The present study examined the expression of transient receptor potential vanilloid subtype 1 (TRPV1) in microglia, and its association with microglial cell death. In vitro cell cultures, RT-PCR, Western blot analysis, and immunocytochemical staining experiments revealed that rat microglia and a human microglia cell line (HMO6) showed TRPV1 expression. Furthermore, exposure of these cells to TRPV1 agonists, capsaicin (CAP) and resiniferatoxin (RTX), triggered cell death. This effect was ameliorated by the TRPV1 antagonists, capsazepine and iodo-resiniferatoxin (I-RTX), suggesting that TRPV1 is directly involved. Further examinations revealed that TRPV1-induced toxicity was accompanied by increases in intracellular Ca2+, and mitochondrial damage; these effects were inhibited by capsazepine, I-RTX, and the intracellular Ca2+ chelator BAPTA-AM. Treatment of cells with CAP or RTX led to increased mitochondrial cytochrome c release and enhanced immunoreactivity to cleaved caspase-3. In contrast, the caspase-3 inhibitor z-DEVD-fmk protected microglia from CAP- or RTX-induced toxicity. In vivo, we also found that intranigral injection of CAP or 12-hydroperoxyeicosatetraenoic acid, an endogenous agonist of TRPV1, into the rat brain produced microglial damage via TRPV1 in the substantia nigra, as visualized by immunocytochemistry. To our knowledge, this study is the first to demonstrate that microglia express TRPV1, and that activation of this receptor may contribute to microglial damage via Ca2+ signaling and mitochondrial disruption.

Transient receptor potential vanilloid subtype 1 (TRPV1)3 is a nonselective, oligomeric cation channel that is activated by vanilloids, capsaicin (CAP) (1, 2, 3, 4, 5, 6, 7), and resiniferatoxin (RTX) (6, 8), and products of lipoxygenases, such as 12-hydroperoxyeicosatetraenoic acid (12-HPETE) (9, 10, 11). TRPV1 was widely expressed in the brain (12, 13), where it plays a significant CNS roles in various brain regions, such as hypothalamus (14), locus ceruleus (15), and hippocampus (16). Activation of TRPV1 excites sensory neurons and induces the accumulation of intracellular Ca2+ ([Ca2+]i) (10, 17). This results in excessive mitochondrial Ca2+ loading and subsequent mitochondrial disruption, leading to neuronal cell death (17, 18). Activation of TRPV1 by treatment with CAP was found to increase glutamate release from nigral slices (19), produce hypokinesia in parallel with decreased nigrostriatal neuron activity (20, 21), and disrupt the blood-brain barrier following ischemia-reperfusion (22). Several lines of evidence have implicated 12-HPETE, an endogenous TRPV1 agonist (10), in the induction of long-term depression in the hippocampus (23) and neurodegeneration in mesencephalic cultures (24). Very recently, we demonstrated that massive TRPV1-induced Ca2+ influxes induced in vivo and in vitro cell death in TRPV1-expressing mesencephalic neurons (5).

Collectively, these observations strongly suggest that TRPV1 plays a significant role in the CNS. However, little is known regarding the expression and function(s) of TRPV1 in microglia. In this study, we sought to determine whether microglia express TRPV1, and whether this receptor mediates microglial cell death in vivo and in vitro. We further assessed whether this toxicity is accompanied by TRPV1-induced Ca2+ influxes and subsequent mitochondrial disruption.

Chemicals were purchased from the following companies: CAP (Sigma-Aldrich), fura-2/AM, Mito-Tracker, Live/Dead Viability/cytotoxicity kit (Molecular Probes), BAPTA-AM (Sigma-Aldrich), capsazepine (CZP), RTX, iodo-RTX (I-RTX) (Tocris), z-DEVD-fmk (caspase-3 inhibitor; Biochemicals), Anti-cytochrome c (Promega or BD Pharmingen), and 12-HPETE (Cayman Chemical). Anti-TRPV1 was obtained from the sensory research center (Seoul National University, Seoul, Korea). The vehicle used to dissolve CAP or 12-HPETE was sterile PBS containing 14% ethanol for stereotaxic injection in brain. RTX was also dissolved in PBS containing ethanol for treatment on cultures, and CZP, BAPTA-AM, or I-RTX were dissolved in DMSO. The final concentration of ethanol or DMSO treated on cultures was 0.1%.

Sprague Dawley rat microglia-enriched cultures were prepared as previously described (25, 26). Cultured rat microglia were plated on 12-mm-round aclar plastic coverslips (5 × 104 cells/coverslip) precoated with 0.1 mg/ml poly-d-lysine housed in 24-well plates, or plated in 24-well plates (5 × 104 cells/well) or 35-mm culture dishes (5 × 105 cells/dish). One hour later, the culture medium was changed to MEM medium containing 5% FBS.

Astrocytes were cultured from the cerebral cortices of 1-day-old Sprague Dawley rats as previously described (27). Briefly, the cortices were triturated into single cells in MEM containing 10% FBS and plated into 75-cm2 T flasks (0.5 hemisphere/flask) for 2 wk. To prepare astrocyte-enriched cultures, microglia were detached by mild shaking and the cells remaining in the flask were harvested with 0.1% trypsin. Astrocytes were plated in 35-mm culture dishes (5 × 105 cells/dish), and incubated in MEM medium containing 5% FBS.

Immortalized human microglial cells (HMO6) were prepared as previously described (28). The permission to use embryonic tissues was granted by the Clinical Screening Committee for Research involving Human Subjects of the University of British Columbia. The cultured HMO6 were plated in 24-well plates (5 × 104 cells/well) or 35-mm culture dishes (5 × 105 cells/dish), and treated with drugs, or processed for Western blot assay, fluorescence staining, or Ca2+ imaging.

As previously described (26, 29), total RNA was extracted, and reverse transcription was conducted. The primers for specific rat TRPV1 were primer 1 (5′-GACATGCCACCCAGCAGG) and primer 2 (5′-TCAATTCCCACACACCTCCC) corresponding to nucleotides 2491–2508 and 2755–2735, respectively, of the published rTRPV1 cDNA sequence (12). The PCR cycles consisted of denaturation at 94°C for 30 s, annealing 55°C for 30 s, and extension at 69°C for 3 min for 30 cycles. GAPDH was also amplified as an internal PCR control using the following primers 1 (5′-TCCCTCAAGATTGTCAGCAA) and primer 2 (5′-AGATCCACAACGGATACATT). The temperature cycling conditions were as follows: 2 min at 94°C, 25 cycles of (94°C for 30 s, 55°C for 30 s, and 72°C for 90 s), and a final extension at 72°C for 10 min. The PCR product was separated by electrophoresis on a 1.6% agarose gel, stained with ethidium bromide, and then detected under UV light.

Expression of TRPV1 in substantia nigra (SN) in vivo or microglia in vitro were assayed as previously described, with modifications (5, 17, 30). Briefly, cells were homogenized with 0.5 ml of ice-cold lysis buffer containing 20 mM Tris-HCl (pH 7.5), 1 mM EDTA, 5 mM MgCl2, 1 mM DTT, 20 μg/ml aprotinin, and 1 mM PMSF. The protein concentration was determined using a BCA kit. Equal amounts of protein were loaded in each lane with loading buffer containing 62.5 mM Tris-HCl (pH 6.8), 2% SDS, 10% glycerol, 50 mM DTT, and 0.1% bromphenol blue. Samples were boiled at 100°C for 2 min before gel loading. Homogenate samples were electrophoresed on 12% SDS-polyacrylamide gel and transferred to polyvinylidene difluoride membranes (Millipore) using an electrophoretic transfer system (Bio-Rad). The membranes were then blocked in TNE buffer (in mM: 10 Tris-HCl (pH 7.5), 50 NaCl, and 2.5 EDTA (pH 8.0)) containing 5% skim milk for 1 h with gentle shaking. Membranes were then incubated overnight at 4°C with antiserum against the C-terminal cytoplasmic domain of TRPV1, as described previously, in TNE containing 5% skim milk. Mitochondrial fractions from cultured microglia treated with CAP or RTX were prepared as described (5), proteins were separated by 12% SDS-PAGE gels, and transferred to membranes. The membranes were immunoblotted with mouse anti-cytochrome c (BD Pharmingen), and proteins were visualized using the ECL kit (Amersham Biosciences).

Cultures and brain tissue were prepared for immunostaining as previously described (5, 29). The primary Abs included those directed against OX-42 (specific for complement receptor type 3; Serotec), tyrosine hydroxylase (TH; Pel-Freez), TRPV1 (Sensory Research Center, Seoul National University, Seoul, Korea) (5, 30), glial fibrillary acidic protein (GFAP; Sigma-Aldrich), cytochrome c (Promega), and cleaved caspase-3 (Cell Signaling). Stained cells were viewed and analyzed under a bright-field microscope (Nikon) or viewed with a confocal laser-scanning microscope (Olympus).

Cultured microglia seeded on 25-mm coverslips were treated with CAP or RTX, and the cultures were stained 48 h later with 2 μM calcein-acetoxymethyl ester (calcein-AM) and 4 μM ethidium homodimer-1 (Eth-1). The calcein-positive live cells (green) and ethidium-positive dead cells (red) were visualized using a confocal laser-scanning microscope (5).

Cell death effect by CAP was determined using the Apoptag Fluorescein Direct in situ Apoptosis Detection kit (Intergen) that detects the 3′-OH region of cleaved DNA. TUNEL staining was followed by kit protocol. Microglia cultures were treated without or with 50 μM CAP for 12 h. And then cultures were fixed in 1% paraformaldehyde in PBS, incubated with a mixture of TdT and reaction buffer containing FITC fluorescein-conjugated-digoxigenin-dUTP (Intergen) in a humidified chamber for 1 h at 37°C, and washed in washing buffer (Intergen). For double staining of TUNEL with anti-OX-42, cultures were incubated with anti-OX-42 Abs for overnight at room temperature, and then applied with Texas Red-conjugated anti-mouse IgG (Vector Laboratories) for 30 min, and viewed using a confocal laser-scanning microscope.

Cultured microglia for Ca2+ imaging were prepared as previously described, with modifications (5). Briefly, [Ca2+]i was determined on the basis of the ratio of fura-2 fluorescence (R) at 340 and 380 nm. Microglia preloaded with fura-2 dye were selected under CCD camera attached to a Zeiss inverted microscope, and changes of fluorescent intensity were performed at 10-s intervals and analyzed using an Ion Application (Empix).

To label mitochondria in cultured microglia seeded on 25-mm coverslips, Mito-Tracker (Molecular Probes) dye was incubated for 30 min at a 250 nM concentration. The cells were washed three times in HBSS containing 1.8 mM CaCl2 and examined under the confocal laser-scanning microscope. Changes of morphology were performed at 10-s intervals with an He-Ne green laser (5).

Female Sprague Dawley rats (230∼250 g) were anesthetized with injection of chloral hydrate (400 mg/kg i.p.), positioned in a stereotaxic apparatus (Kopf Instrument), and received a unilateral administration of drugs into the right SN (anterior and posterior −5.3, medial and lateral −2.3, dorsal and ventral from Bregma −7.6 mm from bregma), according to the atlas of Paxinos and Watson (31). The drugs were injected at a rate of 0.5 μl/min using a 26-gauge Hamilton syringe attached to an automated microinjector (kdScientific). After injection, the needle was left in place for an additional 5 min before being slowly retracted. Intact (noninjected) or vehicle-injected animals were used as controls.

As previously described (32), TL histochemistry was performed to visualize microglia with OX-42. Briefly, some sections were mounted on a gelatin-coated slide incubated in 0.2% Triton X-100 for 30 min, rinsed twice in PBS, and then incubated for 1 h at room temperature with the FITC-labeled TL (Vector Laboratories). Stained cells in tissue were viewed using a confocal laser-scanning microscope.

All values are expressed as mean ± SEM. Statistical significance was assessed by ANOVA using Instat 3.05 (GraphPad Software), followed by Student-Newman-Keuls analyses.

The results of our RT-PCR (Fig. 1,A) and Western blot (Fig. 1,B) analyses revealed that TRPV1 is expressed in cultured rat cortical microglia, but not in astrocytes. Consistent with these findings, double-immunostaining using Abs to TRPV1 (Fig. 1,C, green) and OX-42 (Fig. 1,C, red) for microglia, or GFAP (Fig. 1,D, red) for astrocytes indicated that TRPV1 was expressed in microglia (yellow), but not in astrocytes. In vivo analyses showed that in the SN, TRPV1 (Fig. 1, E and F, green) was expressed in TH-immunopositive (TH-ip) neurons (Fig. 1,E, red) and OX-42-ip microglia (Fig. 1 F, red).

FIGURE 1.

TRPV1 is expressed in microglia. A and B, Analysis of RT-PCR (A) and Western blot (B) show mRNA (265 bp) and protein (∼95 kDa) expression of TRPV1, respectively, in the rat SN and cultured rat microglia (MG), but not in astrocytes (AC). C and D, Cultured rat microglia and astrocytes were immunostained simultaneously with Abs to TRPV1 (green, C and D) and OX-42 (red, C) for microglia, or GFAP for astrocytes (red, D), and the two were merged (yellow), respectively. E and F, Rat SN sections were immunostained simultaneously with Abs to TRPV1 (green, E and F) and TH (red, E) for dopaminergic neurons, or OX-42 (red, F), and the two were merged, respectively. Arrows indicate merged cells (yellow), and arrowheads indicate cells immunostained with Abs to TRPV1 alone (green). The results shown are representative from four independent experiments. Scale bar: C and D, 20 μm; E and F, 50 μm.

FIGURE 1.

TRPV1 is expressed in microglia. A and B, Analysis of RT-PCR (A) and Western blot (B) show mRNA (265 bp) and protein (∼95 kDa) expression of TRPV1, respectively, in the rat SN and cultured rat microglia (MG), but not in astrocytes (AC). C and D, Cultured rat microglia and astrocytes were immunostained simultaneously with Abs to TRPV1 (green, C and D) and OX-42 (red, C) for microglia, or GFAP for astrocytes (red, D), and the two were merged (yellow), respectively. E and F, Rat SN sections were immunostained simultaneously with Abs to TRPV1 (green, E and F) and TH (red, E) for dopaminergic neurons, or OX-42 (red, F), and the two were merged, respectively. Arrows indicate merged cells (yellow), and arrowheads indicate cells immunostained with Abs to TRPV1 alone (green). The results shown are representative from four independent experiments. Scale bar: C and D, 20 μm; E and F, 50 μm.

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Treatment of microglial cultures with 50 μM of the TRPV1 agonist, CAP dramatically increased the number of Eth-1-positive dead cells (Fig. 2,D), and dramatically decreased the number of calcein-AM-positive live cells (Fig. 2,C), compared with vehicle-treated controls (Fig. 2, A and B). Double-immunofluorescence staining with an anti-OX-42 Ab (Fig. 2,D, inset, red) and TUNEL staining (Fig. 2,D, inset, green) indicated that TUNEL/OX-42-positive cells were seen in microglial cultures 12 h after treatment with 50 μM CAP. When microglial cell death was quantitatively assessed by enumeration of Eth-1-positive cells, we found that treatment with 10 μM CAP or 1 μM RTX, another TRPV1 agonist (6, 8, 30), had no effect (data not shown), whereas treatment with 50 μM CAP or 10 μM RTX produced ∼8-fold increases in Eth-1-positive cells compared with vehicle-treated control cultures (Fig. 2 E, p < 0.001). Pretreatment of cultures for 5 min with the TRPV1 antagonists, CZP (10 μM) or I-RTX (1 μM), partially attenuated the accumulation of Eth-1-positive cells induced by 50 μM CAP (32 and 23%, respectively; p < 0.01) or 10 μM RTX (26%, p < 0.01 and 20%, p < 0.05, respectively).

FIGURE 2.

TRPV1 mediates cell death of cultured rat microglia. AD, Microglia cultures were treated with vehicle (MEM medium containing 5% FBS and 0.1% ethanol) (A and B), 50 μM CAP (C and D) for 48 h, and stained with calcein-AM (green for live, A and C) and Eth-1 (red for dead, B and D), respectively. Inset in D shows colocalization of OX-42 and TUNEL in microglial cultures at 12 h after treatment with 50 μM CAP. E, Where indicated, cells were pretreated with 10 μM CZP or 1 μM I-RTX for 5 min before treatment with 50 μM CAP or 10 μM RTX. Cell death was assessed by counting the number of Eth-1-positive dead cells under a confocal microscope. F and G, OX-42 immunostaining of cultures treated with vehicle (F) or 50 μM CAP (G) for 48 h is shown. H, Cultures were pretreated with 10 μM CZP or 1 μM I-RTX for 5 min before treatment with CAP or RTX, or cotreated with 5 μM BAPTA-AM (BAP) and TRPV1 agonists, respectively. Cell death was assessed by counting the number of OX-42-ip cells. The statistical significance of differences was assessed using one-way ANOVA, followed by Student-Newman-Keuls analyses. All values represent the mean ± SEM of triplicate cultures in four separate plating. ∗, p < 0.001, significant from control; #, p < 0.01 and ∗∗, p < 0.05, significant from treatment with CAP or RTX. Scale bar: 100 μm.

FIGURE 2.

TRPV1 mediates cell death of cultured rat microglia. AD, Microglia cultures were treated with vehicle (MEM medium containing 5% FBS and 0.1% ethanol) (A and B), 50 μM CAP (C and D) for 48 h, and stained with calcein-AM (green for live, A and C) and Eth-1 (red for dead, B and D), respectively. Inset in D shows colocalization of OX-42 and TUNEL in microglial cultures at 12 h after treatment with 50 μM CAP. E, Where indicated, cells were pretreated with 10 μM CZP or 1 μM I-RTX for 5 min before treatment with 50 μM CAP or 10 μM RTX. Cell death was assessed by counting the number of Eth-1-positive dead cells under a confocal microscope. F and G, OX-42 immunostaining of cultures treated with vehicle (F) or 50 μM CAP (G) for 48 h is shown. H, Cultures were pretreated with 10 μM CZP or 1 μM I-RTX for 5 min before treatment with CAP or RTX, or cotreated with 5 μM BAPTA-AM (BAP) and TRPV1 agonists, respectively. Cell death was assessed by counting the number of OX-42-ip cells. The statistical significance of differences was assessed using one-way ANOVA, followed by Student-Newman-Keuls analyses. All values represent the mean ± SEM of triplicate cultures in four separate plating. ∗, p < 0.001, significant from control; #, p < 0.01 and ∗∗, p < 0.05, significant from treatment with CAP or RTX. Scale bar: 100 μm.

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To further confirm that TRPV1 agonists can induce microglial cell death, we performed OX-42 immunocytochemical staining. Few of OX-42-ip cells were observed in microglial cultures at 48 h after treatment with 50 μM CAP (Fig. 2,G) or 10 μM RTX (data not shown), but not in vehicle-treated control cultures (Fig. 2,F). CAP- or RTX-induced microglial cell death was also attenuated by pretreatment of cultures for 5 min with 10 μM CZP or 1 μM I-RTX, or cotreatment with 5 μM of the intracellular Ca2+ chelator, BAPTA-AM (Fig. 2 H). When our results were quantified and expressed as a percentage of untreated control values, we found that treatment with 50–100 μM CAP reduced the number of microglia by 51–79% (p < 0.001). The toxicity induced by CAP (100 μM) was partially inhibited by pretreatment with 10 μM CZP (34%, p < 0.01) or 1 μM I-RTX (32%, p < 0.01) or cotreatment with 5 μM BAPTA/AM (24%, p < 0.05), indicating that this toxicity occurs via activation of TRPV1 and increased [Ca2+]i. Treatment of cells with 10 μM RTX also reduced the number of microglia by 59% (p < 0.001), and this toxicity was inhibited by pretreatment with 10 μM CZP (29%, p < 0.05) or 1 μM I-RTX (37%, p < 0.01), or cotreatment with BAPTA-AM (22%, p < 0.05).

To examine whether increases in [Ca2+]i via influx through TRPV1 may account for CAP-induced toxicity (5, 17), we measured changes in [Ca2+]i by monitoring fura-2 fluorescence intensity. We found that administration of 50 μM CAP (Fig. 3,A) increased the fluorescence intensity in cultured rat microglia, indicating elevation of [Ca2+]i. This increase of [Ca2+]i was dramatically attenuated in cultures treated with CAP in the presence of 10 μM CZP (Fig. 3,C) or 1 μM I-RTX (Fig. 3,D), and completely abolished in cells bathed in a Ca2+-free extracellular solution (Fig. 3,B), indicating that the CAP-induced influx of extracellular Ca2+ occurred through TRPV1. In an experiment designed to further determine the source of the [Ca2+]i increase, an endoplasmic reticulum Ca2+ pump inhibitor, thapsigargin (5, 33), was found to have no effects on CAP-induced Ca2+ influx (data not shown). Treatment with BAPTA-AM completely blocked the CAP-induced [Ca2+]i increase (Fig. 3,E) and rescued CAP-treated microglia (Fig. 2,H), suggesting that CAP-induced cell death is associated with increased [Ca2+]i. Treatment with 10 μM RTX also mimicked the effects of CAP (Fig. 3, F and G), and RTX-induced Ca2+ influxes could be blocked by TRPV1 antagonists (10 μM CZP and 1 μM I-RTX; Fig. 3, H and I, respectively) and BAPTA-AM (Fig. 3 J).

FIGURE 3.

Changes of R (F340/380) fluorescence in fura-2-loaded cultured rat microglia. A–J, [Ca2+]i was measured in cultures treated with 50 μM CAP (A–E, arrowhead) or 10 μM RTX (F–J, arrowhead) in the presence (A and F) or absence (B and G) of 1.8 mM extracellular calcium. Response to CAP or RTX after pretreatment with 10 μM CZP (C and H, arrow) or 1 μM I-RTX (D and I, arrow), or cotreatment of 5 μM BAPTA-AM with 50 μM CAP (E and J) in the presence of 1.8 mM extracellular calcium. Data were averaged from 20 to 25 randomly selected cells for each condition, and the results shown are representative from three independent experiments.

FIGURE 3.

Changes of R (F340/380) fluorescence in fura-2-loaded cultured rat microglia. A–J, [Ca2+]i was measured in cultures treated with 50 μM CAP (A–E, arrowhead) or 10 μM RTX (F–J, arrowhead) in the presence (A and F) or absence (B and G) of 1.8 mM extracellular calcium. Response to CAP or RTX after pretreatment with 10 μM CZP (C and H, arrow) or 1 μM I-RTX (D and I, arrow), or cotreatment of 5 μM BAPTA-AM with 50 μM CAP (E and J) in the presence of 1.8 mM extracellular calcium. Data were averaged from 20 to 25 randomly selected cells for each condition, and the results shown are representative from three independent experiments.

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We next examined whether [Ca2+]i elevation via activation of TRPV1 in cultured rat microglia could contribute to mitochondrial disruption, and whether this mitochondrial damage could result in cytochrome c release (5). Untreated control cells showed intact mitochondrial structures, as determined by Mito-Tracker fluorescence (Fig. 4,A). In contrast, mitochondrial disruption was noted 20 min after addition of 50 μM CAP to cell cultures (Fig. 4,B). Consistent with the above-mentioned clearance of cytosolic Ca2+, pretreatment of cells for 5 min with 10 μM CZP (Fig. 4, C and D) or 1 μM I-RTX (data not shown), or cotreatment with 5 μM BAPTA-AM (Fig. 4, E and F), prevented this CAP-induced mitochondrial damage. Also consistent with the above findings, RTX mimicked the effects of CAP in this context (data not shown). Double immunofluorescence staining with Mito-Tracker and anti-cytochrome c Abs revealed that the entire cytochrome c signal was localized to the mitochondria of untreated controls (Fig. 4,G), whereas cells treated with 10 μM RTX (Fig. 4,H) or 50 μM CAP (Fig. 4,I) showed scattered and redistributed spots of cytochrome c into the cytosol, indicating that the protein had been released from the mitochondria. This CAP-induced cytochrome c release was confirmed by Western blot analysis (Fig. 4 J).

FIGURE 4.

Activation of TRPV1 induces mitochondrial disruption of microglia in cultures. A and B, Mitochondrial disruption in rat microglia cultures treated with 50 μM CAP for 20 min in the presence of 1.8 mM extracellular calcium. C and F, Inhibition of CAP-induced mitochondrial disruption by pretreatment with 10 μM CZP (C and D) or cotreatment with 5 μM BAPTA-AM (E and F). Each colored arrow indicates the same cells. GI, Localization of cytochrome c (green) immunoreactivity and Mito-Tracker (red) in intact microglia (G) and cells treated with 10 μM RTX (H) or 50 μM CAP (I) for 12 h. J, Western blot analysis of cytochrome c levels (15 kDa) after treatment of cells with 50 μM CAP or 10 μM RTX for 12 h. Con, nontreated controls; Cyto, cytosolic fraction; Mito, mitochondrial fraction. K–M, Immunochemical localization of cleaved caspase-3 (green) in OX-42-ip (red) cells in untreated cultures (K) and cultures treated with CAP (L) or RTX (M). N, z-DEVD-fmk reduces CAP- or RTX-induced cell death of microglia. The statistical significance of differences was assessed using one-way ANOVA, followed by Student-Newman-Keuls analyses. All values represent the mean ± SEM of triplicate cultures in four separate plating. ∗, p < 0.001, significant from control; #, p < 0.01, and ∗∗, p < 0.05, significant from treatment with CAP or RTX. Scale bars: A–L, 20 μm.

FIGURE 4.

Activation of TRPV1 induces mitochondrial disruption of microglia in cultures. A and B, Mitochondrial disruption in rat microglia cultures treated with 50 μM CAP for 20 min in the presence of 1.8 mM extracellular calcium. C and F, Inhibition of CAP-induced mitochondrial disruption by pretreatment with 10 μM CZP (C and D) or cotreatment with 5 μM BAPTA-AM (E and F). Each colored arrow indicates the same cells. GI, Localization of cytochrome c (green) immunoreactivity and Mito-Tracker (red) in intact microglia (G) and cells treated with 10 μM RTX (H) or 50 μM CAP (I) for 12 h. J, Western blot analysis of cytochrome c levels (15 kDa) after treatment of cells with 50 μM CAP or 10 μM RTX for 12 h. Con, nontreated controls; Cyto, cytosolic fraction; Mito, mitochondrial fraction. K–M, Immunochemical localization of cleaved caspase-3 (green) in OX-42-ip (red) cells in untreated cultures (K) and cultures treated with CAP (L) or RTX (M). N, z-DEVD-fmk reduces CAP- or RTX-induced cell death of microglia. The statistical significance of differences was assessed using one-way ANOVA, followed by Student-Newman-Keuls analyses. All values represent the mean ± SEM of triplicate cultures in four separate plating. ∗, p < 0.001, significant from control; #, p < 0.01, and ∗∗, p < 0.05, significant from treatment with CAP or RTX. Scale bars: A–L, 20 μm.

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We recently reported that TRPV1-facilitated Ca2+ influxes, cytochrome c release and subsequent caspase-3 activation are associated with neurodegeneration (5). Here, we showed that activation of TRPV1 resulted in the release of cytochrome c from mitochondria, leading to degeneration of microglia (Fig. 4, H–J). As previous reports have associated caspase-3 activation with microglial cell death (34, 35, 36), we next examined whether caspase-3 is involved in TRPV1-induced degeneration of microglia. Cultures were treated with the 10 μM of the caspase-3 inhibitor, z-DEVD-fmk (5), together with 100 μM CAP or 10 μM RTX. Treatment of cells with z-DEVD-fmk significantly reduced the microglial cell death induced by 100 μM CAP (39%, p < 0.01) or 10 μM RTX (23%, p < 0.05), compared with their respective controls (Fig. 4,N), whereas 10 μM z-DEVD-fmk alone had no effect on cell viability. Double immunofluorescence staining with Abs against cleaved caspase-3 (Fig. 4, L and M, green) and OX-42 (Fig. 4, K--M, red) showed that activated caspase-3 was present in CAP- (Fig. 4,L), or RTX- (Fig. 4,M) treated microglia, but not in untreated controls (Fig. 4 K).

We next determined the presence and function of TRPV1 in the HMO6, immortalized human microglial cell line (28, 34). Western blot analysis (Fig. 5,A) and immunocytochemical staining with specific Abs (Fig. 5, BE) revealed that TRPV1 is expressed in HMO6 cells. Treatment of HMO6 cultures with 100 μM CAP dramatically increased the number of Eth-1-positive dead cells (Fig. 5,I) and substantially decreased the number of calcein-AM-positive live cells (Fig. 5,H), as compared with vehicle-treated controls (Fig. 5, F and G). Treatment with 10 μM CAP or 1 μM RTX had no effect (data not shown), whereas treatment with 100 μM CAP or 10 μM RTX increased the number of Eth-1-positive dead cells by 57% (p < 0.001) and 43% (p < 0.001), respectively, compared with untreated control cultures (Fig. 5,J). Pretreatment of cultures for 5 min with 10 μM CZP partially attenuated the number of Eth-1-positive cells in cultures exposed to 100 μM CAP (22%, p < 0.05) or 10 μM RTX (21%, p < 0.05). In addition, administration of 50 μM CAP increased the fluorescence intensity of fura-2 fluorescent images taken from HMO6 cells (Fig. 5,K) over 200 s, indicating elevation of [Ca2+]i. Treatment with RTX (10 μM) mimicked the effects of CAP (data not shown), and CAP-induced [Ca2+]i increases were attenuated by pretreatment with 10 μM CZP (Fig. 5 L) or 1 μM I-RTX (data not shown), and were completely abolished in a Ca2+-free extracellular solution (data not shown), indicating that the CAP-induced influx of extracellular Ca2+ occurs through TRPV1 in cultured human microglia.

FIGURE 5.

TRPV1 mediates cell death of immortalized human microglial cell line, HMO6. A, Western blot analysis shows expression of TRPV1 in cultured rat microglia (MG) and HMO6 cells. B–E, Triple staining of TL (B), anti-TRPV1 Ab (C), and Hoechst 33258 dye (D) in the HMO6 cultures. Images were merged (E). F–I, HMO6 cultures were treated with vehicle (MEM medium containing 5% FBS and 0.1% ethanol) (F and G) or 100 μM CAP (H and I) for 48 h and stained with calcein-AM (green for live, F and H) and Eth-1 (red for dead, G and I), respectively. J, Where indicated, cells were pretreated with 10 μM CZP for 5 min before treatment with 100 μM CAP or 10 μM RTX. Cell death was assessed by counting the number of Eth-1-positive dead cells under a confocal microscope. The statistical significance of differences was assessed using one-way ANOVA, followed by Student-Newman-Keuls analyses. All values represent the mean ± SEM of triplicate cultures in four separate plating. ∗, p < 0.001, significant from control; ∗∗, p < 0.05, significant from treatment with CAP or RTX. Scale bar: B–E, 20 μm; F–I, 60μm. K and L, Changes of [Ca2+]i were measured using fura-2 dye in HMO6 cultures treated with 50 μM CAP (arrowhead) (K and L) or pretreated with 10 μM CZP (arrow) (L) in the presence of 1.8 mM extracellular calcium. Data were averaged from 20 to 25 randomly selected cells for each condition.

FIGURE 5.

TRPV1 mediates cell death of immortalized human microglial cell line, HMO6. A, Western blot analysis shows expression of TRPV1 in cultured rat microglia (MG) and HMO6 cells. B–E, Triple staining of TL (B), anti-TRPV1 Ab (C), and Hoechst 33258 dye (D) in the HMO6 cultures. Images were merged (E). F–I, HMO6 cultures were treated with vehicle (MEM medium containing 5% FBS and 0.1% ethanol) (F and G) or 100 μM CAP (H and I) for 48 h and stained with calcein-AM (green for live, F and H) and Eth-1 (red for dead, G and I), respectively. J, Where indicated, cells were pretreated with 10 μM CZP for 5 min before treatment with 100 μM CAP or 10 μM RTX. Cell death was assessed by counting the number of Eth-1-positive dead cells under a confocal microscope. The statistical significance of differences was assessed using one-way ANOVA, followed by Student-Newman-Keuls analyses. All values represent the mean ± SEM of triplicate cultures in four separate plating. ∗, p < 0.001, significant from control; ∗∗, p < 0.05, significant from treatment with CAP or RTX. Scale bar: B–E, 20 μm; F–I, 60μm. K and L, Changes of [Ca2+]i were measured using fura-2 dye in HMO6 cultures treated with 50 μM CAP (arrowhead) (K and L) or pretreated with 10 μM CZP (arrow) (L) in the presence of 1.8 mM extracellular calcium. Data were averaged from 20 to 25 randomly selected cells for each condition.

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Finally, we examined TRPV1-induced microglial cell death in the SN in vivo. Two days after intranigral injection of CAP (500 pmol/3 μl), we observed dramatic reductions in the number of OX-42-positive cells in the SN (Fig. 6, E and F), compared with intact (Fig. 6, A and B) or vehicle-treated controls (Fig. 6, C and D). We also found that 12-HPETE (200 pM/3 μl), an endogenous agonist of TRPV1 (9, 10), mimicked CAP-induced toxicity on microglia (Fig. 6, G and H). To delineate the site and extent of injury induced by a single injection of CAP into the SN over the course of 48 h, we prepared a schematic drawing from tissues immunostained with the anti-OX-42 Ab, using a camera lucida microscope attachment (Fig. 6,I) (32). Fewer OX-42-ip microglia were seen in the ipsilateral SN vs the contralateral SN. Seven days later, a number of OX-42-ip cells with resting morphology were observed (Fig. 6, J and K). Additional experiments revealed that intranigral coinjection of 500 pM CZP prevented the CAP- (Fig. 6, L and M) or 12-HPETE- (Fig. 6, N and O) induced microglial death in the SN, as did coinjection of 500 pM I-RTX (Fig. 6, PS). Treatment of cells with CZP or I-RTX alone had no effect on microglia (data not shown).

FIGURE 6.

TRPV1 mediates cell death of microglia in the SN in vivo. A and B, Intact SN section was immunostained with an OX-42 Ab. C–H, Animals receiving an unilateral injection of vehicle (C and D), 500 pM CAP (E and F) or 200 pM 12-HPETE (G and H) into the SN were sacrificed 2 days later. Brains were removed and tissue sections were immunostained with an anti-OX-42 Ab. I, Schematic drawing of a representative coronal section from rat brain showing distribution of OX-42-ip microglia traced using a camera lucida microscope attachment 2 days after CAP treatment. Note the absence of OX-42-ip microglia within the dotted lines in ipsilateral SN, as shown in E. The damaged area reaches to two-thirds of SN pars compacta (SNpc). Arrow and dotted lines indicate CAP-injected site and SNpc, respectively. J and K, A number of OX-42-ip cells with resting morphology were observed 7 days after CAP injection. L–O, Intranigral cotreatment with 500 pM CAP and 500 pM CZP (L and M), or 200 pM 12-HPETE and 500 pM CZP (N and O) rescued microglia as evidenced by OX-42 immunocytochemistry. PS, Intranigral cotreatment with 500 pM CAP and 500 pM I-RTX (P and Q), or 200 pM 12-HPETE and 500 pM I-RTX (P and Q) also rescued microglia. Six to eight animals were used for each experimental group. Scale bars: (in A) A, C, E, G, J, L, N, P, and R, 100 μm; (in B) B, D, F, H, K, M, O, Q, and S, 20 μm; I, 2 mm.

FIGURE 6.

TRPV1 mediates cell death of microglia in the SN in vivo. A and B, Intact SN section was immunostained with an OX-42 Ab. C–H, Animals receiving an unilateral injection of vehicle (C and D), 500 pM CAP (E and F) or 200 pM 12-HPETE (G and H) into the SN were sacrificed 2 days later. Brains were removed and tissue sections were immunostained with an anti-OX-42 Ab. I, Schematic drawing of a representative coronal section from rat brain showing distribution of OX-42-ip microglia traced using a camera lucida microscope attachment 2 days after CAP treatment. Note the absence of OX-42-ip microglia within the dotted lines in ipsilateral SN, as shown in E. The damaged area reaches to two-thirds of SN pars compacta (SNpc). Arrow and dotted lines indicate CAP-injected site and SNpc, respectively. J and K, A number of OX-42-ip cells with resting morphology were observed 7 days after CAP injection. L–O, Intranigral cotreatment with 500 pM CAP and 500 pM CZP (L and M), or 200 pM 12-HPETE and 500 pM CZP (N and O) rescued microglia as evidenced by OX-42 immunocytochemistry. PS, Intranigral cotreatment with 500 pM CAP and 500 pM I-RTX (P and Q), or 200 pM 12-HPETE and 500 pM I-RTX (P and Q) also rescued microglia. Six to eight animals were used for each experimental group. Scale bars: (in A) A, C, E, G, J, L, N, P, and R, 100 μm; (in B) B, D, F, H, K, M, O, Q, and S, 20 μm; I, 2 mm.

Close modal

To corroborate our finding that TRPV1 mediates microglial cell death, we double-labeled sections adjacent to those used in Fig. 6 with TL (Fig. 7, green) and OX-42 Ab (Fig. 7, red) (32). Consistent with the above findings, TL-positive staining was mainly found in blood vessels, with few of TL- and OX-42-positive microglia noted after 2 days in CAP-treated SN (Fig. 7,B), compared with vehicle-treated controls (Fig. 7,A). Moreover, similar to the result shown in the inset in Fig. 2,D, we found TUNEL-positive microglia in the SN at 12 h after treatment with TRPV1 agonist (Fig. 7,B, inset). By contrast, coinjection of CZP (Fig. 7,C) or I-RTX (Fig. 7 D) with CAP prevented microglial cell death, as evidenced by TL- and OX-42-positive cells (yellow).

FIGURE 7.

Double-label immunofluorescence staining with OX-42 Ab and TL on sections adjacent to those used in Fig. 6. A–D, Colocalization of TL (green) within OX-42-ip microglia (red) in the SN treated with vehicle (A), CAP + CZP (C) or CAP + I-RTX (D). Two images were merged (yellow). Note the absence of TL- and OX-42-positive microglia with only TL-positive blood vessels (BV) in the SN treated with CAP (B). Inset in B shows colocalization of OX-42 and TUNEL in the SN at 12 h after treatment with 500 pM CAP. The yellow arrowheads indicate cells double-merged with an anti-OX-42 Ab and TUNEL, and the white arrowhead indicates cell stained with TUNEL alone. Dotted lines, SNpc. Scale bar: 50 μm.

FIGURE 7.

Double-label immunofluorescence staining with OX-42 Ab and TL on sections adjacent to those used in Fig. 6. A–D, Colocalization of TL (green) within OX-42-ip microglia (red) in the SN treated with vehicle (A), CAP + CZP (C) or CAP + I-RTX (D). Two images were merged (yellow). Note the absence of TL- and OX-42-positive microglia with only TL-positive blood vessels (BV) in the SN treated with CAP (B). Inset in B shows colocalization of OX-42 and TUNEL in the SN at 12 h after treatment with 500 pM CAP. The yellow arrowheads indicate cells double-merged with an anti-OX-42 Ab and TUNEL, and the white arrowhead indicates cell stained with TUNEL alone. Dotted lines, SNpc. Scale bar: 50 μm.

Close modal

Microscopic and RT-PCR analyses previously showed that TRPV1 is widely distributed in various brain regions in rats, mice, and humans (12, 13), including the mesencephalic neurons (5), the small- and medium-sized neurons in the dorsal root and trigeminal ganglia (37, 38, 39, 40). Several lines of evidence have demonstrated that TRPV1 mediates cell death in a variety of cell types, including human neuroblastoma and lymphoma cell lines (41), TRPV1-transfected human kidney cells or sensory neurons (17), and mesencephalic neurons (5). However, little is known about the expression and functional roles of TRPV1 in microglia. To our knowledge, this is the first study to show that TRPV1 is expressed in rat and human microglia. We further demonstrated that treatment of cultured rat and human microglia with TRPV1 agonists (CAP or RTX) and intranigral injection of CAP or 12-HPETE (an endogenous agonist of TRPV1) induced cell death of microglia, as evidenced by cell viability assays, TUNEL staining, OX-42 immunostaining, and TL staining. Moreover, TRPV1 antagonists (CZP and I-RTX) were capable of inhibiting this toxicity, further indicating that TRPV1 mediated the observed microglial cell death. Interestingly, similar to the report of Shin et al. (32), a number of OX-42-ip cells with resting morphology were observed at 7 days post-TRPV1 agonists in the injured site (Fig. 6, J and K), suggesting the possible recruitment of microglia, although the underlying mechanisms are not fully understood. In contrast to microglial cell death, astrocytes were insensitive to TRPV1 agonists-induced cytotoxicity (data not shown). This is in line with our present data showing lack of TRPV1 expression in cortical astrocytes cultures (Fig. 1), although expression of this receptor has been reported in astrocytes around blood vessel in the brain (42) and in a few spinal astrocytes (43).

The present results are quite comparable to our recent data showing TRPV1-mediated neurotoxicity on mesencephalic neurons (5), while other groups found that TRPV1 actually protected against excitotoxicity in vivo (44). This apparent discrepancy (toxicity vs protection) is probably due to differences in experimental design. For example, in our recent report and the present work, we showed the direct actions of TRPV1 by applying its agonists (CAP, RTX, and 12-HPETE) and antagonists (CZP and I-RTX). In contrast, Veldhuis et al. (44) studied the actions of TRPV1 during excitotoxic injury, which might compromise the ability of TRPV1 to prevent secondary damage following excitotoxic insult.

Our previous studies have shown that CAP treatment triggers a large increase in [Ca2+]i via influx through TRPV1 in mesencephalic neurons (5) and sensory neurons (17), leading to the induction of mitochondrial damage and cell death. Other studies have reported that microglial cell death may be induced by elevation of [Ca2+]i (45, 46). Here, we formed a connection between the prior findings by showing that treatment of microglia with CAP- or RTX-increased [Ca2+]i, induced mitochondrial damage and eventually triggered cell death. In addition, CAP- or RTX-induced toxicity was attenuated by CZP, I-RTX, or BAPTA-AM in cultured microglia, indicating that the cell death was associated with increases in [Ca2+]i via TRPV1. This was also true in human microglia (HMO6), which expressed TRPV1, showed CAP- or RTX-induced cell death, and further showed inhibition of this toxicity by CZP and I-RTX (data not shown) in the presence of extracellular calcium. These findings collectively indicate that Ca2+ influx via TRPV1 contributes to rat and human microglial cell death.

The release of cytochrome c from mitochondria occurs in response to various stimuli, and is believed to play an important role in the initiation of cell death (47, 48, 49). It is well-known that cytochrome c and apoptotic protease-activating factor 1 allow for recruitment of procaspase-9 into the apoptosome complex, leading to caspase-9 activation, which in turn activates caspase-3, leading to eventual cell death (50). Accumulating data from both cell culture and animal studies have shown that caspase-3 plays an important role in microglial cell death (34, 35, 36, 51). We recently reported that cytochrome c release from damaged mitochondria and caspase-3 activation are required for TRPV1-mediated neurodegeneration in mesencephalic cultures (5). In the present study, we found that cytochrome c was released from damaged mitochondria, and caspase-3 immunoreactivity appeared within CAP- or RTX-treated microglial cultures. Conversely, treatment with a caspase-3 inhibitor reduced CAP- or RTX-induced microglial cell death. These findings are consistent with previous reports that cytochrome c release and caspase-3 activation resulted in cell death of human neuroblastoma and lymphoma cells via TRPV1 (41). However, cytochrome c release is not always required in TRPV1-mediated cell death; for example, activation of TRPV1 in Jurkat cells induced a mitochondrial Ca2+ overload that triggered mitochondrial damage, leading to cell death without cytochrome c release (52).

The data presented here are the first evidence of TRPV1-mediated microglial cell death in vitro and in vivo. In summary, in addition to our recent report of TRPV1-mediated neurotoxicity (5), the present results suggest that activation of TRPV1 may contribute to microglial damage via Ca2+ signaling and mitochondrial disruption.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by a grant from the BRC of the 21st Century Frontier Research Program of the Korea Ministry of Science (to B.K.J), from the Brain Disease Research Center/Korea Science and Engineering Foundation, and Grant R01-2005-000-10179-0 from the Basic Research Program/Korea Science and Engineering Foundation.

3

Abbreviations used in this paper: TRPV1, transient receptor potential vanilloid subtype 1; CAP, capsaicin; RTX, resiniferatoxin; 12-HPETE, 12-hydroperoxyeicosatetraenoic acid; [Ca2+]i, intracellular Ca2+; CZP, capsazepine; I-RTX, iodo-RTX; SN, substantia nigra; TH, tyrosine hydroxylase; Eth-1, ethidium homodimer-1; TL, tomato lectin; ip, immunopositive; GFAP, glial fibrillary acidic protein; calcein-AM, calcein-acetoxymethyl ester.

1
Huang, S. M., T. Bisogno, M. Trevisani, A. Al-Hayani, L. De Petrocellis, F. Fezza, M. Tognetto, T. J. Petros, J. F. Krey, C. J. Chu, et al
2002
. An endogenous capsaicin-like substance with high potency at recombinant and native vanilloid VR1 receptors.
Proc. Natl. Acad. Sci. USA
99
:
8400
-8405.
2
Ferrer-Montiel, A., C. Garcia-Martinez, C. Morenilla-Palao, N. Garcia-Sanz, A. Fernández-Carvajal, G. Fernández-Ballester, R. Planells-Cases.
2004
. Molecular architecture of the vanilloid receptor: insight for drug design.
Eur. J. Biochem.
271
:
1820
-1826.
3
Garcia-Sanz, N., A. Fernandez-Carvajal, C. Morenilla-Palao, R. Planells-Cases, E. Fajardo-Sanchez, G. Fernandez-Ballester, A. Ferrer-Montiel.
2004
. Identification of a tetramerization domain in the C terminus of the vanilloid receptor.
J. Neurosci.
24
:
5307
-5314.
4
van der Stelt, M., V. Di Marzo.
2004
. Endovanilloids: putative endogenous ligands for transient receptor potential vanilloid 1 channels.
Eur. J. Biochem.
271
:
1827
-1834.
5
Kim, S. R., Y. da Lee, E. S. Chung, U. T. Oh, S. U. Kim, B. K. Jin.
2005
. Transient receptor potential vanilloid subtype 1 mediates cell death of mesencephalic dopaminergic neurons in vivo and in vitro.
J. Neurosci.
25
:
662
-671.
6
Ohta, T., R. Komatsu, T. Imagawa, K. I. Otsuguro, S. Ito.
2005
. Molecular cloning, functional characterization of the porcine transient receptor potential V1 (pTRPV1) and pharmacological comparison with endogenous pTRPV1.
Biochem. Pharmacol.
71
:
173
-187.
7
Wu, Z. Z., S. R. Chen, H. L. Pan.
2005
. Transient receptor potential vanilloid type 1 activation down-regulates voltage-gated calcium channels through calcium-dependent calcineurin in sensory neurons.
J. Biol. Chem.
280
:
18142
-18151.
8
Raisinghani, M., R. M. Pabbidi, L. S. Premkumar.
2005
. Activation of transient receptor potential vanilloid 1 (TRPV1) by resiniferatoxin.
J. Physiol.
567
:
771
-786.
9
Hwang, S. W., H. Cho, J. Kwak, S. Y. Lee, C. J. Kang, J. Jung, S. Cho, K. H. Min, Y. G. Suh, D. Kim, U. Oh.
2000
. Direct activation of capsaicin receptors by products of lipoxygenases: endogenous capsaicin-like substances.
Proc. Natl. Acad. Sci. USA
97
:
6155
-6160.
10
Shin, J., H. Cho, S. W. Hwang, J. Jung, C. Y. Shin, S. Y. Lee, S. H. Kim, M. G. Lee, Y. H. Choi, J. Kim, et al
2002
. Bradykinin-12-lipoxygenase-VR1 signaling pathway for inflammatory hyperalgesia.
Proc. Natl. Acad. Sci. USA
99
:
10150
-10155.
11
Suh, Y. G., U. Oh.
2005
. Activation and activators of TRPV1 and their pharmaceutical implication.
Curr. Pharm. Des.
11
:
2687
-2698.
12
Mezey, E., Z. E. Toth, D. N. Cortright, M. K. Arzubi, J. E. Krause, R. Elde, A. Guo, P. M. Blumberg, A. Szallasi.
2000
. Distribution of mRNA for vanilloid receptor subtype 1 (VR1), and VR1-like immunoreactivity, in the central nervous system of the rat and human.
Proc. Natl. Acad. Sci. USA
97
:
3655
-3660.
13
Roberts, J. C., J. B. Davis., C. D. Benham.
2004
. [3H]Resiniferatoxin autoradiography in the CNS of wild-type and TRPV1 null mice defines TRPV1 (VR-1) protein distribution.
Brain Res.
995
:
176
-183.
14
Sasamura, T., M. Sasaki, C. Tohda, Y. Kuraishi.
1998
. Existence of capsaicin-sensitive glutamatergic terminals in rat hypothalamus.
Neuroreport
9
:
2045
-2048.
15
Marinelli, S., C. W. Vaughan, M. J. Christie, M. Connor.
2002
. Capsaicin activation of glutamatergic synaptic transmission in the rat locus coeruleus in vitro.
J. Physiol.
543
:
531
-540.
16
Huang, S. M., T. Bisogno, M. Trevisani, A. Al-Hayani, L. De Petrocellis, F. Fezza, M. Tognetto, T. J. Petros, J. F. Krey, C. J. Chu, et al
2002
. An endogenous capsaicin-like substance with high potency at recombinant and native vanilloid VR1 receptors.
Proc. Natl. Acad. Sci. USA
99
:
8400
-8405.
17
Shin, C. Y., J. Shin, B. M. Kim, M. H. Wang, J. H. Jang, Y. J. Surh, U. Oh.
2003
. Essential role of mitochondrial permeability transition in vanilloid receptor 1-dependent cell death of sensory neurons.
Mol. Cell. Neurosci.
24
:
57
-68.
18
Olah, Z., T. Szabo., L. Karai, C. Hough, R. D. Fields, R. M. Caudle, P. M. Blumberg, M. J. Iadarola.
2001
. Ligand-induced dynamic membrane changes and cell deletion conferred by vanilloid receptor 1.
J. Biol. Chem.
276
:
11021
-11030.
19
Marinelli, S., V. Di Marzo, N. Berretta, I. Matias, M. Maccarrone, G. Bernardi, N. B. Mercuri.
2003
. Presynaptic facilitation of glutamatergic synapses to dopaminergic neurons of the rat substantia nigra by endogenous stimulation of vanilloid receptors.
J. Neurosci.
23
:
3136
-3144.
20
Di Marzo, V., I. Lastres-Becker, T. Bisogno, L. De Petrocellis, A. Milone, J. B. Davis, J. J. Fernandez-Ruiz.
2001
. Hypolocomotor effects in rats of capsaicin and two long chain capsaicin homologues.
Eur. J. Pharmacol.
420
:
123
-131.
21
de Lago, E., R. de Miguel, I. Lastres-Becker, J. A. Ramos, J. Fernandez-Ruiz.
2004
. Involvement of vanilloid-like receptors in the effects of anandamide on motor behavior and nigrostriatal dopaminergic activity: in vivo and in vitro evidence.
Brain Res.
1007
:
152
-159.
22
Hu, D. E., A. S. Easton, P. A. Fraser.
2005
. TRPV1 activation results in disruption of the blood-brain barrier in the rat.
Br. J. Pharmacol.
146
:
576
-584.
23
Feinmark, S. J., R. Begum, E. Tsvetkov, I. Goussakov, C. D. Funk, S. A. Siegelbaum, V. Y. Bolshakov.
2003
. 12-Lipoxygenase metabolites of arachidonic acid mediate metabotropic glutamate receptor-dependent long-term depression at hippocampal CA3-CA1 synapses.
J. Neurosci.
23
:
11427
-11435.
24
Canals, S., M. J. Casarejos, S. de Bernardo, E. Rodriguez-Martin, M. A. Mena.
2003
. Nitric oxide triggers the toxicity due to glutathione depletion in midbrain cultures through 12-lipoxygenase.
J. Biol. Chem.
278
:
21542
-21549.
25
Chung, E. S., E. H. Joe, J. K. Ryu, J. Kim, Y. B. Lee, K. G. Cho, Y. J. Oh, S. H. Maeng, H. H. Baik, S. U. Kim, B. K. Jin.
2001
. GT1b ganglioside induces death of dopaminergic neurons in rat mesencephalic cultures.
Neuroreport
12
:
611
-614.
26
Lee, D. Y., Y. J. Oh, B. K. Jin.
2005
. Thrombin-activated microglia contribute to death of dopaminergic neurons in rat mesencephalic cultures: dual roles of mitogen-activated protein kinase signaling pathways.
Glia
51
:
98
-110.
27
Giulian, D., T. J. Baker.
1986
. Characterization of ameboid microglia isolated from developing mammalian brain.
J. Neurosci.
6
:
2163
-2178.
28
Nagai, A., E. Nakagawa, K. Hatori, H. B. Choi, J. G. McLarnon, M. A. Lee, S. U. Kim.
2001
. Generation and characterization of immortalized human microglial cell lines: expression of cytokines and chemokines.
Neurobiol. Dis.
8
:
1057
-1068.
29
Choi, S. H., E. H. Joe, S. U. Kim, B. K. Jin.
2003
. Thrombin-induced microglial activation produces degeneration of nigral dopaminergic neurons in vivo.
J. Neurosci.
23
:
5877
-5886.
30
Jung, J., S. Y. Lee, S. W. Hwang, H. Cho, J. Shin, Y. S. Kang, S. Kim, U. Oh.
2002
. Agonist recognition sites in the cytosolic tails of vanilloid receptor 1.
J. Biol. Chem.
277
:
44448
-44454.
31
Paxinos, G., C. Watson.
1998
.
The Rat Brain in Stereotaxic Coordinates
Academic Press, San Diego.
32
Shin, W. H., D. Y. Lee, K. W. Park, S. U. Kim, M. S. Yang, E. H. Joe, B. K. Jin.
2004
. Microglia expressing interleukin-13 undergo cell death and contribute to neuronal survival in vivo.
Glia
46
:
142
-152.
33
Marshall, I. C., D. E. Owen, T. V. Cripps, J. B. Davis, S. McNulty, D. Smart.
2003
. Activation of vanilloid receptor 1 by resiniferatoxin mobilizes calcium from inositol 1,4,5-trisphosphate-sensitive stores.
Br. J. Pharmacol.
138
:
172
-176.
34
Ryu, J. K., A. Nagai, J. Kim, M. C. Lee, J. G. McLarnon, S. U. Kim.
2003
. Microglial activation and cell death induced by the mitochondrial toxin 3-nitropropionic acid: in vitro and in vivo studies.
Neurobiol. Dis.
12
:
121
-132.
35
Jung, D. Y., H. Lee, B. Y. Jung, J. Ock, M. S. Lee, W. H. Lee, K. Suk.
2005
. TLR4, but not TLR2, signals autoregulatory apoptosis of cultured microglia: a critical role of IFN-β as a decision maker.
J. Immunol.
174
:
6467
-6476.
36
Hou, R. C., C. C. Wu, J. R. Huang, Y. S. Chen, K. C. Jeng.
2005
. Oxidative toxicity in BV-2 microglia cells: sesamolin neuroprotection of H2O2 injury involving activation of p38 mitogen-activated protein kinase.
Ann. NY Acad. Sci.
1042
:
279
-285.
37
Guo, A., L. Vulchanova, J. Wang, X. Li, R. Elde.
1999
. Immunocytochemical localization of the vanilloid receptor 1 (VR1): relationship to neuropeptides, the P2X3 purinoceptor and IB4 binding sites.
Eur. J. Neurosci.
11
:
946
-958.
38
Ichikawa, H., T. Sugimoto.
2001
. VR1-immunoreactive primary sensory neurons in the rat trigeminal ganglion.
Brain Res.
890
:
184
-188.
39
Valtschanoff, J. G., A. Rustioni, A. Guo, S. J. Hwang.
2001
. Vanilloid receptor VR1 is both presynaptic and postsynaptic in the superficial laminae of the rat dorsal horn.
J. Comp. Neurol.
436
:
225
-235.
40
Hou, M., R. Uddman, J. Tajti, M. Kanje, L. Edvinsson.
2002
. Capsaicin receptor immunoreactivity in the human trigeminal ganglion.
Neurosci. Lett.
330
:
223
-226.
41
Maccarrone, M., T. Lorenzon, M. Bari, G. Melino, A. Finazzi-Agro.
2000
. Anandamide induces apoptosis in human cells via vanilloid receptors: evidence for a protective role of cannabinoid receptors.
J. Biol. Chem.
275
:
31938
-31945.
42
Toth, A., J. Boczan, N. Kedei, E. Lizanecz, Z. Bagi, Z. Papp, I. Edes, L. Csiba, P. M. Blumberg.
2005
. Expression and distribution of vanilloid receptor 1 (TRPV1) in the adult rat brain.
Brain Res. Mol. Brain Res.
135
:
162
-168.
43
Doly, S., J. Fischer, C. Salio, M. Conrath.
2004
. The vanilloid receptor-1 is expressed in rat spinal dorsal horn astrocytes.
Neurosci. Lett.
357
:
123
-126.
44
Veldhuis, W. B., M. van der Stelt, M. W. Wadman, G. van Zadelhoff, M. Maccarrone, F. Fezza, G. A. Veldink, J. F. Vliegenthart, P. R. Bar, K. Nicolay, V. Di Marzo.
2003
. Neuroprotection by the endogenous cannabinoid anandamide and arvanil against in vivo excitotoxicity in the rat: role of vanilloid receptors and lipoxygenases.
J. Neurosci.
23
:
4127
-4133.
45
Braun, J. S., J. E. Sublett, D. Freyer, T. J. Mitchell, J. L. Cleveland, E. I. Tuomanen, J. R. Weber.
2002
. Pneumococcal pneumolysin and H2O2 mediate brain cell apoptosis during meningitis.
J. Clin. Invest.
109
:
19
-27.
46
Nagano, T., S. H. Kimura, E. Takai, T. Matsuda, M. Takemura.
2006
. Lipopolysaccharide sensitizes microglia toward Ca2+-induced cell death: mode of cell death shifts from apoptosis to necrosis.
Glia
53
:
67
-73.
47
Liu, X., C. N. Kim, J. Yang, R. Jemmerson, X. Wang.
1996
. Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c.
Cell
86
:
147
-157.
48
Li, P., D. Nijhawan, I. Budihardjo, S. M. Srinivasula, M. Ahmad, E. S. Alnemri, X. Wang.
1997
. Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade.
Cell
91
:
479
-489.
49
Zou, H., Y. Li, X. Liu, X. Wang.
1999
. An APAF-1-cytochrome c multimeric complex is a functional apoptosome that activates procaspase-9.
J. Biol. Chem.
274
:
11549
-11556.
50
Hengartner, M. O..
2000
. The biochemistry of apoptosis.
Nature
407
:
770
-776.
51
Lee, P., J. Lee, S. Kim, M. S. Lee, H. Yagita, S. Y. Kim, H. Kim, K. Suk.
2001
. NO as an autocrine mediator in the apoptosis of activated microglial cells: correlation between activation and apoptosis of microglial cells.
Brain Res.
892
:
380
-385.
52
Jambrina, E., R. Alonso, M. Alcalde, M. del Carmen Rodriguezqq, A. Serrano, C. Martinez-A, J. Garcia-Sancho, M. Izquierdo.
2003
. Calcium influx through receptor-operated channel induces mitochondria-triggered paraptotic cell death.
J. Biol. Chem.
278
:
14134
-14145.