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. 2026 Mar 10;241(3):e70159. doi: 10.1002/jcp.70159

TMEM16A Contributes to Calcium‐Activated Chloride Currents and Membrane Potential Changes in the Mouse Oocyte

Sarah Dalati 1, Vanessa J Jones 1, Margot L Day 1,✉
PMCID: PMC12974648  PMID: 41806302

ABSTRACT

Oscillations in intracellular Ca2+ [Ca2+]i are essential for mouse oocyte activation following fertilization. These [Ca2+]i oscillations also induce repetitive hyperpolarizations in the membrane potential (Em). The present study aimed to identify the channels underlying the Em hyperpolarizations. Sulfhydryl reagents such as thimerosal, that oxidize the IP3‐R channel, mimic the physiological changes at fertilization by eliciting simultaneous Em changes and [Ca2+]i oscillations. Thimerosal‐induced Em and [Ca2+]i changes were prevented by the non‐specific Ca2+‐activated Cl– channel (CaCC) inhibitors DIDS and NFA, as well as the TMEM16A/Anoctamin 1 CaCC specific inhibitor, T16Ainh‐01. The K+ channel blocker TEA, and voltage‐gated Cl– channel blocker 9AC failed to inhibit the Em or [Ca2+]i changes. TMEM16A protein was expressed in all stages of mouse preimplantation development, being localized at the plasma membrane in oocytes. Culture of zygotes in the TMEM16A inhibitor prevented development to the blastocyst stage. In summary, we present the first evidence for CaCC channels, namely TMEM16A, being critical for the initiation of Em hyperpolarisations in mouse oocytes.

Keywords: hyperpolarizations, intracellular calcium oscillations, ion channels, membrane potential, oocyte, TMEM16A

1. Introduction

In mammals, mature oocytes are arrested in metaphase of the second meiotic division (MII) at the time of ovulation. Fertilization causes the oocyte to exit from this MII arrest, complete meiosis and progress into interphase (Schultz and Kopf 1995). Fertilization is characterized by a series of transient increases in intracellular Ca2+ ([Ca2+]i) (Cuthbertson et al. 1981; Dumollard et al. 2002; Igusa and Miyazaki 1986), which are essential for completion of the events of oocyte activation; including exit from MII arrest, degradation of cyclin B (Nixon et al. 2002), cortical granule exocytosis (Ducibella et al. 1993) and initiation of mitotic cell divisions (Ducibella et al. 2002). Inhibition of the [Ca2+]i transients in the oocyte after sperm binding prevents all of these events (Jones 2007).

The fertilizing sperm initiates [Ca2+]i changes by delivering phospholipase C (PLC) ζ into the oocyte (Saunders et al. 2007). PLCζ hydrolyses phosphatidylinositol 4,5‐bisphosphate (PIP2) to produce the two signaling molecules inositol 1,4,5‐trisphosphate (IP3) and 1,2‐diacylglycerol (DAG) (Parrington et al. 1998; Rice et al. 2000; Rongish et al. 1999). IP3 causes Ca2+ release from the endoplasmic reticulum (ER) by binding and gating its receptor, the IP3 receptor type 1 (IP3R‐1) (Fissore et al. 1999). The release of Ca2+ into the cytoplasm results in an initial [Ca2+]i spike after sperm binding (Swann and Whitaker 1986). Sequential re‐uptake of Ca2+ into the ER by the sarcoplasmic/endoplasmic Ca2+ ATPase (SERCA; (Wakai et al. 2013)) followed by Ca2+ release results in repetitive [Ca2+]i oscillations that continue until the time of pronuclei formation (Day et al. 2000; Jones et al. 1995).

Accompanying the increases in [Ca2+]i following fertilization are changes in the electrical properties of the oocyte plasma membrane, known as membrane potential (Em) hyperpolarizations. These Em changes were first observed in hamster oocytes, where the hyperpolarizations are caused by Ca2+‐dependent K+ channel activation (Igusa and Miyazaki 1983). In human oocytes, increases in [Ca2+]i, induced by sperm factor and by thimerosal, are also accompanied by Em hyperpolarizations (Homa and Swann 1994). Thimerosal oxidizes the sulfhydryl (SH) groups on the IP3R‐1, resulting in the changes in [Ca2+]i that mimic those generated by fusion of sperm to mammalian oocytes (Cheek et al. 1993). Although hyperpolarizations in Em have been observed in oocytes of a number of species, including humans (Homa and Swann 1994), hamster (Miyazaki and Igusa 1981), and mice (Igusa et al. 1983), the mechanism responsible for initiating these Em changes and their role in fertilization remains unclear.

Ca2+ activated Cl– channels (CaCCs) are expressed in many cell types where they have several physiological functions. CaCCs were first identified in the Xenopus oocyte (Barish 1983; Miledi 1982), where the fertilization‐induced increase in [Ca2+]i activates CaCCs, resulting in Em depolarization and a fast block to polyspermy. Three protein families were originally identified as potential CaCCs: CLCA, which has since been shown to be a chloride channel accessory protein rather than a channel itself (Huang et al. 2012), bestrophins and Anoctamins/TMEM16 (reviewed in (Huang et al. 2012)). The TMEM16 family has 10 members but only TMEM16A/Ano‐1 and TMEM16B/Ano‐2 have been confirmed as CaCCs (Caputo et al. 2008; Schroeder et al. 2008; Yang et al. 2008), while other TMEM16 proteins function as lipid scramblases and, in some cases, both ion channels and scramblases (Kalienkova et al. 2021; Kostritskii and Machtens 2021). Functional characterization of CaCCs has been hampered by the lack of specific inhibitors of these channels, with commonly used compounds such as niflumic acid (NFA), 4,4′‐diisothiocyanatostilbene‐2,2′‐disulfonic acid (DIDS) and 5‐nitro‐2‐(3‐phenylpropylamino)benzoic acid (NPPB) also acting on other classes of Cl– channels and transporters (Verkman and Galietta 2021). Several TMEM16A small‐molecule inhibitors, with nanomolar potency, however, have been identified, including CaCCinh‐A01 (De La Fuente et al. 2008) and T16Ainh‐A01 (Namkung et al. 2011).

In this study, thimerosal was used to activate [Ca2+]i oscillations in mouse oocytes, to mimic those induced by fertilization. The effect of CaCC inhibitors on the [Ca2+]i oscillations and Ca2+‐induced Em hyperpolarisations was analysed by fluorescent imaging and whole‐cell patch‐clamping, respectively. TMEM16A expression in oocytes and early embryos was also examined.

2. Materials and Methods

2.1. Chemicals

All chemicals were obtained from Sigma‐Aldrich except when mentioned. Thimerosal was made as a 200 mM stock solution on each day and stored at room temperature. Stock solutions of 10 M tetraethylammonium (TEA), 100 mM niflumic acid, 100 mM 4,4′‐diisothiocyanatostilbene‐2,2′‐disulfonate (DIDS), 100 mM 4,4′‐Diisothiocyanatostilbene‐2,2′‐disulfonic acid disodium salt hydrate (H2DIDS), 1 M anthracene‐9‐carboxylic acid (9AC) and 10 mM BAPTA‐AM were prepared in DMSO and stored on ice for 1 day's use only. T16Ainh‐A01 (Tocris Bioscience) was made as a 10 or 100 mM stock prepared in DMSO and stored in the fridge for 1 month's use only.

2.2. Oocyte and Embryo Collection and Culture

The use of animals was approved by the University of Sydney Animal Care and Ethics Committee (approval numbers 2008/4838, 2011/5583, 2015/824), and experiments were conducted in accordance with the Australian Code of Practicme for Use of Animals in Research. All experiments were conducted using female Quackenbush Swiss (QS) mice (Animal Resource Centre, Perth and Laboratory Animal Services, University of Sydney) between 3 and 6 weeks old housed in a constant 12 h light/dark cycle environment. Female mice were superovulated by intraperitoneal injection of 10 IU pregnant mares' serum gonadotrophin (PMSG; Intervet, Sydney Australia) and 10 IU human chorionic gonadotrophin (hCG; Intervet, Sydney Australia) 46–50 h later. Mice were euthanized by cervical dislocation 13 h post hCG injection to retrieve unfertilized oocytes. The surrounding cumulus cells and zona pellucida were removed with 0.001% chymotrypsin in Hepes‐modHTF containing 3 mg/mL BSA adjusted to pH 7.4 and 270–280 mOsm/L (Morris et al. 2020). Oocytes were washed and transferred to fresh drops of Hepes mod‐HTF overlaid with mineral oil (Sigma‐Aldrich) and stored at 37°C until needed. For experiments requiring embryos, female mice were mated with a stud male QS mouse (8–24 weeks old) overnight, after hCG injection. Embryos at the zygote, 2‐cell, 4‐cell, 8‐cell, morula and blastocyst stages were isolated at 24, 48, 58, 66, 75 and 96 h after hCG injection, respectively, by flushing the oviduct or uterus as appropriate.

Medium used for embryo culture experiments was potassium simplex optimized medium (KSOM), composed of (mM): NaCl (95), KCl (2.5), KH2PO4 (0.35), MgSO4 (0.2), NaHCO3 (25), Na‐lactate (10), Na‐pyruvate (0.2), glucose (0.2), CaCl2 (1.71), BSA (3 mg/mL) at pH 7.4 and with and osmolality of 260 mOsm/L (Green and Day 2013). Freshly isolated zygotes, 2‐cell, 4‐cell or morula stage embryos were cultured in groups of 10 per 100 µl KSOM in the absence or presence of 10 µM T16inh‐A01 in round bottom 96 well plates (Corning, NY, USA) at 37°C and 5% CO2. Development was scored every 24 h for 5 days.

2.3. Intracellular Calcium Imaging

Changes in intracellular calcium concentrations [Ca2+]i, were examined in zona‐free oocytes which were loaded with the Ca2+‐sensitive ratiometric dye Fura‐2AM (Invitrogen) at 1 µM or the Ca2+ indicator Fluo‐3AM (Invitrogen) at 1 µM in BSA‐free Hepes mod‐HTF perfusion solution for 30 min. Following loading, oocytes were transferred onto a glass coverslip pre‐coated with 0.02% concanavalin‐A, to prevent loss/movement of oocytes during perfusion. The coverslip was attached to the base of a perspex perfusion chamber, which was then filled with BSA‐free Hepes mod‐HTF. Experiments were performed at 37°C by constant perfusion of BSA‐free Hepes mod‐HTF at a rate of 1 ml/minute. Fura‐2AM was excited at 340/380 nm and detected using a 510 nm filter. Fluo‐3 was excited at 480/30 nm and detected using a 535/540 nm filter. Fluorescence intensities were captured by a CCD camera (Pulnix, Sunnyvale, CA), at 5 s intervals using Simple PCI software (Hamamatsu Photonics) and an Olympus IX70 microscope.

2.4. Electrophysiological Recordings

Perforated whole‐cell patch clamping of single oocytes was used to study Em changes induced by thimerosal (200 µM), as described previously (Arnaiz et al. 2013; Li et al. 2009). Pipettes were pulled from borosilicate haematocrit glass tubes (Modulohom, Herlev, Denmark) and had a resistance of 5–10 MΩ. Nystatin (40 µg/ml) was used to perforate the membrane and prepared as a 40 mg/ml stock stolution in DMSO. A K‐glutamate pipette solution consisting of (mM): K‐glutamate (132), KCl (8), MgCl2.6H2O (1), H‐Hepes (10), glucose (10), ethyleneglycol‐tetraacetic acid (EGTA) (0.5) and ethylenediamine‐tetraacetic acid (EDTA) (0.01) adjusted to pH 7.2 with glutamic acid. This enabled influx of Cl to be measured when the Cl–‐rich (HEPES‐modHTF) bath solution was present. A List EPC‐7 amplifier (Darmstadt, Germany) and MacLab/4 data acquisition software (AD Instruments, Sydney, Australia) were used to measure changes in Em in current clamp mode. Patch‐clamping was performed at 37°C by constant perfusion of Hepes mod‐HTF. A stable baseline Em was recorded for approximately 2 min before the addition of desired reagents. Once the reagent was added, the perfusion chamber was continuously perfused to observe membrane potential changes. [Ca2+]i imaging and patch clamp experiments were carried out independent of one another.

2.5. Western Blotting

Samples for SDS‐PAGE were collected by pooling together 100–150 oocytes or embryos and placed in a microfuge tube containing 2 µl Laemmli buffer containing 0.1% (w/v) dithiothreitol and stored at −80°C until use. Samples were heated at 95°C for 5 min before running on a 12% SDS‐polyacrylamide gel. After separation, proteins were transferred onto 0.2 µm nitrocellulose membrane (Hybond, GE healthcare) for 90 min. The membrane was incubated at 4°C in Odyssey blocking buffer (LI‐COR Biosciences) overnight, followed by incubation at 4°C overnight with primary antibodies: rabbit anti‐TMEM16A (Abcam, No. ab53212) and mouse anti‐β actin (Sigma‐Aldrich, No A5316) diluted 1:1000 in blocking buffer + 0.1% Tween‐20. The membrane was then washed in Tris‐buffered saline + 0.05% Tween 20 (TBST) and incubated for 2 h at room temperature with 1:10000 donkey anti‐rabbit IRDye 800CW (Odyssey) and Donkey anti‐mouse IRDye 680LT (Odyssey) secondary antibodies (Green et al. 2015; Green and Day 2013). The membrane was then washed with TBST and imaged on the Odyssey infrared imager.

2.6. Immunofluorescent Staining of TMEM16A in Oocytes

Immunofluorescent staining was performed as described previously (Green et al. 2015; Green and Day 2013). Oocytes were fixed with 4% paraformaldehyde in phosphate buffered saline + 1 mg/ml poly‐vinyl alcohol (PBS + PVA) for 30 min at room temperature and then washed in PBS + PVA and permeabilized with PBS + PVA + 0.3% Triton‐X‐100 for 15 min. Oocytes were then transferred to the blocking solution, PBS + PVA + 0.1% Tween‐20 + 0.7% BSA (PPTB), for 30 min and then incubated in rabbit anti‐TMEM16A (1 in 100 in PPTB; Novus Biologicals, No. NBP1‐19037) overnight at 4°C. Oocytes were then washed in PPTB five times and incubated for 1 h in Alexa Fluor 488‐labeled goat anti‐rabbit IgG (1 in 200 in PPTB) (Invitrogen) at room temperature in the dark. Oocytes were then washed five times in PPTB, left in the final wash for 10 min and mounted on slides in in 5 μL vectashield containing 1.5 µg/ml DAPI (Vector Laboratories). Fluorescence was visualized using a LSM510 Meta confocal microscope (Carl Zeiss) using a 405 nm and Argon 488 nm laser lines at 40 x objective.

2.7. Statistical Analysis

Significance was determined using an unpaired t‐test comparing treated oocytes to control oocytes. p < 0.05 was considered statistically significant. Data are expressed as the mean ± SEM and the number of oocytes from which recordings were made is given in parentheses. All experiments were repeated on at least three separate days. For T16inh‐A01 embryo culture experiments differences were determined using a one‐way ANOVA and Dunnett's post‐hoc test (GraphPad Prism, GraphPad Software, USA).

3. Results

3.1. Thimerosal Induces Em Hyperpolarizations Due to Increases in [Ca2+]i

Chemical stimuli can be used to mimic the sperm induced [Ca2+]i oscillations and are therefore useful tools for investigation of mechanisms underlying the [Ca2+]i oscillations. Thimerosal is a reagent that oxidises sulfhydryl groups on the IP3 receptor, thereby sensitizing the receptor to Ca2+ (Mehlmann and Kline 1994). Thimerosal was used in the present study because it triggers [Ca2+]i oscillations, rather than the single Ca2+ peak triggered by other chemicals such as Ca2+ ionophores (Cheek et al. 1993). Fertilisation with sperm could not be used in this study since movement of the oocyte due to sperm binding interfered with patch‐clamp recording.

Initial experiments were performed to confirm the effect of thimerosal on [Ca2+]i and Em in mouse oocytes (Cheek et al. 1993; Herbert et al. 1995; Homa and Swann 1994). Thimerosal (200 μM) induced repetitive [Ca2+]i oscillations that lasted for the duration of the thimerosal treatment (Figure 1A). Perforated patch‐clamp recordings from oocytes showed that thimerosal induced repetitive hyperpolarizations (HRs) in Em, that were overlayed on an overall depolarization in the resting Em (Figure 1A). The average resting Em at the start of the recording was −27 ± 2.9 mV (n = 15) and the average size of the first hyperpolarization was 10 ± 0.8 mV (n = 15). A slow depolarization of the resting Em was observed before the first hyperpolarization in 8 out of 15 recordings (Figure 1A). The Ca2+ chelator BAPTA‐AM was used to confirm the role of the [Ca2+]i oscillations in activation of the HRs by thimerosal. None of the oocytes that were pre‐loaded with 10 µM BAPTA‐AM showed [Ca2+]i changes (Figure 1B,D; p < 0.001) or Em hyperpolarizations (Figure 1B,E; p < 0.001) in the presence of thimerosal.

Figure 1.

Figure 1

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Thimerosal activates oscillations in [Ca2+]i and Em hyperpolarisations in mouse oocytes that are prevented by BAPTA‐AM but not TEA. Representative tracings showing changes in [Ca2+]i and Em in single oocytes in response to (A) 200 µM thimerosal alone, and thimerosal in the presence of (B) 10 µM BAPTA‐AM or (C) 10 mM TEA. [Ca2+]i is presented as the ratio of fluorescence at 340/380. Arrows indicate addition of reagents to the perfusion solution. Number of [Ca2+]i oscillations in 20 min (D, F) and Em hyperpolarizations (HRs) in 10 min (E, G) in response to thimerosal alone or in the presence of BAPTA‐AM (D, sE) or TEA (F, G). All recordings of [Ca2+]i and Em were obtained from different oocytes. Error bars represent S.E.M and the number of oocytes from which recordings were made is given in parentheses and were obtained from at least 3 separate experiments. ***p < 0.001, NS not significantly different.

3.2. K+ Channel Inhibition Does Not Prevent Thimerosal‐Induced Em and [Ca2+]I Changes

Hyperpolarization of the Em could either be due to K+ efflux or Cl– influx across the oolemma. Since fertilization‐induced HRs in hamster oocytes are caused by activation of a Ca2+‐dependent K+ conductance (Miyazaki and Igusa 1982) the effect of the broad‐spectrum K+ channel blocker TEA (10 mM) on the thimerosal‐induced Ca2+ and Em oscillations in mouse oocytes was investigated. Addition of TEA alone had no effect on resting [Ca2+]i or Em (not shown). Addition of thimerosal in the presence of TEA induced [Ca2+]i and HRs that were similar in appearance and number to those observed in absence of TEA (Figure 1C,F,G). The average size of the first Em hyperpolarization, however, was significantly reduced by the presence of TEA (6 ± 0.5 mV (n = 11) verses 10 ± 0.8 mV (n = 15), p < 0.001).

3.3. DIDS Reduces Thimerosal‐Induced Em and [Ca2+]i Changes

Em changes in Xenopus oocytes and mouse embryos are known to be caused by Ca2+‐activated Cl– channel (CaCC) activity and are prevented by broad specificity Cl– channel blockers, such as DIDS (Barish 1983; Li et al. 2007, 2009). The addition of 500 μM DIDS (or H2DIDS for Ca2+ imaging because of its lower fluorescence) alone had no effect on resting [Ca2+]i or Em in mouse oocytes (not shown). Subsequent addition of thimerosal in the presence of 500 µM DIDS caused [Ca2+]i oscillations and HRs that were significantly reduced in number compared with those induced by thimerosal alone (Figure 2A,C,D; p < 0.001). The size of the first Em HR in the presence of DIDS was 10 ± 4 mV (n = 3), which was not different to the size seen with thimerosal alone (p = 0.518).

Figure 2.

Figure 2

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Effect of the Cl– channel inhibitors, DIDS and 9‐AC, on thimerosal‐induced changes in [Ca2+]i and Em in mouse oocytes. Representative tracings of [Ca2+]i oscillations and Em hyperpolarizations induced by 200 μM thimerosal in the presence of (A) 500 µM DIDS or (B) 1 mM 9‐AC. Recordings of [Ca2+]i and Em were made on separate oocytes. H2DIDS was used for [Ca2+]i experiments because of its lack of autofluorescence. Number of [Ca2+]i oscillations in 20 min (C, E) and Em hyperpolarizations (HRs) in 10 min (D, F). Error bars represent S.E.M and the total number of recordings from at least 3 separate experiments is given in parentheses. ***p < 0.001, *p < 0.05.

3.4. Inhibition of Voltage‐Gated Cl– Channels Has no Effect on Thimerosal Induced [Ca2+]i and Em Changes

The effect of 9AC on Em and [Ca2+]i changes was assessed to eliminate the role of a voltage gated chloride channels (VGCC), which are also present in the mouse oocyte (Arnaiz et al. 2013; Sonoda et al. 2003). Due to the fluorescence of 9AC at 340/380 nm, fluo‐3 was used instead of fura‐2 to monitor changes in [Ca2+]i. 9AC (1 mM) had no effect on resting [Ca2+]i or Em (not shown), or on the number of [Ca2+]i oscillations (Figure 2B,E) or HRs (Figure 2B,F) induced by thimerosal. The average size of the first Em HR in the presence of 9‐AC was 12 ± 3 mV (n = 6), which was not different to the size seen with thimerosal alone (p = 0.373).

3.5. NFA Inhibits Thimerosal Induced [Ca2+]i and Em Changes

Niflumic acid (NFA) is a more selective CaCC blocker (Jentsch et al. 2002) and inhibits CaCCs in a variety of cell types, including Xenopus oocytes (White and Aylwin 1990) and mouse 2‐cell embryos (Li et al. 2009). Addition of 100 µM NFA alone had no effect on resting [Ca2+]i or Em (not shown) but significantly reduced the number of thimerosal‐induced [Ca2+]i oscillations (Figure 3A,B) and HRs (Figure 3C,D) and caused sustained elevation of basal Ca2+ and depolarization of the Em (Figure 3A,C). The size of the first Em HR in the presence of NFA was also significantly reduced (3 ± 1 mV, n = 3 versus 10 ± 0.8 mV, n = 15; p < 0.001).

Figure 3.

Figure 3

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Effect of CaCC inhibitors on thimerosal‐induced changes in [Ca2+]i and Em in mouse oocytes. Representative tracings of [Ca2+]i oscillations and Em hyperpolarizations in response to 200 μM thimerosal in the presence of (A, C) 100 µM NFA or (E, G) 10 µM T16Ainh‐A01. Number of (B, F) [Ca2+]i oscillations in 20 min and (D, H) Em hyperpolarizations (HRs) in 10 min from start of response. Error bars represent S.E.M. and the total number of recordings from at least 3 separate experiments is given in parentheses. ***p < 0.001, *p < 0.05.

3.6. Role of TMEM16A in Thimerosal‐Induced Em Changes

The CaCC Anoctamin‐1 (ANO1) or TMEM16A has been shown to be responsible for the Ca2+‐induced HRs in Xenopus oocytes (Schroeder et al. 2008; Wozniak et al. 2018) and this channel can be blocked by small molecule inhibitors, such as T16Ainh‐A01 (Namkung et al. 2011). The role of TMEM16A in the response of mouse oocytes to thimerosal was therefore investigated. The addition of 10 µM T16Ainh‐A01 alone had no effect on resting [Ca2+]i or Em (not shown). Addition of thimerosal in the presence of 10 µM T16Ainh‐A01 had no effect on the number of [Ca2+]i oscillations induced (Figure 3E,F), while the number of Em HRs was significantly reduced (p < 0.001; Figure 3G,H), as was the size of the first HR (reduced to 4.3 ± 1.1 mV; n = 5; p < 0.05; Figure 3G).

3.7. TMEM16A Is Expressed in the Oocyte and All Stages of the Pre‐Implantation Embryo

Since T16Ainh‐A01 decreased Em HRs, the expression of TMEM16A in mouse oocytes and embryos was examined by western blotting and immunofluorescence. Human prostate cancer cells (PC‐3), which express TMEM16A (Liu et al. 2012), were used as a positive control in western blotting. A band at ~140 kDa was detected in the oocyte and all stages of pre‐implantation development (Figure 4A). Immunofluorescent staining also showed that TMEM16A was expressed in all stages, with the staining in oocytes being concentrated in the cortical region under the oolemma (Figure 4B). In the zygote, 2‐, 4‐ and 8‐cell stages, TMEM16A staining was evenly distributed throughout the cytoplasm (Figure 4B). In the morula and blastocyst, TMEM16A staining was much weaker compared to the other stages (Figure 4B) which corresponded with the decreased band intensity for these stages in the western blot (Figure 4A).

Figure 4.

Figure 4

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Expression of the TMEM16A protein in mouse oocytes and pre‐implantation embryos. (A) Western blot showing expression of TMEM16A (~140 kDa) in oocytes (O) and embryos (1 C: 1‐cell zygote, 2 C: 2‐cell, 4 C: 4‐cell, 8 C: 8‐cell, M Morula, BL blastocyst). The prostate cancer cell line PC3 was used as a positive control. Lower blot indicates β‐actin expression as a loading control. (B) Immunofluorescent staining of TMEM16A in the mouse (i) oocyte, (ii) zygote, (iii) 2‐cell, (iv) 4‐cell, (v) morula and (vi) blastocyst stages. TMEM16A staining is shown in green and DNA in blue (DAPI). Images are representative of 12–30 embryos per developmental stage collected on 3 separate occasions. (C) Dose dependent effect of T16Ainh‐A01 on the development of zygotes to the 2‐cell, 4‐cell, 8‐cell and blastocyst stages. Values are means ± S.E.M from 3 separate experiments with 36–38 oocytes per treatment group. Bars with the same letter in each treatment group are significantly different (p < 0.05) determined by ANOVA.

3.8. Inhibition of TMEM16A Prevents Development of Zygotes to the Blastocyst Stage but Has no Effect on Development When Cultured From the Morula Stage

The effect of culturing zygotes in T16inh‐A01 was investigated to further determine the role of the CaCC in development. Culture in the presence of 2 µM T16Ainh‐A01 reduced development to the morula and blastocyst stages, while 3 µM also reduced development to the 4‐cell stage (p < 0.05; Figure 4C). Culture of zygotes in 5 µM T16Ainh‐A01 reduced division to the 2‐cell stage compared to the control (no drug; p < 0.05) and prevented development to later stages. No embryo development was observed in the presence of 10 µM T16Ainh‐A01 (Figure 4C). When freshly isolated 2‐ and 4‐cell stage embryos were cultured in the presence of 10 µM T16Ainh‐A01 0% of embryos developed to blastocysts (n = 19 and 10, respectively) compared to 90% of controls (n = 20 and 10, respectively). In contrast, 10 µMT16Ainh‐A01 had no effect on development of morula to the blastocyst stage (92% (n = 27) in T16Ainh‐A01 vs 90% development (n = 30) in control medium).

4. Discussion

This study shows that the sulphydryl reagent thimerosal induces Em hyperpolarizations in parallel with [Ca2+]i oscillations in mouse oocytes that mimic those induced by sperm at fertilization (Cheek et al. 1993; Homa and Swann 1994; Igusa et al. 1983). Simultaneous measurements of Em and [Ca2+]i in human oocytes have shown that the hyperpolarizations directly correspond to Ca2+ transients (Homa and Swann 1994) and are due to a Ca2+‐activated outward current, which can be prevented by chelation of intracellular Ca2+. Ca2+‐activated K+ currents are responsible for large, repetitive Em hyperpolarizations in hamster oocytes (Miyazaki and Igusa 1982) and smaller, single hyperpolarizations in human (Dale et al. 1996; Homa and Swann 1994) and bovine (Tosti et al. 2002) oocytes. However, in the present study on mouse oocytes the Ca2+‐activated K+ channel inhibitor TEA did not prevent the repetitive hyperpolarizations induced by thimerosal. Only the size of the first hyperpolarization was reduced by TEA, suggesting that Ca2+‐activated K+ channels may contribute to initiation of the Em hyperpolarizations. There is evidence for a lower density of Ca2+‐activated K+ channels in mouse compared with hamster oocytes (Igusa et al. 1983) suggesting species differences in channel expression in oocytes. Ca2+‐activated K+ channels are expressed at later stages in mouse pre‐implantation development (Lu et al. 2012) and TEA partially inhibits Ca2+‐activated currents in the mouse 2‐cell embryo (Li et al. 2007). Resting membrane potential of mouse oocytes is also dependent on the activity of a large‐conductance K+ channel, but this channel is not Ca2+ sensitive (Day et al. 1993, 1998).

Cl– conductance also plays a major role in regulation of resting membrane potential in mammalian oocytes and embryos (Sonoda et al. 2003). From measurements of intracellular (~9 mM (Baltz et al. 1997)) and extracellular (oviductal; ~102–132 mM (Borland et al. 1980)) Cl– concentrations, the equilibrium potential for Cl– in oocytes is expected to be approximately –70 mV. Thus, the outward current responsible for the thimerosal‐induced hyperpolarisations could be caused by movement of Cl– into the oocyte. Swelling‐activated (Kolajova et al. 2001) and voltage‐gated (Arnaiz et al. 2013; Sonoda et al. 2003) Cl– channels are expressed in pre‐implantation mouse embryos, however the voltage‐gated Cl– channel inhibitor 9AC (Jentsch et al. 2002) had no effect on the thimerosal‐induced Em hyperpolarizations. Instead, the Em hyperpolarizations were prevented by the CaCC blockers DIDS and NFA.

There is limited evidence for CaCC activity in pre‐implantation mammalian embryos. In the mouse 2‐cell embryo Em hyperpolarizations induced by the embryotrophic factor platelet‐activating factor (PAF) are caused by CaCC activation (Li et al. 2007, 2009), although the channel protein responsible has not been determined. The present study shows that TMEM16A CaCCs may be responsible for the Em hyperpolarisations in mouse oocytes during fertilization, which resembles the situation in Xenopus oocytes in which TMEM16A is responsible for Em regulation after fertilization and the fast block to polyspermy (Schroeder et al. 2008). However, the Em hyperpolarizations in the present study were not completely inhibited by T16Ainh‐A01, suggesting that an additional CaCC, such as TMEM16B, which is not blocked by T16Ainh‐A01 and only partially blocked by NFA (Hernandez et al. 2021; Tian et al. 2012), may also be present. The potency of TMEM16A inhibitors, such as T16Ainh‐A01, also appears to be cell type specific, possibly due to expression of different splice variants (O'Driscoll et al. 2011), phosphorylation of the channel and interaction with other cellular proteins (Arreola et al. 2022) and may mean that a higher concentration of the inhibitor is required to completely block the channel in mouse oocytes. The effect of other small molecule inhibitors of TMEM16A (e.g., MONNA and Ani9) could be investigated, although they have also been shown to have off target effects. For example, Ani9 also blocks TMEM16F and changes in intracellular Ca2+ (Centeio et al. 2020), while MONNA is no more potent than T16Ainh‐A01 at blocking TMEM16A in other cells (Boedtkjer et al. 2015) and like T16Ainh‐A01, its potency is also species and splice variant‐dependent (Oh et al. 2013). Alternately, the initiation of the hyperpolarisations may be dependent on Ca2+‐activated K+ channels, since TEA reduced the size of the first hyperpolarisation, while the subsequent repetitive hyperpolarisations are due to TMEM16A.

Western blotting confirmed that TMEM16A protein is expressed in all stages of embryo development, decreasing in levels in the morula and blastocyst stages. The molecular weight detected was ~140 kDa, which is similar to that in other studies on mouse cells (Centeio et al. 2020; Kunzelmann et al. 2019; Sciancalepore et al. 2024) and may be larger than the predicted size (110 kDa) due to glycosylation (Fallah et al. 2011). In oocytes, immunostaining of TMEM16A showed that it was present at the plasma membrane, whereas after fertilization the protein was intracellular and, like CaCCs in other cell types, may be localized to ER or mitochondrial membranes (Barro‐Soria et al. 2010). In vitro culture of mouse embryos in the presence of T16Ainh‐A01 prevented embryo development from the zygote, 2‐ and 4‐cell stages, but had no effect on the development of morulae to blastocysts, suggesting a role for the TMEM16A channel during pre‐compaction stages of development as well as the oocyte.

Both DIDS and NFA also reduced the number of [Ca2+]i oscillations induced by thimerosal and resulted in an overall elevated [Ca2+]i that did not return to basal levels. Normally the return of [Ca2+]i to baseline after release of Ca2+ from stores relies on Ca2+ uptake into the ER by SERCA (Wakai et al. 2013) as well as Ca2+ extrusion across the plasma membrane by Na+‐Ca2+ exchange (Carroll 2000; Pepperell et al. 1999). It is therefore possible that one or both mechanisms was affected by these inhibitors. Cl– plays an important role during the refilling of the ER Ca2+ stores by acting as a counter‐ion for Ca2+ movement (Barro‐Soria et al. 2010; Strauss et al. 2014). Thus, inhibition of CaCCs could alter the gradient for Cl– movement into the ER and thereby prevent Ca2+ re‐uptake, resulting in the sustained elevation of [Ca2+]i observed. Alternately, the overall depolarization of the Em observed in the presence of DIDS and NFA may have caused reversal of electrogenic transport by the Na+‐Ca2+ exchanger, causing Ca2+ influx into the oocyte instead of efflux. These non‐specific CaCC inhibitors may also be causing release of Ca2+ from the ER and/or mitochondrial Ca2+ stores due to direct activation of release mechanisms (Liantonio et al. 2007; O'Neill et al. 2003).

Although it is known that the [Ca2+]i oscillations induced by fertilization cause Em hyperpolarisations, it was not possible to determine the role of TMEM16A in fertilization because this channel is also present in sperm, where it is involved in capacitation and the acrosome reaction (Cordero‐Martínez et al. 2018; Orta et al. 2012). TMEM16A null mice have been produced but die within the first month after birth (Rock et al. 2008). TMEM16A heterozygotes have normal rates of survival and since the null mice were produced by crossing heterozygotes it is possible that fertilization and normal pre‐implantation embryo development were supported in the null mice by the presence of paternal and maternal TMEM16A protein. Further experiments using oocyte‐specific knock‐out of TMEM16A are required to confirm the role of TMEM16A and the hyperpolarizations in fertilization.

In summary, this study provides the first evidence that the CaCC TMEM16A/Ano‐1 is expressed in the QS mouse oocyte where the channel contributes to Em hyperpolarizations in response to oscillations in intracellular Ca2+.

Author Contributions

Sarah Dalati: conceptualization (equal), methodology (equal), investigation, analysis, writing – original draft (lead), formal analysis (lead), writing – review and editing (equal). Vanessa Jones: methodology (equal), investigation, writing – review and editing (equal). Margot Day: conceptualization (lead), supervision, resources, writing – original draft (equal), writing – review and editing (equal).

Ethics Statement

All animal experiments were approved by The University of Sydney Animal Care and Ethics Committee (approval numbers 2008/4838, 2011/5583, 2015/824), and experiments were conducted in accordance with the Australian Code of Practice for Use of Animals in Research.

Conflicts of Interest

The authors declare no conflicts of interest.

Acknowledgements

This work was supported by the Discipline of Physiology and Bosch Institute, University of Sydney. The authors thank Dr. Louise Cole from the Advanced Microscopy Facility, Bosch Institute, University of Sydney for help with confocal microscopy. Open access publishing facilitated by The University of Sydney, as part of the Wiley ‐ The University of Sydney agreement via the Council of Australasian University Librarians.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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