Mutations in Protocadherin 15 and Cadherin 23 Affect Tip Links and Mechanotransduction in Mammalian Sensory Hair Cells

Cochlear hair bundle structure in early postnatal mutants

The functional analysis of the av6J, av3J and v2J mutants described in this paper was performed on cochlear cultures and acute preparations derived from the early postnatal cochlea because the hair cells are known to be particularly suitable for studying mechanotransduction at this stage of development [23], [24]. Figure 2 compares confocal images of phalloidin-stained cochlear hair bundles from av6J/av6J (Fig. 2A–C), av3J/av3J (Fig. 2D–F), v2J/v2J (Fig. 2G–I) and heterozygous +/av3J (Fig. 2J–L) mice at P3. The hair bundles of the av6J/av6J mutant (Fig. 2A–C) showed the least signs of disruption, whilst those of the v2J/v2J mouse (Fig. 2G–I) were the most severely disorganised. In all three mutants, hair-bundle structure was disrupted throughout the length of the cochlea (Fig. 2A–I).

Reduced Pcdh15 ectodomain staining in av6J/av6J mutants

Previously published immunocytochemical data have shown that the av3J/av3J and v2J/v2J mice are effective nulls for Pcdh15 and Cdh23 respectively [11], [25]. To determine whether PCDH15 is expressed in the hair cells of the av6J/av6J mutant, whole-mount preparations of P3 cochleae from +/av6J and av6J/av6J mice were stained with antibodies directed against the intracellular domain of the three major transmembrane forms of PCDH15 that are known to be expressed in the mouse ear, CD1, CD2 and CD3 [11]. Cochlear cultures were also stained with HL5614, an antibody that was raised to a large recombinant fragment encompassing the first two cadherin repeats of the ectodomain of PCDH15, a region that is common to many of the 24 different Pcdh15 mRNA isoforms that have been detected by RT-PCR in the early postnatal cochlea [11]. All these antibodies have been previously validated [11].

At P3, the antibodies to CD1 and CD3 stain the hair bundles of basal-coil hair cells more intensely than those at the apex of the cochlea, whilst antibodies to CD2 and the ectodomain show the reverse pattern, staining hair bundles in the apex more intensely than those in the base. At any given point along the cochlea, the hair-bundle staining observed in the +/av6J and av6J/av6J mice with antibodies to CD1 (Fig. 3A, B), CD2 (Fig. 3C, D) and CD3 (Fig. 3E, F) is similar, both in terms of intensity and pattern. Anti-CD1 and anti-CD2 label the entire bundle and anti-CD3 labels punctae at the tips of stereocilia (Fig. 3 A–F). In contrast, the intensity of hair-bundle staining observed with the ectodomain antibody HL5614 was much reduced in the av6J/av6J mutant relative to that observed in the +/av6J heterozygote, and concentrated on the stereocilia lying in proximity to the kinocilium (Fig. 3G–H). These results suggest that the av6J mutation may lead to degradation or loss of the ectodomain from the transmembrane variants, or specifically reduce the expression of the 4^th isoform class that is predicted to lack the transmembrane and intracellular domains and may be secreted.

FM1-43 loading is reduced in av6J/av6J mutants and abolished in av3J/av3J & v2J/v2J mutants

FM1-43 is an amphipathic styryl dye that is known to rapidly accumulate in sensory hair cells via the mechanotransducer channels that are open at rest in non-stimulated hair bundles [26], [27]. Cochlear cultures from av6J (Fig. 4A–D), av3J (Fig. 4E–H) and v2J (Fig. 4I–L) mice were examined for their ability to load with FM1-43 in response to a brief, 10 second exposure to the dye. Hair cells from av6J/av6J mice showed dye loading both in the apical and the basal turns of the cochlea (Fig. 4B, D). Quantitative analysis indicated a 20–25% reduction in dye loading in both the apical and basal coils of the av6J/av6J mutants relative to that observed in the heterozygous +/av6J controls. In the apical coils the mean grey scale levels were 45.9±3.0 (n = 11) in the +/av6J heterozygotes versus 34.9±2.8 (n = 6) in the av6J/av6J mutants, and in the basal coils the mean grey scale levels were 87.4±4.6 (n = 11) in the +/av6J heterozygotes versus 69.6±7.1 (n = 6) in the av6J/av6J mutants. The differences in grey scale levels between heterozygotes and mutants were not statistically significant (ANOVA plus Tukey test). In contrast, loading of FM1-43 dye was not detected at all in hair cells from either the apical or the basal coils of cochlear cultures prepared from av3J/av3J (Fig. 4F, H) and v2J/v2J mice (Fig. 4J, L). These results indicate that hair cells from av6J/av6J mice, but not those from av3J/av3J or v2J/v2J mice, have mechanotransducer channels that are open at rest.

Gentamicin loading and aminoglycoside sensitivity in Pcdh15 mutants

Hair cells in cochlear cultures prepared from shaker1 mouse mutants that have a mutation in myosin VIIa and have all transducer channels closed at rest [26], [28], fail to load with FM1-43 [26], do not load with [³H]-gentamicin and are resistant to aminoglycoside-induced ototoxicity [29]. Furthermore, there is evidence that the aminoglycoside antibiotics are, like FM1-43, permeant blockers of the hair cell's mechanotransducer channels [30]. [³H]-gentamicin accumulation and sensitivity to aminoglycoside antibiotics were therefore examined and compared in the av6J/av6J and av3J/av3J mutants. Following exposure to 0.1 mM [³H]-gentamicin for a period of 2 h at 37°C, the selective accumulation of radioactive label was observed in the inner and outer hair cells of the av6J/av6J mutant but reduced relative to that seen in the heterozygous +/av6J control (Fig. 5A, B). Labelling was not observed in the hair cells of the av3J/av3J mutant relative to that observed in the heterozygous +/av3J control (Fig. 5C, D). As expected from these data, the hair cells in the av6J/av6J mutant were damaged and extruded from the epithelium when exposed to 1 mM gentamicin for 3 h at 37°C (Fig. 5E, F), whilst those of the av3J/av3J mutant were not (Fig. 5G, H).

Transducer currents are abnormal in av6J/av6J, av3J/av3J and v2J/v2J mutant outer hair cells

To determine whether the mechanotransducer channels could be activated by hair bundle displacements in the mutants, currents were measured in the whole cell mode whilst stimulating the hair bundles with a fluid jet (Fig. 6A–I). Heterozygous +/av6J, +/av3J and +/v2J outer hair cells (OHCs) all responded with large transducer currents when the hair bundles were deflected in the positive direction (towards the kinocilium) using saturating force stimuli (Fig. 6A, D, G). At −104 mV the size of the transducer currents was −970 pA for the +/av6J OHCs (range −700 to −1230 pA), −540 pA for the +/av3J OHCs (range −370 to −850 pA) and −300 pA for the +/v2J OHCs (range −240 to −350 pA), showing considerable variability as reported before for rat OHCs [31]. Such variability may reflect differences between OHCs in the numbers of intact tip-link – transducer channel complexes surviving experimental procedures, a variable that may also differ between the strains of mice used. When the bundles were moved in the negative direction (away from the kinocilium), any transducer channels open in the absence of mechanical stimulation (normally about 5–10%) were closed, resulting in a reduction of the inward current. Starting from −104 mV and stepping the membrane potential to more depolarized values, the transducer currents decreased in size at first and then reversed at close to 0 mV. For more positive membrane potentials, fluid jet stimuli in the positive direction caused outward currents and the fraction of transducer channels open without stimulation increased, probably as a consequence of reduced adaptation due to a decreased driving force for Ca²⁺ ions through the transducer channels [32].

In the av6J/av6J mutants, transducer currents could be elicited over the range of membrane potentials tested (Fig. 6B). The current-voltage curves (Fig. 6C), averaged for two +/av6J and three av6J/av6J OHCs, show that the amplitudes were smaller in the homozygotes by about 65% at −104 mV, gradually reducing to 37% at +96 mV. Also, a small resting transducer current was evident at negative potentials that increased at positive potentials. When the same voltage step protocol was applied to homozygous av3J/av3J OHCs the transducer current was absent or small at all but the most positive potentials (Fig. 6E), where clear current responses could be seen. These currents were, surprisingly, in response to the negative phase of the sinusoidal force stimulus that would move the bundle away from the kinocilium. Figure 6F shows the averaged current-voltage curves for the peak-to-peak transducer currents of five +/av3J and five av3J/av3J OHCs. The amplitudes of the transducer currents were considerably smaller in the homozygous mutant OHCs than those measured in the heterozygous controls, by 93% at −104 mV. The reduction in current decreased gradually at positive membrane potentials, down to 53% at +96 mV. One of the five av3J/av3J OHCs had, when stimulated with larger sinusoidal force stimuli (45 V driver voltage amplitude), larger transducer currents in response to stimuli directed away from the kinocilium, also at negative membrane potentials. This suggests that large transducer currents can be elicited, but they are not activated at rest and large forces are required to evoke them. The homozygous v2J/v2J mutants had transducer currents at all membrane potentials (Fig. 6H), but again mostly in response to force directed away from the kinocilium. Currents at negative membrane potentials were smaller than those at positive membrane potentials and reached a peak earlier, suggestive of strong adaptation. No resting transducer current occurred at any potential. The current-voltage curves (averaged from two +/v2J and three v2J/v2J OHCs) show the relatively small transducer currents observed at negative potentials for the homozygous mutant OHCs, which were 81% smaller than those of the heterozygous controls at −104 mV. Note that at positive potentials the currents are actually larger in the homozygotes, most likely because the heterozygous control currents happened to be rather small in this sample of two OHCs from a single mouse.

One concern about the unusual electrical responses of the av3J/av3J and v2J/v2J OHCs is that they might not be transducer currents but some artefact, for example, of modulating the seal resistance between the patch pipette and the cell membrane. The currents in one of these mutants, v2J/v2J, were therefore investigated further, using mechanical step stimuli. At a hyperpolarized holding potential of −84 mV a considerably larger driver voltage to the fluid jet, and hence force on the hair bundle, was needed to elicit currents from the mutant v2J/v2J OHC than was required for the control +/v2J OHC (Fig. 7). Whereas the control cell responded maximally to force stimuli in the positive direction (Fig. 7A, C), the mutant OHC responded best to negatively directed stimuli (Fig. 7B, D). Of the five mutant OHCs tested using force steps, one responded exclusively to positive, one exclusively to negative, and three mainly to negative steps, but with smaller responses also to positive steps. The currents in all five mutant OHC showed strong adaptation at −84 mV during the force steps (example in Fig. 7B), and considerably more so than in the heterozygous controls (Fig. 7A). Adaptation was no longer evident when the membrane potential was stepped to +86 mV (Fig. 7A, B), at which potential the currents in the mutant OHCs decayed quite slowly upon termination of the force step (Fig. 7B). Currents also developed more slowly in the mutant OHCs, with onset kinetics speeding up with the larger steps (Fig. 7B). In three mutant OHCs, bundle displacements were measured and it was found that a bundle displacement of 57±16 nm (in the negative direction) was required to elicit a transducer current at −84 mV. At a depolarized membrane potential of +86 mV the relation between currents and driver voltage shifted towards smaller driver voltages for both the control +/v2J OHCs and the mutant v2J/v2J OHCs (Fig. 7C, D), an observation usually interpreted as being due to abolished Ca²⁺-induced adaptation upon approaching the Ca²⁺ equilibrium potential [32]. For the +/v2J OHCs this shift resulted in an increased fraction of the transducer channels open at rest. However, the shift was not sufficient to establish a resting transducer conductance at +86 mV for the v2J/v2J OHCs. Although the currents of the v2J/v2J OHCs had several unusual features, including abnormal directional sensitivity and slow onset kinetics, the disappearance of adaptation at positive potentials cannot be explained by a seal-resistance artefact and identifies the currents as mechano-electrical transducer currents.

Tip link status in mutants

Electron microscopy was used to assess the status of tip links in av6J, av3J and v2J mice. Difficulties are, however, encountered locating tip links or potential tip link sites in disorganised hair bundles and ambiguities can be caused by the transient lateral links that are present in abundance at the early stages of development. Several types of tissue were therefore examined. Cochlear hair bundles were studied by field emission scanning electron microscopy (FESEM) in the av6J mouse at P9, and in both the av3J and v2J mice at P4. Utricular bundles were examined by transmission electron microscopy (TEM) in the av6J and av3J mice at P15 and P26, and in the v2J mice at P11. In scanning electron micrographs, links were categorized as tip links if they emanated from the tip of a stereocilium in any of the first 3–4 tallest rows and attached to the side of a taller stereocilium in the next row. In immature bundles (i.e., at P3 and P4), such tip links could be directed to either or both of the two nearest taller stereocilia [33], unlike in the mature bundle where tip links primarily are directed to only one taller stereocilium and usually along the excitatory axis [5]. Links categorized as ‘tip links’ therefore do not conform exclusively to the adult form and may represent a combination of transient lateral and/or tip links that are subsequently refined to remove extraneous ones.

Tip-link like links were present in both +/av6J (Fig. 8A, B) and av6J/av6J cochlear hair cells at P9 (Fig. 8C–G). These tip links included single filaments, forked filaments, broken filaments and multiple filaments (Fig. 8A–E) similar to those described previously in various rodents [5], [33], [34], [35], [36], [37]. Double links were also detected around the tips of some stereocilia (Fig. 8F) and lateral links were seen close to the tips (Fig. 8G). To determine whether the number of these tip-link like links was reduced in av6J/av6J mice, 10 randomly selected hair bundles were assessed. The links were assigned to one of four categories: ‘unforked’, ‘forked’, ‘stumps’, or ‘possible’, the latter where a link was morphologically unclear. Expressed as a proportion of possible tip link sites (i.e. tip-side appositions) the homozygote had fewer occupied sites (30%) compared with the heterozygote (46%), fewer unforked links (23% vs 47%) and fewer stumps (5% vs 9%) but more ambiguous links (65% vs 38%). Since most of the hair bundles had some abnormality in row structure, the number of possible sites was smaller in the homozygote than the heterozygote.

Clearly identifiable tip links, including forks, were visible on the OHCs and IHCs from the +/av3J (Fig. 9A–C) +/v2J mice (Fig. 10A–C) at P4. Because these are early stages, the heterozygotes tended to have spoke-like links typical of immature hair cells [33]. The av3J/av3J hair cells had tip-link like links (Fig. 9D–F), whilst the v2J/v2J hair cells had no unambiguous full or forked tip links but there were occasional shortened stumps and random filaments at the tips (Fig. 10D–F). Counts from two av3J/av3J mice showed 17% and 6% respectively of sites were occupied by some form of link-like structure compared with 67% in the +/av3J control. In a v2J/v2J mouse the number of occupied sites was 3% compared with 30% in the control. Ranking of the stereocilia was also apparent in both the av3J/av3J and the v2J/v2J homozygotes although bundle defects were evident.

Using TEM, tip links could also be documented in mature vestibular hair cells of av6J/av6J mice (Fig. 11A, B), but not in av3J/av3J (Fig. 11C, D) or v2J/v2J (Fig. 11E, F) mice. The number of potential tip link sites that were occupied by tip links (including putative tip links) was evaluated quantitatively. In the +/av6J mouse 86% (12/14) of potential sites were occupied by a tip link, whereas only 46% (14/30) were found to have tip links in the av6J/av6J mutant. For the +/av3J mouse 80% (32/40) sites were occupied whilst 0% (0/43) was occupied in the av3J/av3J mutant. For the +/v2J mouse 43% (23/53) sites were occupied whereas 4.3% (2/46) were occupied in the v2J/v2J mice. The top of the shorter stereocilium at tip-link sites was often found to be wedge-shaped in the +/av6J and av6J/av6J mice, as well as in the +/av3J and the +/v2J mice (Fig. 11A–C, E), possibly as a consequence of mechanotransducer channel activity regulating actin polymerisation near the channel. In contrast, the tops of the shorter stereocilia at the unoccupied tip-link sites in the av3J/av3J or v2J/v2J mice were usually more rounded and dome-shaped (Fig. 11D, F).

Mice

The use of mice at Case Western Reserve University (CWRU) was approved under protocol number 2010-0074 (entitled ‘Characterization of mouse models of deafness’) by the Institutional Animal Care and Use Committee (IACUC). The main objective of protocol 2010-0074 is to use various tools and techniques to understand the molecular basis of hearing and deafness using mouse models. At the University of Sussex, all animal procedures were performed under a UK Home Office Project licence (PPL70/6721 entitled Molecular, Cellular and Physiological Basis of Hearing and Deafness) and with the approval of The University of Sussex Ethical Review Committee.

To study changes associated with mutations in Pcdh15, two alleles of Ames waltzer (av) were used: Pcdh15^av6J [21] and Pcdh15^av3J [48]. Most of the data from the av mice was compared to that obtained from hair cells from mice with a mutation in Cdh23, the waltzer v2J allele. Mice homozygous for the av6J allele (av6J/av6J) were crossed to mice heterozygous for av6J allele (+/av6J) and offspring were genotyped using a PCR-based approach [21]. A similar breeding scheme was used to obtain mice heterozygous (+/av3J) or homozygous (av3J/av3J) for the av3J allele of Pcdh15, and mice that were heterozygous (+/v2J) or homozygous (v2J/v2J) for the v2J allele of Cdh23. Offspring from the av3J and av6J lines were genotyped as described previously by Pawlowski et al and Zheng et al respectively [21], [49]. Offspring from the v2J line were genotyped using a scheme that takes advantage of the point mutation that results in the loss of HpyCH4 III restriction site in the target sequence. Genomic DNA (∼500 ng/reaction) was amplified under standard reaction conditions using Platinum Taq DNA polymerase (Invitrogen, CA). The reaction mix was incubated at 94°C for 120 s followed by 35 cycles of 94°C for 30 s, 55°C for 20 seconds and 72°C for 30 s, 72°C for 45 s. Primer pairs and product size are as follows: Primer KA1063 – 5′CGA GAC CAA GAC CAG CTA CC 3′ and KA1064 – 5′ACC AGA CTG ACT GGC TTT CA, 412 bp. The PCR product was digested with HpyCH4 III (NEB, MA) according to manufacturer's protocol followed by heat inactivation at 80°C for 15 minutes. The digested sample was resolved on a 3% agarose gel. Three bands (284, 88 and 40 bp) were observed with wild-type allele and two bands (372 and 40 bp) were observed for the v2J allele.

Cochlear culture preparation

Cochlear organ cultures were prepared from av3J, av6J and v2J mice at 0–3 days of age as described previously [23] and maintained on collagen-coated glass coverslips in Maximow side assemblies at 37°C in a medium containing 93% DMEM/F12, 7% foetal bovine serum and 10 µg/ml ampicillin for 24 h before use.

FM1-43 labelling

FM1-43 dye loading experiments were performed as described previously [26]. In brief, coverslips with adherent cochlear cultures were washed once with HEPES Buffered (10 mM, pH 7.2) Hanks' Balanced Salt Solution (HBHBSS), dipped for 10 seconds in HBHBSS containing 3 µM FM1-43, and washed immediately three times in a large volume of HBHBSS (10 seconds for each wash). The coverslips were then placed in a glass-bottomed Perspex chamber containing 0.5 ml HBHBSS and viewed with an upright microscope equipped with epifluorescence optics and FITC filters (excitation 488 nm, emission 520 nm) using a 63× water immersion or a 40× dry objective. Images were captured at fixed time points after dye application using a 12-bit cooled CCD camera (SPOT-JNR). FM1-43 levels were quantified for +/av6J and av6J/av6J hair cells (see Fig. 4 A–D). Images were imported into Adobe Photoshop CS3 and grey levels were measured in the cytoplasm between the cuticular plate and hair-cell nuclei for 23 hair cells for each genotype, for both the apical and basal coils, using a 400 pixel (20×20) region of interest. Statistical significance was assessed using one-way ANOVA followed by the Tukey test, with P<0.05 as the criterion. The number of cochlear cultures used, each of which contained two apical-coil and two basal-coil explants, was: +/av3J (n = 4), +/av6J (n = 11), av3J/av3J (n = 4), av6J/av6J (n = 6). Similarly, apical- and basal-coil explants from +/v2J (n = 5) and v2J/v2J (n = 6) were cultured for FM1-43 labeling.

[³H]-gentamicin labelling

[³H]-gentamicin labelling was performed as described previously [29]. Briefly, coverslips with adherent cochlear cultures were removed from Maximow slide assemblies, placed in 35 mm diameter plastic culture dishes, washed three times with HBHBSS, and incubated in HBHBSS containing 0.1 mM [³H]-gentamicin (Amersham, UK) for 2 h at 37°C. Cultures were then placed on ice, washed three times with 4 ml of ice cold HBHBSS over a 10 minute period and fixed in cold 2.5% glutaraldehyde buffered with 0.1 M sodium cacodylate, pH 7.2 for 1 h. Following glutaraldehyde fixation, cultures were washed thrice in 0.1 M cacodylate buffer, fixed for 1 h with 1% osmium tetroxide, washed in buffer, dehydrated with ethanol and embedded in Epon 812 resin (TAAB, Berks, UK). One-micrometer thick sections of the apical and basal-coil cochlear cultures were cut from the plastic blocks with glass knives, mounted on glass slides and either stained with Toluidine blue or coated with Ilford L4 Nuclear Research Emulsion (Ilford Imaging UK Limited, Mobberley, Cheshire) for autoradiography. Emulsion-coated slides were exposed in light-tight boxes in the presence of dessicant at 4°C for 1 to 3 weeks, and developed in Ilford Phenisol (diluted 1+4 with H₂O) for 4 min at 18°C. Developed autoradiographs were washed, fixed and dried, mounted under glass coverslips in Histomount, and photographed with an Axioplan 2 light microscope using phase contrast optics.

Aminoglycoside treatment

To test for sensitivity to aminoglycoside exposure, coverslips with adherent cochlear cultures were placed in 35 mm plastic petri dishes, washed once with 3 ml of HBHBSS and then incubated in 3 ml of HBHBSS or 3 ml of HBHBS containing 1 mM neomycin or 1 mM gentamicin at 37°C for 2–4 hours. Following a brief wash, cultures were fixed as described above for grain density autoradiography. Effects of aminoglycosides were evaluated by light microscopy using 1 micron-thick, Toluidine blue-stained sections of basal-coil cultures.

Recording mechanotransducer currents

Mechano-electrical transducer currents from apical-coil outer hair cells were studied in organotypic cochlear cultures of av3J mice (P2 plus two days in vitro) and v2J mice (P1 plus 2 or P0 plus 3 days in vitro), and from the cochleae of av6J mice that were acutely dissected at P4 and P5. All experiments were at room temperature (22–25°C). Extracellular solution was continuously bath-applied at a rate of 6 ml/h. It contained (in mM): 135 NaCl, 5.8 KCl, 1.3 CaCl₂, 0.9 MgCl₂, 0.7 NaH₂PO₂, 2 Na-pyruvate, 5.6 D-glucose, 10 HEPES, with amino acids and vitamins for Eagle's minimum essential medium (MEM) added from concentrates (Invitrogen, UK). The pH was adjusted to 7.5 and the osmolality was about 308 mmol kg⁻¹. The organs of Corti were observed with an upright microscope (Zeiss ACM or Leica DMLFSA, Germany) with Nomarski optics.

Transducer currents were elicited using fluid-jet stimulation (45 Hz sinewaves or steps filtered at 1.0 or 3.0 kHz, 8-pole Bessel) and recorded under whole-cell voltage clamp (HEKA EPC8, Germany or Cairn Optopatch, UK) as previously described [24], [50]. The maximum driver voltage to the fluid jet was set at levels (35 to 40 V) that elicited large, near-saturating transducer currents in control cells. Patch pipettes (resistance in the bath 2–3 MΩ) were pulled from soda glass capillaries and coated with wax. Intracellular solutions contained (in mM): 147 CsCl, 2.5 MgCl₂, 1 EGTA-CsOH, 2.5 Na₂ATP, 5 HEPES; pH adjusted to 7.3 with 1 M CsOH. Data were filtered at 2.5–3.0 kHz, sampled at 5 kHz and stored on computer for off-line analysis using Origin software. Membrane capacitance (C_m) was 5.9±0.3 pF and series resistance after electronic compensation of up to 70% (R_s) was 2.5±0.3 MΩ, resulting in voltage-clamp time constants of 15±2 µs (n = 21). Membrane potentials were corrected for a −4 mV liquid junction potential between pipette and bath solutions. No correction was made for the voltage drop across this series resistance, which was at most 2 mV at extreme potentials. Voltage clamp protocols are referred to a holding potential of −84 mV. Mean values are quoted ± s.e.m. in text and figures.

Immunofluorescence microscopy

Cochleae or cochlear cultures were fixed by immersion in 3.7% formaldehyde in 0.1 M sodium phosphate buffer for 1 hour at RT, and washed 3 times in PBS. After removal of the cartilaginous capsule and the lateral wall, cochlear coils were preblocked and permeabilised in TBS containing 10% heat inactivated horse serum and 0.1% TX-100 for 1 hour, and incubated overnight in the same solution containing a 1∶100 dilution of each of the following affinity purified rabbit antibodies directed against PCDH15 (a kind gift from Dr. Tom Friedman, NIDCD): PB303, 464-2B and PB375 recognising the cytoplasmic domains of the CD1, CD2 and CD3 isoforms classes of PCDH15 respectively, and HL5614 directed against an N-terminal ectodomain sequence common to all classes. Antibody HL5614 was used in the presence of 2 mM EDTA. Samples were washed 3× with TBS, and stained with FITC conjugated swine anti-rabbit Ig (1∶100 dilution) and rhodamine conjugated phalloidin (1∶1000 dilution) for 2 h, washed in TBS, mounted in Vectashield and viewed with a Zeiss LSM 510 confocal microscope using a 100× oil immersion lens NA1.4. To examine hair bundle morphology in av6J mice, cochlear coils were stained only with Texas Red conjugated phalloidin.

Transmission electron microscopy

Utricles from av3J mice at P15 and P26 and av6J mice at P15 were dissected in HBHBSS and fixed immediately in glutaraldehyde followed by osmium tetroxide and embedded in resin as described above for grain density autoradiography. For v2J mice the utricular maculae were dissected in 70% ethanol from labyrinths that had been immersion fixed in glutaraldehyde followed by osmium. Thin (∼90 nm thick) sections were cut with a diamond knife, mounted on copper grids, double stained with uranyl acetate and lead citrate [51]. Sections were viewed in a Hitachi 7100 transmission electron microscope and images were captured with a Gatan Ultrascan 1000 CCD camera. Sections from the utricles of 2 +/av6J, 4 av6J/av6J, 6 +/av3J, 7 av3J/av3J, 4 +/v2J and 2 v2J/v2J mice were examined by transmission electron microscopy.

Scanning electron microscopy

Cochleae from heterozygous and homozygous av6J mice at P9, v2J at P4 and av3J mice at P3 were fixed by intra-labyrinthine perfusion with 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer containing 2 mM CaCl₂ (pH 7.4) and then immersed in the fixative for 2 h. They were stored in 1/10^th concentration fixative (0.25% glutaraldehyde) diluted with buffer at 4°C until further processing, subsequently washed in buffer, dissected to reveal the organ of Corti, and post-fixed with cacodylate-buffered 1% OsO₄ for 1 h. After washing in cacodylate buffer, samples were impregnated with osmium using the osmium-thiocarbohydride (OTOTO) method; for details see [52]. After OTOTO they were dehydrated through a series of increasing ethanol concentrations up to 100% ethanol dried over molecular sieve, and critical point dried from liquid CO₂ using a Polaron critical point dryer. Cochleae were affixed to specimen stubs using silver conducting paint (Agar Scientific, Stansted, UK) and examined in a Hitachi S4500 field emission SEM operated at 5 kV. Specimens from the cochleae of 4 +/av6J, 2 av6J/av6J, 2 +/v2J, 2 v2J/v2J, 2 +/av3J and 2 av3J/av3J mice were examined by scanning electron microscopy.