The general anesthetic ketamine has been repurposed by physicians as an anti-depressant and by the public as a recreational drug. However, ketamine use can cause extensive pathological changes, including ketamine cystitis. The mechanisms of ketamine’s anti-depressant and adverse effects remain poorly understood. Here we present evidence that ketamine is an effective L-type Ca2+ channel (Cav1.2) antagonist that directly inhibits calcium influx and smooth muscle contractility, leading to voiding dysfunction. Ketamine prevents Cav1.2-mediated induction of immediate early genes and transcription factors, and inactivation of Cav1.2 in smooth muscle mimics the ketamine cystitis phenotype. Our results demonstrate that ketamine inhibition of Cav1.2 signaling is an important pathway mediating ketamine cystitis. In contrast, Cav1.2 agonist Bay k8644 abrogates ketamine-induced smooth muscle dysfunction. Indeed, Cav1.2 activation by Bay k8644 decreases voiding frequency while increasing void volume, indicating Cav1.2 agonists might be effective drugs for treatment of bladder dysfunction.
The anesthetic ketamine has been increasingly used for treatment of depression and pain, and increasingly subject to recreational use and abuse for its mood-elevating effects over the last 20 years1,2. Ketamine is well established as a N-methyl-d-asparate receptor (NMDAR) antagonist, buts its antidepressant and mood-elevating effects remain poorly understood. Ketamine can inhibit NMDAR-dependent bursting activity of lateral habenula neurons to elevate mood3, while the ketamine metabolite, hydroxynorketamine, activates the α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPAR)4. Although both NMDAR and AMPAR are highly expressed in neurons, neither NMDAR inhibition nor AMPAR activation likely explain the broad spectrum of acute and chronic adverse effects of ketamine, including cognitive and memory impairment, hypertension, renal and urological dysfunction, hepatobiliary abnormalities, gastrointestinal symptoms, and many other recently recognized disorders5,6,7,8. Thus, ketamine’s adverse effects might reflect action on one or more previously unrecognized receptors.
Ketamine cystitis is one of the most common among severe ketamine-induced pathologies, affecting up to 30% of long-term users5,8,9,10. Since first reported in 200711, thousands of subsequently reported cases have left uncounted many more unreported and/or unrecognized ketamine abusers throughout the world5,8,10. The major manifestation of ketamine cystitis is intolerable bowel or bladder pain, accompanied by urinary urgency, frequency, nocturia, dysuria, and even hematuria. Urodynamic and ultrasonic studies reveal a contracted bladder with detrusor instability and decreased bladder capacity (as low as 10–20 ml per void), resulting in severe urge incontinence with napkin-dependence. Fifty percent of ketamine cystitis patients have hydronephrosis with impaired kidney function.
The molecular mechanism of ketamine-induced pathological changes remains unclear. Cystoscopy reveals inflammation with lymphocyte infiltration, mucosal ulceration, urothelial atypia and thinning. Pathophysiological studies have shown increased inflammation, dedifferentiation, and apoptosis in patient bladder, in human cell lines, and in animal models12,13,14,15,16. Proposed pathological mechanisms have included direct damage to urothelial barrier function, increased autoimmune response, ischemic microvascular toxicity, smooth muscle myocyte vacuolation, and neuronal degeneration or hyperplasia17. Although urothelium is in direct contact with high concentrations of ketamine in the urine, and disruption of the urothelial barrier has been reported in both patients and animal models, we recently observed no effect on urothelial structure and permeability from 12 weeks of ketamine therapy that nonetheless reduced bladder compliance, void volume, and voiding intervals18. Similar findings in human patients and animal models suggest that disrupted urothelial barrier function may not be the direct cause of ketamine cystitis14,18,19.
The few current approaches for treatment of ketamine cystitis reflect the lack of a generally accepted mechanism. Abstinence from ketamine use is the most effective treatment, whereas anti-inflammatory drugs, anticholinergics, pentosan polysulfate, intravesical glycosaminoglycan preparations, and other supportive therapies have been of limited benefit. This lack of effective medical therapy can lead to cystectomy and bladder reconstruction. Thus, mechanistic understanding of ketamine cystitis and other adverse effects of ketamine constitutes an urgent unmet need, and is a prerequisite for development of effective therapies.
Here, we present evidence that ketamine causes functional and pathological changes in smooth muscle by inhibiting the L-type voltage-gated calcium channel Cav1.2, and that activation of this channel can reverse these pathological changes.
Ketamine inhibits BSM contractility
Ketamine and its metabolite norketamine are both excreted by the kidney, with reports of urinary ketamine/norketamine concentrations up to ~50 µg ml−1 after therapeutic use or recreational abuse20,21,22,23. Although ketamine cystitis has been attributed to urinary ketamine toxicity, this hypothesis has never been tested by direct ketamine instillation into the bladder. We therefore performed cystometrogram (CMG) studies during ketamine infusion into mouse bladder. Ketamine instillation induced acute voiding dysfunction phenotypes typical of ketamine cystitis, including decreased voiding pressure, decreased voiding interval, decreased bladder compliance, and sometimes bladder spasm (Supplementary Fig. 1). These smooth muscle-related abnormalities suggested direct inhibition of ketamine on bladder smooth muscle (BSM) function. Indeed, our myographic studies detected inhibition of BSM contraction at sub-µg ml−1 concentrations of ketamine, with complete inhibition between 100 and 500 µg ml−1 (Fig. 1a, b). These data are consistent with plasma ketamine concentrations in both ketamine abusers and patients, with plasma concentrations of 0.1 μg ml−1 producing analgesia, 0.05–0.2 μg ml−1 drowsiness and perceptual distortions, and 2–3 μg ml−1 for general anesthesia, with reported anesthetic concentrations as high as 26 μg ml−1 24,25. In chronic ketamine abusers, ketamine consumption up to 28 g per day has been reported, with urine ketamine/norketamine concentrations up to 25/50 µg ml−1 23,26. BSM contraction was also inhibited by the major ketamine metabolite, norketamine, but not by hydroxynorketamine (Supplementary Fig. 2). Norketamine exhibits lower potency than ketamine as an NMDAR antagonist.
To rule out ketamine action via inhibition of neurotransmitter release from local nerve fibers, ketamine was tested in the presence of direct BSM contractile agonists carbachol (an M2/M3 cholinergic receptor ligand) and α,β-meATP (a P2X1 purinergic receptor ligand). BSM contraction elicited by these direct agonists was inhibited by ketamine in a dose-dependent manner (Fig. 1c–h), consistent with ketamine’s direct inhibition of BSM contraction in response to electrical field stimulation (EFS).
In addition to severe urinary tract symptoms, ketamine induces dysfunction of gastrointestinal and hepatobiliary tracts, and the vasculature as each is surrounded by smooth muscle. We therefore performed myographic studies on gastric and jejunal smooth muscle, and found inhibition by ketamine of enteric smooth muscle contraction in a dose-dependent manner (Supplementary Figs. 3, 4), suggesting smooth muscle as an important target for ketamine-induced pathology.
Ketamine inhibition of BSM contraction is NMDAR-independent
Ketamine is a noncompetitive NMDAR antagonist, with a sub-µg ml−1 IC50. NMDAR subunit mRNA and protein are expressed in male lower urogenital tract27, but expression has not been reported in BSM. We therefore characterized the pharmacology of BSM strip contraction. The NMDAR agonists (NMDA and RS-Tetrazol-5-yl glycine) at concentrations up to 100 µM had no effect on EFS-stimulated BSM contractile force (Supplementary Fig. 5a, b). Moreover, NMDAR antagonists D-AP5, CGS 19755, and MK 801 had little or no effect on BSM contraction (Supplementary Fig. 5c–e).
To further test a possible role of NMDAR in BSM contraction, we created a smooth muscle-specific knockout mouse strain for the functionally essential NMDAR subunit, NR1 (SMNR1KO) by mating NR1f/f mice (B6.129S4-Grin1tm2Stl/J) with Sm22α-cre mice (B6.Cg-Tg(Tagln-cre)1Her/J). These smooth muscle-specific NR1 knockout mice were grossly normal, with wild type voiding function as indicated by voiding spot assay (VSA) (Fig. 2a–c). Furthermore, these mice exhibit normal BSM contraction force in response to EFS, carbachol, α,β-meATP, and KCl (Fig. 2d–g). EFS-elicited contractile force in the SMNR1KO mouse retained dose-dependent inhibition by ketamine (Fig. 2h). These data suggest that NMDAR plays no role in BSM contractility, and thus is not likely a major mediator of ketamine cystitis.
Ketamine inhibits Cav1.2-mediated calcium channel activity
Ketamine’s dose-dependent, NMDAR-independent inhibition of BSM contraction in response to EFS and to muscarinic/purinergic agonists (Figs. 1 and 2) suggests ketamine blockade of a critical contractile pathway target downstream of both muscarinic and purinergic signaling. The L-type calcium channel Cav1.2 was shown to be essential for urinary BSM contractility, and mouse BSM deficient in Cav1.2 lacked contractile responses to stimulation by carbachol and by high K+28,29. As with ketamine inhibition of BSM contraction, the Cav1.2-specific antagonist, nifedipine, inhibited BSM contraction induced by EFS and by muscarinic and purinergic agonists (Supplementary Fig. 6). We therefore hypothesized that ketamine might act as a Cav1.2 antagonist to inhibit BSM contraction, leading to pathological consequences. Indeed, both ketamine and nifedipine inhibited BSM contraction elicited by depolarization with 50–100 mM extracellular KCl, to a degree adequate to activate voltage-dependent Cav1.2. (Supplementary Fig. 6). Furthermore, Cav1.2 agonist Bay k8644-potentiated BSM contraction was dose-dependently inhibited by ketamine as well as by nifedipine. Moreover, prior inhibition of BSM contraction by nifedipine or by ketamine was dose-dependently reversed by subsequent exposure to Cav1.2 agonist Bay k8644 (Fig. 3). These data indicate mutual antagonism of ketamine and Bay k8644, suggesting ketamine inhibits BSM contraction through its action as a Cav1.2 antagonist.
To further explore the hypothesis that ketamine directly inhibits Cav1.2, we tested ketamine’s inhibition of Cav1.2 expressed in Xenopus oocytes (from cRNAs encoding Cav1.2 subunits mouse α1, rat α2δ1, and rat β3). Seventy-two hours post-cRNA injection, oocytes were subjected to two-microelectrode voltage clamp in a Ca2+-free bath containing 10 mM Ba2+ solution30. As shown in Fig. 4a–e, whereas water-injected oocytes exhibited no apparent inward Ba2+ current, oocytes previously injected with Cav1.2 subunit cRNAs displayed significant inward Ba2+ current, indicating robust functional expression of Cav1.2 sensitive to inhibition by both ketamine and nifedipine (Fig. 4d, e). To further test ketamine inhibition of Cav1.2, freshly isolated mouse BSM cells were subjected to whole cell patch clamp recording. Ketamine consistently inhibited Cav1.2-mediated Ba2+ currents (Fig. 4f–h). Inward currents induced in freshly isolated BSM cells by exposure to 100 nM Cav1.2 agonist Bayk8644 were completely inhibited by 500 µg ml−1 ketamine added in the continued presence of Bayk8644 (Suppl. Fig. 7a, d), consistent with ketamine effects on Xenopus oocytes (Fig. 4), on BSM strip contraction force (Fig. 1), and on Bayk8644-induced muscle contraction (Fig. 3e). Ketamine inhibition of BSM cell Cav1.2 activity is mediated in part by inducing a left-shift in voltage-dependent inactivation (Supplementary Fig. 7h) without significant change in time constant of inactivation (Supplementary Fig. 7e, h).
Furthermore, in primary cultured mouse BSM (mBSM) cells, Bay k8644-induced Ca2+ influx and intracellular Ca2+ signaling were similarly blocked by both ketamine and by nifedipine (Fig. 5). Ketamine and nifedipine also inhibited Ca2+ influx and intracellular Ca2+ signaling induced by treatment of mBSM cells with KCl, carbachol, or ATP (Fig. 5). Ketamine and nifedipine also inhibited Ca2+ influx and intracellular Ca2+ signaling elicited by Bay k8644, by KCl depolarization, by carbachol, or by ATP in primary cultured human BSM (hBSM) cells (Supplementary Fig. 8). These data conclusively identify ketamine as a Cav1.2 antagonist which can block BSM cell Ca2+ influx and inhibit BSM contractility.
Ketamine disrupts Cav1.2-regulated transcription
Cav1.2 channels play central roles in regulating biochemical and electrical signaling of neurons and muscle cells31. Ca2+ influx through Cav1.2 activates neuronal Ca2+-dependent signaling proteins which transmit signals into the nucleus to regulate transcription factors such as CREB, MEF, and NFAT. Ca2+ entering through Cav1.2 channels can also diffuse to the nucleus to activate nuclear calcium-dependent enzymes such as CaMKIV, which in turn regulate activities of transcription factors and transcriptional coregulators. Neuronal c-fos and c-jun are well documented immediate early genes (IEGs) upregulated in response to Cav1.2 activation32,33,34,35,36. c-fos and c-jun in urinary bladder are up-regulated in response to mechanical stretch, suggesting a role of these IEGs in maintaining normal bladder function during cyclic filling and emptying37. We thus hypothesized that ketamine induced pathological changes by inhibition of Cav1.2-mediated transcription. Indeed, Cav1.2 activation in mouse BSM strips by 50 mM KCl depolarization increased BSM mRNA and protein levels of both c-fos and c-jun. These changes were blocked by 15 min pretreatment with either ketamine or nifedipine before KCl depolarization (Fig. 6a–d), and were replicated in primary cultured hBSM cells (Supplementary Fig. 9). Further immunostaining studies confirmed that ketamine and nifedipine inhibited nuclear translocation of c-fos and c-jun in hBSM cells, and that Bay k8644 rescued this inhibition (Fig. 6e, f). KCl depolarization-mediated activation of BSM cell Cav1.2 also upregulated CREB, MEF, NFAT, and FoxO transcription factors in manners sensitive to inhibition by pretreatment with ketamine or nifedipine (Supplementary Figs. 11–14). Ketamine and nifedipine also inhibited stretch-induced upregulation of c-fos and c-jun in mouse BSM cells (Supplementary Fig. 10)37, suggesting that mechanical stretch during bladder filling and voiding modulates bladder function in a Cav1.2-dependent manner.
IEGs and transcription factors modulated by Cav1.2 signaling are important for cell survival, plasticity, proliferation, and differentiation. Ketamine cystitis patients commonly present with smaller bladders and decreased body weight, findings reproduced in many ketamine cystitis animal model studies. We thus hypothesize that ketamine-inhibited Cav1.2 signaling decreases BSM proliferation, in part through inhibition of c-fos and c-jun action. To test this hypothesis, hBSM cells were incubated for up to 7 days in the presence of ketamine or nifedipine at a range of concentrations, then assessed for proliferation. Control cells proliferated until reaching plateau at day 4–5. In contrast, ketamine and nifedipine both inhibited hBSM cell proliferation, and at high concentrations led to cell death. (Fig. 7a, b). This inhibition of BSM proliferation was validated by the observation that ketamine and nifedipine each reduced Ki67-positive cell numbers in a manner reversed by Bay k8644 treatment (Fig. 7c), supporting the importance of Cav1.2 signaling in ketamine toxicity.
Cav1.2 inactivation mimics the ketamine cystitis phenotype
Cav1.2 is essential for BSM contractility, and mice BSM deficient in Cav1.2 lacked contractile responses to carbachol and to high K+ depolarization28,29. In ketamine cystitis patients, urine (or plasma) ketamine concentrations likely inhibit Cav1.2 function partially. To mimic partial inhibition of Cav1.2, we generated SMCav1.2+/− mice with heterozygous knockout of Cav1.2 in smooth muscle by crossing mice carrying Cre recombinase under the control of the smooth muscle transgelin promoter (SM22α-creKI) with Cav1.2fl/fl. SMCav1.2+/− BSM exhibited reduced Cav1.2 mRNA and protein (Supplementary Fig. 15), and responded to individual stimulation by EFS, carbachol, and α,β-meATP with decreased contractile force partially reversible by Bay k8644 (Fig. 8h–m). SMCav1.2+/− BSM cells in primary culture showed reduced [Ca2+]i in response to stimulation by high K+, carbachol, ATP, or Bay K8644 (Supplementary Fig. 16). The reduced size of SMCav1.2+/− mouse bladders, accompanied by reduced bladder wall thickness and smooth muscle cell size (Fig. 8a–g), resembled human patients and animal models with ketamine cystitis in bladder size and histopathology, and in decreased c-fos and c-jun mRNA and protein levels (Supplementary Fig. 17). In vivo VSA and CMG studies in SMCav1.2+/− mice indicated bladder overactivity with reduced bladder volume, contractile pressure and compliance (Fig. 9). Thus, Cav1.2 haploinsufficiency in smooth muscle decreased BSM contractile force and growth, leading to an overactive bladder phenotype recapitulating ketamine cystitis in humans.
To rule out possible developmental compensations to the phenotype of the constitutively haploinsufficient SMCav1.2+/− mouse model, we created a tamoxifen-inducible smooth muscle-specific Cav1.2 knockout (TISMCav1.2−/−) mouse model by crossing smooth muscle myosin heavy chain promoter-Cre (SMMHC-CreERT2, Jackson laboratory, Maine) males with Cav1.2fl/fl females. Cav1.2 genes were selectively deleted in smooth muscle by intraperitoneal injection of 4-hydroxytamoxifen (OHT, 40 mg/kg/day) for five consecutive days. Mice were phenotyped two weeks after injection for Cav1.2 mRNA (qPCR) and protein (immunoblot) to verify the BSM genotype (Supplementary Fig. 15d–f). TISMCav1.2−/− mice consistently had lower body and bladder weights, resembling ketamine cystitis patients and animal models. TISMCav1.2−/− mouse bladders also show reduced muscle mass and significantly smaller BSM cell size. These bladders exhibited dramatically reduced contractile force in response to EFS, KCl, and agonists carbachol and α,β-meATP, consistent with their diminished Ca2+ influx in response to stimulation by Bay k8644 or by KCl (Supplementary Fig. 18). The data conclusively indicate a critical role of Cav1.2 in BSM physiology and recapitulate aspects of the human ketamine cystitis phenotype.
Cav1.2 agonist Bay k8644 rescues ketamine-induced pathology
Although Cav1.2 blockers such as nifedipine have been widely used to treat cardiac ischemia and hypertension, trials of Cav1.2 inhibitors have been unsuccessful in treatment of urinary dysfunction38,39,40,41. In remarkable contrast, our findings demonstrate that Cav1.2 inhibition by ketamine or by nifedipine causes bladder dysfunction. Moreover, Cav1.2 agonist Bay k8644 reverses these in vitro effects of ketamine and nifedipine. We thus hypothesized that Bay k8644 might be used to treat ketamine-induced bladder dysfunction in vivo. As shown in Fig. 10 and Supplementary Figs. 19, 20, intravesical infusion of ketamine or of nifedipine during CMG significantly increased voiding frequency and decreased peak bladder pressure. Moreover, ketamine and nifedipine each occasionally produced ketamine cystitis-like “bladder spasm” (defined as continuously elevated basal pressure, increased contraction frequency and decreased peak pressure, as demonstrated in Supplementary Fig. 1). All these abnormalities were rapidly abrogated upon instillation of Bay k8644 into the bladder lumen (Fig. 10, in the absence or presence of Cav1.2 antagonists). As described above, heterozygous inactivation of Cav1.2 in smooth muscle also produced a ketamine cystitis-like bladder phenotype, and intravesical infusion of Bay k8644 during CMG corrected the associated voiding abnormalities (Fig. 9d). Intraperitoneal injection of Bay k8644 also corrected these voiding abnormalities (Fig. 9c), including decreased voiding frequency and increased voiding spot size (Fig. 9e–g). In summary, these data confirm Cav1.2 dysregulation as a major mediator of ketamine cystitis, and also confirm Cav1.2 agonist Bay k8644 as a highly effective treatment for experimental ketamine cystitis and potentially other ketamine-associated smooth muscle pathologies.
We have described the Cav1.2 antagonist action of ketamine, and further demonstrated that ketamine inhibition of Cav1.2-mediated signaling plays a major role in the pathogenesis of ketamine cystitis. Ketamine inhibited Cav1.2 expressed in Xenopus oocytes, and similarly inhibited L-type Ca2+ channel activity in freshly isolated mouse bladder smooth muscle cells. Indeed, ketamine was previously reported to inhibit L-type Ca2+ channel activity in bullfrog atrial myocytes42, guinea pig ventricular myocytes43, in smooth myocytes of pig tracchea44, rabbit portal vein45, and to inhibit KCl depolarization induced [Ca2+]i elevation in PC12 cells46. The broad tissue distribution of Cav1.2 is consistent with the broad spectrum of ketamine’s physiological and pathological effects, suggesting that Cav1.2 may serve as a therapeutic target not only in ketamine cystitis, but also for treatment of other smooth muscle-related adverse effects of ketamine.
The antihypertensive properties of L-type calcium channel blockers (CCB) such as nifedipine reflect their inhibition of vascular smooth muscle contraction. As we have shown, CCBs also inhibit contraction of BSM. Indeed, nifedipine was tested as a treatment for overactive bladder patients >40 years ago. These and subsequent tests of additional CCBs to treat bladder hyperactivity or urinary incontinence revealed similar lack of efficacy. Here, we show that although CCB inhibit the magnitude of BSM contractile force, they also promote bladder overactivity and urinary symptoms. This counter-intuitive idea opposes current dogma in clinical urology. However, nifedipine instillation into mouse bladder during CMG not only reduced peak pressure (contractile force), but also caused voiding frequency and even bladder spasm (Fig. 10 and Supplementary Fig. 1), with no instances of increased voiding interval or increased voiding volume at any concentration tested. Indeed, hypertensive patients treated with nifedipine reported increased nocturia averaging four times/night, which ceased or improved after nifedipine was discontinued47,48. More recent large-scale clinical studies revealed increased nocturia and urinary urgency and intermittency in CCB-treated patients49,50,51. Consistent with these results, we demonstrated that Cav1.2 agonist Bay k8644 not only reversed the in vitro effects of ketamine and of nifedipine on BSM, but also rapidly and completely reversed ketamine-induced and nifedipine-induced voiding dysfunction in intact mice. Our findings provide mechanistic understanding for the problem of ketamine cystitis, and suggest an approach to clinical therapeutics for ketamine-induced pathologies. These findings also introduce the idea that Cav1.2 agonists of appropriate tissue specificity, rather than (as previously believed) Cav1.2 antagonists such as nifedipine, might effectively treat lower urinary tract syndromes (LUTS). The tissue specificity of alternative splicing of Cav1.2 subunit mRNAs in smooth muscle and cardiac myocytes suggests potential for development of smooth muscle-specific Cav1.2 agonists, perhaps even those selective for bladder smooth muscle over other smooth muscle types52.
As Cav1.2 plays crucial roles in behavior and nociception, it is also possible that Cav1.2 modulation contributes to the antinociceptive and antidepressant effects of ketamine. Recent studies revealed that mice haploinsufficient for Cav1.2 in prefrontal cortex exhibited antidepressant-like behavior53,54, while human patients with Cav1.2 gain-of-function mutants developed depression-related symptoms55,56. Indeed, CCBs such as nifedipine can have antidepressant effects57,58,59, whereas treatment of mice with Cav1.2 agonist Bay k8644 induced a depressive-like phenotype with self-injury behavior60,61. These reports strongly suggest that our identification of ketamine as an antagonist for Cav1.2 might also provide insight into the antidepressant and antinociceptive actions of ketamine.
Unless otherwise specified, all chemicals were obtained from Sigma (St. Louis, MO, USA) and were of reagent grade or better. Agonists and antagonists, including α, β-methylene adenosine 5′-triphosphate trisodium (α, β-meATP), Bay K 8644, nifedipine and carbachol were all purchased from Tocris (Minneapolis, MI, USA). Ketamine were purchased from Patterson (Saint Paul, MN, USA). Fluo-4 AM and Pluronic F127 were purchased from Invitrogen (Carlsbad, CA, USA).
Animal care and use
Mature female Xenopus laevis were obtained from the Dept. of Systems Biology, Harvard Medical School. Frogs were maintained in the AAALAC-accredited animal research facility at Beth Israel Deaconess Medical Center, and subjected to partial ovariectomy under hypothermic tricaine anesthesia. Smooth muscle GluN1 (NR1) homozygous knockout mice (SMNR1KO) and smooth muscle Cav1.2 α1 subunit heterozygous knockout mice (SMCav1.2+/−) were generated by crossing mice carrying Cre recombinase under control of the smooth muscle transgelin promoter, (SM22α-creKI, Stock 006878; Jackson Laboratory). Tamoxifen-inducible smooth muscle Cav1.2 knockout mice (TISMCav1.2−/−) were generated by crossing male smooth muscle myosin heavy chain promoter-Cre male mice (SMMHC-CreERT2, Jackson laboratory, Maine) with Cav1.2fl/fl female mice. Since no differences were observed among the NR1fl/fl, Cav1.2fl/fl, and wild type mice, the results from these control mice were pooled. All mice used in this study were on the C57bl/6j background and were 12–16 weeks’ old of age. All animal studies were performed in adherence to U.S. National institutes of Health guidelines for animal care and use, and with the approval of the Beth Israel Deaconess Medical Center Institutional Animal Care and Use Committee.
Voiding spot assay (VSA)
Individual mice were gently placed in a standard mouse cage with Blicks Cosmos Blotting Paper (Catalog no. 10422-1005) placed in the bottom for 4 h, during which time water was withheld and standard dry mouse chow was available. Mice were then returned to their home cages and the filter paper was recovered. Filters were imaged under ultraviolet light at 365 nm in a UVP Chromato-Vue C-75 system (UVP, Upland, CA) with an onboard Canon digital single lens reflex camera (EOS Rebel T3, 12 megapixels). Overlapping voiding spots were visually examined and manually separated by outlining and copying, then pasting to a nearby empty space using Fiji software (http://fiji.sc/wiki/index.php/Fiji). Images were analyzed by UrineQuant software developed by us in collaboration with the Harvard Imaging and Data Core. The results table containing the area of each individual voiding spot and the total number of spots, was imported into Excel for statistical analysis. A volume:area standard curve on this paper determined that a 1 mm2 voiding spot represents 0.283 μl of urine. Voiding spots with an area ≥80 mm2 were considered primary voiding spots (PVS), based on voiding spot patterns from hundreds of mice18,62.
Bladders were pinned on a small Sylgard block, and muscle was dissected free of the mucosal tissue. BSM strips were then cut longitudinally (2–3 mm wide and 5–7 mm long). Full thickness stomach strips of 5 mm width were cut from the fundus region in the direction of the longitudinal muscle. Full thickness jejunum was cut into ~3 mm rings. Muscle strips were mounted in an SI-MB4 tissue bath system (World Precision Instruments, Sarasota, FL, USA). Force sensors were connected to a TBM 4 M transbridge (World Precision Instruments), and the signal amplified by PowerLab (AD Instruments, Colorado Springs, CO, USA) and monitored through Chart software (AD Instruments). BSM strips were gently prestretched to optimize contraction force, then pre-equilibrated for at least 1 h. All experiments were conducted at 37 °C in physiological saline solution (in mM: 120 NaCl, 5.9 KCl, 1.2 MgCl2, 15.5 NaHCO3, 1.2 NaH2PO4, 11.5 Glucose and 2.5 mM CaCl2) with continuous bubbling of 95% O2/5% CO2. Contraction force was sampled at 2000/s using Chart software. BSM tissue was treated with agonists or antagonists, and/or subjected to electrical field stimulation (EFS).
Electrical field stimulation
BSM strip EFS was carried out by a Grass S48 field stimulator (Grass Technologies, RI, USA) using standard protocols63. EFS stimulation parameters were set as follows: voltage 50 V; duration: 0.05 ms; trains of stimuli: 3 s; frequencies: 1, 2, 5, 10, 20 and 50 Hz. For stomach muscle strips, EFS stimulation parameters were set as follows: voltage 10 V; duration: 5 ms; trains of stimuli: 8 s; frequencies: 1, 5 10, and 20 Hz.
CMG was performed with PBS infusion (25 μl/min)64. Mice were anesthetized by a subcutaneous injection of urethane (1.4 g/kg from 250 mg ml−1 solution in PBS) 30–60 min before surgery. At time of surgery, the mouse was further anesthetized with continuous flow isoflurane (3% induction, 1.0% maintenance). Once the pedal reflex was absent, a 1-cm midline abdominal incision was performed. Flame-flanged polyethylene-50 tubing sheathing a 25Gx1.5 in. needle was implanted through the dome of the bladder, then sutured in place (8-0 silk purse string). The incision site was sutured around the tubing using sterile 5-0 silk, and mice were then placed into a Bollman mouse restrainer for 30–60 min stabilization. The catheter was connected to a pressure transducer (and syringe pump by side arm) coupled to data-acquisition devices (WPI Transbridge and AD Instruments Powerlab 4/35) and a computerized recording system (LabChart software). Bladder filling then commenced, after which voiding occurred naturally through the urethra. Repeated voiding cycles were assessed for change of voiding interval (time between peak pressures), basal pressure (minimum pressure after voiding), threshold pressure (pressure immediately before onset of voiding contraction), peak pressure (maximum voiding pressure minus basal pressure) and compliance (volume, in μl required to increase pressure by 1 cm H2O).
Primary smooth muscle cell isolation and culture
All procedures were carried out in a laminar flow hood using sterile techniques. Mouse urinary bladder was placed in a Petri dish (6 cm diameter) containing DMEM supplemented with antibiotics. The adjacent connective and fatty tissues were removed. Urinary bladder was opened longitudinally and bladder neck including trigone and urethra were resected. The detrusor smooth muscle was cut into tiny pieces using surgical scissors. Tissue fragments were washed five times with PBS and digested in 1% collagenase (Gibco, Carlsbad, USA) at 37 °C for 30 min on a shaker. After digestion, enzymes were neutralized by addition of an equal volume of DMEM medium containing 10% FBS. Resulting suspensions were filtered through 100 μm nylon cell strainers (BD, USA) and centrifuged at 180 × g for 5 min. The cell pellet was resuspended in culture medium, and cells were cultured at 37 °C in 5% CO2 and 95% humidity. Growth medium was changed every 2–3 days. Human bladder primary smooth muscle cells (Catalog: FC-0043) were purchased from LifeLine Cell Technology (Frederick, MD) and were cultured in VascuLife SMC culture medium (Catalog: LL-0014) with 5% FBS, 10 mM l-glutamine, 30 µg ml−1 ascorbic acid, 5 ng ml−1 recombinant human EGF and FGF basic at 37 °C in 5% CO2 and 95% humidity. We used primary cultured mouse bladder smooth muscle cells (1st passage) and primary cultured human smooth muscle cells (3rd–4th passage) for Ca2+ imaging and for detection of mRNA and protein. The functional contractile phenotype of these primary cultured smooth muscle cells was confirmed before each experiment.
Primary cultured BSM cells were seeded at 5 × 103 cells cm−2 onto collagen-coated glass coverslips and maintained for 24–48 h. Before imaging, cultures were washed with physiological saline solution, then loaded with 10 μM Fluo-4 (AM) and 0.02% Pluronic acid for 60 min. After washing cultures twice in PSS, coverslips were placed in a perfusion chamber (Warner instruments, supplied by Harvard Apparatus Ltd., Edenbrige, UK) on the stage of a fluorescent microscope (Olympus, Tokyo, Japan). The chamber was perfused with an automated pump at a flow rate of 1.5 ml min−1. Cells were excited at 494 nm (Lambda LS light source; Sutter Instruments, Novato, CA, USA), and the fluorescence emission signal proportional to [Ca2+]i was measured at 506 nm and images were recorded twice per second. Changes in Fluo-4 fluorescence intensity in all cells in the visual field were acquired with MetaFluor Imaging Software (Molecular Devices). Data were presented as fluorescence intensity. Drug application was regulated by a VC-6 channel valve controller (Warner Instruments) with a Peri-Star perfusion system (WPI).
Cav1.2 expression in Xenopus oocytes
Plasmids encoding mouse Cav1.2 subunits α1 (Catalog no. 26572), rat α2δ1 (Catalog no. 26575), and rat β3 (Catalog no. 26574) were purchased from Addgene and sub-cloned into transcription plasmid pXT7 for cRNA transcription and oocyte expression. Mature female Xenopus were anesthetized in a 0.3% tricaine solution. Ovary lobes were surgically isolated from anesthetized frogs and cut into ~1 cm diameter fragments in a standard oocyte solution (in mM: 100 NaCl, 2 KCl, 1.8 CaCl2, 1 MgCl2, 5 HEPES, 2.5 pyruvic acid, and 50 μg ml−1 gentamicin; pH 7.6). Follicle membranes were removed by shaking of oocyte clusters treated with collagenase (10 mg ml−1, Gibco-BRL, Gaithersburg, MD, USA) in a Ca2 +-free OR2 solution (in mM: 82.5 NaCl, 2.5 KCl, 1 MgCl2, 5 HEPES; pH 7.6) for 30 min. Stage V-VI oocytes were isolated. Each oocyte was then injected with 6 ng of α1 subunit cRNA, 3 ng of α2δ1 cRNA, and 2 ng of β3 cRNA to achieve a 1:1:1 molar ratio in a volume of 50 nl using a Drummond Nanoject pipette injector (Parkway, PA, USA) attached to a Narishige micromanipulator (Tokyo, Japan) under a stereo microscope.
Electrophysiological recording in oocytes
Oocyte Ba2+ currents were recorded at room temperature by two-electrode voltage clamp 3–4 days after cRNA injection, using a GeneClamp 500 two-electrode voltage clamp amplifier (Molecular Devices). Glass microelectrodes of resistance ~1.0 MΩ were pulled from capillaries (Warner Instruments, Hamden, CT, USA) using a P97 pipette puller (Sutter Instrument Co.) and filled with 3 M KCl. Prior to recording, oocytes were injected with ~50 nl of 50 mM BAPTA (1, 2-bis (o-aminophenoxy) ethane-N,N,N,N′-tetraacetic acid, buffered to pH 7.2) to minimize Ca2+-activated Cl− currents native to oocytes. The bath recording solution contained 10 mM Ba(OH)2, 90 mM NaOH, 1 mM KOH, and 5 mM HEPES (pH 7.4 adjusted with methanesulfonic acid). The currents were sampled at 5 kHz, low pass-filtered at 1 kHz (Digidata 1322 A) using pClamp 8 (Molecular Devices, Palo Alto, CA, USA), and analyzed in Clampfit (Molecular Devices).
Electrophysiological recording in fresh isolated BSM cells
Mucosa removed bladder smooth muscle tissue was cut into small strips (5–8 mm long by 2–3 mm wide) and incubated in dissecting solution (in mM: 80 Na Glutamate, 55 NaCl, 6 KCl, 10 glucose, 2 MgCl2, 10 HEPES, final pH 7.3) supplemented with 1 mg ml−1 bovine serum albumin (BSA), 1 mg ml−1 papain (Worthington Biochemical Corp., NJ), and 1 mg ml−1 DL-dithiothreitol (DTT) for ~15 min at 37 °C. Tissues were then transferred to dissecting solution containing 1 mg ml−1 BSA, 1 mg ml−1 collagenase type II (Life Technologies, CA), 0.5 mg ml−1 trypsin inhibitor, and 100 µM CaCl2 for 9–15 min incubation at 37 °C. After digestion, tissues were washed three times with dissecting solution containing 1 mg ml−1 BSA and gently triturated with a fire-blunted Pasteur pipette to generate single dissociated BSM cells. Drops of BSM cell suspension were placed into the recording chamber, and cells were allowed to adhere to the bottom for ~20 min before experiment.
Inhibition of Cav1.2 by ketamine was tested in freshly isolated mouse BSM cells by the nystatin-permeabilized cell-attached patch technique for whole cell recording. Patch pipettes were pulled (Sutter Flaming-Brown P97 puller) from borosilicate glass capillaries to tip resistance >3 MΩ when filled with pipette solution. Pipette tips were fire-polished on a microforge (Narashige) before use. For permeabilized cell-attached whole cell recording, the bath solution contained (in mM) 120 NaCl, 20 TEACl, 10 BaCl2, 4 KCl, 10 Hepes, pH 7.4. The pipette solution contained (in mM) 140 CsCl2, 20 KCl, 1 MgATP, 1 EGTA, 10 Hepes, pH 7.2, with addition of 100 μg ml−1 nystatin freshly prepared from a 5 mg ml−1 stock in methanol. Nystatin permeabilization of the patch membrane after attaining an on-cell gigohm seal established whole cell configuration. Whole cell currents were recorded using an Axopatch 200B amplifier with a Digidata 1440A converter (Axon Instruments/Molecular Devices), using the following voltage pulse protocol. From a holding potential of −70 mV, cells were clamped for 200 ms intervals to test potentials ranging from −80 to +50 mV, then returned to holding potential. Currents were low-pass filtered with a cutoff frequency of 1 kHz, digitized at 2.5 kHz, and stored on a computer. Current densities (pA/pF) were obtained for each cell by normalization of whole cell current to cell capacitance to account for differences in cell membrane surface area. For each cell, capacitative currents measured during 10 ms pulses from a holding potential of −80 mV to a test potential of −75 mV were filtered at a low-pass cutoff frequency of 5 kHz. Data acquisition and analysis was with pCLAMP 11.0.3 software (Axon Instruments/Molecular Devices). Voltage-dependent inactivation was measured from peak inward current during a 500 ms step depolarization to +20 mV following a 4 s prepulse at membrane potentials ranging from −80 to +50 mV in 10 mV increments. Relative peak current (I/Imax) was fit to a Boltzmann equation.
Total RNA was extracted using the RNeasy kit (Qiagen, Valencia, CA). First-strand cDNA was synthesized from 1 μg total RNA using a SuperScript III First-Strand Synthesis System (Invitrogen). RT-PCR for gene detection was performed with Bio-Rad T100 Thermal Cycler. Quantitative RT-PCR was performed using SYBR Green PCR Mix kits and a 7300 real-time PCR system (Applied Biosystems). SDHA (Succinate dehydrogenase complex, subunit A) was used as an internal control. Primer details are provided in Supplemental Table 1.
Rabbit anti-α1C antibody (1:1000, #AB5156) was from Millipore. Rabbit monoclonal anti-c-fos (1:1000, #2250) and rabbit monoclonal anti-c-jun (1:1000, #9165) antibodies were from Cell Signaling Technology (Danvers, MA, USA), and rabbit anti-NR1 antibody was from Alomone Lab (#AGC-001). Goat antirabbit Ig-HRP (1:5000, #1705046) was from Bio-Rad (Hercules, CA, USA). Bladder samples, oocytes and cells were lysed in RIPA buffer (150 mM NaCl, 50 mM Tris, 1% v/v NP-40, 0.5% deoxycholic acid, and 0.1% w/v SDS, pH 7.4) containing protease inhibitors (Complete Mini; Roche Applied Science, Indianapolis, IN, USA). Proteins were separated by SDS-PAGE and transferred to PVDF membranes. Membranes were incubated with primary antibody at 4 °C for 1 h after overnight block with 5% nonfat milk. β-actin (Sigma) was used as the internal control. Bands were visualized with Amersham ECL reagent (Arlington Heights, IL, USA). Exposed and developed film was scanned, and the image contrast was corrected with Photoshop (Adobe Systems, San Jose, CA, USA). Images were imported into Adobe Illustrator CS4 (Adobe) for generation of figures. The blots were quantified using FIJI software. Full blots are shown in source data.
Immunofluorescence staining and confocal microscopy
Excised bladders were fixed in 4% w/v paraformaldehyde dissolved in 100 mM sodium cacodylate (pH 7.4) buffer for 2 h at room temperature. Fixed tissue was cut into small pieces with a razor blade, cryoprotected, frozen, and sectioned. Cells grown on collagen-coated glass coverslips were fixed for 20 min with 4% (w/v) paraformaldehyde. Tissue or cells were incubated with primary antibodies (1:100) for 2 h at room temperature. After washing away unbound primary antibody, sections were incubated with an Alexa 488-conjugated secondary antibody (diluted 1:100), and actin was stained with rhodamine phalloidin (Cytoskeleton, PHDR1). Tissue or cells were mounted with SlowFade Diamond Antifade Reagent containing DPAI (Thermo Fisher Scientific) to stain nuclei. Images acquired by Zeiss LSM-510 confocal microscope were saved as TIFF files, and imported into Adobe Illustrator CS3. Additional antibodies included rat anti-mouse β1 integrin antibody (BD Bioscience, #550531), anti-c-fos (Cell Signaling, #2250), anti-c-jun (Cell Signaling, #9165), and anti-Ki67 (Abcam, #ab15580).
All data are expressed as means ± SD, or presented as boxes and whiskers (whiskers extending from minimum to maximum values). Data were analyzed by Student’s t-test between two groups or by one-way analysis of variance (ANOVA) for comparison among groups using GraphPad Prism 8 software. If possible, paired t-test was used. Tukey HSD (independent) or Bonferroni (paired) multiple comparison post-hoc tests were used where necessary, and P < 0.05 was considered significant.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
Nucleotide sequences of mouse Cav1.2 subunits α1 (Catalog no. 26572), rat α2δ1 (Catalog no. 26575), and rat β3 (Catalog no. 26574) are available at www.addgene.org. The datasets generated and/or analysed during the current study are available from the corresponding author on reasonable request. Source data are provided with this paper.
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We thank Drs. Warren Hill (Beth Israel Deaconess Medical Center, Boston), Marcelo Carattino (University of Pittsburgh, Pittsburgh), Robert Moldwin (Northwell Health, Hofstra North Shore-LIJ School of Medicine, New York), Retnagowri Rajandram, and Teng Aik Ong (University Malaya Medical Center, Kuala Lumpur, Malaysia), and Chi Fai Ng (The Chinese University of Hong Kong, Hong Kong) for useful discussions. We thank generous support from Beth Israel Deaconess Medical Center Department of Medicine for this work. We are deeply grateful for the participation of all subjects contributing to this research.
The authors declare no competing interests.
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Chen, H., Vandorpe, D.H., Xie, X. et al. Disruption of Cav1.2-mediated signaling is a pathway for ketamine-induced pathology. Nat Commun 11, 4328 (2020). https://doi.org/10.1038/s41467-020-18167-4