|Home | About | Journals | Submit | Contact Us | Français|
The heat and capsaicin receptor, TRPV1, is required for the detection of painful heat by primary afferent pain fibers (nociceptors), but the extent to which functional TRPV1 channels are expressed in the central nervous system (CNS) is debated. As previous evidence is based largely on indirect physiological responses to capsaicin, here we genetically modified the TrpV1 locus to reveal, with excellent sensitivity and specificity, the distribution of TRPV1 in all neuronal and non-neuronal tissues. In contrast to reports of widespread and robust expression in the CNS, we find that neuronal TRPV1 is largely restricted to nociceptors in primary sensory ganglia, with minimal expression in a few discrete brain regions, most notably in a contiguous band of cells within and adjacent to the caudal hypothalamus. We confirm hypothalamic expression in the mouse using several complementary approaches, including in situ hybridization, calcium imaging, and electrophysiological recordings. Additional in situ hybridization experiments in rat, monkey and human brain demonstrate that the restricted expression of TRPV1 in the CNS is conserved across species. Outside of the CNS, we find TRPV1 expression in a subset of arteriolar smooth muscle cells within thermoregulatory tissues. Here, capsaicin increases calcium uptake and induces vasoconstriction, an effect that likely counteracts the vasodilation produced by activation of neuronal TRPV1.
TRPV1 is a non-selective cation channel that is activated by noxious heat, protons, the vanilloid compounds capsaicin (CAP) and resiniferatoxin (RTX), and membrane-derived lipids, including anandamide (Caterina et al., 1997; Caterina and Julius, 2001). Studies of Trpv1 knockout mice demonstrated a critical contribution of TRPV1 to the cellular and behavioral responses to noxious heat (Caterina et al., 2000; Davis et al., 2000). Although expression of TRPV1 was originally reported to be restricted to primary afferent nociceptors of the dorsal root (DRG), trigeminal (TG) and nodose ganglia (NG) (Szallasi et al., 1995; Caterina et al., 1997; Tominaga et al., 1998), subsequent studies argued for a much wider distribution, both in the CNS and in non-neuronal tissues.
Expression of TRPV1 in the CNS has been reported using a variety of methods, including pharmacological characterization (Steenland et al., 2006), immunohistochemistry (Toth et al., 2005; Cristino et al., 2006), in situ hybridization (Mezey et al., 2000), radioligand binding (Acs et al., 1996; Roberts et al., 2004), and RT-PCR (Mezey et al., 2000). Despite this abundance of studies, the existence of TRPV1 in the brain remains a controversial topic, largely because the extent and localization of TRPV1 varies considerably among studies, even those using very similar assays. For example, RTX binding studies reported results ranging from a lack of binding in the CNS (Szallasi et al., 1995), to widespread binding throughout the brain (Roberts et al., 2004). In part, the lack of consensus reflects the limitations of traditional approaches to determining gene expression, including variable sensitivity, poor signal to noise, and lack of specificity.
There is also controversy as to the functional contribution of TRPV1 in the CNS. For example, several groups reported alterations in hippocampal synaptic function in Trpv1 knockout mice (Marsch et al., 2007; Gibson et al., 2008). However, these reports did not provide anatomical or biophysical evidence of TRPV1 expression in these cells. Furthermore, other groups found no effect of TRPV1 agonists on hippocampal synaptic function (Kofalvi et al., 2006; Benninger et al., 2008). These results illustrate the importance of definitively resolving the distribution of TRPV1. This is especially true because TRPV1 expression in the CNS has important implications for the potential side effects of TRPV1 agonists and antagonists being developed for the treatment of chronic pain.
Here we used gene targeting to modify the Trpv1 genetic locus and generated two lines of reporter mice, which allow for a highly sensitive readout of TRPV1 expression patterns. Consistent with our early reports(Tominaga et al., 1998), we demonstrate robust expression of TRPV1 in primary afferent neurons. However, with the exception of very low-level expression in a few discrete brain areas, most notably within and adjacent to the caudal hypothalamus, we could not confirm previous findings of a widespread TRPV1 distribution in the CNS. Calcium imaging, whole-cell recording and in situ hybridization experiments demonstrated that the expression patterns revealed in the reporter mice reflected functional TRPV1 expression that is conserved across multiple mammalian species. Finally, the sensitivity and cellular resolution of our genetic marking strategy revealed functional TRPV1 expression in a subset of arteriolar smooth muscle cells in thermoregulatory tissues.
Animal experiments were approved by the Institutional Animal Care and Use Committee and conducted in accordance with the NIH Guide for the Care and Use of Laboratory Animals and the recommendations of the International Association for the Study of Pain.
A genomic clone containing the last exon of the Trpv1 gene (Wellcome Trust Sanger Institute; Geneservice, Ltd.) was used to generate a TRPV1 targeting vector. We used site-directed mutagenesis to engineer an AscI site 3bp downstream of the TRPV1 stop codon, and constructs containing IRES-PLAP-IRES-nlacZ (TRPV1PLAP-nlacZ mice) or IRES-mycCre (TRPV1Cre mice) were inserted into this targeting vector. Homologous recombinant ES clones were used in blastocyst injections to obtain chimeric mice. R26R-lacZ (Soriano, 1999), R26R-YFP (Srinivas et al., 2001), and Trpv1 knockout (Caterina et al., 2000) were described previously.
Intrathecal injections were performed as previously described (Cavanaugh et al., 2009). For RTX injections, mice were anesthetized with 1.5% isoflurane, and injected subcutaneously with escalating doses of RTX on consecutive days (30 mg/ml, 70 mg/ml, 100 mg/ml, and 200 mg/ml), followed 7 d later with 200 mg/ml. Histology was performed 7–10 d following this final RTX injection.
Adult mice (6–8 weeks old) were perfused with 10mL Hepes-buffered saline (HBS) followed by 20mL of ice cold 3.7% formaldehyde. Brain, SC, peripheral tissue, DRG and TG were dissected out, post-fixed 3–4 (for PLAP staining) or 1.5 (for nlacZ staining) hr at 4°C and cryoprotected overnight in 30% sucrose. For brain and SC, 40–80μm sections were processed as free-floating sections. For DRG, TG, and tongue, 14μm cryostat sections were processed on slides. Ear, dura, and cremaster muscle were processed as a whole mounts. PLAP and nlacZ staining was carried out as previously described (Shah et al., 2004).
Following nlacZ histochemistry, tissue was processed for immunohistochemistry as previously described (Tominaga et al., 1998). Primary antisera were as follows: guinea pig anti-TRPV1 (1:1000; Julius Lab, UCSF), mouse anti-reelin (1:500; Chemicon), mouse anti-SMA, (1:800; Sigma).
Adult mice were perfused with 10 mL PBS followed by 10 mL RNA later (Ambion). TG, liver, bladder, and cremaster were dissected out and placed in RNA later on ice. For brain regions, 500 μm brain sections were cut with a McIlwain Tissue Chopper into ice cold PBS, and brain regions were microdissected out with sharp tissue forceps and put into RNA later on ice. To avoid contamination, separate forceps were used for dissection of each tissue/brain region.
RNA isolation was performed with TRIzol reagent (Sigma) according to the manufacturers specifications. 1 μg of each RNA was reverse transcribed with SuperScript II Reverse Strand Synthesis System (Invitrogen) using an oligo (dT) primer. Control reactions omitted reverse transcriptase. PCR amplification (94°C for 5 min; 35 cycles of 94°C for 30 s, 58°C for 30 s, 72°C for 1 min; 72°C for 10 min) was performed with 1 μl of cDNA using the following primer pair: 5′-gtttgtggacagctacagtg-3 ′ and 5 ′-gaagccacatactccttgcg-3′. Control PCR amplification (94°C for 5 min; 25 cycles of 94°C for 30 s, 58°C for 30 s, 72°C for 1 min; 72°C for 10 min) was performed with a GAPDH primer pair (5′-accacagtccatgccatcac-3′ and 5′-tccaccaccctgttgctgta-3′) to ensure cDNA integrity. Amplification products were visualized on a 1% agarose gel with ethidium bromide and sequenced to confirm identity.
For in situ hybridization histochemistry (ISHH), mice were killed by decapitation under deep anesthesia. Brain and L4/5 DRGs were dissected out, rapidly frozen in powdered dry ice, and cut on a cryostat (5 μm for DRG and 10μm for brain). Sections were thaw-mounted onto slides and fixed in 4% formaldehyde (pH 7.4) for 20 minutes. ISSH was performed as described previously (Kobayashi et al., 2005) with an 35S-labeled cRNA probe directed against bases 1,155–1,908 of the full-length TRPV1 cDNA (antisense or sense; 1×107 cpm/ml). For autoradiography, sections were coated with NTB emulsion (Eastman Kodak, Rochester, NY) diluted 1:1 with distilled water at 45°C and exposed for 2 months in light-tight boxes at 4°C. After development in D-19 (Eastman Kodak) and fixation in 24% sodium thiosulfate, the sections were stained with hematoxylin-eosin (H&E), air dried, cleared in xylene, and coverslipped.
For combined IHC with ISHH, mice were deeply anesthetized with sodium pentobarbital and perfused transcardially with 20 ml of PB, followed by 50 ml of 4% formaldehyde. DRGs were dissected outand postfixed in the same fixative at 4°C O/N, followed by immersion in 20% sucrose in PB at 4°C for 3 days. Tissue was frozen in powdered dry ice and cut on a cryostat at a 5 μm thickness. Sections were preincubated in TBS containing 5% normal goat serum (NGS) for 30 minutes, followed by incubation in guinea pig anti-TRPV1 (1:1000; Julius Lab, UCSF) in TBS containing 5% NGS for 24 hr at 4°C, washed in TBS and then processed for DAB immunohistochemistry, as described (Tominaga et al., 1998). Sections were then fixed in 4% formaldehyde for 5 min, and immediately processed for ISHH, as described above, but emulsion was stopped after 3 weeks.
Frozen human tissues (with postmortem delay of 7 hrs) were acquired from the Brain and Tissue Bank for Developmental Disorders (University of Maryland, Baltimore, MD, USA). Monkey tissues were obtained from an adult male Macaca Fascicularis (ITR Laboratories Canada, Montreal, Canada). Rat brain was acquired from adult male Sprague–Dawley rats (Charles River, Saint-Constant, Quebec, Canada).
Brains were rapidly dissected, snap-frozen at −40°C in isopentane for 20 s and stored at −80°C. Frozen tissues were cryosectioned at 14 μm and mounted onto Superfrost Plus slides (VWR Cananda, Montreal, Quebec, Canada). Slides were stored at −80°C until further use.
Species specific antisense riboprobes for rat, mokey and human TRPV1 were transcribed in vitro using SP6 RNA polymerase (Promega) and radiolabeled with 35S-UTP and 35S-CTP (800 Ci/mmol; Amersham Biosciences, Inc). Following transcription, the TRPV1 DNA templates were digested with DNAse I (Promega) and subsequently purified using G-50 Sepharose microspin columns (GE Healthcare). The quality of labeled riboprobes was verified by polyacrylamide-urea gel electrophoresis and scintillation counting.
For ISHH, tissue sections were fixed with 4% paraformaldehyde, rinsed 3 times in 2× standard sodium citrate buffer (2× SSC), equilibrated in 0.1 M triethanolamine (TEA), and treated with 0.25% acetic anhydride in 0.1 M TEA. After a rinse in 2× SSC and dehydration through an ethanol series (50–100%), hybridization was performed in buffer containing 75% formamide (Sigma), 600 mM NaCl, 10 mM Tris-HCl (pH 7.5), 1 mM EDTA, 1× Denhardt’s solution (Sigma), 50 μg/ml denatured salmon sperm DNA (Sigma), 10% dextran sulfate (Sigma), 20 mM dithiothreitol, and [35S]-labeled cRNA probe (20 × 106 cpm/ml) at 55°C overnight in chambers humidified with 75% formamide. Following hybridization, slides were rinsed twice in 2× SSC at room temperature, treated with 20 μg/ml RNase IA in RNase buffer (25 mM NaCl, 5 mM Tris-HCl pH 7.5, 0.5 mM EDTA pH 8.0) for 45 minutes at 37°C, and washed to a final stringency of 0.1× SSC at 70°C. Sections were then dehydrated and exposed to Kodak Biomax MR-2 film. After exposure to film, the slides were dipped in Kodak NTB2 emulsion and exposed for 8 weeks at 4°C prior to development and counterstaining with H&E. Film autoradiograms were digitized with a high-resolution Xillix Microimager digital camera via the MCID image analysis system (Imaging Research, St-Catharines, Ontario, Canada). Bright- and darkfield photomicrographs of emulsion-dipped tissue sections were captured using a Leica (DMRBE/DM 4000B) microscope equipped with a Leica DFC490 digital camera. Digital images were transferred to Adobe Photoshop 7.0 for minimal image processing.
DRG neurons were isolated from 6 week old mice, dissociated and subjected to ratiometric calcium imaging the next day as previously described (Caterina et al., 2000). Cells were stimulated with 1 μM CAP, followed by 150mM K+ (HiK) Ringer’s Solution, then fixed and stained for nlacZ histochemistry. All neurons that responded to HiK solution were included for analysis.
200 μm coronal slices of caudal hypothalamus were cut from TRPV1Cre/R26R-EYFP mice, aged P21-26, using a DTK-100 Microslicer (Ted Pella, Inc.). Slices were cut in an ice-cold, high-sucrose solution consisting of (mM): NaCl 87, NaH2PO4 1.25, NaHCO3 25, KCl 2.5, CaCl2 0.5, MgCl2 7, glucose 25 and sucrose 75, saturated with 95% O2/5% CO2. Freshly cut slices were placed in an incubating chamber containing artificial cerebrospinal fluid (ACSF), containing (mM): NaCl 125, KCl 2.5, NaHCO3 25, Na2PO4 1.25, glucose 15, CaCl2 2, MgCl2 1, saturated with 95% O2/5% CO2, and allowed to recover at 35°C for ~30 min, then 0.5 to 1 hr at room temperature.
Slices were loaded with 10 μm Fura-2-AM (Molecular Probes) and 0.2% pluronic acid at 22–25 °C for 60 m in ACSF, saturated with 95% O2/5% CO2. For imaging, slices were perfused with ACSF, saturated with 95% O2/5% CO2 and containing TTX (500 nM), picrotoxin (100 μM), APV (100 μM) and NBQX (10 μM). CAP (1–10 μM) and HiK solution (140mM) were added to the same supplemented ACSF solution and applied using a bath perfusion system. Image acquisition and analysis were performed as detailed above for DRG neurons.
Following recovery, slices were transferred to a submersion chamber on an upright Olympus BX51 microscope, and perfused with ACSF, saturated with 95% O2/5% CO2. EYFP+ cells in the supramammillary nucleus and posterior hypothalamus were identified by epifluorescence microscopy and recorded with 3- to 5-MΩ borosilicate glass pipettes, filled with an internal solution consisting of (mM): CsMeSO3 140, HEPES 10, EGTA 10, NaCl 2, Mg-ATP 2, QX-314 1, TEA-Cl 5 and CaCl2 1, pH 7.3. Following the formation of a seal and achieving whole-cell configuration, membrane potential was held in voltage-clamp mode at −60 mV, and stable baseline holding current was established. The bath solution was then switched to a control ACSF solution containing TTX (500 nM), picrotoxin (100 μM), APV (100 μM) and NBQX (10 μM) for several min, followed by ACSF containing 10 μM CAP for ~5 min, and then back to ACSF to wash out responses. In some cases, 20 μM Ruthenium Red (RR) (final concentration) was added directly to the bath.
Coronal slices of caudal hypothalamus were cut as described and transferred to a dissociation solution consisting of 2.5 mg/ml protease XXIII (Sigma) dissolved in the sucrose cutting solution. Slices were incubated at 35°C for 10 min, rinsed several times in cold sucrose solution, and placed in cold (4°C) sucrose solution containing 1 mg/ml trypsin inhibitor (Sigma) and 1 mg/ml bovine serum albumin (Sigma). The SuM was isolated with iridectomy scissors and gently triturated using a series of fine-bore Pasteur pipettes. Dissociated neurons were imaged as described for DRG neurons.
200 μm coronal slices of P19 hippocampus were cut from wildtype mice as detailed above for caudal hypothalamic slices. Slices were loaded with 100 μm Fura-2-AM (Molecular Probes) and 0.2% pluronic acid at 22–25 °C for 60 m in ACSF, saturated with 95% O2/5% CO2, and calcium imaging was performed as detailed above for hypothalamic slices.
Third order arterioles were dissected out of ears, bathed in ice cold physiological salt solution consisting of (mM): NaCl 137, KCl 5.6, MgCl2 1, Na2HPO4 .42, NaH2P04 .44, 4.2 NaHCO3 4.2, HEPES 10, 1 mg/ml BSA, pH 7.4, cut into ~2mm pieces and cultured in poly-D-lysine coated wells in low-glucose DMEM with 10% fetal bovine serum and pen–strep. The next day, arterioles were subjected to ratiometric calcium imaging as described above. Cells were stimulated with 1 μM and 10 μM CAP, followed by 10 μM RR to block responses, and finally with HiK+ Ringer’s Solution.
We used gene targeting to create a line of mice (TRPV1PLAP-nlacZ) in which placental alkaline phosphatase (PLAP) and nuclear lacZ (nlacZ), are expressed under the control of the TRPV1 promoter. We inserted an IRES-PLAP-IRES-nlacZ cassette immediately 3′ of the TRPV1 stop codon, which permits the transcription and translation of PLAP and nlacZ in cells expressing TRPV1, without disrupting the Trpv1 coding region (Fig. 1A). In addition to providing a very sensitive detection method, background levels of PLAP and nlacZ are negligible in the mouse (Shah et al., 2004), which allows for discrete localization of TRPV1.
As expected, TRPV1PLAP-nlacZ mice robustly expressed nlacZ in nuclei of primary afferent neurons of the DRG and TG (Fig. 1B). PLAP histochemistry labeled both cell bodies and axonal processes in the DRG (Fig. 1C), as well as primary afferent terminals of the spinal cord (SC) dorsal horn (Fig. 1D). The SC staining was eliminated by intrathecal injection of high-dose CAP (not shown), which selectively destroys TRPV1 fibers (Cavanaugh et al., 2009), indicating that the PLAP staining indeed arises from TRPV1 nociceptors.
To establish the specificity of nlacZ expression for TRPV1 neurons, we investigated the CAP-responsiveness of nlacZ+ DRG neurons cultured from adult TRPV1PLAP-nlacZ mice, using live-cell calcium imaging (Fig. 1F-J). Importantly, we found that >99% of nlacZ+ neurons responded to 1 μM CAP, and >90% of CAP-responsive neurons were nlacZ+ (n = 568). We also compared the expression of nlacZ and TRPV1 in DRG sections (Fig. 1E). Consistent with the calcium imaging results, we found that >95% of TRPV1-immunoreactive neurons were nlacZ+. These results demonstrate that nlacZ histochemistry provides a sensitive and accurate correlate of functional TRPV1 expression, in vivo as well as in vitro.
We found no PLAP+ cell bodies in the brain of TRPV1PLAP-nlacZ mice. We did, however, find limited nlacZ expression in a few brain areas, primarily along midline structures of the rostral midbrain and caudal hypothalamus (Fig. 2). Prominent among these brain areas were the intrafascicular (IF) (Fig. 2A) and supramammillary nuclei (SuM) (Fig. 2D), as well as the dorsomedial (DMH) and posterior hypothalamus (PH) (Fig. 2G). We also detected scattered nlacZ+ cells in the entorhinal cortex (Fig. 2B), the rostral linear raphe nucleus (RLi) (Fig. 2C) and the periaqueductal gray (PAG) (Fig. 2E). Finally, nlacZ expression marked a discrete subset of (non pyramidal) hippocampal neurons (Fig. 2F) that co-labeled for reelin, a marker of Cajal-Retzius cells (Fig. 2F, inset), as well as reelin+ cells in the accessory olfactory bulb (AOB) (Fig. 2H). It is likely that our failure to detect PLAP in these brain regions reflects the greater sensitivity of nlacZ versus PLAP histochemistry.
We used several complementary approaches to verify the expression pattern revealed in the Trpv1 knock-in mice. We could not detect staining in any of these brain areas using TRPV1 antisera, most likely because TRPV1 is expressed at levels below the detection threshold of immunohistochemistry. Furthermore, with the exception of the caudal hypothalamus (which included the SuM and IF), we failed to detect TRPV1 mRNA by RT-PCR in any of the brain regions where we saw nlacZ expression (Fig. 5J). Finally, radioactive in situs confirmed the restricted expression suggested by the Trpv1 reporter mice, with positive signals in the IF, SuM, DMH and PH (Fig. 3M-O). Together with the lack of PLAP staining, these results indicate that TRPV1 in the brain is limited to very low-level expression in a small subset of cells of a few discrete brain regions.
The distribution of TRPV1 in the DRG of the mouse differs from that of the rat (Woodbury et al., 2004), which suggests that brain expression patterns might also differ across species. To investigate this, we performed radioactive in situ hybridization in several species, including rat, monkey, and human. Importantly, using several different high affinity probes directed against rat TRPV1, we found a near identical pattern of CNS staining compared to that obtained in the mouse. Most notably, specific signals were found in caudal hypothalamic regions, including the PH, SuM, and IF (Figure 3E-L). Positive signals were also found in the dorsal motor nucleus of the vagus (DMV), mesencephalic trigeminal nucleus, parabrachial nucleus, rostral linear raphe nucleus, and lateral hypothalamus of the rat (not shown). In monkey (not shown) and human brain sections (Fig. 3R-S), we could not detect any TRPV1+ signal, however, we did not have access to sections that included the areas of the caudal hypothalamus that had positive signals in rat and mouse. Nevertheless, the lack of signal in other brain areas, despite the strong specific signal in DRG neurons (Fig. 3Q), is consistent with the highly restricted pattern we observed in mouse and rat, thus demonstrating that this property is conserved across multiple mammalian species.
As this highly restricted CNS expression pattern contrasted with reports of widespread TRPV1 distribution, we created a second line of reporter mice (TRPV1Cre mice), in which Cre recombinase is expressed under the control of the TRPV1 promoter (Figure 4A). Crosses of this line with Cre-dependent reporter lines provide a highly sensitive fate map of TRPV1 expression, which reveals all loci of TRPV1 expression, no matter how transient. When we crossed TRPV1Cre mice with the Rosa-lacZ reporter line (TRPV1Cre/R26R-lacZ) (Soriano, 1999), we observed the expected pattern of DRG and SC staining (Fig. 4B, C).
Importantly, all the nlacZ+ brain areas in the TRPV1PLAP-nlacZ mice were also stained by lacZ in the TRPV1Cre/R26R-lacZ mice (Fig. 4D-K). The staining in TRPV1Cre/R26R-lacZ mice was more widespread than that in TRPV1PLAP-nlacZ mice, with the exception of the reelin+ cells of the hippocampus and AOB, likely because expression of the Rosa locus is limited in these cells (not shown). The more extensive labeling in the TRPV1Cre/R26R-lacZ mice compared to TRPV1PLAP-nlacZ shows that the cellular distribution of TRPV1 in these brain areas undergoes a developmental restriction. We additionally noted several regions in which expression is observed in the TRPV1Cre/R26R-lacZ mice, but not in the TRPV1PLAP-nlacZ mice (Fig. 5). These are likely areas where TRPV1 is transiently expressed during the embryonic or early postnatal periods. However, because only a few molecules of Cre are necessary to induce recombination, it is possible that some of these areas represent regions where TRPV1 is expressed at extremely low levels in the adult, below the detection threshold of the TRPV1PLAP-nlacZ mice. This could be the case for the DMV, where we observed in situ signals in both the mouse and rat despite a lack of staining in sections from TRPV1PLAP-nlacZ mice (Fig. 3.H, L and P).
As lacZ expression in the IF, SuM, DMH and PH of reporter mice was confirmed by RT-PCR and radioactive in situ, we sought to validate functional TRPV1 expression in these brain areas using calcium-imaging and whole-cell recordings in brain slices from TRPV1Cre mice crossed to EYFP reporter mice (TRPV1Cre/R26R-YFP mice) (Srinivas et al., 2001). Importantly, EYFP staining in these mice was identical to lacZ expression in TRPV1Cre/R26R mice (data not shown).
We carried out calcium-imaging experiments in caudal hypothalamic neurons in intact slices (Fig. 6A-D) and acutely dissociated cells (Fig. 6E-H). We consistently observed calcium responses in EYFP+ cells of hypothalamic slices in response to bath application of 10 μM CAP, in the presence of the fast synaptic transmission blockers picrotoxin, NBQX, and APV, and the voltage-gated sodium channel blocker TTX. In acutely dissociated cells, CAP elicited robust responses at doses as low as 100 nM. CAP responses were blocked by pre-incubation with 10 μM Ruthenium Red (RR) a non-selective antagonist that blocks many calcium-permeant ion channels, including TRPV1 (Fig. 6J), and were absent in Trpv1 knockout mice (data not shown). CAP-induced calcium increases were limited to EYFP+ cells; ~97% (30/31) of CAP-responsive cells were EYFP+. However, only ~59% (30/51) EYFP+ cells tested were CAP-responsive, consistent with a developmental downregulation of TRPV1.
Finally, we carried out whole-cell patch-clamp recordings from acute hypothalamic slices, containing the SuM and PH, prepared from P21-26 TRPV1Cre/R26R-YFP mice. We performed voltage-clamp recordings from visually identified EYFP+ neurons, held at a membrane potential of −60 mV, while bath applying 10 μM CAP in the presence of TTX and synaptic blockers. We found that ~53% (10/19) of EYFP+ neurons responded to CAP with inward current greater than −5 pA; the average amplitude was −52.9 ± 14 pA (n=10) (Fig. 6K). CAP responses generally desensitized during application, slowly reversed following washout and could be blocked by RR (not shown).
As a further test of the accuracy and selectivity of our reporter mice, we performed calcium imaging experiments in regions of the hippocampus where others have argued for a functional contribution of TRPV1 (Marsch et al., 2007; Gibson et al., 2008, Chavez et al., 2010), but where we see no anatomical evidence for TRPV1 expression. In intact slices of the dentate gyrus, we never observed calcium increases in response to administration of 10 μM capsaicin (Fig. 6L-O). We also failed to detect capsaicin responses in the CA1 region of the hippocampus (not shown), although pyramidal cell loading in these preparations was sub-optimal. However, we did not observe capsaicin-induced calcium increases in E19 rat hippocampal cultures, which included CA1 and CA3 pyramidal cells (not shown). These results are consistent with our anatomical data, and demonstrate that the lack of reporter molecule expression parallels what, in our hands, is a functional absence of TRPV1.
Unexpectedly, when examining peripheral tissue for PLAP+ axons, we observed discrete, striated staining running in proximity to blood vessels (Fig. 7A, C, E and F). We found comparable patterns of nlacZ staining (Fig. 7B, D and G), demonstrating that the PLAP derived from TRPV1+ cells that are intrinsic to these tissues. In agreement with this observation, RT-PCR analysis revealed TRPV1 mRNA in cremaster muscle, which has extensive nlacZ+ labeling (Fig. 5J). Furthermore, the nlacZ staining co-localized with immunostaining for alpha smooth muscle actin (SMA), a filament that is specifically associated with smooth muscle cells (SMCs) (Fig. 7D).
The SMC label was not ubiquitous, but was concentrated in thermoregulatory tissues, including the cremaster muscle (Fig. 7A, B), dura (Fig 7C), tongue (Fig. 7D), trachea (Fig. 7E), skin (Fig. 7F), and ear (Fig. 7G). By contrast, we found minimal to no SMC labeling in many other organs, including the liver, lung, kidney, pancreas, SC, brain, and aorta. SMC expression was restricted to certain small to medium diameter vessels (>100 μm in diameter), which, given their size and location, are most likely arterioles. Importantly, the SMC label was substantially reduced following systemic RTX, which kills TRPV1+ cells (Fig. 7G, H). In addition, we saw similar patterns of blood vessel staining in TRPV1Cre/R26R-lacZ mice. (Fig. 7I and J).
As a further test of the functionality of the TRPV1 expression in SMCs, we performed live-cell calcium-imaging (Fig. 8A-F). Pieces of arteriole were dissected out of the ear and cultured overnight to ensure that any vessel-associated nerves were rendered non-functional during imaging. CAP (1 and 10 μM), evoked arteriole constriction that was accompanied by increases in intracellular calcium (Fig. 8C, G and I). Importantly, RR (10 μM) reversed the calcium responses (Fig. 8G and I), and neither calcium influx nor constriction was recorded in Trpv1 knockout mice (Fig. 8D, H, I). We conclude that TRPV1 is functional and mediates vessel constriction in arteriolar SMCs.
To unequivocally resolve the disagreement as to the extent of TRPV1 expression outside of the primary afferent nociceptor, we used a genetic approach to label TRPV1+ cells with excellent sensitivity and precision. In contrast to previous reports, we conclude that TRPV1 in the CNS is limited to very low-level expression in a few discrete brain regions. Most prominent among these is a contiguous band of cells centered along the midline of the posterior hypothalamus and rostral midbrain. We confirmed expression using a number of independent techniques, including RT-PCR, radioactive in situ hybridization, calcium-imaging, and slice recording, thus providing multifaceted evidence that nlacZ in these brain areas represents bona fide TRPV1 expression.
We also performed in situ hybridization in rat, primate and human tissue, and found that the restricted expression pattern revealed by the Trpv1 reporter mice was conserved across these species. Notably, the areas of the caudal hypothalamus that have TRPV1+ in situ hybridization signals in both the mouse and rat are the only areas in the rat brain where injection of a high dose of CAP induces cell body degeneration (Ritter and Dinh, 1988), which demonstrates that these signals reflect functional TRPV1 expression in the rat, as we have shown for the mouse. The physiological function of TRPV1 in these brain areas is unclear, however, given the proximity of these regions to the third ventricle, it is possible that TRPV1 could detect changes in CSF temperature.
Several lines of evidence suggest that TRPV1 is present at extremely low levels in these brain areas. First, we were unable to detect TRPV1 by immunohistochemistry or PLAP staining. Second, we detected only a very faint TRPV1 band with by RT-PCR analysis, and needed an extended (2 month) exposure period to observe signals with radioactive in situ. Finally, the average currents elicited by CAP in caudal hypothalamic neurons are roughly 40 times smaller than those observed in DRG neurons (Caterina et al., 2000). Based on the single channel conductance of TRPV1 at −60 mV (Tominaga et al., 1998), we estimate that the smallest CAP-induced currents (−5 pA) that we observed in EYFP+ cells of the caudal hypothalamus arise from just a few functional channels, highlighting the remarkable sensitivity of the Trpv1 knock-in mice.
Several groups used Trpv1 knockout mice to suggest a functional contribution of this channel outside of the DRG, including urothelial cells of the bladder (Birder et al., 2002), osmosensitive neurons of the supraoptic and paraventricular regions of the hypothalamus (Ciura et al., 2006; Sharif-Naeini et al., 2006; Sharif-Naeini et al., 2008), striatal neurons (Maccarrone et al., 2008), dentate gyrus neurons (Chavez et al., 2010), medium spiny neurons of the nucleus accumbens (Greuter et al., 2010) and hippocampal pyramidal cells (Marsch et al., 2007; Gibson et al., 2008). However, despite using what we consider to be the most sensitive measure yet for detecting TRPV1, our analysis of Trpv1 reporter mice does not support the conclusions of these reports. We did not detect lacZ in any of these cell populations, even in the TRPV1Cre/R26R-lacZ mice, where embryonic expression greatly exceeds that observed in the adult and in which only a few molecules of Cre expression are sufficient to drive recombination. In agreement with the lack of reporter molecule expression, capsaicin did not elicit calcium increases in either intact hippocampal slices, or in cultured hippocampal neurons. Our observations are thus more consistent with studies that found no evidence for TRPV1 in several candidate regions (Kofalvi et al., 2006; Benninger et al., 2008; Taylor et al., 2008; Everaerts et al., 2010).
One caveat is that the use of the IRES in our reporter constructs could have led to spurious reporter molecule expression in areas that lack Trpv1 transcription, or to artificially low expression in certain brain areas that do express TRPV1. However, given the consistency of our results using multiple independent assays, and across various species, we do not believe this to be the case. It is also possible that our reporters are not expressed in cells that transcribe alternate isoforms of Trpv1 that lack the final exon, though such splice variants have not been reported. Finally, although many of the reports of functional TRPV1 in the brain argue for a postsynaptic localization, some of the phenotypes described in these knockout papers could reflect a loss of presynaptic TRPV1 that arises from axonal projections of neurons that we demonstrate do express TRPV1. For example, SuM neurons are known to project heavily to the hippocampus (Vertes, 1992).
Classic studies have shown that CAP-evoked release of neuropeptides from the peripheral nociceptor terminals promotes vasodilation. More recently, a direct action of CAP on vascular smooth muscle was reported, but whether this mechanism is widespread or regionally restricted has not been determined (Donnerer and Lembeck, 1982; Duckles, 1986; Kark et al., 2008). Although TRPV1 immunoreactivity has been observed in rat SMCs (Kark et al., 2008) we did not see similar staining in the mouse (data not shown), likely due to high non-specific antibody staining in peripheral tissues. On the other hand, PLAP and nlacZ provided strong signals in SMCs with minimal background. Importantly, nlacZ staining was reduced in SMCs following systemic RTX injection, confirming functional TRPV1 expression. Finally, calcium imaging of isolated arterioles from the ear showed that genetically marked SMCs respond to CAP application, leading to arteriole constriction. Similar responses could not be evoked in Trpv1 knockout mice, demonstrating that the direct CAP effects on SMCs require TRPV1.
Interestingly, the SMC labeling was restricted to a subset of arterioles in thermoregulatory tissues, including the tongue, skin, trachea, and cremaster. Whereas activation of TRPV1 on sensory nerve endings leads to local vasodilation, vascular TRPV1 mediates vasoconstriction. Thus, activation of TRPV1 on SMCs could counteract nerve-related changes in vascular tone in response to physiological TRPV1 agonists, such as heat and pH. TRPV1 expression in SMC may, therefore, contribute to the thermoregulatory effects induced by TRPV1 antagonists, many of which are being developed to treat chronic pain conditions (Gavva, 2008). On the other hand, the limited TRPV1 expression in the brain indicates that untoward side effects of these drugs are unlikely to be CNS-mediated.
This work was supported by NIH grants NS14627 to AIB, R01NS049488, DP1OD006425 to NMS and R37NS047723 to DJ and by NIMH grants to RAN. ATC is supported by an NIH postdoctoral training grant from the UCSF Cardiovascular Research Institute. ACJ is supported by a Ruth L. Kirschstein National Research Service Award from the NIMH (F32MH081430). We thank Cindy Yang for the IRES-mycCre construct and Nidhi Nuwal for help with rat hippocampal cultures. RG & DOD are employees of AstraZeneca and declare that they have competing financial interest.