K_V2.1 and electrically silent K_V channel subunits control excitability and contractility of guinea pig detrusor smooth muscle

Abstract

Voltage-gated K⁺ (K_V) channels are implicated in detrusor smooth muscle (DSM) function. However, little is known about the functional role of the heterotetrameric K_V channels in DSM. In this report, we provide molecular, electrophysiological, and functional evidence for the presence of K_V2.1 and electrically silent K_V channel subunits in guinea pig DSM. Stromatoxin-1 (ScTx1), a selective inhibitor of the homotetrameric K_V2.1, K_V2.2, and K_V4.2 as well as the heterotetrameric K_V2.1/6.3 and K_V2.1/9.3 channels, was used to examine the role of these K_V channels in DSM function. RT-PCR indicated mRNA expression of K_V2.1, K_V6.2–6.3, K_V8.2, and K_V9.1–9.3 subunits in isolated DSM cells. K_V2.1 protein expression was confirmed by Western blot and immunocytochemistry. Perforated whole cell patch-clamp experiments revealed that ScTx1 (100 nM) inhibited the amplitude of the K_V current in freshly isolated DSM cells. ScTx1 (100 nM) did not significantly change the steady-state activation and inactivation curves for K_V current. However, ScTx1 (100 nM) decreased the activation time-constant of the K_V current at positive voltages. Although our patch-clamp data could not exclude the presence of the homotetrameric K_V2.1 channels, the biophysical characteristics of the ScTx1-sensitive current were consistent with the presence of heterotetrameric K_V2.1/silent K_V channels. Current-clamp recordings showed that ScTx1 (100 nM) did not change the DSM cell resting membrane potential. ScTx1 (100 nM) increased the spontaneous phasic contraction amplitude, muscle force, and muscle tone as well as the amplitude of the electrical field stimulation-induced contractions of isolated DSM strips. Collectively, our data revealed that K_V2.1-containing channels are important physiological regulators of guinea pig DSM excitability and contractility.

detrusor smooth muscle (DSM), which makes up the bladder wall, relaxes during bladder filling and contracts phasically during voiding (2). The underlying cause of the spontaneous phasic DSM contractions is the spontaneous action potential and corresponding Ca²⁺ transients (19–22, 25–27, 47). Initiation of the action potential in guinea pig DSM arises from the opening of L-type voltage-gated Ca²⁺ channels followed by an increase in the intracellular Ca²⁺ concentration (25). The repolarization phase of the DSM action potential is initiated by activation of large-conductance Ca²⁺-activated K⁺ (BK) channels, voltage-gated K⁺ (K_V) channels (56), and negative feedback Ca²⁺-mediated inhibition of the L-type voltage-gated Ca²⁺ channels (7, 11, 19, 26, 35, 46, 48), whereas small-conductance Ca²⁺-activated K⁺ (SK) channels and K_V channels are responsible for the slow afterhyperpolarization (16, 20, 56). The resting membrane potential and action potential frequency are considered to be regulated by K_ATP, BK, and K_V channels (7, 16, 19, 46, 47, 56). However, there remains a substantial gap in our knowledge regarding the cellular and functional roles, expression, and regulation of different subtypes of K⁺ channels in DSM.

Of all ion channels that control DSM excitability, the K_V channels are the most diverse superfamily (45, 60). These channels have been identified in many different types of smooth muscle (5, 50, 53), including DSM (9, 13, 56). The expression of K_V2.1 and K_V2.2 channels has been demonstrated in rat DSM (9, 17, 41). Expression of K_V1 channels has been shown in human DSM (13) and of K_V2.1, K_V5.1, K_V6.1, K_V6.2, and K_V6.3 subunits in mouse DSM (14, 56).

The structural diversity of the K_V channels contributes to the variety of their biophysical and pharmacological properties and makes them potentially attractive targets for pharmacological treatment of diseases such as overactive bladder (OAB) and urinary incontinence (UI) (3, 9, 45, 60). Further investigation in this area may lead to development of selective pharmacological or genetic therapies for OAB and UI.

In the human genome, the members of the K_V superfamily are encoded by 40 genes and comprise 12 K_V channel families (45, 60). Only K_V1–K_V4, K_V7, K_V11 (also known as hERG), and K_V12 (also known as elk) channel families form pore-conducting channels, homo- or heterotetramers, whereas the other members of the K_V families, K_V5.1, K_V6.1–6.4, K_V8.1–8.2, and K_V9.1-K_V9.3, are electrically silent and do not form functional channels when expressed alone (8, 36, 42, 44, 45, 54, 55). Instead, they coassemble with K_V2 family, thus increasing K_V channel diversity (45, 52, 60). Those channels form heterotetrameric channels with K_V2 subunits, and their activation, deactivation, inactivation, and recovery from inactivation properties differ from those reported for homotetrameric K_V2 channels (8, 36, 42, 44, 51, 52). For example, the K_V9.3 subunit causes an important alteration of the channel biophysical properties and increases the single-channel conductance of the heterotetrameric K_V2.1 channel (44). This modulatory K_V subunit combines with K_V2.1 to form a functional heterotetrameric K_V2.1/9.3 channel consisting of three K_V2.1 subunits and one K_V9.3 subunit (33). Expression of the electrically silent K_V6.3 and K_V9.3 subunit has been reported in vascular smooth muscle cells (4, 12, 23, 38–40, 43, 44, 49, 58, 59, 61). It is suggested that the heterotetrameric K_V2.1/9.3 channel contributes to the excitability in oxygen-sensitive pulmonary myocytes, and it is involved in hypoxic pulmonary vasoconstriction (44). Structural, biophysical, and pharmacological properties of the heterotetrameric K_V2.1/silent K_V channels have been extensively studied (14, 32–34, 40, 44), but their physiological roles in smooth muscle cells, including DSM cells, remain unclear.

In the past, the study of K_V channels' function has been hampered by the lack of selective inhibitors. Stromatoxin-1 (ScTx1), a spider venom peptide isolated from the African tarantula (Stromatopelma calceata), was recently identified as a highly selective inhibitor of the K_V2.1, K_V2.2, and K_V4.2 homotetramers as well as the K_V2.1/6.3 and K_V2.1/9.3 heterotetramers (14, 15, 40). Studies in neurons, cerebral arteries, urethra, and DSM have confirmed that ScTx1 is a useful pharmacological tool to study the functional role of these K_V channels (1, 6, 9, 37, 40, 61).

The aim of this study was to examine the expression and function of K_V2.1 and electrically silent K_V subunits in guinea pig DSM excitability and contractility. We applied a multidisciplinary approach to this investigation, using molecular biological techniques, patch-clamp electrophysiology, isometric tension recordings of isolated guinea pig DSM strips, and pharmacological protocols using ScTx1.

MATERIALS AND METHODS

Animal care and tissue collection.

All animal studies were reviewed and approved by the University of South Carolina Institutional Animal Care and Use Committee and carried out in accordance with the Animal Use Protocol 1747. Eighty-five adult Harley albino guinea pigs (Charles River Laboratories) were used in this study (54 males and 31 females, average weight 372.8 ± 2.1 g). Guinea pigs were euthanized with CO₂, followed by thoracotomy. The urinary bladder was quickly removed and placed in an ice-cold, Ca²⁺-free dissection solution (see Solutions and drugs) or in RNAlater solution (Qiagen, Hilden, Germany) for the RT-PCR experiments. The urinary bladder was then cut open, the entire mucosal and urothelium layers were carefully removed, and only the smooth muscle tissue from the detrusor was used. For RT-PCR and Western blot experiments, the guinea pig brains were also removed and placed in RNAlater solution.

DSM single-cell isolation.

DSM single cells for RT-PCR, immunocytochemistry, and electrophysiology were freshly isolated and collected as previously described (7, 9, 10, 31). Briefly, isolated DSM was cut in strips (5–8 mm long, 2–4 mm wide), and 1–2 strips were placed in dissection solution (2 ml) supplemented with 1 mg/ml bovine serum albumin (BSA), 1 mg/ml papain (Worthington Biochemical, Lakewood, NJ), and 1 mg/ml dl-dithiothreitol and incubated for 25–30 min at 37°C. The DSM strips were transferred to dissection solution (2 ml) containing 1 mg/ml BSA, 1 mg/ml collagenase (type II from Sigma-Aldrich, St. Louis, MO), and 100 μM CaCl₂ and incubated for 5–6 min at 37°C. DSM strips were then washed with fresh BSA-containing dissection solution. The individual DSM cells were dispersed when the enzyme-treated tissue was passed through the tip of a Pasteur pipette and then used for electrophysiology, RT-PCR, and immunocytochemistry studies. DSM cells for patch-clamp experiments were stored at 4°C for further use.

RNA extraction, RT-PCR, and sequencing.

For RT-PCR experiments, whole DSM tissue and freshly isolated single DSM cells were immediately used for RNA extraction. RT-PCR experiments were conducted as previously described, and the single-cell RT-PCR experiments were carefully controlled for contaminations from non-DSM cells (9, 10). Specific primers for K_V2.1, K_V2.2, K_V4.2, K_V5.1, K_V6.1–6.3, K_V8.1–8.2, and K_V9.1–9.3 were designed based on the sequences of multiple species present in GenBank and aligned using Primer Premier 5 software (PREMIER Biosoft International, Palo Alto, CA). Specific primer pair sequences are listed in Table 1. The cDNA production was amplified using GoTaq green master mix (Promega) and specific primers for K_V channel subunits. The PCR annealing temperature for each primer pair was optimized using a Mastercycler gradient thermocycler (Eppendorf, Hamburg, Germany). Negative control experiments for PCR were performed in the absence of the reverse transcriptase (−RT). Guinea pig brain tissue, which is known to express the studied K_V channels, was used as a positive control. PCR products were purified using the GenElute PCR clean-up kit (Sigma-Aldrich) and sequenced at the University of South Carolina Environmental Genomics Facility for sequence confirmation.

Table 1.

RT-PCR primers for subunits of the K_V channels sensitive to ScTx1 and for the silent K_V channel subunits

	Sense	Antisense	Product Size, bp
KV2.1	CTCCACCATTGCCCTGTC	TCCGCTTGATTGCTTTCTC	695
KV2.2	TCCCAGGAACAGATGAGC	GAGTGGTGAGCGGAAAGT	324
KV4.2	CTTCACTATCCCCGCCAT	GTTTCCACCACATTCGCG	308
Kv5.1	GCGAAGACATTGAGATCGTG	CGTCCAAGATGAGCTGCAC	393
Kv6.1	CTGGACAGCGAGGATCAAG	TACCATGTCTCCGTAGCCT	731
Kv6.2	CGAACGTGTACTGTCATCA	GCTCGTGCACCTGGCTG	309
Kv6.3	ATGACGACGGTGGGCTAT	CTGCTCCTGCTCCCTCTTG	256
Kv8.1	ACCTCCTTGCCATCTTGC	TTGCCTGTCGTGGTGTCT	385
Kv8.2	GAGGAGCAGCGGGACAGCAA	AGCGGCGGAGAAGGCAAAG	181
Kv9.1	CCCCATCACCATCATCTTCAA	CGTCAACGCTACTCAGCAAGTC	119
Kv9.2	TGTGCGTCTTCTCCTTCA	CTGGTCACTCTGCTCGTC	137
KV9.3	ACAGTGGGCTATGGAGAC	GATGCCAATGGAAGACAG	273

ScTx1, stromatoxin-1.

Western blot analysis.

Protein extraction and Western blot analysis were performed as previously described (9, 10). The blots were incubated with the affinity-purified rabbit polyclonal primary antibodies anti-K_V2.1 (DRK1, 1:400), anti-K_V2.2 (CDRK, 1:2,000), and anti-K_V4.2 (Kcnd2, 1:200; Alomone Labs, Jerusalem, Israel) overnight at 4°C. The membrane was then incubated with goat anti-rabbit IgG conjugated with horseradish peroxidase diluted to 1:3,000 in the blocking buffer for 1 h at room temperature. Bound antibodies were detected using an ECL substrate kit (Amersham, Piscataway, NJ) according to the manufacturer's instructions. Staining specificity was verified by preincubation of the antibodies with a competing peptide.

Immunocytochemistry.

Immunocytochemistry was performed as previously described (9, 10). DSM cells were incubated with the rabbit polyclonal primary antibodies anti-K_V2.1, anti-K_V2.2, and anti-K_V4.2 (DRK1, CDRK, and Kcnd2, 1:100; Alomone Labs) at 37°C for 1 h. Next, cells were labeled with secondary antibodies (Cy3-conjugated anti-rabbit IgG, 1:200 in PBS/3% normal donkey serum/0.01% Triton X-100; Jackson ImmunoResearch, West Grove, PA) for 1 h in a dark room. After labeling, DSM cells were washed with PBS and incubated with phalloidin for 2 h in the dark. Cells were then washed two more times, incubated with 4′,6-diamidino-2-phenylindole for 15 min and washed again, and then mounted onto slides with DABCO (1,4-diazabicyclo-2–2-2-octane). Control treatments were carried out as follows: 1) omission of the primary antibody for confirming the specificity of the secondary antibody and 2) absorption of the primary antibody by a competing peptide for confirming the specificity of the primary antibody. Images at ×63 magnification were acquired with an LSM 510 META confocal microscope (Carl Zeiss, Göttingen, Germany). The slides for each group were imaged with the same laser power, gain settings, and pinhole for the controls and antibody treatment.

Electrophysiological patch-clamp recordings.

To preserve the physiological environment of the DSM cells, the whole cell perforated patch-clamp technique was applied in all electrophysiological experiments (18, 29). Freshly isolated DSM cells were used for patch-clamp experiments within 12 h after isolation. Isolated DSM cells were placed in an experimental chamber for at least 20 min to allow them to adhere to the glass bottom and were then washed with extracellular solution (see Solutions and drugs). Only DSM cells with a spindle shape, bright appearance, and well-defined cell edges were used. The ionic currents were recorded using an Axopatch 200B amplifier, Digidata 1440A, and pCLAMP 10.2 software (Molecular Devices, Sunnyvale, CA) and then filtered using an eight-pole Bessel filter 900CT/9L8L (Molecular Devices). The glass pipettes used for the patch-clamp experiments were made from borosilicate glass (Sutter Instruments, Novato, CA) and pulled using a Narishige PP-830 vertical puller (Narishige Group, Tokyo, Japan). The pipettes were polished with a Micro Forge MF-830 fire polisher (Narishige Group) and coated with dental wax to reduce capacitance.

In all voltage-clamp experiments, the BK channel was blocked with 100 nM paxilline, a BK channel-selective inhibitor. To study the K_V currents, we applied various voltage-step protocols. The leak current was not subtracted during the experiments. The average current value during the last 50 ms of the 400-ms depolarization step was taken for the K_V current. For the controls, current traces of at least 5 pulses within 10 min were recorded and averaged. Only cells with stable control currents in response to depolarization steps within at least 10 min were used to study ScTx1 effects. All patch-clamp experiments were carried out at room temperature.

Isometric DSM tension recordings.

Isometric tension recordings of guinea pig DSM isolated strips were performed as previously described (7, 9, 31). Briefly, small DSM muscle strips (5–8 mm long, 2–3 mm wide) were placed between a stationary mount and a force-displacement transducer in thermostatically controlled (at 37°C) 10-ml tissue baths filled with Ca²⁺-containing physiological salt solution (see Solutions and drugs). Strips were stretched to 1 g of tension and were then washed every 15 min during an equilibration period of 45–60 min. Electrical field stimulation (EFS) was generated using a PHM-152I stimulator (Med Associates, Georgia, VT). DSM contractile responses were recorded using MyoMed software (Med Associates).

Solutions and drugs.

The Ca²⁺-free dissection solution had the following composition (in mM): 80 monosodium glutamate, 55 NaCl, 6 KCl, 10 glucose, 10 HEPES, and 2 MgCl₂, pH 7.3, adjusted with NaOH. The Ca²⁺-containing physiological salt solution was prepared daily and contained (in mM) 119 NaCl, 4.7 KCl, 24 NaHCO₃, 1.2 KH₂PO₄, 2.5 CaCl₂, 1.2 MgSO₄, and 11 glucose and was aerated with 95% O₂-5% CO₂ to obtain pH 7.4. The extracellular solution used in the electrophysiological experiments contained (in mM) 134 NaCl, 6 KCl, 1 MgCl₂, 2 CaCl₂, 10 glucose, and 10 HEPES; pH was adjusted to 7.4 with NaOH. The pipette solution contained (in mM) 110 potassium aspartate, 30 KCl, 10 NaCl, 1 MgCl₂, 10 HEPES, and 0.05 EGTA, pH adjusted to 7.2 with NaOH, and was supplemented with freshly dissolved 200 μg/ml amphotericin-B. ScTx1 was purchased from Alomone Labs. dl-Dithiothreitol, paxilline, tetrodotoxin (TTX), carbachol, and collagenase type II were obtained from Sigma-Aldrich. BSA and amphotericin-B were obtained from Fisher Scientific. Papain was obtained from Worthington Biochemical.

Data analysis and statistics.

Clampfit version 10.2 software (Molecular Devices) was used for electrophysiological data analyses, MiniAnalysis software (Synaptosoft, Decatur, GA) was used for data analyses of DSM contraction, and GraphPad Prism 4.03 software (GraphPad Software, La Jolla, CA) was used for statistical analyses. For data presentation, CorelDraw Graphic Suite X3 software (Corel, Ottawa, ON, Canada) was used. Results are summarized as means ± SE for the number of DSM strips or cells (n) and the number of guinea pigs (N). Data were compared using ANOVA or paired Student's t-test. P values <0.05 were considered statistically significant.

RESULTS

RT-PCR.

RT-PCR experiments were conducted in both whole DSM tissues and freshly isolated single DSM cells to determine the expression of mRNA messages for the subunits of the known ScTx1-sensitive K_V channels (K_V2.1, K_V2.2, and K_V4.2) as well as all K_V silent subunits (K_V5.1, K_V6.1–6.3, K_V8.1–8.2, and K_V9.1–9.3) that associate with the K_V2 family. The expression of mRNA message for all these K_V channel subunits was detected in guinea pig brain, which was used as a positive control (Fig. 1). However, the whole DSM tissue expressed detectible mRNA messages for K_V2.1, K_V6.2, K_V6.3, K_V8.2, and K_V9.1–9.3 but not for K_V2.2, K_V4.2, K_V5.1, K_V6.1, and K_V8.1 channel subunits (Fig. 1). The presence of other cell types within the DSM layer, such as neurons, fibroblasts, and vascular myocytes, may lead to the detection of K_V channel subunits expressed in cell types other than DSM cells. To eliminate the possible contamination from other cell types, we applied single-cell RT-PCR experiments to freshly isolated DSM cells. Again, the single-cell RT-PCR confirmed the expression of mRNA messages for K_V2.1, K_V6.2, K_V6.3, K_V8.2, and K_V9.1–9.3 but not for K_V2.2, K_V4.2, K_V5.1, K_V6.1, and K_V8.1 subunits (Fig. 1). A lack of genomic DNA contamination in mRNA isolated from single DSM cells was also confirmed by using the negative control reactions lacking the reverse transcriptase. All purified PCR products were sequenced to confirm their identity. Results were verified in 18 different preparations, obtained from 16 guinea pigs. We next applied Western blot analysis and immunocytochemistry to detect whether proteins for the three main channel subunits that can form their own homotetramers (K_V2.1, K_V2.2, and K_V4.2) are expressed in DSM.

Fig. 1.

Detection of mRNA message for K_V channel subunits in detrusor smooth muscle (DSM) whole tissue (WT) and freshly isolated DSM single cells (SC). mRNA message was detected for K_V2.1, K_V6.2, K_V6.3, K_V8.2, and K_V9.1–9.3 but not for K_V2.2, K_V4.2, K_V5.1, K_V6.1, and K_V8.1 channel subunits. The base pair sizes for each product are included in Table 1. No products were observed in the negative controls in which reverse transcriptase (RT) was left out of the reaction. Guinea pig brain tissue was used as a positive control.

Western blot analysis.

Using commercial K_V channel subunit-specific antibodies, we confirmed protein expression of K_V2.1 subunit in whole DSM tissue with Western blot experiments (Fig. 2). Preabsorption of the primary antibody with its antigenic competing peptide indicated the specificity of the antibodies for their intended epitope. Consistent with our RT-PCR data, no K_V2.2 and K_V4.2 proteins were detected in whole DSM tissue (data not shown). The protein expression for each separate K_V channel subunit was verified in three separate Western blot reactions using proteins isolated from three guinea pigs.

Fig. 2.

Western blot detection of K_V2.1 channel protein expression in whole DSM tissues. The immunoreactive band in DSM tissue was eliminated by a competing peptide (+CP). Guinea pig brain tissue was used as a positive control.

Immunocytochemistry.

To further verify the presence of K_V2.1 channel subunits in guinea pig DSM, we immunocytochemically labeled freshly isolated DSM cells. Freshly isolated DSM cells had bright, distinct edges under phase-contrast optics. Anti-K_V2.1 antibody specifically labeled the freshly isolated cells (Fig. 3). However, immunofluorescence staining was not detected for K_V2.2 and K_V4.2 channel subunits in isolated DSM cells (data not shown), which is consistent with our RT-PCR and Western blot experiments. The immunocytochemical experiments were carefully controlled for specificity by the omission of the primary antibody or absorption of the primary antibody by a competing peptide (Fig. 3). Results were verified in eight cells, obtained from three guinea pigs.

Fig. 3.

Immunocytochemical detection of K_V2.1 channel in freshly isolated single DSM cells using channel-specific antibody. Red staining (bottom left panels) indicates detection of K_V2.1 subunit (A); no staining was visible after absorption of the primary antibody with a competing peptide (B; +CP control) or after the primary antibody was omitted and cells were incubated with the secondary antibody only (C; 2° control). Cells nuclei are shown in blue (top left panels); F-actin is shown in green (top right panels). The merged images are also shown (bottom right panels). Images at ×63 magnification were obtained via confocal microscopy.

Voltage dependence and ScTx1 sensitivity of guinea pig DSM whole cell K_V current.

All voltage-clamp experiments were conducted in the presence of 100 nM paxilline to eliminate the contribution of the BK channels to the total whole cell current and to study the remaining K_V current. The average DSM cell capacitance was 28.2 ± 1.2 pF (n = 155 cells; N = 61 guinea pigs). In the first experimental series, we applied a 400-ms voltage-step protocol, illustrated in Fig. 4A. A series of potentials between −70 and +60 mV were applied from a holding potential (V_h) of −70 mV in 10-mV increments, and then cells were repolarized to −40 mV for another 800 ms. The K_V current amplitude increased steeply with increasing the membrane potential, and small deactivating tail currents were recorded on repolarization to −40 mV (Fig. 4A). Application of ScTx1 (100 nM) caused a significant inhibition of the whole cell K_V current amplitude (n = 12; N = 9; P < 0.05; Fig. 4). The remaining ScTx1-insensitive current is possibly determined by other types of K_V channels. The inhibitory effect of ScTx1 (100 nM) on the current-voltage relationship in guinea pig DSM cells is shown in Fig. 4B (n = 12; N = 9; P < 0.05).

Fig. 4.

Voltage-dependent activation of stromatoxin-1 (ScTx1)-sensitive K_V currents in guinea pig DSM cells. A: representative recording of the effect of ScTx1 on the whole cell K_V current elicited by a short (400 ms) voltage-step protocol (top). ScTx1 (100 nM) significantly decreased the voltage step-induced whole cell K_V current. ScTx1-sensitive current was obtained after subtraction of the remaining whole cell current from the control current. The scale bar applies to all recordings. B: the current-voltage relations show that ScTx1 (100 nM) decreased voltage-dependent whole cell K_V current in guinea pig DSM cells (n = 12; N = 9). *P < 0.05; **P < 0.005; ***P < 0.001. The current-voltage relation of the subtracted ScTx1-sensitive current is also shown. C: steady-state activation curves of the K_V current in the absence (control) or presence of 100 nM ScTx1 (ScTx1-insensitive current). Data were obtained from the current recordings shown in A. The tail currents recorded after stepping back to −40 mV for 800 ms were used to build the normalized conductance (G/G_max) curve. The curves were fitted using a single-component Boltzmann equation. The steady-state activation curve of the ScTx1-sensitive current is also shown. Values are means ± SE. All experiments were performed in the presence of the selective BK channel inhibitor paxilline (100 nM).

In the second experimental series, a long-lasting double-pulse protocol was applied (Fig. 5A). The cell membrane potential was stepped from a V_h of −70 mV to potentials between −120 and +40 mV for 15 s, before stepping to +60 mV for 250 ms to determine the current availability at the end of each presiding step. As is evident from Fig. 5A, the K_V currents at various voltages showed a slow pattern of inactivation. Typical current traces of K_V current elicited by this protocol before and after application of 100 nM ScTx1 are shown in Fig. 5A (n = 7; N = 5).

Fig. 5.

Voltage-dependent inactivation of ScTx1-sensitive K_V currents in guinea pig DSM cells. A: representative recording of the effect of ScTx1 on the whole cell K_V current elicited by a 15-s long-lasting double-pulse voltage-step protocol (top). The scale bar applies to all recordings. The expanded scale at right shows the currents during the 250-ms test pulse to +60 mV. B: steady-state K_V current inactivation curves show voltage dependence of inactivation in the absence (control) or presence of 100 nM ScTx1 (n = 7; N = 5). The inactivation curves were obtained by plotting the normalized peak currents (I/I_max) recorded during the 250-ms test pulse to +60 mV as a function of the voltage of the 15-s prepulse. Data were fitted using a single-component Boltzmann equation. The steady-state inactivation curve of the subtracted ScTx1-sensitive current is also shown. Values are means ± SE. All experiments were performed in the presence of the selective BK channel inhibitor paxilline (100 nM).

Effects of ScTx1 on the steady-state activation and inactivation of the K_V current.

We examined whether ScTx1 (100 nM) can shift the steady-state activation and inactivation curves of the K_V current. Voltage-dependent activation and inactivation of ScTx1-sensitive current was studied with the voltage protocols shown in Figs. 4A and 5A, respectively. Activation curves of the K_V current in the presence or absence of 100 nM ScTx1 (Fig. 4C) were obtained from the tail currents recorded after stepping back to −40 mV (Fig. 4A). The activation curves are plotted as normalized conductance (G/G_max) against voltage and are fitted using a single-component Boltzmann equation (Fig. 4C). ScTx1 (100 nM) did not significantly affect the half-maximal activation potential (V_0.5) of the steady-state activation. The V_0.5 was −11.7 ± 0.8 mV in the absence of ScTx1 and −14.5 ± 2.2 mV in the presence of 100 nM ScTx1 (Fig. 4B; Table 2). The slope factor (k) of the steady-state activation curve remained unchanged by 100 nM ScTx1, and it was 10.1 ± 0.8 mV under control conditions and 11.5 ± 2.0 mV in the presence of 100 nM ScTx1 (Fig. 4B).

Table 2.

Comparison of steady-state properties of total whole cell K_V, ScTx1-sensitive, ScTx1-insensitive currents and published data on the homotetrameric K_V2.1 and heterotetrameric K_V2.1/silent K_V channel currents

	Steady-State Activation		Steady-State Inactivation
	V0.5, mV	Ref.	V0.5, mV	Ref.
Control current in guinea pig DSM	−11.7 ± 0.8		−63.6 ± 1.7
ScTx1-insensitive current in guinea pig DSM	−14.5 ± 2.2		−65.3 ± 1.7
ScTx1-sensitive current in guinea pig DSM	−6.9 ± 1.4		−50.1 ± 3.7
KV current in mouse DSM	1.1 ± 1.3	56	−61.4 ± 1.2	56
ScTx1-sensitive current in rabbit urethra	−7.0 ± 5.0	37	−55.0 ± 3.0	37
KV2.1 current	−1.7	36	−30.1	36
	+12.2 ± 1.4	42	−15.9 ± 2	42
	+1.0 ± 1.0	34	−18.4 ± 2.8	6
	+7.8	12a	−28.3	12a
	+20.4	44	−34.7 ± 0.6	34
			−20.5 ± 0.9	52
KV2.1/6.2 current	−10	62	NA
KV2.1/6.3 current	−13 ± 1.8	52a	−55.6 ± 1.1	42
KV2.1/8.2 current	−2.8	12a	−30.6	12a
KV2.1/9.1 current	NA		−33.3 ± 0.2	52
KV2.1/9.2 current	NA		−35.7 ± 3.2	52
KV2.1/9.3 current	−9.5 ± 2	34	−54.8 ± 1.8	34
	+7.2	14	−47.2 ± 0.7	6
	+3.2	44	−44.9	44

Values are means ± SE of half-maximal potentials (V_0.5) of steady-state activation and inactivation from our experiments in guinea pig detrusor smooth muscle (DSM), as well as data from the literature for currents in other species and for homotetrameric K_V2.1 and heterotetrameric K_V2.1/silent K_V channel currents. NA, not available.

We next evaluated the effect of ScTx1 (100 nM) on the steady-state inactivation curve. Using the protocol shown in Fig. 5A, steady-state inactivation curves were obtained from the maximum value of the normalized end-pulse current amplitudes (I/I_max) recorded at +60 mV and plotted against test potentials. K_V current steady-state inactivation curves in the presence and absence of 100 nM ScTx1, fitted by a single-component Boltzmann equation, are shown in Fig. 5B. There was no significant difference in the V_0.5, which was −63.6 ± 1.7 mV in the absence of ScTx1 and −65.3 ± 1.7 mV in the presence of 100 nM ScTx1 (n = 7; N = 5; P > 0.05; Fig. 5B). This V_0.5 value is consistent with data from mouse DSM cells (56). The slope factor k of the steady-state inactivation curves also remained unchanged, and it was −24.5 ± 1.7 mV under control conditions and −22.1 ± 1.8 mV in the presence of 100 nM ScTx1 (n = 7; N = 5; P > 0.05; Fig. 5B). We also found that the total K_V current of guinea pig DSM cells did not completely inactivate (Fig. 5B).

The ScTx1-sensitive current was evaluated after subtraction from the total K_V current and the remaining current after application of 100 nM ScTx1 (Figs. 4 and 5). All parameters of the ScTx1-sensitive current were elucidated and are shown in Tables 2 and 3. The half-maximal activation of the ScTx1 current was V_0.5 = −6.9 ± 1.4 mV, and the slope factor k was 10.2 ± 1.2 mV (Fig. 4C). The half-maximal steady-state inactivation was V_0.5 = −50.1 ± 3.7 mV, and the slope factor k was −23.4 ± 3.9 mV (n = 7; N = 5; P > 0.05; Fig. 5B). Comparisons of the steady-state activation and inactivation parameters of the ScTx1-sensitive current are shown in Table 2. Table 2 also presents published data on homotetrameric K_V2.1 and heterotetrameric K_V2.1/silent K_V channels in heterologous expression systems and native tissues such as neurons, arterial and urethral smooth muscle, and mouse DSM. Most importantly, as shown in Table 2, the steady-state properties of the ScTx1-sensitive K_V current correspond to a channel population of homotetrameric K_V2.1 and heterotetrameric K_V2.1/silent K_V channels.

Table 3.

Comparison of time constants of total whole cell K_V, ScTx1-sensitive, and ScTx1-insensitive currents and published data on the homotetrameric K_V2.1 and heterotetrameric K_V2.1/silent K_V channel currents

	Time Constant of Activation		Time Constant of Inactivation
	τ, ms	Ref.	τ, s	Ref.
Control current in guinea pig DSM	25.5 ± 2.1 (+40 mV)		5.4 ± 0.8 (+40 mV)
ScTx1-insensitive current in guinea pig DSM	23.7 ± 1.2 (+40 mV)		5.4 ± 1.3 (+40 mV)
ScTx1-sensitive current in guinea pig DSM	35.3 ± 3.6 (+40 mV)		5.9 ± 1.6 (+40 mV)
KV2.1 current	15.6 ± 0.9 (+30 mV)	52	7.0 (+40 mV)	34
	21 ± 1 (+20 mV)	34
	7 (+10 mV)	44
	74 ± 3 (−10 mV)	34
	68.4 ± 6.8 (+30 mV)	42
KV current in mouse DSM	29.9 ± 2.0 (+30 mV)	56	3.4 ± 0.2 (+20 mV)	56
KV2.1/KV6.3	22.6 ± 3.2 (+30 mV)	42	NA
KV2.1/9.1 current	14.3 ± 0.8 (+30 mV)	52	NA
KV2.1/9.2 current	20.3 ± 1.3 (+30 mV)	52	NA
KV2.1/9.3 current	20.1 ± 1 (+20 mV)	34	16.2 ± 1.8 (+40 mV)	34
	79 ± 2 (−10 mV)	34
	4.9 (+10 mV)	44
	7.3 ± 0.6 (+60 mV)	31a

Values are means ± SE of the time constants (τ) of activation and inactivation (with action potential given in parentheses) from our experiments in guinea pig DSM, as well as data from the literature for homotetrameric K_V2.1 and heterotetrameric K_V2.1/silent K_V channel currents.

ScTx1 affects the kinetics of K_V current activation.

We investigated the effect of 100 nM ScTx1 on the time constants of activation (τ-activation) and inactivation (τ-inactivation) of the K_V current. Figure 6A shows representative K_V current traces evoked by depolarization steps to +40 mV (400 ms), which is close to the peak of the action potential in DSM. The data analysis showed that ScTx1 (100 nM) decreased the time constant of total outward current activation at +40 mV from 25.5 ± 2.1 to 23.7 ± 1.2 ms (n = 12; N = 9; P < 0.05; Fig. 6A). The time constant of activation of the ScTx1-sensitive current was 35.3 ± 3.6 ms at +40 mV (Fig. 6B).

Fig. 6.

Effects of ScTx1 on time constants (τ) of activation and inactivation of the K_V current. A: representative whole cell K_V current traces evoked by depolarization steps to +40 mV from a holding potential (V_h) of −70 mV in the presence or absence of 100 nM ScTx1. The subtracted ScTx1-sensitive current is also shown. Bar graphs show the time constants of activation of the control current, ScTx1-insensitive current, and ScTx1-sensitive current at 0 and +40 mV. ScTx1 decreased significantly the time constant of activation at +40 mV (n = 12; N = 9). *P < 0.05. B: ScTx1 (100 nM) did not affect time constant kinetics of inactivation of K_V current elicited by a voltage-step depolarization to +40 mV (protocol as shown in Fig. 5A; n = 7; N = 5; P > 0.05). Values are means ± SE. All experiments were performed in the presence of the selective BK channel inhibitor paxilline (100 nM).

The time constant of inactivation was determined using the stimulation protocol shown in Fig. 5A. ScTx1 did not affect the kinetics of inactivation of K_V current. The time constant of inactivation was 5.4 ± 0.8 s in the absence of ScTx1 and 5.4 ± 1.3 s in the presence of 100 nM ScTx1 at +40 mV (n = 7; N = 5; P > 0.05; Fig. 6B). The time constant of inactivation of the subtracted ScTx1-sensitive current was 5.9 ± 1.6 s at +40 mV (Fig. 6B).

Collectively, our findings indicate that at positive voltages, ScTx1 has a statistically significant effect on the kinetics of the K_V current activation. The time constants of activation and inactivation of the ScTx1-sensitive current obtained experimentally in our study compared with published data on homotetrameric K_V2.1 and heterotetrameric K_V2.1/silent K_V channels are summarized in Table 3.

ScTx1 does not affect the resting membrane potential in guinea pig DSM cells.

We used the amphotericin-perforated, current-clamp mode of the patch-clamp technique to test the effect of ScTx1 on the resting membrane potential in guinea pig DSM cells in the absence of paxilline. In current-clamp mode, we recorded at least 7–10 min of stable control before applying 100 nM ScTx1 for another 10 min. ScTx1 (100 nM) did not affect the resting membrane potential. The values of the resting membrane potential were −32.2 ± 2.3 mV under control conditions and −32.9 ± 3.2 mV in the presence of 100 nM ScTx1 (n = 10; N = 6; P > 0.05).

ScTx1 increases spontaneous and nerve-evoked contractions of isolated DSM strips.

We studied the effect of ScTx1 on physiologically relevant spontaneous (myogenic) phasic and tonic contractions of guinea pig DSM isolated strips. To minimize any possible effects caused by neurotransmitter release, we performed all experiments with the exception of the EFS-induced contractions in the presence of 1 μM TTX, a neuronal Na⁺ channel blocker. DSM strips were preincubated with 1 μM TTX before the experiments. Isometric DSM tension recordings of isolated guinea pig DSM strips showed that ScTx1 (100 nM) increased spontaneous phasic contraction amplitude, muscle force integral (area under the curve), and muscle tone without any effect on the phasic contraction frequency (n = 13; N = 7; P < 0.05; Fig. 7). Both ScTx1-induced and spontaneous phasic contractions were completely inhibited by 1 μM nifedipine, a selective blocker of the L-type voltage-gated Ca²⁺ channels (n = 4, N = 4).

Fig. 7.

ScTx1 increases spontaneous phasic contraction amplitude, muscle force integral, and muscle tone in guinea pig DSM strips. A: representative original trace of spontaneous phasic DSM contractions and the effect of 100 nM ScTx1. B: summary data showing a statistically significant increase in DSM spontaneous phasic contraction amplitude, muscle force integral, and muscle tone. For comparison of the phasic contraction parameters of the DSM strips, data were normalized to the spontaneous contractions, which were taken to be 100%, and then expressed as percentages. DSM force is presented as muscle force integral and was determined by integrating the area under the curve of the phasic contractions component. The tone of the DSM strips was determined by measuring the phasic contraction baseline curve. To evaluate the effect of the ScTx1 on the phasic contractions, we analyzed a 5-min period before ScTx1 application for the controls and another 5-min period 30 min after ScTx1 application (100 nM). Values are means ± SE (n = 13; N = 7). *P < 0.05; **P < 0.01. Tetrodotoxin (1 μM) was present throughout the experiments.

In the next experimental series, we studied the effect of ScTx1 on DSM contractions under conditions of sustained membrane depolarization, induced by either 20 mM KCl or 1 μM carbachol. ScTx1 (100 nM) slightly increased only the carbachol-induced phasic contraction amplitude (n = 9; N = 9; P < 0.05; Fig. 8; Table 4) but had no effect on any other parameters of the DSM contractions induced by 1 μM carbachol (n = 9; N = 9; P > 0.05; Fig. 8; Table 4) or 20 mM KCl (n = 12; N = 10; P > 0.05; Fig. 8; Table 4). After application of ScTx1 (100 nM), nifedipine (1 μM) completely inhibited the DSM contractility of strips precontracted with 1 μM carbachol (n = 8; N = 7) or 20 mM KCl (n = 4; N = 4).

Fig. 8.

Representative original traces showing the effect of ScTx1 on the phasic contractions induced by 1 μM carbachol (top trace) and 20 mM KCl (bottom trace) in DSM isolated strips. ScTx1 (100 nM) did not substantially affect the parameters of the 1 μM carbachol- or 20 mM KCl-induced phasic contractions in DSM isolated strips (see also Table 4).

Table 4.

Effects of ScTx1 on DSM contraction parameters under conditions of KCl- or carbachol-induced membrane depolarization

	Amplitude, %change	Force, %change	Frequency, %change	Tone, %change
Carbachol (1 μM)-induced contractions	136.8 ± 14.70*	106.0 ± 9.8	100.1 ± 3.3	97.8 ± 1.8
K+ (20 mM)-induced contractions	108.3 ± 10.6	105.9 ± 11.7	92.7 ± 6.7	99.9 ± 2.4

Values are means ± SE of %change in DSM contraction parameters under conditions of membrane depolarization induced by 1 μM carbachol or 20 mM KCl (K⁺) in the presence of 100 nM ScTx1. Carbachol results are from 9 DSM strips obtained from 9 guinea pigs; KCl results are from 12 DSM strips obtained from 10 guinea pigs.

^*P < 0.05, ScTx1 slightly increased phasic contraction amplitude only under conditions of depolarization with carbachol. ScTx1 had no other statistically significant effect on DSM contraction parameters.

Our next goal was to investigate whether ScTx1-sensitive K_V channels modulate the nerve-evoked (EFS-induced) contractions of guinea pig DSM. These experiments were conducted in the absence of TTX. Figure 9 shows original EFS-induced DSM contraction recordings and EFS frequency-response curves created by measuring the amplitude of the EFS-induced contraction at stimulation frequencies of 5, 7.5, 10, 12.5, 15, 20, 30, 40, and 50 Hz. An initial EFS frequency-response curve was generated under control conditions, and then DSM strips were preincubated for 30 min with 100 nM ScTx1. After this procedure, a second EFS frequency-response curve was generated using the same EFS parameters as indicated above. The results showed that EFS frequency-responses of guinea pig DSM strips were significantly increased in the presence of 100 nM ScTx1 at all stimulation frequencies (n = 6; N = 5; P < 0.05; Fig. 9). To confirm the stability of the guinea pig DSM strips, we also performed time controls by generating two consecutive EFS frequency-response curves in a 30-min interval without applying ScTx1. There was no statistically significant difference between the two EFS frequency-response curves in the absence of ScTx1 (n = 4; N = 4; P > 0.05). The EFS experiments suggest that ScTx1-sensitive K_V channels also work to oppose the nerve-evoked contractions in guinea pig DSM.

Fig. 9.

ScTx1 increases the amplitude of the electrical field stimulation (EFS)-induced contractions in DSM isolated strips. A: original traces of EFS-induced contractions (5–50 Hz) in the absence and presence of 100 nM ScTx1. B: summary data showing a significant increase in the amplitude of the EFS (5–50 Hz)-induced contractions following 30-min application of 100 nM ScTx1. The maximal EFS contraction amplitude at a stimulation frequency of 50 Hz under control conditions was taken to be 100%, and the contractions were normalized. Values are means ± SE (n = 6; N = 5). *P < 0.05, **P < 0.01.

DISCUSSION

In this study, we revealed that K_V2.1-containing channels are physiologically relevant regulators of guinea pig DSM excitability and contractility. On the basis of our data obtained from molecular biology, patch-clamp, and functional studies on DSM contractility, we can draw three important conclusions. First, K_V2.1 and several silent K_V channel subunits (K_V6.2–6.3, K_V8.2, and K_V9.1–9.3) are expressed in guinea pig DSM. Second, ScTx1-sensitive K_V channels contribute to the total K_V current in isolated DSM cells. Third, ScTx1-sensitive K_V channels regulate DSM contractility by opposing both myogenic and nerve-evoked contractions.

In a recent article (9), we reported that ScTx1-sensitive K_V channels are expressed and functional in rat DSM. We have since extended our study and hypothesized that a whole population of ScTx1-sensitive K_V2.1-containing channels assembled by various silent K_V channel subunits are key regulators of DSM excitability and contractility of guinea pig DSM. To test this hypothesis, we applied a multitechnique molecular biology approach to elucidate the presence of all known ScTx1-sensitive K_V channels in guinea pig DSM. The RT-PCR experiments showed mRNA expression of K_V2.1, K_V6.2–6.3, K_V8.2, and K_V9.1–9.3 in both whole DSM tissue and single DSM cells and ruled out the expression of two other ScTx1-sensitive channels, the K_V2.2 and K_V4.2 homotetramers, along with K_V5.1, K_V6.1, K_V8.1 silent subunits (Fig. 1). Using Western blot and immunocytochemistry, we confirmed the expression of the K_V2.1 channel protein in guinea pig DSM (Figs. 2 and 3). Therefore, the ScTx1-sensitive K_V current in guinea pig DSM cells is determined by K_V2.1-containing channels. Consistent with our findings, the K_V2.1 subunit has been reported to be the major K_V subunit in rat and mouse DSM (9, 17, 41, 56). From the electrically silent subunits that we identified (K_V6.2–6.3, K_V8.2, and K_V9.1–9.3), only K_V6.3 and K_V9.3 have been reported to contribute to the ScTx1-sensitive current by forming K_V2.1/6.3 and K_V2.1/9.3 ScTx1-sensitive K_V heterotetramers (14, 15, 40). It is also well known that the K_V9.3 subunit combines only with the K_V2.1 subunit to form a functional heterotetramer and remains electrically silent if expressed alone in artificial systems (14, 32–34, 44, 45). Although ScTx1 sensitivity has not yet been reported for all combinations of K_V2.1 and K_V silent subunits, the most plausible assumption is that all heterotetrameric K_V2.1/K_V silent subunits will be ScTx1-sensitive, since the K_V2 subunit must be part of the channel.

Using the amphotericin-perforated patch-clamp technique, we investigated the properties of the guinea pig DSM K_V current under conditions of paxilline-blocked BK current, which is the main voltage-dependent K⁺ current in DSM cells (7, 25, 46, 48). The application of 100 nM ScTx1 inhibited ∼22% (at 0 mV) of the remaining K_V current amplitude in guinea pig DSM cells (Fig. 4). Most likely, the residual ScTx1-insensitive current is encoded by other types of K_V channels. Also, ScTx1 affected the K_V current time constant of activation (Fig. 6A).

Our molecular biology studies did not detect the expression of K_V2.2 or K_V4.2 channel subunits (Fig. 1), and therefore we excluded those channels as possible candidates determining the observed ScTx1-sensitive K_V current in guinea pig DSM. Because we identified the mRNA expression of only K_V2.1 and electrically silent K_V channel subunits, we hypothesized that the ScTx1-sensitive K_V current could be attributed to homotetrameric K_V2.1 channels and/or heterotetrameric K_V2.1/silent K_V channels. Our detailed analyses of the biophysical properties of the ScTx1-sensitive K_V current in guinea pig DSM demonstrate substantial similarities to those reported for heterotetrameric K_V2.1/silent K_V channels (Tables 2 and 3). Specifically, our data indicate that the V_0.5 of the steady-state inactivation of the ScTx1-senstitive current was about −50 mV, which is more negative than the reported values for homotetrameric K_V2.1 channels and consistent with the reported values for heterotetrameric K_V2.1/silent K_V channels (Table 2). In addition, this V_0.5 value of −50 mV is consistent with the recently reported V_0.5 for the ScTx1-sensitive current in smooth muscle cells isolated from rabbit urethra (37). Therefore, heterotetrameric assembly of K_V2.1 and silent K_V channel subunits accounts for the ScTx1-sensitive inhibition of the K_V current in guinea pig DSM. Although we do not exclude the possible involvement of the K_V2.1 homotetrameric channel, we assume that a population of heterotetrameric K_V2.1/silent K_V channels most likely underlies the K_V current in guinea pig DSM because all these silent subunits cannot form functional channels and associate only with the K_V2 family. This assumption is also supported by findings of Thorneloe and Nelson (56) suggesting that K_V2.1 does not form its own homotetramer in mouse DSM. They further suggest that in mouse DSM, K_V2.1 can potentially coassemble with K_V5.1 and K_V6 subunits (56). In the present study, however, we did not find K_V5.1 or K_V6.1 subunit but confirmed the expression of K_V6.2–6.3 subunits (Fig. 1). This may be due to species differences. We have also further identified the expression of K_V8.2 and K_V9.1–9.3 silent subunits in guinea pig DSM cells (Fig. 1).

It has been suggested that K_V2.1/9.3 heterotetramer contributes to the excitability in pulmonary and cerebral arteries (1, 44, 61). We observed in current-clamp mode that ScTx1 did not affect the cell resting membrane potential of guinea pig DSM cells. Analysis of steady-state activation and inactivation properties of the K_V channels (Figs. 4 and 5) showed that the ScTx1-sensitive channels are likely active at voltages close to the level of the recorded resting membrane potential in guinea pig DSM cells (approximately −32 mV), although their activity at these voltages was relatively low based on the steady-state activation curve. The lack of ScTx1 effect of the resting membrane potential could be explained by the relatively minor contribution of the ScTx1-sensitive current to the total whole cell outward current in DSM cells, as shown in the current-voltage relationship in Fig. 4B. Furthermore, our voltage-clamp experiments were performed in the presence of paxilline to eliminate the BK channel, which is the main contributor to the total whole cell outward current in DSM cells (30, 31, 48), whereas the current-clamp recordings were performed in the absence of paxilline with physiologically active BK channels. On the basis of these results, we conclude that under physiological conditions, ScTx1-sensitive channels are not the main contributors to setting the resting membrane potential in guinea pig DSM cells. However, they regulate excitability at more positive potentials, during the peak of an action potential (Fig. 4). Consistently, Hayase et al. (24) reported no effect of ScTx1 on the resting membrane potential in mouse DSM recorded by intracellular microelectrodes. Our data support the hypothesis that K_V2.1-containing channels most likely control the repolarization phase of the action potential and perhaps also the slow afterhyperpolarization in guinea pig DSM.

In the bladder filling phase, spontaneous contractions do not increase substantially intravesical pressure and thus do not initiate voiding (2, 3, 28). During bladder filling, spontaneous contractions occur locally and do not spread throughout the tissue, probably due to a weak electrical coupling between the DSM cells (21, 22). Voiding contractions occur after excitation of the parasympathetic nerves, which causes synchronized contraction of the whole bladder, followed by elevation of the intravesical pressure (2, 3, 26). In guinea pig DSM, a burst of action potentials triggers a single phasic contraction (21). Blocking the K_V2.1-containing channels with ScTx1 is expected to increase the action potential frequency within a burst of action potential, and therefore the amplitude of the phasic contraction, without any effect on the phasic contraction frequency (Fig. 7). Our studies on DSM contractility revealed that ScTx1 significantly increased the spontaneous phasic contraction amplitude, muscle force integral, and muscle tone in isolated guinea pig DSM strips (Fig. 7), consistent with previous findings in rat DSM (9). However, ScTx1 had no effect on the DSM contractions induced by 1 μM carbachol or 20 mM KCl with the exception of slightly increased phasic contraction amplitude in the presence of 1 μM carbachol (Table 4; Fig. 8). These results demonstrate that under experimental conditions of membrane depolarization, the K_V2.1-containing channels have no further contribution to the membrane excitability and contractility (Table 4; Fig. 8). The K_V2.1-containing channels, by decreasing membrane excitability, work to oppose spontaneous contractions under physiological conditions. Our results also showed that ScTx1 significantly increased the amplitude of the EFS-induced contractions over a range of stimulation frequencies, 5–50 Hz (Fig. 9). This indicates that K_V2.1-containing channels, by decreasing membrane excitability, also oppose the DSM contractility in response to excitatory neurotransmitters.

Under pathological conditions, such as some forms of OAB, spontaneous myogenic contractions can also cause a rise of intravesical pressure, initiating involuntary voiding (3). These pathological conditions are poorly understood, and the current pharmacological treatment is limited to antimuscarinics that have many adverse effects (3). Our study shows that pharmacological block of K_V2.1-containing channels with ScTx1 increases DSM contractility (Figs. 7 and 9). Therefore, genetic mutations causing K_V2.1 channel malfunction would be expected to cause DSM dysfunction. Consistently, other studies also suggest the implication of the K_V2.1 channels in OAB (17, 41). Thus a better understanding of K_V2.1-containing channels is an important step for developing new therapeutic strategies for treatment of OAB and UI. The K_V2.1-containing channels may represent new targets for pharmacological or genetic control of OAB.

GRANTS

This study was supported by National Institutes of Health Grants DK084284, DK083687, and UL1RR029882 (to G. V. Petkov).

DISCLAIMER

The contents are solely the responsibility of the authors and do not necessarily represent the official views of the National Institutes of Health.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: K.L.H., M.C., and G.V.P. conception and design of research; K.L.H., M.C., R.P.S., S.P.P., Q.C., and W.F.K. performed experiments; K.L.H., M.C., R.P.S., Q.C., and W.F.K. analyzed data; K.L.H., M.C., R.P.S., S.P.P., and G.V.P. interpreted results of experiments; K.L.H., M.C., R.P.S., Q.C., W.F.K., and G.V.P. prepared figures; K.L.H. and G.V.P. drafted manuscript; K.L.H., M.C., R.P.S., S.P.P., Q.C., W.F.K., and G.V.P. edited and revised manuscript; K.L.H., M.C., R.P.S., S.P.P., Q.C., W.F.K., and G.V.P. approved final version of manuscript.