All experiments were performed in accordance with Home Office regulations under the Animals (Scientific Procedures) Act 1986 and following approval by the University of Sheffield Ethical Review Committee. Tmc2 knockout mice were obtained from The Jackson Laboratory (B6.129-Tmc2^tm1.1Ajg), maintained on a C57BL/6J background, and genotyped as previously described (Pan et al., 2013).

OHCs were studied in acutely dissected organs of Corti from mice of postnatal day (P) 3 to P14. This wide age-range was used to investigate the role of TMC2, which is mainly expressed during the first postnatal week, in determining the biophysical and developmental properties of the MET current, and how these compare when TMC1 is present (mainly expressed from the second postnatal week) (Kawashima et al., 2011). We have also investigated whether the absence of TMC2 affected the normal progression of OHC development by measuring their basolateral membrane properties at immature (P6) and mature (P13–P14) stages.

In the mouse, experiments were performed on OHCs positioned in the apical coil of the cochlea, corresponding to a frequency range in adult animals of about 6–11 kHz (Müller et al., 2005). This cochlear region corresponds to about the position Ac-2 in Figure 2. We focused our experiments on apical OHCs because this cochlear region is easier to dissect in the mouse. In a few experiments, P7–P8 OHCs were also recorded from the apical region of the Mongolian gerbil cochlea (Figure 6), which is tuned to a characteristic frequency of ~0.35 kHz (Müller, 1996).

Mice and gerbils of either sex were killed by cervical dislocation, the cochlea removed and the organ of Corti dissected in extracellular solution composed of (in mM): 135 NaCl, 5.8 KCl, 1.3 CaCl₂, 0.9 MgCl₂, 0.7 NaH₂PO₄, 5.6 D-glucose, 10 HEPES-NaOH, 2 Na-pyruvate. Amino acids and vitamins (Eagle’s MEM) were added from concentrate (pH 7.5, 308 mOsmol/kg). Once dissected, the apical coils of the organs of Corti were transferred to a microscope chamber filled with extracellular solution and viewed using a long working-distance 63× water-immersion objective on a Leica DMLFS microscope.

Soda glass capillaries were used to pull patch pipettes of resistances 2–4 MΩ and the shank of the electrode was coated in surf wax (Mr. Zoggs Sex Wax, CA, USA). Pipettes were filled with an intracellular solution of composition (in mM): 106 L-glutamic acid, 20 CsCl, 10 Na₂phosphocreatine, 3 MgCl₂, 1 EGTA-CsOH, 5 Na₂ATP, 5 HEPES and 0.3 GTP (pH adjusted to 7.3 with CsOH, 294 mOsmol/kg) for most MET current recordings except those measuring Ca²⁺ permeability. For basolateral recordings, pipettes were filled with the following solution (in mM): 131 KCl, 3 MgCl₂, 1 EGTA-KOH, 5 Na₂ATP, 5 Hepes-KOH, 10 Na-phosphocreatine (pH 7.28, 298 mmol/kg). An Optopatch amplifier (Cairn Research Ltd., UK) was used to obtain MET current recordings at room temperature (20–25°C). pClamp software (Molecular Devices, USA: RRID:SCR_011323) and a Digidata 1440A were used to acquire data. Data were filtered at either 2.5 or 5 kHz (8-pole Bessel) and sampled at 10 or 50 kHz. Origin software (OriginLab, USA: RRID:SCR_002815) was used to perform offline data analysis. Membrane potentials were corrected for a measured liquid junction potential of −11 mV and −4 mV for Cs-glutamate and KCl intracellular solutions, respectively.

A 25 mm diameter piezoelectric disc was used to direct a fluid jet via a pipette onto the stereociliary bundle, displacement of which elicited the MET currents (Kros et al., 1992; Corns et al., 2014; Corns and Marcotti, 2016). The tip of the fluid jet pipette had a diameter of 8–10 μm and was positioned at about 8 μm from the bundles. Bundle displacement was calculated from previously calibrated recordings (10.1 nm/V: Corns et al., 2014). Mechanical stimuli were applied as steps or 50 Hz sinusoids (filtered at 1 kHz, 8-pole Bessel).

The Ca²⁺ selectivity of the MET channel was determined under conditions where Ca²⁺ was the only permeant cation in the extracellular solution, and Cs⁺ was present in the intracellular solution. The reversal potential of the MET current, which was interpolated by fitting the data around the 0-current level with a linear regression line, was used to calculate the relative permeability, P_Ca/P_Cs (see also Kim and Fettiplace, 2013). For these experiments, MET currents were recorded in response to sinusoidal 50 Hz force stimulation of hair bundles combined with a voltage ramp (288 mV/s) from −129 mV to +91 mV (see Corns et al., 2016). To measure the calcium reversal potential, a CsCl based intracellular solution of the following composition (mM): 135 CsCl, 3 MgATP, 10 Tris phosphocreatine, 1 EGTA-CsOH, 10 HEPES-CsOH (pH 7.2, 293 mOsmol/kg) and a high Ca²⁺ extracellular solution containing (mM): 100 CaCl₂, 20 N-methylglucamine, 6 Tris, 10 D-glucose (pH 7.4), were applied as described above. A liquid junction potential of −9 mV was used to correct the reversal potentials. To calculate the relative permeability, P_Ca/P_Cs, the Goldman-Hodgkin-Katz equation was applied: P_Ca/P_Cs = {a₁[Cs⁺]/4a₂[Ca²⁺]} × {exp(V_revF/RT)} × {1 + exp (V_revF/RT)}, where RT/F has its usual meaning with a value at room temperature of 25.7 mV, [Cs⁺] and [Ca²⁺] are the concentrations of Cs⁺ intracellularly (140 mM) and Ca²⁺ extracellularly (100 mM) and a₁ (0.711) and a₂ (0.519) are the published activity coefficients for Cs⁺ (Partanen, 2010) and Ca²⁺ (Rard and Clegg, 1997), respectively.

Stock solutions of 100 mM DHS (Sigma, UK; molecular weight = 730.7) were prepared in water and diluted to the final concentration in either extracellular or intracellular solution on the experimental day. During the recordings, all test solutions containing DHS were superfused via a pipette positioned orthogonally to the axis of mechanical sensitivity of the hair bundle and care was taken to ensure they were present in the fluid jet (for more details see: Corns et al., 2014; Corns and Marcotti, 2016).

The voltage-dependent block of the transducer current by extracellular DHS was quantitatively described by a two-barrier one-binding-site model previously described in more detail (Marcotti et al., 2005; van Netten and Kros, 2007), according to the reaction scheme: C+D0k1⇔k−1CDk2⇔k−2C+Di

where C represents the unblocked MET channels, CD the blocked channels, D₀ and D_i the extra- and intracellular blocker. The forward (k₁, k₂) and backward (k₋₁, k₋₂) rate constants are voltage dependent. In brief, the voltage dependence of the block is expressed by four parameters: E_b, the free energy of the drug binding site at zero membrane potential; δ_b, the site’s fractional electrical distance across the membrane from the extracellular side; ΔE = E₂ − E₁, the difference between the intracellular and extracellular free energy levels of the two barriers at zero membrane potential; Δδ = δ₂ − δ₁, the fractional electrical distance between the intra- and extracellular barriers. Fits using this model with a Hill coefficient, n_H, of one and a valence for DHS of +2 (Marcotti et al., 2005) yield values for ΔE, E_b, Δδ and δ_b, where the binding site is located at a relative electrical distance δ_b of 0.79 and Δδ is 0.91 (Marcotti et al., 2005). The forward rate constant k₁ equals the slope of τ⁻¹ vs. [D]₀, allowing calculation of the absolute values of the energy barriers E₁ and E₂. This in turn enabled us to calculate k₂ and the entry rate of drug molecules into the OHCs.

Cochlear cultures from homozygous Tmc2 knockout mice, their heterozygous littermate controls and wild-type C57BL/6J mice were prepared as described previously (Richardson and Russell, 1991). Briefly, cochleae were dissected from 3 day postnatal pups in HEPES buffered (10 mM, pH 7.2) Hanks’ balanced salt solution (HBHBSS), placed onto collagen-coated glass coverslips, fed one drop of complete medium (containing 93% DMEM/F12 (Sigma, UK), 7% fetal bovine serum (FBS: Biosera, UK) and 10 μg/ml ampicillin (Sigma)), sealed into Maximow slide assemblies and maintained at 37°C for 1 day.

After a day at 37°C (see above, “Cochlear Culture Preparation” section), cochlear cultures were transferred into a sterile 35 mm-diameter petri dish (Corning) and incubated in a medium (100% DMEM/F12 and 10 μg/ml ampicillin) containing different concentrations of the aminoglycoside antibiotic streptomycin (Sigma) for 24 h at 37°C. A stock solution of streptomycin (100 mM; molecular weight 728.7) was prepared in HBHBSS. After 24 h, the coverslips with adherent cultures were washed three times with 3 ml of HBHBSS, fixed with 4% paraformaldehyde in phosphate buffered saline (PBS) for 1 h at 22°C, washed three times with PBS and stained with a solution containing Alexa Fluor 488 phalloidin (1:300: Life Technologies, RRID:AB_2315147), 0.7% FBS and 0.01% Triton -X100 for 2 h. The coverslips were then washed another three times in PBS, and the collagen with the attached cochleae was peeled off from the coverslips and mounted in Vectashield mounting medium (Vector Laboratoris, RRID:AB_2336789). Cochleae were imaged with an Olympus BXB61 with 10× or 20× dry objectives, and images were captured using the Volocity 3D Image Analysis Software (RRID:SCR_002668). The number of hair cells along the four different cochlear regions (see “Results” section) was measured over a 200 μm length region using Photoshop.

Texas Red conjugated neomycin was prepared using a method similar to that described (Steyger et al., 2003) for producing a Texas Red conjugated derivative of gentamicin. In brief, 4.5 ml of neomycin sulfate (Sigma, 50 mg/ml 100 mM in K₂CO₃ buffer pH 8.5) was added to 0.5 ml of Texas Red succimidyl esters (mixed isomers, Invitrogen T6134, 2 mg/ml in dry dimethyl formamide) and the solution was mixed overnight on a rotator at 4°C. Aliquots were then snap frozen and used without further purification. Cultures were incubated in HBHBSS containing ~0.1 mM Texas Red neomycin for 5 min at 22°C, fixed in 4% PFA for 1 h, washed three times in PBS and labeled with phalloidin as described above. Images were captured using an Olympus BXB61 (see above). Texas Red neomycin was used for these experiments rather than Texas Red streptomycin as it produced more reproducible hair-cell loading during acute application. As a control for the stability of neomycin during conjugation, 4.5 ml of the neomycin solution was mixed with 0.5 ml dimethylformamide (DMF) overnight, and as a control for the specificity of the Texas Red neomycin conjugate, 0.5 ml of the Texas Red ester was mixed with 4.5 ml of the K₂CO₃ buffer. The neomycin retained toxicity after overnight incubation in the presence of 10% DMF, and labeling of hair cells was not observed with Texas Red alone. Hair-cell labeling was only observed with the Texas Red neomycin conjugate. In total, six control and four mutant cochlear cultures were examined from five mice.

Statistical analysis was performed with Prism (GraphPad Software, RRID:SCR_002798). Comparisons of means were made by Student’s two-tailed t test or deduced from the lower and upper confidence limits. For multiple comparisons ANOVA was used (one-way ANOVA followed by Tukey’s test; two-way ANOVA followed by Bonferroni’s test). P < 0.05 was selected as the criterion for statistical significance. All values are quoted as mean ± SEM. For most of the experiments the number of OHCs (or cochleae) and mice used is listed in the Figure legend. Note that in most cases both cochleae were used from each mouse.

Saturating MET currents from apical-coil OHCs of control (Tmc2^+/−) and littermate mutant (Tmc2^−/−) mice were elicited by displacing their hair bundles with sinewave stimuli from a piezoelectric fluid jet stimulator (Kros et al., 1992; Corns et al., 2014; Corns and Marcotti, 2016). At negative membrane potentials, bundle displacement in the excitatory direction (i.e., towards the taller stereocilia) elicited large inward MET currents from postnatal day 5 (P5) OHCs of both genotypes (Figures 1A,B). By this age, the expression of Tmc2 mRNA in apical OHCs has slightly declined to ~75% of its maximum, which occurs at P4 (Kawashima et al., 2011). Membrane depolarization caused the MET current to decrease in size at first and then reverse near 0 mV to become outward at positive potentials (Figures 1A,B). As previously reported (Kim and Fettiplace, 2013), the maximum amplitude of the MET current was significantly reduced (P < 0.0001) in apical OHCs of Tmc2^−/− compared to Tmc2^+/− mice at P4 (by 61%) and P5 (by 28%; Figure 1E). Taking a value of 62 pS as the conductance of the MET channel for both genotypes (Kim et al., 2013), we infer the presence of some 156 MET channels in the Tmc2^+/− OHCs at P4, but only 60 in the Tmc2^−/− OHCs, suggesting a limiting supply of TMC1 protein at this stage. However, we also found that from P6 onward the MET current amplitude was comparable between the two genotypes (Figures 1C–E); this is because from around the end of the first postnatal week TMC1 is becoming the main TMC protein expressed in OHCs (Kawashima et al., 2011; Kim and Fettiplace, 2013) and the absence of TMC2 did not impair the normal development of the MET current. In contrast to recent observations in OHCs from mice with a point mutation in Tmc1 (Corns et al., 2016), the resting open probability of the MET channel at both negative and positive membrane potentials was not affected by the absence of TMC2 (Figure 1F), indicating that the latter is not required for regulating the Ca²⁺ sensitivity of the MET channel’s adaptation sensor. We also found that during the second postnatal week, the Ca²⁺ permeability (see also “Materials and Methods” section for details) of the MET channel was similar in Tmc2^+/− (P_Ca/P_Cs: P6–P7 3.37 ± 0.17, n = 7; P9 3.10 ± 0.13, n = 4) and Tmc2^−/− (P6–P7 2.98 ± 0.12, n = 6; P9 3.27 ± 0.16, n = 5) OHCs. Despite this similarity at older ages, the Ca²⁺ permeability was significantly reduced in Tmc2^−/− P3–P4 OHCs (Tmc2^+/−: 4.40 ± 0.24, n = 4; Tmc2^−/−: 2.52 ± 0.13, n = 5, P < 0.0005), in agreement with previous findings (Kim and Fettiplace, 2013; Pan et al., 2013; Beurg et al., 2015).

We further tested the MET channel properties in Tmc2^−/− OHCs by evaluating the permeation of larger molecules such as aminoglycoside antibiotics, which are known to behave as permeant blockers of the channel (Marcotti et al., 2005) leading to hair cell apoptosis (Forge and Schacht, 2000; Schacht et al., 2012). Initially, we tested the effect of streptomycin on cochlear hair cells from P3 C57BL/6J wild-type mice (Figures 2A–C), in order to determine the optimal concentration to be used on OHCs from Tmc2^−/− mice. Following 1 day in culture, we exposed the organs of Corti of C57BL/6J wild-type mice for 24 h to three different concentrations of streptomycin and found that while 0.1 mM had very little effect on hair-cell survival, 0.6 mM almost completely abolished all apical-coil OHCs (Figures 2A–C). Therefore we decided to use an intermediate concentration of streptomycin (0.3 mM) for the following experiments on Tmc2^−/− mice (Figures 2D–G). Streptomycin was tested on apical-coil OHCs at different postnatal ages (P0, P2, P4, P5, P6 and P8) in order to correlate the effect of streptomycin with the temporal expression of TMC2 and TMC1. We found that during a time when Tmc2 is the major subunit expressed in the cochlea (P2–P4: Kawashima et al., 2011), OHCs from Tmc2^−/− mice were protected from the ototoxic effects of the aminoglycoside (Figures 2D,F) compared to those from Tmc2^+/− mice (Figures 2D,E). This protection was most likely due to, at least in part, the reduced number of MET channels, and hence reduced resting MET current in Tmc2^−/− OHCs. This interpretation was also supported by the reduced Texas Red-neomycin hair-cell labeling of Tmc2^−/− compared to that of Tmc2^+/− organs of Corti at this early age (Figure 2G). Although the reduced labeling in Tmc2^−/− mice was not quantified, the same result as that shown in Figure 2G was seen in a total of 12 cochleae from six Tmc2^+/− mice and 10 cochleae from five Tmc2^−/− mice (both genotypes: P4–P5 + 1 day in vitro). Below, we investigate whether different permeation properties of the MET channels for aminoglycoside antibiotics might also contribute to this marked difference in susceptibility. From about P6 onwards, a time when TMC1 expression rapidly increases in the cochlea (Kawashima et al., 2011), OHCs from Tmc2^−/− mice became gradually more susceptible to streptomycin, and similar to that observed in Tmc2^+/− cells (Figures 2D–F).

The aminoglycoside antibiotic DHS is a well characterized permeant blocker of the MET channel (Marcotti et al., 2005). The degree of block of the MET channel by DHS varies with extracellular Ca²⁺ and membrane potential (Kroese et al., 1989; Ricci, 2002; Marcotti et al., 2005), with high Ca²⁺ concentrations reducing both the entry of the aminoglycoside into the MET channel and the channel’s affinity for the drug, and positive membrane potentials reducing the degree of block of the current by DHS. These results led to the proposal that the DHS-binding site sits within the channel’s permeation pathway (Kroese et al., 1989; Marcotti et al., 2005; van Netten and Kros, 2007). Therefore, we assessed whether TMC2 directly affected the ability of DHS to block the MET channel. MET currents were recorded by stepping the membrane potential between −161 mV and +99 mV in 20 mV increments while displacing the hair bundles of Tmc2^+/− and Tmc2^−/− OHCs before and during the application of 30 μM extracellular DHS (only voltage steps to −81 mV and +99 mV are shown in Figures 3A,B). Extracellular DHS caused a voltage-dependent block of the MET current, with positive membrane potentials relieving the block in both genotypes (Figures 3A–D), as previously shown (Marcotti et al., 2005). The dose-dependence of the block of the MET current by DHS is shown in Figure 3E. At all concentrations tested, the degree of block of the MET current (I_DHS/I_control) at −81 mV in Tmc2^−/− OHCs was significantly stronger than in Tmc2^+/− OHCs (3 μM: P < 0.05; 10 μM: P < 0.005; 30 μM: P < 0.02). The concentration for half block (K_D) at −81 mV in Tmc2^−/− OHCs (5.9 μM) was significantly lower (P < 0.001) than that measured in Tmc2^+/− OHCs (9.1 μM). We further investigated the voltage dependence of the block of the MET channel by DHS in OHCs by plotting the MET current in the presence of the aminoglycoside as a fraction of the control current (I_DHS/I_control, Figure 3F). The block of the OHC MET channel by DHS was consistently more effective in Tmc2^−/− mice compared to that in Tmc2^+/− mice and a partial relief of the block for large negative membrane potentials (negative to −80 mV) was seen in both genotypes (Figure 3F). This relief of the block is consistent with DHS being a permeant blocker of the MET channel when sufficient electrical driving force is present (Marcotti et al., 2005). The fits through the data are according to a two-barrier one-binding-site model (Marcotti et al., 2005).

To determine whether the rate of entry of DHS molecules into the OHCs changed in the presence of TMC2, we measured the time constant of MET current decline when exposed to DHS. To ensure that the MET channels are initially free from DHS, the hair bundles are first deflected in the inhibitory direction, closing all MET channels so that DHS is driven out of the permeation pathway, as the DHS molecules cannot reside in the closed channel (Marcotti et al., 2005). A saturating excitatory stimulus is then given prior to and during the application of different concentrations of extracellular DHS. In the presence of DHS the MET current is initially large, and then reduces to a steady level (Figures 4A,B) as the DHS enters and blocks the MET channel (open-channel block: Marcotti et al., 2005). Plotting the inverse of the time constant of DHS binding kinetics against the DHS concentration allows us to calculate the rate constant k₁ (see “Materials and Methods” section) of DHS entry into the OHCs of both Tmc2^+/− and Tmc2^−/− mice (Figure 4C). Figure 4D shows that the absence of TMC2 increases the strength of E_b, the DHS binding site in the channel pore. Because of the different numbers of MET channels between the two genotypes, and to take any possible difference in resting open probability out of the equation, we calculated entry rates of DHS per open channel, rather than per OHC. In the first instance we calculated entry for a 1 μM DHS concentration, as this is the approximate aminoglycoside concentration in the endolymph that just leads to ototoxicity in vivo (Tran Ba Huy et al., 1981), and for comparison with data from wild-type OHCs (Marcotti et al., 2005). Assuming a driving force of −150 mV and using 1.3 mM extracellular Ca²⁺, we estimated the rate of DHS molecules entering the OHC via the open MET channels to be 75.7 molecules/open channel/s for Tmc2^+/− and 63.9 molecules/channels/s for Tmc2^−/− OHCs. At −55 mV, near the resting potential of neonatal OHCs (Marcotti and Kros, 1999; Marcotti et al., 1999), the rates were 18.9 molecules/channel/s and 26.6 molecules/channel/s, respectively. These rates were comparable, but with some voltage dependence: in the absence of TMC2 they were somewhat lower at −150 mV and somewhat higher at −55 mV. At higher concentrations of DHS, when the entry was found to approach saturation, rates were consistently lower in the absence of TMC2. For 300 μM DHS, the same concentration that we used for the streptomycin experiments shown in Figure 2, entry rates reached 1078 molecules/channel/s for Tmc2^+/− and 586 molecules/channel/s for Tmc2^−/− OHCs at −150 mV. At −55 mV, rates were 315 molecules/channel/s for Tmc2^+/− and 74 molecules/channel/s for Tmc2^−/− OHCs. The lower saturation level for aminoglycoside entry in the absence of TMC2 is thus likely to contribute to the protection from ototoxic damage in the Tmc2^−/− mice during the first postnatal week, in addition to their smaller MET currents.

In order to determine whether the ability of DHS to block the MET current from the intracellular side was also affected in Tmc2^−/− OHCs, we added DHS in concentrations ranging from 30 μM to 3 mM into the intracellular solution. For these experiments the block of the MET current is only observed at positive potentials in OHCs from both genotypes (Figures 5A–C) but, as previously described (Marcotti et al., 2005), with a much reduced potency compared to extracellular DHS (Figure 3). Because of the difficulty of obtaining drug-free control recordings before and after intracellular drug application, we normalized the unaffected currents at −101 mV as a control to calculate the degree of block at +99 mV. We found that in P4–P5 OHCs the K_D for intracellular DHS block was significantly lower in Tmc2^−/− (124 μM) compared to that measured for Tmc2^+/− OHCs (717 μM, P < 0.00001) (Figure 5D). We also found that in Tmc2^+/− OHCs, the degree of block of the MET channel by 300 μM intracellular DHS (I_DHS/I_c), which corresponds to the steeper part of the dose-response curve, was significantly (P < 0.001) lower at P4–P5 (closed circles: Figure 5D) when compared to P8 (red triangle: Figure 5D). However, in Tmc2^−/− OHCs the degree of block was similar between P4–P5 (open circles: Figure 5D) and P8 (blue triangle: Figure 5D). The overlapping symbols at P8 also show that by this time the block was the same for Tmc2^+/− and Tmc2^−/− OHCs. This highlights that TMC2 confers a lower affinity of binding for DHS within the MET channel pore only during the first postnatal week when it is preferentially expressed over TMC1.

Vestibular hair cells, which, different from mouse cochlear hair cells, transduce low frequency signals, have been shown to retain TMC2 throughout adulthood (Kawashima et al., 2011). We hypothesized that TMC2 could be important for low frequency MET and as such it could remain in the low frequency region of the mature cochlea. Although the mouse cochlea is widely used for hearing research, it is mainly tuned to high frequencies (hearing frequency range: ~2–100 kHz, Ehret, 1975; Greenwood, 1990). To test our hypothesis, we performed MET current recordings from OHCs positioned in the apical coil of the gerbil cochlea, which is tuned to frequencies well below 1 kHz (mean in vivo characteristic frequency of ~0.35 kHz: Müller, 1996). Saturating bundle displacement of apical-coil gerbil OHCs (Figure 6A) elicited large MET currents similar to those obtained from mouse OHCs (Figure 1). In order to investigate whether TMC2 might be retained in older low-frequency gerbil OHCs, we tested the degree of block of the MET current by 300 μM intracellular DHS. At +99 mV a substantial block of the MET current was observed in apical OHCs of the gerbil (Figure 6B). The degree of block at +99 mV by intracellular DHS in gerbil P7–P8 OHCs (0.52 ± 0.07, n = 4) was not significantly different from that of mouse P8 OHCs of either Tmc2^+/− (0.57 ± 0.02, n = 7) or Tmc2^−/− (0.57 ± 0.04, n = 7) mice (Figure 6C). This suggests that, as for P8 mouse apical OHCs, TMC2 is not expressed in more mature gerbil OHCs, even in low-frequency regions of the cochlea.

The absence or mutation of Tmc1 (deafness and Beethoven mice, respectively) has been shown to prevent the normal progression in hair cell maturation at the onset of hearing, such that they retain immature biophysical properties (Marcotti et al., 2006). Given the similarity between the Beethoven (Bth) mutation in Tmc1 and the Tmc2 knockout in terms of causing a reduction in MET channel Ca²⁺ permeability (see “Results” section: also see Pan et al., 2013; Corns et al., 2016), we investigated whether the absence of Tmc2 affected the development of basolateral K⁺ currents in OHCs (Figure 7). Membrane currents in OHCs were elicited by applying depolarizing voltage steps in 10 mV increments from −124 mV starting from a holding potential of −84 mV. At P6, OHCs from both genotypes showed a delayed rectifier outward K⁺ current (I_K,neo) (Figures 7A–C). I_K,neo exhibited a normal time course and voltage dependence as previously reported in normal mice (Marcotti and Kros, 1999), and its size, measured at 0 mV, was similar between Tmc2^+/− (2.69 ± 0.21 nA, n = 5) and Tmc2^−/− (2.74 ± 0.09 nA, n = 9, P = 0.82) OHCs (Figure 7C). At around P8, I_K,neo is gradually replaced by I_K,n, which becomes the major K⁺ current expressed in mature mouse OHCs (Marcotti and Kros, 1999). I_K,n is an outward K⁺ current activated at hyperpolarized membrane potentials (Housley and Ashmore, 1992; Marcotti and Kros, 1999) and carried by KCNQ4 channels (Kubisch et al., 1999). We found that I_K,n was present in both Tmc2^−/− and Tmc2^+/− P13–P14 OHCs (Figures 7C,E). The size of I_K,n, which was measured as the deactivating tail currents at −124 mV from the holding potential of −84 mV (difference between instantaneous and steady state inward currents: Marcotti and Kros, 1999), was similar between Tmc2^+/− (396 ± 45 pA, n = 10, P13–P14) and Tmc2^−/− (395 ± 53 pA, n = 9, P13–P14, P = 0.99). The total outward current measured at 0 mV (Figure 7F) was also similar between the two genotypes (Tmc2^+/−: 2.6 ± 0.1 nA, n = 10; Tmc2^−/−: 2.2 ± 0.2 nA, n = 8, P13–P14, P = 0.10). Moreover, all of the other biophysical properties of OHCs, including the resting membrane potential (V_m, Figure 7G) and cell membrane capacitance (C_m, Figure 7H) did not differ between Tmc2^+/− and Tmc2^−/− mice. This indicates that a loss of TMC2 appears to have no detrimental effect on the maturation of the basolateral membrane currents of OHCs, in contrast to what has previously been shown for the M412K point mutation in TMC1 (Beethoven mice: Marcotti et al., 2006).

Increasing evidence, including the localization of TMC1 and TMC2 to the site of the MET channel (Kawashima et al., 2011; Kurima et al., 2015), points to roles for both proteins as components of the MET channel. The M412K point mutation in Tmc1 in IHCs (Pan et al., 2013) and OHCs (Beurg et al., 2015; Corns et al., 2016) reduces the MET channel’s permeation for Ca²⁺ ions. By contrast, TMC2 confers higher Ca²⁺ permeability (see also Kim and Fettiplace, 2013; Pan et al., 2013; Beurg et al., 2015) to the MET channel. Moreover, both proteins directly affect the strength of the binding site for the aminoglycoside DHS within the MET channel pore, with TMC2 (Figure 4D) and the M412K point mutation in Tmc1 (Corns et al., 2016) both reducing the affinity of the DHS binding site in the channel pore. While the point mutation in Tmc1 strongly affected the binding site in the permeation pathway for extracellular DHS, resulting in a reduced entry rate of the drug molecules into the hair cells, the effect of TMC2 was more subtle, with no clear consistent effect on drug entry for low concentrations of DHS. For higher, saturating DHS concentrations the presence of TMC2 increased drug entry during the first postnatal week compared to when the MET channels contained a greater proportion of TMC1 (from ~P5 onwards: Kawashima et al., 2011), pointing to different MET pore properties between the two phenotypes. The saturating DHS entry rate for the MET channel in OHCs from P5–P8 wild-type CD1 mice was 1527 molecules/open channel/s (calculated at −150 mV for 300 μM DHS and using 1.3 mM extracellular Ca²⁺: Marcotti et al., 2005). Despite the fact that at P5–P8 the normal MET channel contains little or no TMC2, the DHS entry, per open channel, was much larger than when TMC2 was not present in the first postnatal days (586 molecules/s: see “Results” section), but closer to when TMC2 was present (1078 molecules/s). Wild-type controls (P6–P8, apical coil) for the Bth mutant mice (Tmc1^+/+: C3HeB/FeJ), gave a similar result to the CD1 mice, at 1462 molecules/s (calculated from data in Corns et al., 2016). The M412K point mutation in Tmc1^Bth/Bth mice (P6–P9) reduced the entry of 300 μM DHS to 521 molecules/s, similar to the TMC2-lacking channels in the first postnatal week. These findings suggest that there is a developmental change in the pore properties of MET channels containing TMC1 but not TMC2 subunits between the first and second postnatal weeks. This change, revealed by a low saturation level of DHS entry in the first postnatal week, is partly compensated for by TMC2.

Block by intracellular DHS, which appeared non-permeant over the voltage range that we could test, was more strongly reduced by the presence of TMC2 (Figure 5) than by the Tmc1 point mutation, suggesting that structural differences between TMC2 and TMC1 are located at different positions along the proteins than the point mutation. The ability of TMC2 and TMC1 to directly alter the permeation properties, conductance, DHS binding site and resting open probability of the MET channel (for TMC1: Pan et al., 2013; Corns et al., 2016), not only supports a direct role for both in MET, but also the conclusion that these two proteins contribute to the properties of the permeation pore of the MET channel.

Different from the mammalian cochlea, vestibular hair cells retain TMC2 in their stereociliary bundle throughout adulthood, such that mechanoelectrical transduction in vestibular organs relies on both TMC1 and TMC2 (Kawashima et al., 2011). Tmc2 transcripts are also observed in the adult inner ear of zebrafish (Maeda et al., 2014). From a functional point of view, one of the main differences between the auditory and the vestibular system is that vestibular end organs, such as the semicircular canals and utricle, respond to subacoustic stimuli up to a few tens or hundreds of Hertz, respectively, in both mammals and lower vertebrates (see: Wilson and Jones, 1979; Eatock and Lysakowski, 2006; Abbas and Whitfield, 2010). We found that the sensitivity to intracellular DHS of the MET current recorded from apical OHCs of the P8 gerbil cochlea (tuned in adult gerbil to ~350 Hz: Müller, 1996), the frequency range of which is near the upper limit of that of the vestibular end organs, was indistinguishable from that measured in aged-matched mouse OHCs (Figure 6). By this time TMC1 is already the main MET channel subunit in mice (Kawashima et al., 2011). This indicates that the expression of TMC2 is not a specific characteristic of hair cells detecting low frequency stimuli. Therefore, there must be another reason as to why TMC2 is no longer expressed in the functionally mature MET channel of the mammalian cochlea.

One of the characteristic features of the mature cochlea is the presence of a large endocochlear potential (~+90 mV) between the endolymph, which surrounds the mechanotransducer apparatus, and the perilymph present around the hair cell basolateral membrane (Békésy, 1952; Bosher and Warren, 1971). This potential is linked to the different ionic composition between the endolymph (K⁺: ~150 mM; Wangemann and Schacht, 1996; Ca²⁺: ~20 μM in the adult; Bosher and Warren, 1978) and the perilymph (Na⁺: ~150 mM, Ca²⁺: 1.3 mM; Wangemann and Schacht, 1996). However, in rodents, the endocochlear potential is absent or very small during the first postnatal week when the endolymph and perilymph are still similar in composition, and it only starts to increase to ~+15 mV during the second postnatal week (Bosher and Warren, 1971). This early low endocochlear potential is comparable to the endovestibular potential measured in mature vestibular end organs (a few mV: Schmidt, 1963). The physiological consequence of this small potential is that the electrochemical gradient that drives Ca²⁺ entry into hair cells is substantially reduced compared to that observed in the mature cochlea. The increased Ca²⁺ permeability conferred to the MET channel by TMC2 in vestibular and early postnatal cochlear hair cells is likely to compensate for this reduced driving force. This could be essential to enable sufficient Ca²⁺ to enter the hair cells to drive Ca²⁺-dependent processes such as adaptation of the MET channel (Corns et al., 2014; Marcotti et al., 2016), which is known to first appear in mice during a time when TMC2 is the main MET channel subunit (<P6). Calcium-induced adaptation in early postnatal OHCs is quite slow and comparable to that recorded in vestibular hair cells (Eatock, 2000; Géléoc and Holt, 2003). Conversely, in the adult mammalian cochlea, the combined endocochlear potential and the higher Ca²⁺ permeability conferred by TMC2 might produce excessive Ca²⁺ entry through the MET channel, reducing its resting open probability and the ability to function at its optimal operating range. This excessive Ca²⁺ influx into mature cochlear hair cells through the MET channels might then in turn lead to cytotoxicity and cell degeneration (Orrenius et al., 2003). An additional or alternative role for the increased Ca²⁺ permeability conferred by TMC2 might be to promote the initial growth of the transducing stereocilia (shorter rows) (Vélez-Ortega et al., 2017), which is known to occur within the first few postnatal days (Tilney et al., 1992).

We also found that different from TMC1 (Marcotti et al., 2006; Kawashima et al., 2011; Kim and Fettiplace, 2013), TMC2 was not required for the maturation of mechanoelectrical transduction and the basolateral properties of OHCs, which parallels previous evidence showing that type II cells of the utricle develop normally in its absence (Kawashima et al., 2011). This suggests that the presence of a functional MET channel during the first few postnatal days, when TMC2 expression predominates, has no bearing on hair cell maturation; whereas the ability of cochlear hair cells to mechanotransduce during the second postnatal week, when TMC1 expression predominates, is essential for their correct maturation into mature sensory receptors (Marcotti et al., 2006).

Recent studies have demonstrated that the auditory part of the inner ear evolved from the vestibular part (Gacek, 2009; Duncan and Fritzsch, 2012). Therefore, we propose that although TMC2 is present during the initial assembly of the MET channel in early postnatal cochlear hair cells, it may be an evolutionary remnant that is not required for the highly specialized hair cells of the mature mammalian cochlea.