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The Journal of Physiology logoLink to The Journal of Physiology
. 2018 Nov 22;597(1):137–149. doi: 10.1113/JP276806

Oxytocin can regulate myometrial smooth muscle excitability by inhibiting the Na+‐activated K+ channel, Slo2.1

Juan J Ferreira 1, Alice Butler 2, Richard Stewart 2, Ana Laura Gonzalez‐Cota 1, Pascale Lybaert 1,3, Chinwendu Amazu 1, Erin L Reinl 1,4, Monali Wakle‐Prabagaran 1, Lawrence Salkoff 2, Sarah K England 1, Celia M Santi 1,2,✉
PMCID: PMC6312452  PMID: 30334255

Abstract

Key points

  • At the end of pregnancy, the uterus transitions from a quiescent state to a highly contractile state. This transition requires that the uterine (myometrial) smooth muscle cells increase their excitability, although how this occurs is not fully understood.

  • We identified SLO2.1, a potassium channel previously unknown in uterine smooth muscle, as a potential significant contributor to the electrical excitability of myometrial smooth muscle cells.

  • We found that activity of the SLO2.1 channel is negatively regulated by oxytocin via Gαq‐protein‐coupled receptor activation of protein kinase C. This results in depolarization of the uterine smooth muscle cells and calcium entry, which may contribute to uterine contraction.

  • These findings provide novel insights into a previously unknown mechanism by which oxytocin may act to modulate myometrial smooth muscle cell excitability. Our findings also reveal a new potential pharmacological target for modulating uterine excitability.

Abstract

During pregnancy, the uterus transitions from a quiescent state to a more excitable contractile state. This is considered to be at least partly a result of changes in the myometrial smooth muscle cell (MSMC) resting membrane potential. However, the ion channels controlling the myometrial resting membrane potential and the mechanism of transition to a more excitable state have not been fully clarified. In the present study, we show that the sodium‐activated, high‐conductance, potassium leak channel, SLO2.1, is expressed and active at the resting membrane potential in MSMCs. Additionally, we report that SLO2.1 is inhibited by oxytocin binding to the oxytocin receptor. Inhibition of SLO2.1 leads to membrane depolarization and activation of voltage‐dependent calcium channels, resulting in calcium influx. The results of the present study reveal that oxytocin may modulate MSMC electrical activity by inhibiting SLO2.1 potassium channels.

Keywords: SLO2.1 Potassium channels, Oxytocin, Smooth muscle, Myometrium

Key points

  • At the end of pregnancy, the uterus transitions from a quiescent state to a highly contractile state. This transition requires that the uterine (myometrial) smooth muscle cells increase their excitability, although how this occurs is not fully understood.

  • We identified SLO2.1, a potassium channel previously unknown in uterine smooth muscle, as a potential significant contributor to the electrical excitability of myometrial smooth muscle cells.

  • We found that activity of the SLO2.1 channel is negatively regulated by oxytocin via Gαq‐protein‐coupled receptor activation of protein kinase C. This results in depolarization of the uterine smooth muscle cells and calcium entry, which may contribute to uterine contraction.

  • These findings provide novel insights into a previously unknown mechanism by which oxytocin may act to modulate myometrial smooth muscle cell excitability. Our findings also reveal a new potential pharmacological target for modulating uterine excitability.

Introduction

At the end of pregnancy, the uterus transitions from a quiescent non‐contractile state to an active contractile state to expel the fetus. Uterine quiescence depends on maintaining the human myometrial smooth muscle cell (hMSMC) resting membrane potential at a sufficiently negative voltage to prevent excessive calcium (Ca2+) influx through voltage‐dependent Ca2+channels (VDCCs). In hMSMCs, the membrane potential is determined by a balance between a hyperpolarizing force generated by an outward potassium (K+) leak current and opposing depolarizing forces generated, in part, by an inward sodium (Na+) leak current, which is probably carried by Na+ leak channel, non‐selective (NALCN) (Reinl et al. 2015). Through most of pregnancy, the K+ leak current is dominant and maintains the cell at a sufficiently negative membrane potential to prevent premature uterine contraction. At the end of pregnancy, the delicate balance of K+ and Na+ leak currents must be modified to allow depolarization and uterine contraction (Casteels & Kuriyama, 1965; Parkington et al. 1999). One important mechanism for modifying this balance is hormonally‐regulated inhibition of the K+ leak current, thus permitting the Na+ leak current to gain prominence, depolarizing the resting membrane potential and opening VDCCs. Such a K+ leak channel must be able to be open at the cell resting membrane potential and to be closed by hormones present at the end of pregnancy. Although hMSMCs have been reported to express several types of K+ channels, none have been identified with the required properties (Anwer et al. 1993; Knock et al. 1999; Coleman et al. 2000; Brainard et al. 2007; McCloskey et al. 2014).

Another important regulator of the transition from uterine quiescence to contractility is the peptide hormone oxytocin (OXT). In the canonical activation pathway, OXT binds to the oxytocin receptor (OXTR), which is a Gαq‐protein‐coupled receptor that signals through phospholipase C (PLC) (Arthur et al. 2007; Arrowsmith & Wray, 2014). Activation of PLC leads to an increase in diacylglycerol andinositol 3‐phosphate (IP3), which activates the IP3 receptor and causes Ca2+ release from intracellular stores. This increased intracellular Ca2+ activates myosin light chain kinase to phosphorylate myosin, subsequently leading to myometrial cell contraction (Wray, 2007; Aguilar & Mitchell, 2010). Additionally, it has been proposed that OXT induces membrane depolarization and Ca2+ influx through VDCCs. However, the mechanism by which OXTR activation causes depolarization is unknown (Mironneau, 1976).

In the present study, we present evidence that the K+ channel SLO2.1 is a key regulator of myometrial excitability and links membrane potential and OXT‐dependent depolarization. We show that SLO2.1 is expressed in hMSMCs, is open at resting membrane potential, and is inhibited by OXT signalling through the Gαq‐protein‐coupled receptor pathway, resulting in membrane depolarization, VDCC activation and Ca2+ influx. Given the increase in OXTR activation at the end of pregnancy in women (Kimura et al. 1996; Cook et al. 2000), inhibition of SLO2.1 might be a key mechanism of regulating uterine contraction at this stage.

Methods

Ethical approval

The present study conformed with the Declaration of Helsinki and was approved by the Institutional Review Board at Washington University School of Medicine (approval no. 201108143) except for registration in a database. All subjects signed written consent forms approved by the Washington University in St Louis Internal Review Board.

Human myometrial tissue samples from the lower uterine segment were obtained from non‐labouring women at term (≥37 weeks of gestation) during elective Caesarean section under spinal anaesthesia. The recruited subjects had a history of repeat Caesarean sections with no spontaneous or induced labour. Samples were stored in ice‐cold PBS and processed for explants expansion and smooth muscle cell isolation within 60 min of acquisition.

Cell culture

Briefly, myometrial tissue was washed in cold Dulbecco's PBS containing 50 μg mL−1 gentamicin and 5 μg mL−1 fungizone (Gibco‐BRL, Gaithersburg, MD, USA). The tissue was then cut into 2–3 mm pieces and cultured in Dulbecco's modified Eagle's medium:nutrient mixture F‐12 (DMEM:F12) supplemented with 5% fetal bovine serum (FBS), 0.2% fibroblast growth factor‐β, 0.1% epidermal growth factor, 0.05% insulin, 0.05% gentamicin and 0.05% fungizone. Once explant colonies formed, they were expanded. Primary hMSMCs and human telomerase reverse transcriptase‐transformed human myometrial (hTERT‐HM) cells (Condon et al. 2002) were incubated at 37°C and 5% CO2 in DMEM:F12 medium with 10% FBS, 100 units mL−1 penicillin and 100 μg mL−1 streptomycin. For all experiments, hMSMCs were used at passage 1 or 2.

RNA isolation, reverse transcription and PCR

The Qiagen RNeasy Kit (catalogue no. 74124; Qiagen, Valencia, CA, USA) was used to isolate total RNA from primary hMSMCs, uterine tissue obtained from pregnant and non‐pregnant patients, and hTERT‐HM cells. cDNAs from these RNA preparations and from total human brain RNA (Ambion catalogue no. AM7962; Thermo Fisher Scientific Inc., Waltham, MA, USA) were generated with Superscript III Reverse Transcriptase (catalogue no. 18080‐044; Invitrogen, Carlsbad, CA, USA) and random hexanucleotide primers from Sigma‐Aldrich (catalogue no. H0268‐1UN; Sigma‐Aldrich, St Louis, MO, USA). The PCR primers were designed with the help of Oligo 7 software from Molecular Biology Insights (Colorado Springs, CO, USA) and made by Integrated DNA Technologies (Coralville, IA, USA). Human SLO2.1 PCR primers (S4S:5′‐ATGATCTACACAGAGCCATTCAGCGTACACAG‐3′; RCK1AS:5′‐GATGATGTCCTATCCACTTCACAACGGCTACT‐3′) were designed to amplify a 640 bp cDNA amplicon and SLO2.2 primers (S4S:5′‐CATGATTAATGACTTCCACCGTGCCATCCT‐3′; RCK1AS:5′‐GTCCACCTCGTTCCTGCTGCTGA‐3′) were designed to amplify a 657 bp cDNA amplicon. To reduce amplification of genomic DNA, both primer sets were designed to amplify a region that spans four intron splice junctions and the S4S primer spanned an intron splice junction. PCR was performed with the KAPA2G Fast HotStart PCR Kit (KAPA Biosystems, Wilmington, MA, USA).

Small interfering RNA

hTERT‐HM cells and hMSMCs were transfected with scrambled small interfering RNAs (siRNA) (control) or siRNA targeting SLO2.1. Briefly, each plate was transfected with a combination of three 27‐mer siRNAs from OriGene (catalogue no. SR317593; Origene, Rockville, MD, USA) targeting the human SLO2.1 gene, KCNT2, at a final concentration of 25 nm each, or with 75 nm scrambled siRNA. Lipofectamine 2000 transfection reagent (catalogue no. 11668‐027; Invitrogen) was used in accordance with the manufacturer's instructions. As a marker for transfection, cells were also co‐transfected with 10 μg mL−1 of a vector encoding green fluorescent protein. Experiments were performed 36–72 h after transfection.

Immunocytochemistry

Primary myometrial cells from pregnant non‐laboring women were cultured in DMEM with 10% FBS. Cells were fixed in 4% paraformaldehyde in PBS for 15 min at room temperature, washed with PBS and 10 mm glycine, then stored at 4°C. Briefly, cells were permeabilized with 0.2% saponin in Tris buffered saline (TBS) for 10 min, incubated in blocking buffer (TBS with 2% FBS) for 1 h, incubated with primary antibodies diluted in blocking buffer for 1 h at room temperature, washed for ∼30 min in three or four changes of TBS, incubated with secondary antibodies diluted in blocking buffer for 1 h, washed, incubated with 1 μg mL−1 of 4′,6‐diamidino‐2‐phenylindole (DAPI), and then mounted in Fluoro‐shield (Sigma‐Aldrich). The antibodies used were: rabbit polyclonal anti‐SLO2.1 (catalogue no. APC‐126; Alomone Labs, Jerusalem, Israel; dilution 1:1000), mouse monoclonal anti‐SLO1 (catalogue no. 73‐022; clone L6/60; NIH/NeuroMabs, Davis, CA, USA; dilution 1:50), monoclonal anti‐actin α‐smooth muscle (catalogue no. A2547; Sigma‐Aldrich) and appropriate fluorophore‐coupled secondary antibodies. Mounted cells were imaged with a 60× oil objective on an E800 microscope (Nikon, Tokyo, Japan) with a CoolSnapEZ camera (Qimaging, Surrey, BC, Canada). Raw images were minimally adjusted with ImageJ (NIH, Bethesda, MD, USA). To assess the specificity of SLO2.1 immunostaining, cells were stained without any primary antibody or with antibody that was pre‐incubated with SLO2.1 peptide antigen. All controls were processed with the same parameters employed for the SLO2.1 antibody.

Electrophysiology

Cells were serum starved in plain DMEM for at least 2 h before the patch clamp experiments. Borosilicate glass pipettes (Warner Instruments, Hamden, CT, USA) with 0.8–1.8 megaohm resistance were used for inside‐out macro‐patch and whole‐cell recordings, and pipettes with 4–6 MΩ resistance were used for single‐channel recordings.

Whole‐cell recordings of hMSMCs and hTERT‐HM cells were performed in asymmetrical K+ solutions (unless specified) with the compositions (in mm): external solution, 150 NaCl (replaced by 150 choline Cl, pH 7.4, in the Na‐free or 0 Na+ solution), 5 KCl, 5 Hepes, 2 MgCl2; internal solution, 140 KCl, 5 Hepes, 0.5 MgCl2, 5 ATPMg (0.6 mm free Mg2+) and 10 EGTA (0 Ca2+ solution), or 1 EGTA and 100 nm Ca2+ free (Ca2+‐containing solution). Inside‐out macro patch and single‐channel patch clamp recordings were performed in symmetrical K+ solutions. The internal solution contained (in mm): 80, 140 or 160 KCl, 80 NaCl (or 80 choline Cl for the 0 sodium condition) and 10 Hepes. The pipette solution comprised (in mm): 80, 140 or 160 KCl, 80 NaCl, 2 MgCl2 and 10 Hepes. The pH of all solutions was adjusted to 7.2. Any alterations in solutions are indicated, as appropriate.

Traces were acquired with Axopatch 200B (Molecular Devices, Sunnyvale, CA, USA), digitized at 10 kHz for whole‐cell and macro‐patches recordings and at 100 kHz for single‐channel recordings. Records were filtered at 2 kHz for macro currents (whole‐cell and macro‐patches) and at 20 kHz for single‐channel recordings. Data were analysed with pClamp, version 10.6 (Molecular Devices) and SigmaPlot, version 12 (Systat Software Inc., Chicago, IL, USA). During electrophysiological experiments, cells and the intracellular side of the membrane were continuously perfused.

Two‐electrode voltage clamp experiments in Xenopus oocytes

Oocytes were harvested from female Xenopus laevis as described in Yuan et al. (2000). Defolliculated oocytes were injected with 46 and 92 ng of cRNA with a nano injector (Drummond Scientific, Broomall, PA, USA). Injected oocytes were kept at 18°C in ND96 medium containing (in mm): 96 NaCl, 2 KCl, 1.8 CaCl2, 1 MgCl2 and 5 Hepes, pH 7.5 (with NaOH). Two‐electrode voltage clamp experiments were performed 3–5 days after oocyte injection, as described previously (Wei et al. 1994). Whole‐cell current recordings from Xenopus oocytes were performed when the oocytes were being perfused with ND96 + 2 μm 4,4′‐diisothiocyanatostilbene‐2,2′‐disulphonic acid (catalogue no. D3514; Sigma‐Aldrich) to block endogenous chloride currents. All records were obtained with the voltage protocols specified. Data were acquired with a Digidata 1440 (Molecular Devices). Phorbol 12‐myristate 13‐acetate (PMA) (CAS number 16561‐29‐8; Sigma‐Aldrich) or OXT (catalogue no. 1910; Tocris Bio‐Techne Corporation, Minneapolis, MN, USA) were applied to the recording chamber by continuous perfusion.

Statistical analysis

Statistical analysis was performed using Sigmaplot, version 12.0 (Systat Software Inc.). An unpaired Student's t test was used to determine significant differences between independent samples tested. A paired t test was used to determine significant differences in case–control studies performed in the same individuals. Results are expressed as the mean ± SD. P < 0.05 was considered statistically significant.

Results

Human myometrial cells conduct a Na+‐activated K+ current (KNa)

To determine the types of K+ currents present in hMSMCs, we performed whole‐cell patch clamp experiments in which we applied depolarizing voltage clamp steps to the cells, revealing the presence of a time‐dependent, non‐inactivating outward K+ current (Fig. 1 A, Control). Because we suspected that part of this current was conducted by the large‐conductance Ca2+‐activated K+ channel SLO1 (BKCa, MaxiK), which is expressed in hMSMCs (Khan et al. 1993; Khan et al. 1998; Brainard et al. 2009; Lorca et al. 2014; Lorca et al. 2014; Wakle‐Prabagaran et al. 2016; Lorca et al. 2017; Lorca et al. 2018), we treated the cells with the SLO1 blocker tetraethylammonium (TEA) (5–10 mm). These experiments revealed two components of the macroscopic current: a TEA‐sensitive current and a TEA‐resistant current (Fig. 1 A). Approximately 53% of the total whole‐cell K+ current under these conditions was TEA resistant. Consistent with the whole‐cell recordings, single‐channel analysis of inside‐out patch clamp recordings revealed the presence of two predominant types of high‐conductance K+ channels in hMSMCs. One was activated by 50 μm intracellular Ca2+ in the absence of Na+ and had a single channel conductance of ∼241pS. The other was activated by 80 mm intracellular Na+ in the absence of Ca2+ and had a single channel conductance of ∼78 pS (Fig. 1 B). The larger, Ca2+‐activated conductance was characteristic of the SLO1 current in hMSMCs.

Figure 1. hMSMCs conduct a TEA‐resistant and Na+‐activated K+ current.

Figure 1

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A, left, schematic of whole‐cell recording set‐up. Middle, representative whole‐cell currents (V h = −70 mV, with step pulses from −80 to +150 mV) from hMSMC recorded in the absence (control) and presence of 5 mm TEA in the external solution. The TEA‐sensitive component was obtained by subtracting traces (control − TEA resistant). Right: TEA‐resistant component of the current at different external TEA concentrations (n = 13 cells, data plotted as the mean ± SD). The curve is fitted with an exponential decay function [f = y0 + a *exp(–b * x)]. B, left: schematic of inside‐out single‐channel recording set‐up. Middle: representative single‐channel recordings from a hMSMC under the indicated conditions. Right: single channel current I–V plots for the (1) Ca2+‐dependent (241 ± 9.9 pS, n = 4) and (2) Na+‐dependent (78 ± 6.3 pS, n = 5) channels. Data are plotted as the mean ± SD. [Color figure can be viewed at wileyonlinelibrary.com]

To further characterize the Na+‐activated current, we examined the Na+ sensitivity of the TEA‐resistant current in inside‐out macro patches in both the immortalized human myometrial cell line hTERT‐HM (Condon et al. 2002) and in hMSMCs. In both cell types, a substantial fraction of the TEA‐resistant outward K+ current was Na+‐dependent (Fig. 2 A). Similarly, in single‐channel patch clamp experiments in both cell types, we measured little channel activity at 0 mm Na+, whereas perfusing the bath with 80 mm Na+ (Fig. 2 B) resulted in a four‐ to five‐fold increase in K+ channel openings. The percentage of Na+‐sensitive K+ (KNa) current in macropatches (Fig. 2 A) and the open probabilities of these channels in single‐channel recordings (Fig. 2 B) were similar at −60 and +80 mV, indicating that the KNa current was not voltage‐dependent and that the channels were open at the physiological resting membrane potential of hMSMCs (−70 mV) (Parkington et al. 1999). Together, these results indicate that the KNa channel we measured could contribute to the myometrial resting membrane potential. The experiments shown in Fig. 2 and previous studies reporting the properties of KNa channels in inside‐out patch clamp experiments (Dryer et al. 1989; Yuan et al. 2003; Salkoff et al. 2006; Santi et al. 2006; Budelli et al. 2009; Kaczmarek, 2013) indicate that KNa channels are activated by Na+ interacting with the intracellular surface of the channel. However, whole‐cell patch clamp experiments in which the intracellular solution contained no Na+ demonstrated that influx of Na+ from the extracellular solution through channels that carry a persistent inward sodium leak, can activate KNa channels (Budelli et al. 2009; Hage & Salkoff, 2012). If KNa channel activation in myometrial cells depended on a persistent Na+ leak influx, we expected to measure a decrease in K+ currents when we decreased the extracellular Na+ from 150 to 0 mm (with no Na+ in the intracellular pipette solution) in whole‐cell patch clamp experiments. Indeed, in both hTERT‐HM cells and hMSMCs, the total K+ current was reduced markedly when the external solution contained no Na+; the current in 0 mm external Na+ was ∼40% of the current in 150 mm external Na+ (Fig. 3). These results indicate that the outward K+ current substantially depended on Na+ influx in both hTERT‐HM cells and hMSMCs.

Figure 2. Na+‐dependent K+ current in hTERT‐HM cells and hMSMCs.

Figure 2

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A, representative traces of Na+‐dependent K+ currents obtained from inside‐out macropatches (schematic on left) of hTERT‐HM cells and hMSMCs. Currents were recorded with 5–10 mm TEA in the pipette (extracellular) and in the presence and absence of intracellular Na+. Currents were elicited with V h = 0 mV and step pulses from −90 to +150 mV. The graphs show the percentage of Na+‐dependent current in hTERT‐HM cells (50 ± 12.7%, n = 5 and 64 ± 28.2%, n = 5 at −60 and +80 mV, respectively) and in hMSMCs (48 ± 13.9%, n = 3 and 44 ± 30.5%, n ± 3 at −60 and +80 mV, respectively). B, representative single‐channel recordings (schematic on left) of Na+‐dependent K+ currents recorded from hTERT‐HM cells and hMSMCs. Showing the ratios of the open probability of the channel in 0 mm Na+ vs. 80 mm Na+ in hTERT‐HM cells (24.6 ± 14.1%, n ± 3 and 17.8 ± 18.6%, n ± 10 at −60 and +80 mV, respectively) and in hMSMCs (21.3 ± 31%, n ± 3 and 31 ± 26.8%, n ± 3 at −60 and +80 mV, respectively). * P < 0.05, ** P < 0.001 comparing nPo at 0 mm and 80 mm Na+. A paired t test was used to determine statistical significance. [Color figure can be viewed at wileyonlinelibrary.com]

Figure 3. Na+ influx activates Na+‐dependent K+ channels in hTERT‐HM cells and hMSMCs.

Figure 3

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Whole‐cell recordings (schematics on left) of K+ currents from hTERT‐HM cells (A) and hMSMCs (B) in the presence of 150 mm and 0 mm extracellular Na+. Whole‐cell currents were elicited at V h ± −70 mV and step pulses from −90 to +150 mV. Na+‐dependent K+ currents were obtained by subtracting the current recorded in 0 mm Na+ from the current recorded at 150 mm Na+. Showing the percentage of Na+‐dependent K+ current in hTERT‐HM cells at −60 mV (43 ± 29%, n ± 7) and +80 mV (33 ± 17%, n ± 7) and in hMSMCs at −60 mV (37 ± 21%, n ± 4) and +80 mV (31 ± 18%, n ± 6).

SLO2.1 underlies the KNa currents in human myometrial cells

KNa currents with the properties that we have shown in the present study could be carried by either SLO2.1 or SLO2.2 channels (Yuan et al. 2003; Santi et al. 2006; Kaczmarek, 2013). To distinguish between these alternative KNa channel types, we used RT‐PCR to examine the expression of these channels in hMSMCs, hTERT‐HM cells and uterine tissue. RT‐PCR performed with primers specific to SLO2.1 showed that SLO2.1 mRNA was expressed in human uterine tissue, hMSMCs isolated from both pregnant and non‐pregnant women (Fig. 4 Aa to Ac) and hTERT‐HM cells (Fig. 4 Ad). By contrast, SLO2.2 was not detected in hMSMCs from pregnant or non‐pregnant women, although it was expressed in the uterine tissue (Fig. 4 Ba and Bb). Consistent with the RT‐PCR data, immunocytochemistry with a rabbit polyclonal antibody against SLO2.1 showed that the SLO2.1 protein was expressed in hMSMCs (Fig. 4 C). These cells also expressed SLO1 and beta‐actin, which are expressed in hMSMCs but not in fibroblasts (data not shown). Confirming that the SLO2.1 antibody effectively recognizes SLO2.1 channels, it was found that we could detect a robust signal in Chinese hamster ovary cells transfected with SLO2.1 but not in non‐transfected cells (data not shown).

Figure 4. SLO2.1 expression in hTERT‐HM and hMSMCs.

Figure 4

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A, representative RT‐PCR of SLO2.1 in hMSMCs from pregnant (Aa) and non‐pregnant (Ac) women, human term pregnant uterine tissue (UT), and hTERT‐HM cells (Ad). Negative (no reverse transcriptase; RT) and positive (human brain cDNA and SLO2.1 cDNA) controls are also shown (Ab and Ad). B, representative RT‐PCR of SLO2.2 in hMSMCs from pregnant women (Ba), non‐pregnant women (Bb), and human term pregnant (Ba) and non‐pregnant (Bb) uterine tissue. Negative (no RT) and positive (human brain cDNA and SLO2.2 cDNA) controls are also shown. C, immunolocalization of SLO2.1 (red) and DAPI (blue) in hMSMCs. Scale bars = 10 μm.

To determine whether the Na+‐dependent current and the TEA‐resistant current were carried by SLO2.1 channels, we transfected hTERT‐HM cells with either scrambled siRNA or SLO2.1‐targeted siRNAs and then performed whole‐cell patch clamp experiments in the presence or absence of extracellular Na+. In cells transfected with scrambled siRNA, the addition of 80 mm Na+ increased the K+ current as it did in non‐transfected cells. However, the addition of extracellular Na+ did not increase the K+ current in cells transfected with SLO2.1 siRNAs (Fig. 5 A). Similarly, TEA blocked 50% of the current in cells transfected with scrambled siRNA, whereas TEA blocked 80% of the K+ current in cells transfected with SLO2.1 siRNA (Fig. 5 B). Thus, we conclude that SLO2.1 is expressed in myometrial cells and is required for the Na+‐dependent and TEA‐resistant K+ current in these cells.

Figure 5. SLO2.1 channels carry the Na+‐sensitive and the TEA‐resistant K+ current in hTERT‐HM cells.

Figure 5

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A, representative whole‐cell K+ currents (schematics on left) at 80 mm or 0 mm extracellular Na+ in hTERT‐HM cells transfected with scrambled siRNA or SLO2.1 siRNA. Whole‐cell currents were elicited at V h ± 0 mV and step pulses from −90 to +150 mV. Na+‐dependent K+ currents were obtained by subtracting traces (80 mm − 0 mm Na+). Showing the Na+‐dependent current as a percentage of the control in cells transfected with scrambled siRNA (75 ± 31%, n ± 3) and cells transfected with SLO2.1 siRNA (−10 ± 21%, n ± 6). B, representative whole‐cell K+ currents (schematics on left) from hTERT‐HM cells transfected with scrambled siRNA or SLO2.1 siRNA in the presence or absence of 10 mm TEA in the external solution. TEA‐sensitive currents were obtained by subtracting traces (control − TEA resistant). Showing the TEA‐resistant current as a percentage of control in cells transfected with scrambled siRNA (52 ± 0.17%, n ± 8) and cells transfected with SLO2.1 siRNA (18 ± 0.09%, n ± 17). ** P ± 0.002, *** P < 0.001. An unpaired t test was used to determine statistical significance.

Oxytocin inhibits SLO2.1 currents in hMSMCs

In neurons, SLO2.1 is inhibited by protein kinase C (PKC), which is activated by binding of neuromodulators to Gαq‐protein‐coupled receptor (Santi et al. 2006; Chen et al. 2009; Kaczmarek, 2013). Given that OXT binds and activates the Gαq‐protein‐coupled receptor OXTR, leading to PKC activation (Kimura et al. 1996; Arthur et al. 2007; Arrowsmith & Wray, 2014), we considered whether OXT inhibits SLO2.1 activity. We first tested this possibility in the Xenopus oocyte heterologous expression system. We expressed SLO2.1 alone or co‐expressed SLO2.1 and OXTR and then measured whole‐cell currents under voltage clamp before and after applying 200 nm OXT. Application of OXT substantially reduced SLO2.1 whole‐cell currents in oocytes expressing OXTR and SLO2.1 (Fig. 6 A) but not in oocytes expressing only SLO2.1 (Fig. 6 B). Additionally, we bypassed the Gαq‐protein‐coupled receptor signalling cascade by adding the PKC activator PMA and found that it reduced SLO2.1 currents in oocytes expressing only SLO2.1 (Fig. 6 B). Finally, we determined that OXT had no effect on currents in Xenopus oocytes co‐expressing SLO1 and OXTR (Fig. 6 C). We conclude that the effect of OXT on SLO2.1 was specific.

Figure 6. Oxytocin inhibits SLO2.1 but not SLO1.

Figure 6

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A, whole‐cell currents recorded from Xenopus oocytes co‐injected with human SLO2.1 and OXTR cRNAs. Currents are shown before (control) and after the addition of 200 nm OXT. The OXT‐sensitive current was obtained by subtracting traces (control − OXT). Showing the percentage of SLO2.1 current that is sensitive to OXT (78 ± 8.1%, n ± 3) or PMA (75 ± 15.4%, n ± 3) in oocytes expressing both SLO2.1 and OXTR. B, whole‐cell currents from oocytes injected with only SLO2.1cRNA, under control conditions and in the presence of 200 nm OXT or 1 μm PMA. The OXT‐ and PMA‐sensitive currents were obtained by subtracting traces. Showing the percentages of SLO2.1 currents that were sensitive to OXT (1 ± 14.5%, n ± 5) or PMA (72 ± 11.3%, n ± 5) in oocytes expressing only SLO2.1. C, whole‐cell currents from oocytes co‐injected with SLO1 and OXTR cRNAs. Currents were recorded under control conditions and in the presence of 400 nm OXT. The OXT‐sensitive current was obtained by subtracting traces. Showing the percentage of SLO1 current that was sensitive to OXT (1.8 ± 17.5%, n ± 8) and PMA (8.8 ± 32.7%, n ± 4). In all cases, whole‐cell currents were elicited with V h ± −80 mV and step pulses from −100 to +80 or +100 mV and all data were collected at +80 mV. *** P < 0.001. An unpaired t test was used to determine statistical significance.

To explore the effect of OXT on SLO2.1 in myometrial cells, we first measured single‐channel currents in the cell‐attached configuration in hMSMCs in the presence and absence of OXT. In the presence of TEA (to inhibit SLO1 channels), single channels were inhibited by 200 nm OXT (Fig. 7 A). We then undertook similar experiments in the whole‐cell configuration in hTERT‐HM cells (Fig. 7 B). These experiments showed that K+ channel activity was inhibited by treatment with both OXT and PMA. To confirm that OXT‐sensitive current was carried by SLO2.1 channels, we transfected hTERT‐HM cells with either scrambled siRNA or SLO2.1 siRNA. OXT reduced the K+ current in cells transfected with scrambled siRNA (Fig. 8 A), whereas it had almost no effect on K+ current in cells transfected with SLO2.1 siRNA (Fig. 8 B). Together, these experiments strongly support the hypothesis that OXT inhibits SLO2.1 in the tested cell types.

Figure 7. OXT inhibits TEA‐resistant K+ currents in hMSMCs and hTERT‐HM cells.

Figure 7

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A, representative single‐channel SLO2.1 recordings obtained from hMSMCs in the cell‐attached configuration (schematic on left) under control conditions and in the presence of 200 nm OXT in the external solution. Recordings were obtained at −60 mV. B, representative whole‐cell (schematic at left) recordings of the TEA‐resistant K+ currents in hTERT cells (5 mm TEA in extracellular solution) under control conditions, after the application of 200 nm OXT, wash, and after application of 1 μm PMA. The OXT‐ and PMA‐ sensitive currents were obtained by subtracting traces. Showing the percentage of currents sensitive to OXT (90 ± 5.2%, n ± 4) and PMA (87 ± 2.7%, n ± 3) at +80 mV. In all cases, whole‐cell currents were elicited at V h ± −70 mV and step pulses from −90 to +150 mV.

Figure 8. SLO2.1 channels conduct the OXT‐sensitive current in hTERT‐HM cells.

Figure 8

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Representative whole‐cell currents (schematic on left) from hTERT‐HM cells transfected with scrambled siRNA (A) or SLO2.1 siRNA (B). Currents were recorded under control conditions and after the addition of 200 nm OXT to the bath solution. The OXT‐sensitive current was obtained by subtracting traces. In both cases, whole‐cell currents were elicited with V h ± 0 mV and step pulses from −90 to +150 mV. C, showing the percentage of OXT‐sensitive currents at +80 mV in cells transfected with scrambled siRNA (28 ± 12.4%, n ± 4) and SLO2.1 siRNA (−11.5 ± 18.7%, n ± 5). ** P ± 0.009. An unpaired t test was used to determine statistical significance.

OXT signalling reduces SLO2.1‐dependent Ca2+ influx through voltage‐dependent Ca2+ channels

Previous work (Mironneau, 1976; Parkington et al. 1999; Arthur et al. 2007) has suggested that, in addition to triggering Ca2+ release from intracellular stores, OXT can trigger Ca2+ entry through VDCCs by causing membrane depolarization. Given that SLO2.1 activity hyperpolarizes the membrane and OXT inhibits SLO2.1, we considered whether OXT depolarized the membrane by inhibiting SLO2.1. To test this model, we used the Ca2+ indicator Fluo4‐AM to measure intracellular Ca2+ in hMSMCs. First, to confirm that membrane depolarization activates VDCCs to allow Ca2+ influx in these cells, we directly depolarized the cells with 80 mm KCl. This treatment caused an increase in intracellular Ca2+ (Fig. 9 Aa) that did not occur in the absence of external Ca2+ or in the presence of the VDCC blocker verapamil (Fig. 9 Ab and Ac). Next, we treated hMSMCs with PMA and observed an increase in intracellular Ca2+ (Fig. 9 Ba). This PKC‐induced increase in intracellular Ca2+ was not observed in the absence of external Ca2+, indicating that it was a result of Ca2+ entry (Fig. 9 Bb). In addition, 1 μm verapamil prevented an increase in intracellular Ca2+, indicating that Ca2+ influx was through VDCCs (Fig. 9 Ac). Finally, to confirm that the Ca2+ entry induced by PMA was the result of inhibition of SLO2.1 and did not occur via another pathway, we examined the effect of PMA in cells transfected with scrambled siRNA or siRNA directed against SLO2.1. It was found that 67% of non‐transfected (Fig. 9 Ba) and 58% of cells transfected with scrambled siRNA showed an increase in intracellular Ca2+ upon PMA treatment, whereas only 14% of cells transfected with SLO2.1 siRNA showed this Ca2+ increase (Fig. 9 Bd).

Figure 9. SLO2.1 inhibition with PMA induces an increase in intracellular Ca2+ in hMSMCs.

Figure 9

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A, representative fluorescence traces (Aa to Ac) and images (Ad) from 10 μm Fluo‐4 AM‐loaded hMSMCs in response to membrane depolarization with 80 mm external KCl under different conditions: (Aa) 2 mm external Ca2+ (control), (Ab) 0 mm external Ca2+ and (Ac) 1 μm verapamil. 5 μm Ca2+ ionophore ionomycin (+2 mm Ca2+ if necessary) was added at the end of each trace as a positive control. Percentage of cells with traces similar to the representative examples and number of cells recorded are indicated. All values were normalized to the fluorescence in 5 μm ionomycin and 2 mm extracellular Ca2+. B, representative fluorescence traces (Ba to Bd) and images (Be) from 10 μm Fluo‐4 AM‐loaded hMSMCs in response to 500 nm PMA under different conditions: (Ba) 2 mm Ca2+ external solution (control), (Bb) 0 mm external Ca2+, (Bc) 1 μm Ca2+ channel blocker verapamil and (Bd) cell treated with SLO2.1 siRNA. 5 μm Ca2+ ionophoreionomycin (+2 mm Ca2+ if necessary) was added at the end of each trace as a positive control. Percentage of cells with traces similar to the representative examples and number of cells recorded are indicated. All values were normalized to the fluorescence in 5 μm ionomycin and 2 mm extracellular Ca2+.

Discussion

Taken together, our experiments support the working model shown in Fig. 10, in which we propose that OXT could contribute to an increase in myometrial excitability, by depolarizing hMSMCs and increasing intracellular Ca2+, through inhibition of SLO2.1 channels. We propose that, throughout most of pregnancy, and in the presence of low levels of OXT, SLO2.1 conducts an outward K+ current that opposes the inward Na+ current through the NALCN present in these cell types (Reinl et al. 2015). Together, the balanced activity of these inward and outward leak currents maintains the myometrial resting potential at a more positive voltage than the negative equilibrium potential of K+. By counteracting the outward Na+ current, SLO2.1 current ensures that the membrane potential remains sufficiently hyperpolarized to prevent Ca2+ entry through VDCCs, thus helping to maintain myometrial quiescence. At the end of pregnancy, when OXT levels are elevated, OXT binds to the Gαq‐protein‐coupled receptor OXTR, activating PKC and thus inhibiting SLO2.1. As a result of the reduced outward K+ current, the membrane becomes depolarized; VDCCs become active, Ca2+ influx occurs, actin‐myosin cross‐bridging begins, and this mechanism could potentially contribute to myometrial cell contraction.

Figure 10. Proposed model.

Figure 10

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During the hMSMC resting state, the K+ leak current is dominant and maintains the cell sufficiently negative so the voltage‐dependent Ca2+ channels (VDCC) are closed and uterine contraction does not occur. Our data suggest that a Na+‐activated K+ current, carried by SLO2.1 channels, contributes a substantial percentage of the outward current at negative voltages and is therefore a major contributor to the K+ leak currents in MSMCs at rest. In the active state, OXT binds OXTR, activating phospholipase C beta (PLCβ), which in turn hydrolyzes phosphatidyl inositol 4,5‐bisphosphate (PIP2) to diacylglycerol (DAG) and inositol trisphosphate (IP3). DAG activates protein kinase C (PKC), which inhibits SLO2.1. Closing of SLO2.1 channels depolarizes the membrane, thus opening VDCCs, allowing Ca2+ influx to increase intracellular Ca2+ and activate myosin to cause muscle contraction.

To our knowledge, this is the first observation and characterization of a Na+‐activated K+ channel in hMSMCs. High‐conductance K+ channels activated by Na+ (KNa channels) were originally observed in inside‐out membrane patches pulled from guinea pig cardiomyocytes (Kameyama et al. 1984). KNa channels may have been previously overlooked in myometrium because of their resemblance to high conductance SLO1 (BK) channels, which have been well characterized in MSMCs but are not considered to contribute to the cell resting membrane potential, becoming active only after an intracellular rise in Ca2+. KNa channels are also high conductance K+ channels but are insensitive to Ca2+ and, instead, appeared to require high concentrations of intracellular Na+ to be active (Kameyama et al. 1984; Bhattacharjee et al. 2003; Yuan et al. 2003; Kaczmarek, 2013; Thomson et al. 2015; Hite & MacKinnon, 2017).

Because of the high intracellular Na+ needed to activate KNa channels in excised patches, it was originally assumed that KNa channels were principally active under ischaemic and other low oxygen conditions. Under such conditions, KNa channels could counter toxicity in cardiomyocytes when an increase in intracellular Na+ concentration resulted from inhibition of the Na+/K+‐ATPase (Luk & Carmeliet, 1990). However, Budelli et al. (2009) found that KNa channels are commonly active and prominent under normal physiological conditions. Because of their low voltage sensitivity, KNa channels are active over a wide voltage range. Indeed, these channels are active even at very low intracellular Na+ concentrations, and Na+ influx can activate current through these channels in the absence of intracellular Na+. In some instances, KNa channels may become transiently active in neurons in response to an intracellular build‐up of Na+ during periods of intense electrical activity. In neurons (Budelli et al. 2009; Hage & Salkoff, 2012) (and, as reported in the present study, in myometrial cells), as long as there is a small but steady Na+ leak current into cells, the KNa current is active to some extent. Hence, in both neurons and myometrial cells, KNa current can be observed as a non‐inactivating delayed outward current similar to the Hodgkin‐Huxley ‘delayed rectifier’ current (Yuan et al. 2000; Yuan et al. 2003; Santi et al. 2006), although with lower voltage sensitivity.

KNa currents are carried by two members of the SLO K+ channel family: SLO2.1 and SLO2.2 (also known a Slick and Slack, respectively) (Yuan et al. 2003; Kaczmarek, 2013). We found that SLO2.1 but not SLO2.2 is present in both hTERT‐HM cells and hMSMCs. Because SLO2.1 has low voltage dependence and can be activated by Na+ leak currents, and thus can be significantly active at physiological intracellular Na+ concentration (Budelli et al. 2009; Hage & Salkoff, 2012), this channel probably contributes to setting the hMSMC resting membrane potential. As a result of their high conductance, SLO2.1 channels could be pivotal even at a relatively low open channel probability.

The inter‐relatedness of the Na+ leak and KNa current suggests that the control of cell membrane resting potential and cell excitability can be regulated by modulating either the inward Na+ or the counterbalancing outward K+ current. Although NALCN is a good candidate to carry the Na+ leak current that activates SLO2.1, other sources of Na+ entry into MSMCs cannot be ruled out.

Lastly, our results shed light on conflicting data regarding the contribution of VDCC activity to oxytocin stimulation in myometrial cells (Inoue et al. 1992; Arnaudeau et al. 1994). We show here that PKC causes calcium entry by possibly activating VDCC indirectly through inhibition of SLO2.1 channels and membrane depolarization (Fig. 10).

In summary, we demonstrate the presence of a KNa current in hMSMCs and provide data suggesting that this current is carried by SLO2.1 channels. Because of their biophysical properties, SLO2.1 channels appear to be important determinants of hMSMC resting membrane potential. In addition, SLO2.1 inhibition by PKC activation caused membrane depolarization, activation of VDCC and an increase in intracellular Ca2+. We propose that this could be a mechanism for oxytocin to trigger membrane depolarization and contribute to contraction at the end of pregnancy when the OXTR expression increases in the MSMC. The finding that SLO2.1 channel activity is regulated by OXT maybe an important step towards understanding the molecular basis of uterine excitability and could lead to the development of effective approaches for enhancing or inhibiting uterine quiescence to regulate timing of parturition. This mechanism of action could be relevant in other cells that express SLO2.1 and OXTR, such as neurons.

Additional information

Competing interests

The authors declare that they have no competing interests.

Author contributions

CMS, SKE and LS conceived, planned, supervised the experiments and wrote the manuscript. JF performed experiments, analysed the data, prepared the final figures and contributed to the writing. AB, RS, AGC, PL, CA, ELR and MW performed experiments and reviewed the manuscript. All authors approved the final version of the manuscript submitted for publication.

Funding

This work was supported by Eunice Kennedy Shriver National Institute of Child Health and Human Development (NICHD) R01 HD088097 to CMS and SKE as well as (NIGMS and NIMH) R01 GM114694 and R21MH107955 to LS, respectively. A CONACyT Postdoctoral Fellowship EPE‐2016291121 and EPE‐2017 291231 to AG also funded this work.

Acknowledgments

We thank Dr Deborah Frank for critical review of the manuscript and the Clinical Research Nurses in the Department of Obstetrics and Gynecology for gaining the consent of patients and acquiring myometrial biopsies.

Biography

Juan J. Ferreira is currently a PhD student at Washington University in St Louis. He obtained his Bachelor degree in Human Biology in 2016 at the Universidad de la República, Uruguay. From 2016 to July 2018, he was working as a Research technician in Dr C. M. Santi's laboratory, where he studied the role of SLO2 channels in myometrial smooth muscle excitability. In 2018, he was admitted as a PhD student in the Neuroscience Program at Washington University in St Louis. He has experience in patch clamp recordings and calcium imaging techniques.

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Edited by: Laura Bennet & Suzanne Miller

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