation or adaptation of NMDARs.
To determine if adaptation following NMDAR potentiation is unique to Tat, we conducted a time course experiment using interleukin-1β (IL-1β). IL-1β is a pro-inflammatory cytokine that potentiates NMDAR-mediated currents (Liu et al. 2013) and NMDA-evoked [Ca2+]i increases via activation of a SFK (Viviani et al. 2003). We replicated this result and found that 5-min exposure to IL-1β (50 pg/mL) potentiated NMDA-evoked [Ca2+]i responses. IL-1β-induced NMDAR potentiation persisted for longer than 24 h then NMDA-evoked [Ca2+]i responses adapted, gradually reversing toward baseline by 48 h (Figure S1). Pre-treatment for 1 h with the soluble guanylate cyclase inhibitor ODQ had no effect on IL-1β-induced NMDAR potentiation and failed to prevent adaptation. Thus, while adaptation of potentiated NMDA-evoked Ca2+ responses may be common, it appears that the mechanism of adaptation is stimulus specific.
Tat does not affect surface expression of GluN2B
We hypothesized that the reduction in the amplitude of NMDA-evoked [Ca2+]i increases following 48-h exposure to Tat resulted from NMDAR internalization. We first determined the GluN2 subtype mediating the NMDA-evoked response using a pharmacological approach (Figure S2). We recorded an initial NMDA-evoked control response (response #1), allowed the cell to recover for 5 min in the presence of ifenprodil (10 μM), a selective GluN2B antagonist (Williams 1993), and then evoked a second response in the continued presence of ifenprodil (response #2). The majority (74 ± 1%) of the NMDA-evoked increase in [Ca2+]i was mediated by ifenprodil-sensitive GluN2B-containing NMDARs, consistent with previous reports indicating that GluN2B preferentially localizes to extrasynaptic sites (Tovar and Westbrook 1999), including the neuronal cell body (She et al. 2012). In addition, the response amplitude in the presence of ifenprodil was not different in untreated control cells compared to cells treated with Tat for 48 h (Figure S2), suggesting that the ifenprodil-insensitive component was not participating in the Tat-induced adaptive changes. Thus, even though both GluN2A- and GluN2B-containing NMDARs are susceptible to internalization (Lavezzari et al. 2004), we focused on the surface expression of GluN2B subunits because the NMDA-evoked increase in [Ca2+]i was predominantly mediated by GluN2B-containing NMDARs, the surface expression of GluN2B is more dynamic than GluN2A (Groc et al. 2006), and our region of interest focused on the neuronal cell body.
Next, we determined whether 48-h exposure to Tat affected GluN2B surface expression in primary hippocampal cultures using a biotinylation approach. To confirm the selectivity of our antibody for GluN2B, we first transfected HEK293 cells with plasmids encoding GluN1 with GluN2A or GluN2B and quantified protein expression via Western blot. The antibody selectively labeled GluN2B, but not GluN2A (data not shown). As shown in Fig. 7, GluN2B surface expression was unaffected by 48-h exposure to Tat (50 ng/mL). Addition of KT5823 (10 μM) 1 h prior to and during treatment with Tat for 48 h also had no effect on GluN2B surface expression. We conducted a time course experiment in which cultures were treated with Tat (50 ng/mL) for 0, 8, 24, and 48 h. GluN2B surface expression was unaffected by Tat exposure at all time points (data not shown). Furthermore, treating cultures with 100 nM Tat for 1 h did not affect surface expression of GluN2B-containing NMDARs (data not shown). To confirm that the assay was sufficiently sensitive to detect changes in NMDAR surface expression, we induced NMDAR internalization by a 5-min pre-treatment with glycine followed by 5-min treatment with glycine + NMDA (Nong et al. 2003). Treatment with glycine + NMDA evoked a 62 ± 33% decrease in GluN2B surface expression while 48-h treatment with Tat (50 ng/mL) tested in parallel had no effect (data not shown). These data indicate that adaptation following Tat-induced NMDAR potentiation is not because of internalization of GluN2B.
NMDARs are implicated in many neurodegenerative disorders including HAND (Young et al. 1988; Kaul et al. 2001; Snyder et al. 2005; Potter et al. 2013; Rossi et al. 2013; Spalloni et al. 2013). The HIV-1 protein Tat is a neurotoxin involved in the neuropathogenesis of HIV (Nath 2002; King et al. 2006). Here, we used fura-2-based Ca2+ imaging to investigate the effects of Tat on NMDA-evoked [Ca2+]i responses in rat hippocampal neurons in vitro. Tat potentiated NMDA-evoked increases in [Ca2+]i by LRP-dependent activation of Src kinase. Intriguingly, NMDA-evoked responses adapted following Tat-induced potentiation by activation of the NOS/sGC/PKG pathway. Adaptation of NMDA-evoked responses may be a novel neuroprotective mechanism to prevent excessive Ca2+ influx through NMDARs and could identify new targets for the treatment of HAND. The signaling pathways responsible for the biphasic modulation of NMDARs during exposure to Tat are summarized in Fig. 8.
Tat has been detected in the brains of patients with HIV (Wiley et al. 1996; Del Valle et al. 2000; Hudson et al. 2000). Tat is shed by HIV-infected cells (Steinaa et al. 1994; Chang et al. 1997) and can affect NMDAR function directly and indirectly. Previous work showed that Tat directly activates the NMDAR (Song et al. 2003) resulting in neurotoxicity (Li et al. 2008). Other reports indicate that Tat indirectly enhances NMDAR function via binding to and relieving the inhibitory effect of Zn2+ (Chandra et al. 2005) or via tyrosine kinase-mediated phosphorylation of NMDARs (Haughey et al. 2001). We found that Tat indirectly potentiated NMDA-evoked increases in [Ca2+]i as indicated by prevention of potentiation by the LRP antagonist, RAP. Tat also induces synapse loss (Kim et al. 2008; Shin et al. 2012) and cell death (Eugenin et al. 2007) via an LRP-dependent mechanism (Liu et al. 2000). LRP1 recognizes over 30 ligands with high affinity (Herz and Strickland 2001). Some of these agents alter NMDAR function. For example, activated α2-macroglobulin (Qiu et al. 2002) and amyloid-β (Snyder et al. 2005) inhibit NMDA-evoked currents and apolipoprotein-E4 potentiates NMDAR function (Qiu et al. 2003). We used a combined pharmacological and genetic approach to show that the previously described potentiation of NMDA-evoked [Ca2+]i responses by Tat (Haughey et al. 2001) was mediated by LRP-dependent activation of Src. The mechanism by which Tat activates Src following binding to LRP remains unclear. Ligand binding to LRP activates SFKs (Bock and Herz 2003; Shi et al. 2009). Neurons from animals with an inactivating knock-in mutation in the Lrp1 gene exhibit reduced phosphorylation of GluN2B tyrosine-1472, although this particular mutation increased both LRP1 and GluN2B surface expression (Maier et al. 2013). Thus, it is possible that Tat binding to the LRP activates a signaling cascade resulting in phosphorylation of NMDARs. Alternatively, following internalization, Tat causes fundamental changes in endolysosomal structure and function (Hui et al. 2012); thus, it is possible that Tat disrupts endolysosomal membrane integrity and escapes the endosome to activate Src. NMDAR function could be directly affected by Src-mediated phosphorylation (Salter and Kalia 2004) or indirectly via activation of other Src substrates.
Tat-induced potentiation of the NMDA-evoked increase in [Ca2+]i followed a biphasic time course. The Tat-induced NMDAR potentiation peaked at 8 h and then NMDA-evoked responses returned to baseline by 24 h eventually dropping below control by 48 h. Down-regulation of NMDAR function over the course of hours to days has been described previously following ethanol exposure (Wu et al. 2011) and for excitatory conditions such as benzodiazepine withdrawal (Shen and Tietz 2011). Adaptation following Tat-induced NMDAR potentiation has not been previously described, although it does correlate with adaptive changes in synapse number induced by Tat. Twenty-four-hour exposure to 50 ng/mL Tat produced a 50 ± 7% loss of glutamatergic synapses and a simultaneous 38 ± 3% increase in GABAergic synapses (Kim et al. 2008; Hargus and Thayer 2013). We suggest that Tat-induced synaptic changes and attenuation of NMDA-evoked [Ca2+]i responses are part of a neuroprotective response orchestrated by the cell to reduce excess excitatory input.
Adaptation of NMDA-evoked [Ca2+]i responses resulted from the activation of a NOS/sGC/PKG signaling pathway. A series of pharmacological and genetically expressed inhibitors of this pathway prevented adaptation without affecting potentiation of NMDARs or NMDA-evoked responses in cells that were not exposed to Tat. PKG-mediated phosphorylation of the vasodilator-stimulated phosphoprotein, a protein expressed in neurons and used to assess PKG activity (Butt et al. 1994; Wang and Robinson 1997), increased following 24-h exposure to Tat (Shin, A. H. and Thayer, S. A., personal communication) providing further support for Tat-induced activation of an NO-PKG pathway. Thus, this pathway was only activated after exposure to Tat, possibly resulting from potentiation. Sustained Tat-induced NO production results following the formation of a macromolecular complex composed of LRP, PSD95, NMDAR, and nNOS (Eugenin et al. 2007). NO inhibits NMDA receptor function (Manzoni et al. 1992), although this feedback inhibition is generally thought to result from the direct nitrosylation of NMDARs (Lei et al. 1992; Lipton and Stamler 1994) rather than activation of PKG. Activation of PKG leads to suppression of NMDAR-mediated currents (Furukawa and Mattson 1998); however, the mechanism by which PKG attenuates NMDAR function remains unclear.
Adaptation of potentiated NMDA-evoked [Ca2+]i increases is not unique to Tat. IL-1β potentiated NMDA-evoked responses which subsequently adapted following prolonged exposure, although it appears that the mechanism of adaptation is stimulus specific. Adaptation of Tat-induced NMDAR potentiation does not appear to result from attenuation of Src activity. The SFK inhibitor, PP2, suppressed NMDA-evoked increases in [Ca2+]i below control in fully adapted, Tat-treated cells, although it had no effect on NMDA-evoked responses in naive cells, suggesting that Src remained active even in cells in which NMDA-evoked responses had returned to control levels. Sustained Src-mediated phosphorylation of Tyr 1472 on the C-terminal tail of GluN2B could disrupt AP-2 binding and prevent clathrin-mediated endocytosis (Lavezzari et al. 2003), which might explain the sustained NMDAR surface expression indicated by the biotinylation experiments. Biotinylation experiments indicated that adaptation did not result from internalization of GluN2B-containing NMDARs. Thus, an additional process was activated to counteract the sustained Src-mediated potentiation of the NMDA-evoked increase in [Ca2+]i. One possibility that could account for the reduction in NMDA-evoked responses without a concomitant decrease in GluN2B surface expression is Ca2+-induced cytoskeletal changes. Depolymerization of actin protects from excitotoxicity by reducing glutamate receptor-mediated Ca2+ influx (Furukawa et al. 1995). PKG is known to alter actin polymerization via Rho-associated kinase (Sunico et al. 2010) and such cytoskeletal changes reduce Ca2+ influx through the NMDAR (Rosenmund and Westbrook 1993; Lei et al. 2001). Perhaps prolonged exposure to Tat alters the gating or ligand binding properties of the NMDAR by affecting the physical interaction between the actin cytoskeleton and NMDARs resulting in attenuated NMDA-evoked responses.
The biphasic change in NMDA-evoked [Ca2+]i responses induced by Tat raises interesting questions about the balance between optimal NMDAR function and neuronal survival. Pharmacologic inhibition of NMDARs protects neurons from Tat-induced cell death (Shin et al. 2012); thus, the adaptation described here would seem likely to improve neuronal survival. However, Tat-induced cell death and NMDAR adaptation are both prevented by inhibition of NOS with L-NAME (Eugenin et al. 2007). Does the timing, duration, and amount of NO production distinguish protective from toxic effects? Intriguingly, NO can be neuroprotective or neurotoxic (Lipton et al. 1993). NO confers neuroprotection by several mechanisms. It directly attenuates NMDAR-mediated Ca2+ influx via s-nitrosylation resulting in improved neuronal survival (Choi et al. 2000; Jaffrey et al. 2001). Furthermore, NO can stimulate the sGC/cGMP/PKG pathway to produce neuroprotective proteins such as the transcription factor cAMP-response element binding protein and the kinase Akt (Contestabile and Ciani 2004). Alternatively, the neuroprotective effects of NO may be overwhelmed by nitrosative stress following prolonged, persistent, and excessive production of NO, ultimately resulting in neurotoxicity and cell death.
Although NO production is necessary for Tat-induced cell death to occur (Eugenin et al. 2007), impaired brain function from HIV initially results from synaptodendritic injury that precedes overt neuronal death. Proper NMDAR function is essential for normal cognition. For example, over-expression of GluN2B-containing NMDARs enhances learning and memory function (Tang et al. 1999) while inhibition of NMDARs impairs these processes (Malhotra et al. 1996; Newcomer and Krystal 2001). Does the cognitive decline observed in patients with HAND result, in part, from excessive attenuation of NMDAR function? If so, then the pathways that potentiate and attenuate NMDARs during a neurotoxic challenge may prove useful targets for the pharmacological treatment of HAND. In addition, if the complex time course of NMDAR modulation observed during in vitro exposure to Tat is extrapolated to HAND, then different therapeutic approaches may be required for different stages of the disease.
In summary, we have shown that exposure to Tat evokes a biphasic modulation of NMDA-evoked [Ca2+]i responses. An LRP/Src pathway initially potentiates NMDA-evoked increases in [Ca2+]i and then activation of the NOS/sGC/PKG signaling pathway attenuates NMDA-evoked responses. Future experiments to determine how these pathways balance NMDAR-dependent cognitive functions with NMDAR-dependent toxicity in vivo may inform the targeting and timing of neuroprotective agents of potential use in treating HAND.
Acknowledgments and conflict of interest disclosure
We thank Dr Colin Campbell (University of Minnesota, Minneapolis, MN) for assistance with the generation of the G1αR-DsRed2 expression construct. The NIH AIDS Research and Reference Reagent Program provided HIV-1 Tat protein from Dr John Brady and DAIDS, NAID. We thank Dr Stephen Traynelis (Emory University, Atlanta, GA) for providing GluN1, GluN2A, and GluN2B expression constructs and Dr Darren Browning (Georgia Regents University, Augusta, GA) for providing the DN-PKG expression construct. The National Institutes of Health (DA07304 and DA035663 to SAT, DA034696 and MH061933 to KW) supported this work. National Institute on Drug Abuse Training Grants supported KK (DA007097) and NW (DA007234).
All experiments were conducted in compliance with the ARRIVE guidelines. The authors have no conflicts of interest to declare.