Harmonin enhances voltage-dependent facilitation of Ca_v1.3 channels and synchronous exocytosis in mouse inner hair cells

Abstract

Ca_v1.3 channels mediate Ca²⁺ influx that triggers exocytosis of glutamate at cochlear inner hair cell (IHC) synapses. Harmonin is a PDZ-domain-containing protein that interacts with the C-terminus of the Ca_v1.3 α₁ subunit (α₁1.3) and controls cell surface Ca_v1.3 levels by promoting ubiquitin-dependent proteosomal degradation. However, PDZ-domain-containing proteins have diverse functions and regulate other Ca_v1.3 properties, which could collectively influence presynaptic transmitter release. Here, we report that harmonin binding to the α₁1.3 distal C-terminus (dCT) enhances voltage-dependent facilitation (VDF) of Ca_v1.3 currents both in transfected HEK293T cells and in mouse inner hair cells. In HEK293T cells, this effect of harmonin was greater for Ca_v1.3 channels containing the auxiliary Ca_vβ₁ than with the β₂ auxiliary subunit. Ca_v1.3 channels lacking the α₁1.3 dCT were insensitive to harmonin modulation. Moreover, the ‘deaf-circler’dfcr mutant form of harmonin, which does not interact with the α₁1.3 dCT, did not promote VDF. In mature IHCs from mice expressing the dfcr harmonin mutant, Ca_v1.3 VDF was less than in control IHCs. This difference was not observed between control and dfcr IHCs prior to hearing onset. Membrane capacitance recordings from dfcr IHCs revealed a role for harmonin in synchronous exocytosis and in increasing the efficiency of Ca²⁺ influx for triggering exocytosis. Collectively, our results indicate a multifaceted presynaptic role of harmonin in IHCs in regulating Ca_v1.3 Ca²⁺ channels and exocytosis.

Key points

• Ca_v1.3 Ca²⁺ channels mediate sound transmission by triggering presynaptic exocytosis of glutamate from cochlear inner hair cells (IHCs).

• Harmonin is a PDZ-domain-containing protein in IHCs that is altered in Usher syndrome, a form of deaf–blindness in humans.

• We show that harmonin enhances Ca_v1.3 voltage-dependent facilitation (VDF) in transfected HEK293T cells in a manner that depends on the identity of the auxiliary Ca²⁺ channel β subunit.

• Ca_v1.3 VDF is impaired, and synchronous exocytosis and the Ca²⁺ efficiency of exocytosis are reduced, in IHCs from deaf-circler mice expressing a mutant form of harmonin (dfcr) that cannot interact with Ca_v1.3.

• We conclude that harmonin regulates presynaptic function in mouse IHCs, which adds to our understanding of the factors that may influence hearing impairment in Usher syndrome.

Introduction

Voltage-gated Ca_v1.3 Ca²⁺ channels are highly expressed in cochlear inner hair cells (IHCs), where their activity is tightly coupled to the exocytic release from specialized ‘ribbon’ synapses (Platzer et al. 2000; Brandt et al. 2003). Mice lacking Ca_v1.3 are deaf (Platzer et al. 2000; Dou et al. 2004) as are humans with loss-of-function mutations in the CACNA1D gene encoding the pore-forming Ca_v1.3 α₁ subunit (α₁1.3) (Baig et al. 2011). Ca_v1.3 channels are subject to diverse forms of regulation, which can strongly impact neuronal and cardiac signalling (Mangoni et al. 2003; Olson et al. 2005; Hetzenauer et al. 2006; Zhang et al. 2006; Chan et al. 2007; Navedo et al. 2007). Therefore, characterization of the factors that modulate Ca_v1.3 channels in IHCs is essential for understanding the dynamics of presynaptic Ca²⁺ signals and sound transmission by IHCs.

Like other Ca_v1 channels, Ca_v1.3 can interact directly with various proteins (Calin-Jageman & Lee, 2008). The distal C-terminus (dCT) of the Ca_v1.3 α₁1.3 contains a consensus site for binding to PDZ (PSD-95 (postsynaptic density-95)/Discs large/ZO-1 (zona occludens-1)) domains (Songyang et al. 1997). Interactions with PDZ-domain-containing proteins affect the localization and function of Ca_v1.3 in neurons (Olson et al. 2005; Zhang et al. 2005, 2006). One such protein, erbin, binds to the α₁1.3 dCT and potentiates Ca_v1.3 currents in response to depolarizing stimuli through a process known as voltage-dependent facilitation (VDF; Calin-Jageman et al. 2007). Densin-180 also interacts with the α₁1.3 dCT but does not enhance Ca_v1.3 VDF. Rather, densin-180 tethers calmodulin-dependent protein kinase II to the Ca_v1.3 channel complex, which mediates Ca²⁺-dependent facilitation of Ca_v1.3 currents in response to high-frequency repetitive stimuli (Jenkins et al. 2010).

Like erbin and densin-180, harmonin is a PDZ-domain-containing protein expressed in the brain, but is additionally localized in IHCs (Verpy et al. 2000; Reiners et al. 2005). The gene encoding harmonin corresponds to the USH1C locus for Usher Type 1 syndrome (Verpy et al. 2000), an autosomal recessive sensory disorder characterized by deafness, vestibular dysfunction, and late-onset retinitis pigmentosa (Kimberling & Moller, 1995). Harmonin is concentrated in the apical hair bundles of cochlear and vestibular hair cells (Adato et al. 2005), where it interacts with multiple proteins and regulates mechanotransduction channels that convert mechanical stimuli into changes in hair cell membrane potential (Grillet et al. 2009; Michalski et al. 2009). In mature IHCs, harmonin is also localized to a subset of ribbon-type active zones. Harmonin binds to the α₁1.3 dCT, which enhances proteosomal degradation of Ca_v1.3 and controls Ca_v1.3 channel density at mouse IHC synapses (Gregory et al. 2011). It is not known whether, like erbin and densin-180, harmonin has other modulatory actions on Ca_v1.3 that could impact presynaptic function in IHCs. In addition, given the strong localization of harmonin at IHC synapses, harmonin may also play a role in glutamate exocytosis, which has also not been investigated.

To address these open questions, we tested if harmonin influenced additional properties of Ca_v1.3, and probed the impact of harmonin on exocytosis in IHCs. We found that like erbin, harmonin enhances Ca_v1.3 VDF, which depends on the interaction of harmonin with the α₁1.3 dCT and the identity of the Ca_vβ subunit. Moreover, we established that harmonin regulates Ca_v1.3 VDF and exocytosis in mouse IHCs. Our results highlight new roles for harmonin as a regulator of presynaptic function in IHCs.

Methods

Ethical approval

All procedures involving mice were approved by the Institutional Animal Care and Use Committee at the University of Iowa in accordance with National Institutes of Health guidelines and animal welfare guidelines at the University of Göttingen and the State of Lower Saxony. After mice were killed by decapitation (for mice less than 10 days old) or isoflurane overdose and/or decapitation (for mice greater than 10 days old), the skull was opened, and the cochlea was removed and opened at the apex so that the apical coil could be harvested for electrophysiological experiments.

Constructs and molecular biology

The following Ca_v subunit cDNAs were used: α₁1.3 containing exon 42 (GenBank no. AF370009 and AF370010 for additional sequence encoded by exon 42), β_1b (GenBank no. NM017346), α₁1.2 (GenBank no. M67515), β_2A (GenBank no. NM053851), and α₂δ-1 (GenBank no. M21948). Expression constructs for FLAG-tagged α₁1.3, α₁1.3_L-A, α₁1.3_exon42A, and green fluorescent protein (GFP)- and myc-tagged harmonin, dfcr mutant were previously described (Calin-Jageman et al. 2007; Gregory et al. 2011).

Cell culture and transfection

Human embryonic kidney cells transformed with SV40 T-antigen (HEK293T) were maintained in Dulbecco's modified Eagle's medium with 10% fetal bovine serum (Life Technologies, Grand Island, NY, USA) at 37°C in a humidified atmosphere under 5% CO₂. Cells were grown to ∼60–90% confluence in 100 mm plates and transfected using GenePorter Reagent (Gene Therapy Systems, San Diego, CA, USA). For immunoprecipitation experiments, HEK293T cells were transfected with cDNAs encoding Ca_v1.3 (FLAG-α₁1.3, FLAG-α₁1.3_exon42A, or FLAG-α₁1.3_L-A (6 μg), β_1b and α₂δ (2 μg each)) and myc-harmonin (4 μg). For electrophysiological experiments, cells were plated on 35 mm culture dishes and transiently transfected using Fugene transfection reagent (Promega, Fitchburg, WI, USA). A total of ∼3 μg total DNA was transfected: α₁1.3, 1.5 μg; β, 0.8 μg; α₂δ, 0.8 μg; ±GFP-tagged harmonin or dfcr mutant, 0.5 μg; or GFP expression plasmid, 0.01 μg.

Coimmunoprecipitation

HEK293T cells were harvested and lysed 48 h after transfection. Lysates were incubated with ANTI-FLAG M2-Agarose Affinity Gel (all reagents from FLAG Immunoprecipitation Kit, Sigma-Aldrich, St Louis, MO, USA) for 2.5 h, rotating at 4°C. After three washes with wash buffer (provided in the FLAG Immunoprecipitation Kit), proteins were eluted with SDS-containing sample buffer and subjected to SDS-PAGE. Coimmunoprecipitated proteins were detected by Western blotting with antibodies against α₁1.3 (Ab144; Gregory et al. 2011) or myc epitopes (Sigma-Aldrich). Following incubation with appropriate secondary antibodies, WesternC reagent (Bio-rad, Hercules, CA, USA) was utilized for development and the Geldoc Imager for image collection (Bio-rad). Quantification was performed densitometrically with Quantity One software (Bio-rad). To obtain the fraction of harmonin that coimmunoprecipited with α₁1.3, Western blot signals corresponding to bands for harmonin were divided by those representing the FLAG-immunoprecipitated α₁1.3. For each experiment, these values for harmonin coimmunoprecipitated with α₁1.3_ex42A or α₁1.3_L-A were normalized to that for α₁1.3 to determine the percentage change in harmonin that coimmunoprecipitated with α₁1.3_ex42A or α₁1.3_L-A relative to that for α₁1.3. Data from three independent experiments were averaged (mean ± SEM).

Electrophysiological recordings

Ba²⁺ currents (I_Ba) were recorded 48–72 h after transfection at room temperature using the whole-cell patch clamp electrophysiology technique from transiently transfected HEK293T cells. The internal solution contained (in mm): 140 NMDG, 5 EGTA, 10 Hepes, 2 MgCl₂, 2 Mg-ATP, pH 7.3 (with methanesulfonate) and adjusted to ∼290 mosmol l⁻¹ with glucose. The external solution contained (in mm): 150 Tris, 2 MgCl₂, 10 BaCl₂, pH 7.35 (with methanesulfonate) and adjusted to ∼310 mosmol l⁻¹ with glucose. Pipettes of 3–5 MΩ resistance were used. Voltage clamp recordings were performed with an EPC-9 or EPC-10 amplifier under control of PULSE or Patchmaster software (HEKA Elektronik, Lambrecht, Germany). Currents were filtered at 2 kHz and sampled at 10–20 kHz. A P/4 protocol was used to subtract leak currents.

For whole-cell patch clamp recordings of mouse IHCs, cochlear tissue was dissected from mice (postnatal days (p) 6–8 or p16–18) in Minimum Essential Medium (MEM)/Glutamax-1 (Invitrogen, Gaithersburg, MD, USA) supplemented with 10 mm Hepes at room temperature and kept up to 18 h at 37°C prior to recording. IHCs in the apical cochlear turn were visualized on an upright microscope (BX51WI, Olympus) with a ×40 water-immersion objective with DIC optics. The basolateral membrane of IHCs was patch-clamped with electrodes pulled from thick-walled borosilicate glass capillaries (1B150F, Warner Instruments, Camden, CT, USA). The internal solution contained (in mm): 120 caesium gluconate, 80 CsCl, 0.1 CaCl₂, 4 MgATP, 5 Hepes and 5 EGTA; pH was adjusted to 7.35 with CsOH; osmolarity ∼305 mosmol l⁻¹. External solution contained (in mm): 105 NaCl, 5.8 KCl, 10 CsCl, 55 TEA-Cl, 10 BaCl₂, 1 MgCl, 10 glucose and 10 Hepes supplemented with MEM Vitamins and Amino Acids (Invitrogen, Gaithersburg, MD, USA) at 1X; pH was adjusted to 7.4 with TEA-OH; osmolarity ∼320 mosmol l⁻¹. On the day of recording, 4-aminopyridine (4 mm), apamin (0.3 mm) and TTX (0.5 mm, for p6–8 IHCs) were added to the external solution. Electrode resistances were 3.5–6.2 MΩ in the external solution. Data were acquired with HEKA EPC-9 or EPC-10 amplifiers controlled by Patchmaster software (HEKA Elektronik, Lambrecht, Germany). Leak subtraction was done online with a P/6 protocol. Series resistance was compensated with the patch clamp circuitry (50–70%); average uncompensated series resistance was 12.9 ± 0.6 (n= 89 IHCs). Currents were low-pass filtered at 5 kHz and sampled at 20 kHz except for VDF measurements, where currents were filtered at 2.9 kHz and sampled at 10 kHz. Voltages were not corrected for the liquid junction potential of −7 mV in the external recording solution.

Electrophysiological data were analysed with custom routines in IgorPro software (Wavemetrics, Portland, OR, USA). Average data are expressed as mean ± SEM. Statistical comparisons were done using SigmaPlot software (Systat, Chicago, IL, USA).

Confocal Ca²⁺ imaging

Confocal Ca²⁺ imaging was performed in IHCs from dfcr mice and their wild-type littermates as described previously (Frank et al. 2009; Gregory et al. 2011). In brief, synaptic Ca²⁺ microdomains were identified as hotspots of Fluo-5N fluorescence (low affinity Ca²⁺ indicator) in XY scans using long (200–254 ms) step depolarizations. We then positioned the laser at the peak pixel of each Ca²⁺ microdomain as identified in the XY scan (spot detection) and invoked Ca²⁺ influx by 20 ms step depolarizations to −32 mV. We then applied a 50 ms depolarization to +63 mV preceding the second 20 ms depolarization to −32 mV in order to facilitate the Ca²⁺ influx. The internal solution contained (in mm): 115 caesium glutamate, 13 TEA-Cl, 1 MgCl₂, 1 CaCl₂, 10 EGTA, 2 ATP-Mg, 0.3 GTP-Na, 20 Hepes (pH adjusted with CsOH to 7.2, osmolarity ∼295 mosmol l⁻¹) and 0.4 Fluo-5N (penta-potassium salt; Invitrogen). The external solution contained (in mm): 102 NaCl, 35 TEA-Cl, 2.8 KCl, 5 CaCl₂, 1 MgCl₂, 10 Hepes, 1 CsCl, 11.1 d-glucose (pH adjusted with NaOH to 7.2, osmolarity ∼300 mosmol l⁻¹). Data are presented as mean ± SEM, unless otherwise stated.

Capacitance recordings

For capacitance recordings, IHCs from dfcr mice and their wild-type littermates (p13–19) were subjected to perforated and ruptured patch-clamp recording as described previously (Moser & Beutner, 2000). The internal solution for perforated-patch experiments contained (in mm): 130 caesium gluconate, 10 TEA-Cl, 10 4-aminopyridine, 1 MgCl₂, 10 Hepes (pH adjusted with HCl to 7.17, osmolarity ∼290 mosmol l⁻¹) and 300 μg ml⁻¹ amphotericin B. The internal solution for EGTA experiments contained (in mm): 115 caesium glutamate, 13 TEA-Cl, 1 MgCl₂, 4 EGTA, 2 CaCl₂, 20 Hepes, 2 Mg-ATP, 0.3 Na-GTP (pH adjusted with CsOH to 7.2, osmolarity ∼290 mosmol l⁻¹). The external solution contained (in mm): 100–104 NaCl, 35 TEA-Cl, 2.8 KCl, 10 CaCl₂, 1 MgCl₂, 10 Hepes, 1 caesium gluconate or CsCl, 5 4-aminopyridine, 11.1 d-glucose (pH adjusted with NaOH to 7.2, osmolarity ∼300 mosmol l⁻¹). For most perforated-patch experiments the external solution also contained apamin (0.1 mm). An EPC-9 amplifier controlled by Pulse software (HEKA Elektronik) was used for measurements. All voltages were corrected for liquid junction potentials. Currents were sampled at 20 kHz and low-pass filtered at 2 kHz. Cells that displayed a holding current exceeding −50 pA were discarded from analysis. Ca²⁺ currents were further isolated using a P/n protocol. Series resistance (R_S) measured at the beginning of the perforated- patch recording was 26.3 ± 1.5 MΩ, n= 15, for control, and 26.9 ± 1.0 MΩ, n= 26, for dfcr. For ruptured-patch recordings, R_S was 8.5 ± 0.5 MΩ, n= 10, for control, and 10.4 ± 1.3 MΩ, n= 10, for dfcr. To calculate ΔC_m, the initial 30 ms following the voltage step were ignored due to a non-exocytic capacitance artifact (Neef et al. 2007) and the average C_m measured over at least 40 ms. No R_S compensation was used.

Immunofluorescence labelling of mouse organ of Corti

Double-labelling for CtBP2/RIBEYE and GluA2/3 was performed as previously described (Khimich et al. 2005). The following antibodies were used: mouse IgG1 anti-CtBP2 (BD Biosciences, 1:200), rabbit anti-GluR2/3 (Chemicon, 1:200) and secondary AlexaFluor488- and AlexaFluor568-labelled antibodies (Molecular Probes, 1:200). Confocal images were acquired using a laser scanning confocal microscope (Leica TCS SP5, Leica MicrosystemsCMS GmbH, Mannheim, Germany) with 488 nm (Ar) and 561 nm (DPSS) lasers for excitation and a ×63 oil immersion objective (NA = 1.4–0.7). For 3-D reconstructions of the specimen, z-axis stacks of 2-D images were taken with a step size of 0.5 μm. Image stacks represent maximum z projections, done in ImageJ. The CtBP-2/RIBEYE and GluA2/3 immunofluorescence spots were counted in the z-stacks and divided by the number of IHCs (number of nuclei in the field of view) in order to yield the number of synapses per IHC. Juxtaposed pre- and postsynaptic spots were considered as mature synapses (Khimich et al. 2005).

Results

Harmonin enhances VDF of Ca_v1.3 in HEK293T cells

We have previously shown that cotransfection of harmonin with Ca_v1.3 channels in HEK293T cells increases ubiquitination of α₁1.3 and decreases Ca_v1.3 current density through enhanced proteosomal degradation of α₁1.3 (Gregory et al. 2011). While determining if harmonin affected other parameters of Ca_v1.3 function, we noted that harmonin had a particularly prominent effect of enhancing VDF in HEK293T cells. We measured VDF with a triple-pulse voltage protocol in which test current amplitudes are compared before (P1) and after (P2) a conditioning prepulse to various voltages (Fig. 1A). Ba²⁺ was used as the charge carrier to increase resolution of VDF by limiting the competing effects of Ca²⁺-dependent inactivation (CDI). With this protocol, VDF is evident as an increase in the ratio of the P2:P1 current (F_ratio) with depolarized prepulse voltages. Cotransfection of harmonin with Ca_v1.3 (α₁1.3, β_1b and α₂δ subunits) caused a significant increase in maximal VDF seen with a +50 mV prepulse (F_ratio,+50= 1.18 ± 0.02 for Ca_v1.3 alone vs. 1.52 ± 0.08 for Ca_v1.3 + harmonin; P < 0.01, by Mann–Whitney rank sum test; Fig. 1B). Increased magnitude of I_Ba with harmonin was not secondary to shifts in the conductance–voltage profile, since Boltzmann fits of tail current activation curves revealed similar parameters (V_1/2=−19.6 ± 3.0; k= 7.9 ± 0.8 for Ca_v1.3 alone vs. V_1/2=−18.6 ± 7.5; k= 7.6 ± 1.1 for Ca_v1.3 + harmonin, P= 0.98 for V_1/2 and P= 0.52 for k; Fig. 1C). These results confirm that harmonin increases Ca_v1.3 VDF, which is consistent with its interaction with Ca_v1.3 channels in the plasma membrane.

Figure 1. Harmonin enhances VDF of Ca_v1.3 channels in transfected HEK293T cells.

A, voltage protocol for VDF (top) and representative traces showing I_Ba evoked by 10 ms steps from −90 to −20 mV before (P1) and after (P2) a 20 ms conditioning prepulse (Pre) to +50 mV. B, ratio of P2/P1 current amplitudes (F_ratio) is plotted against prepulse voltage for Ca_v1.3 alone (α₁1.3, β_1b, α₂δ; n= 12) or +harmonin (n= 12). C, normalized tail current–voltage relationships obtained for Ca_v1.3 alone (n= 14) or +harmonin (n= 10). I_Ba was evoked by test pulses from −90 mV to various voltages. Tail currents measured upon repolarization to −70 mV were normalized to the maximal tail current amplitude and plotted against test voltage.

Ca_vβ subunits modulate effects of harmonin on Ca_v1.3 VDF

Although we found that harmonin modulated the VDF of Ca_v1.3 channels containing the auxiliary Ca_vβ_1b subunit (Ca_v1.3(β_1b), Fig. 1), Ca_vβ₂ is the major Ca_vβ subunit contributing to Ca_v1.3 function in mouse IHCs (Neef et al. 2009). Therefore, we characterized the impact of harmonin on VDF in HEK293T cells transfected with Ca_v1.3 subunits containing Ca_vβ_2A (Ca_v1.3(β_2A)). While Ca_v1.3(β_2A) exhibited marginal VDF (F_ratio,+50= 1.17 ± 0.02) that was not significantly different from Ca_v1.3(β_1b) (F_ratio,+50= 1.18 ± 0.02; P= 0.22, by t test; Fig. 2A–C), cotransfection with harmonin caused a smaller increase in VDF of Ca_v1.3(β_2A) than Ca_v1.3(β_1b). While Ca_v1.3(β_1b) VDF was increased 28.5 ± 7.2% by harmonin, this increase was only 10.6 ± 1.6% for Ca_v1.3(β_2A) (P < 0.05, by t test; Fig. 2D). These results confirm the importance of Ca_vβ subunits in modulating responsiveness of Ca_v1.3 VDF to PDZ-domain-containing proteins (Calin-Jageman et al. 2007), and suggest that native Ca_v1.3(β₂) in IHCs may undergo VDF enhancement by harmonin, although to a lesser extent than Ca_v1.3(β_1b).

Figure 2. The extent of Ca_v1.3 VDF due to harmonin depends on the identity of the Ca_vβ subunit.

A and B, voltage protocol, representative I_Ba, and F_ratio were as described in Fig. 1A and B except for channels containing the Ca_vβ_2A subunit. n= 12 cells for Ca_v1.3(β_2A) alone, n= 12 cells for +harmonin. C, comparison of facilitation obtained at +50 mV prepulse voltage (F_ratio,+50) for β_1b- or β_2A-containing channels. *P < 0.001 compared to Ca_v1.3 alone, by t test. Number of cells is indicated in parentheses. D, percentage increase in F_ratio,+50 due to harmonin (%F_harmonin) for channels with β_1b (n= 12 cells) or β_2A (n= 12 cells). %F_harmonin was calculated as (F_ratio,+50 for Ca_v1.3 alone/mean F_ratio,+50 for +harmonin) × 100. *P < 0.05 by t test.

Harmonin binding to α₁1.3 dCT is required for VDF enhancement

To test if increased VDF was due to harmonin binding to the α₁1.3 dCT, we took advantage of a short splice variant of the rat α₁1.3 lacking the dCT in which substitution of exon 42A for exon 42 eliminates much of the C-terminal domain including the PDZ-binding sequence in the dCT (Xu & Lipscombe, 2001). We also used α₁1.3 constructs in which the final leucine residue in the dCT was mutated to alanine (Ca_v1.3_L-A; Fig. 3A). This mutation disrupts the consensus PDZ-binding sequence and prevents binding of harmonin to a C-terminal fragment of α₁1.3 in vitro (Gregory et al. 2011). To verify that Ca_v1.3_42A and Ca_v1.3_L-A have limited interaction with harmonin, we compared their abilities to coimmunoprecipitate with harmonin with that of the wild-type Ca_v1.3 in transfected HEK293T cells. While harmonin still coimmunoprecipitated with both Ca_v1.3_42A and Ca_v1.3_L-A, quantitative analyses revealed a consistent reduction in the amount of harmonin that associated with Ca_v1.3_42A and Ca_v1.3_L-A (66.5 ± 8.8% and 82.1 ± 3.9%, respectively) compared to wild-type Ca_v1.3 (Fig. 3B and C). Evidently, the dCT contributes to, but is not the sole determinant for, harmonin binding to Ca_v1.3. Yet, electrophysiological recordings revealed that Ca_v1.3_42A and Ca_v1.3_L-A underwent VDF that was not changed by cotransfection with harmonin. As we have shown previously, Ca_v1.3_42A VDF was greater than for exon 42-containing channels, since the dCT contains a module that normally inhibits VDF (Calin-Jageman et al. 2007). However, there was no difference in maximal VDF in cells transfected with Ca_v1.3_42A alone (F_ratio,+50= 1.37 ± 0.06) and those cotransfected with harmonin (F_ratio,+50= 1.44 ± 0.08; P= 0.48, by t test; Fig. 4A). Similarly, harmonin did not affect VDF of Ca_v1.3_L-A (F_ratio,+50= 1.14 ± 0.03 for Ca_v1.3_L-A alone vs. 1.18 ± 0.04 for Ca_v1.3_L-A+ harmonin, P= 0.75, by t test; Fig. 4B). These results demonstrate that, despite other potential harmonin interaction sites within the Ca_v1.3 channel complex, harmonin binding to the dCT is required for VDF modulation.

Figure 3. Harmonin binding is reduced by disruption of the PDZ-binding site in the α₁1.3 dCT.

A, schematic diagram of harmonin and α₁1.3. The long C-terminal domain encoded by exon 42 (Ex42) includes the PDZ-binding motif (ITTL) that interacts with the second of three PDZ domains of harmonin. The coiled-coil domain (CC) that is deleted in the dfcr harmonin mutant is indicated. The ‘x’ marks approximate location of truncation of the dCT due to inclusion of exon 42A. B, coimmunoprecipitation of harmonin with Ca_v1.3 channels in transfected HEK293T cells. Cells were transfected with harmonin alone (control) or cotransfected with Ca_v1.3 subunits including α₁1.3 with full-length (Ex42) or truncated (Ex42A) dCT or α₁1.3 with terminal leucine substituted with alanine (L-A). Western blotting detected α₁1.3 (upper panel) and coimmunoprecipitated harmonin (lower panel). C, quantification reflecting percentage change in harmonin coimmunoprecipitated with α₁1.3_Ex42A and α₁1.3_L-A compared to that for α₁1.3_Ex42 (%co-ip harmonin). See Methods for details.

Figure 4. Disruption of the PDZ-binding site in α₁1.3 prevents modulation of VDF by harmonin.

Same as in Fig. 1A and B except for cells transfected with Ca_v1.3_42A alone (n= 10) or +harmonin (n= 6) (A), or Ca_v1.3_L-A alone (n= 12) or +harmonin (n= 9) (B).

The dfcr harmonin mutant does not regulate Ca_v1.3 VDF

‘Deaf-circler’ (dfcr) mice harbour a mutation in the gene encoding harmonin which deletes 132 amino acids between PDZ domains 2 and 3. Similar to humans with Usher syndrome, dfcr mice are deaf and exhibit vestibular defects (Johnson et al. 2003). The dfcr mutant form of harmonin still contains the PDZ domain 2, which interacts with the α₁1.3 dCT, but the internal deletion removes the coiled-coil domain, which disrupts binding of PDZ ligands, including α₁1.3 (Gregory et al. 2011). The dfcr mutant does not bind to the α₁1.3 dCT in vitro and cannot modulate Ca_v1.3 current density like the wild-type harmonin in transfected cells and in mouse IHCs (Gregory et al. 2011). Since the effects of the dfcr mutant on Ca_v1.3 VDF have not been characterized, we first compared the effects of co-transfecting wild-type or dfcr harmonin with Ca_v1.3 in HEK293T cells.

Consistent with the importance of harmonin interactions with the α₁1.3 dCT for modulation of Ca_v1.3 VDF (Figs 1–4), dfcr harmonin did not augment VDF of Ca_v1.3(β_2A) (F_ratio,+50= 1.15 ± 0.02 for Ca_v1.3(β_2A) alone vs. 1.14 ± 0.02 for Ca_v1.3(β_2A) +dfcr; P= 0.99, by t test; Fig. 5A) or Ca_v1.3(β_1b) (F_ratio,+50= 1.21 ± 0.02 for Ca_v1.3(β_1b) alone vs. 1.27 ± 0.04 for Ca_v1.3(β_1b) +dfcr; P= 0.18, by t test; data not shown). To compare a second metric for VDF, we measured the activation kinetics of I_Ba evoked before and after the conditioning prepulse. Since VDF involves enhanced channel opening in response to depolarization, VDF should manifest as faster activation of the P2 current relative to the P1. To test this, we obtained the time constants (τ) for activation with exponential fits of I_Ba for P1 and P2, such that enhanced VDF due to harmonin should be measurable as a larger difference in τ for P1 and P2 compared to the dfcr mutant (Δτ, Fig. 5B). While P2 currents activated significantly faster (∼36%) than P1 currents in cells cotransfected with harmonin, no such difference was observed with the dfcr mutant (Fig. 5B). VDF by this metric was significantly greater in cells cotransfected with harmonin than with dfcr mutant (∼70%; Fig. 5B), which further confirms the reduced ability of the dfcr mutant to modulate Ca_v1.3.

Figure 5. The dfcr mutant of harmonin does not enhance Ca_v1.3 VDF.

A, same as in Fig. 1A and B except for cells transfected with Ca_v1.3(β_2A) alone (n= 12) or +dfcr (n= 5). Dashed line in graph represents data redrawn from Fig. 2B for Ca_v1.3(β_2A) + harmonin. B: top, representative current traces for P1 and P2 overlaid for comparison; bottom, time constants measured from exponential fits of P1 or P2 current traces for Ca_v1.3(β_2A) + harmonin or Ca_v1.3(β_2A) +dfcr. Δτ represents difference in τ for P1 and P2 currents. P values for τ were determined by t test and for Δτ from paired t test.

Neither wild-type nor dfcr harmonin modulate CDI

To determine if harmonin regulates other Ca_v1.3 properties in transfected HEK293T cells, we measured the effects of harmonin on inactivation. Like other Ca_v channels, Ca_v1.3 undergoes inactivation due to Ca²⁺- or voltage-dependent mechanisms (CDI or VDI, respectively). CDI is due to calmodulin, which senses local Ca²⁺ influx due to its direct association with the channel (reviewed in Christel & Lee, 2012). Since Ba²⁺ ions bind poorly to calmodulin (Wang, 1985), I_Ba exhibits primarily VDI. Depending on the identity of the auxiliary β subunit, I_Ca can show both CDI and VDI such that CDI can be isolated as the difference in inactivation of I_Ca and I_Ba. With 300 ms step-depolarizations, we measured inactivation as the ratio of the residual current amplitude at the end of the pulse (Ires) and the peak current (Ipk) amplitude (I_res/I_pk), and CDI as the difference in I_res/I_pk for I_Ca and I_Ba (Fig. 6A). As expected, I_Ca inactivated significantly faster than I_Ba in cells transfected with Ca_v1.3 alone or cotransfected with harmonin or the dfcr mutant. However, there was no difference in CDI between the three groups (Fig. 6A). Since our metric for CDI may not have detected effects of harmonin or dfcr on the kinetics of CDI, we compared parameters from double exponential fits of the I_Ca. There was no significant difference in the fractional contribution or time constants for fast or slow inactivation (Fig. 6B). These results indicate that harmonin does not affect CDI in transfected HEK293T cells.

Figure 6. Ca_v1.3 CDI is not affected by harmonin or the dfcr mutant.

A, top, representative traces for I_Ca and I_Ba normalized and overlaid for comparison in cells transfected with Ca_v1.3(β_2A) alone or cotransfected with harmonin or dfcr. Currents were evoked by 300 ms pulses to −20 mV from −90 mV. Bottom, inactivation was calculated as the current amplitude at the end of the pulse normalized to the peak current amplitude (I_res/I_pk) and is shown for I_Ca and I_Ba. CDI represents the difference in I_res/I_pk for I_Ca and the mean I_res/I_pk for I_Ba. For Ca_v1.3(β_2A) alone, n= 5 for I_Ca, n= 10 for I_Ba; for +harmonin, n= 7 for I_Ca, n= 11 for I_Ba; for +dfcr, n= 9 for I_Ca, n= 9 for I_Ba. *P < 0.001 by t test. B, parameters obtained from double exponential fits of current traces obtained as in A.

Impaired VDF in a mouse model of Usher syndrome

To test the physiological relevance of Ca_v1.3 VDF modulation by harmonin, we performed whole-cell patch clamp recordings of IHCs from mice expressing the dfcr mutant. We showed previously that Ca_v1.3 current density is abnormally elevated in IHCs from mature dfcr mice, consistent with decreased proteosomal degradation of Ca_v1.3 (Gregory et al. 2011). If harmonin also enhances VDF of Ca_v1.3, we would expect VDF to be reduced in IHCs from dfcr mice. Since the presynaptic localization of harmonin and functional interactions of harmonin with Ca_v1.3 are characteristic of IHCs from mice at ages (older than p12) after hearing onset (Gregory et al. 2011), modulation of Ca_v1.3 VDF would be expected in mouse IHCs at p16–18 but not p6–8.

To test these predictions, we compared Ca_v1.3 VDF in IHCs from heterozygous control and homozygous mutant dfcr mice. Since Ca_v1.3 accounts for ∼90% of the IHC whole-cell Ca²⁺ current (Platzer et al. 2000; Brandt et al. 2003), recording solutions were designed to isolate Ca_v1.3 I_Ba from other ionic currents without the addition of Ca_v channel blockers. To determine the optimal conditions for measuring VDF in IHCs, we measured the onset of VDF by comparing test currents evoked before (P1) and after (P2) a conditioning prepulse of varying durations. VDF increased exponentially with prepulse duration with a time constant of ∼11 ms (Fig. 7A). Since maximal VDF was obtained with a 50 ms prepulse duration, we used 50 ms prepulses for the comparisons of VDF in control and dfcr IHCs. Consistent with our findings in HEK293T cells, I_Ba exhibited modest VDF in mature (p16–18) control IHCs (F_ratio,+50= 1.15 ± 0.01) that was significantly weaker in IHCs from dfcr mice (F_ratio,+50= 1.07 ± 0.01, P < 001, by t test; Fig. 7B). This difference in VDF was not observed in immature IHCs (p6–8; F_ratio,+50= 1.08 ± 0.01 for control vs. 1.07 ± 0.01 for dfcr, P= 0.59, by t test; Fig. 7C). VDF in mature IHCs was associated with significantly faster activation kinetics of I_Ba evoked after (P2) compared to before (P1) the prepulse (in control IHCs, τ_P1= 0.60 ± 0.04 ms vs.τ_P2= 0.37 ± 0.02 ms for +50 mV prepulse; P < 0.001, by paired t test; Fig. 8A and B). By this metric, VDF was still evident in dfcr IHCs (τ_P1= 0.66 ± 0.05 ms vs.τ_P2= 0.41 ± 0.03 ms for +50 mV prepulse; P < 0.001, by paired t test; Fig. 8C). However, in dfcr IHCs, the prepulse-induced acceleration of I_Ba activation was 30–68% weaker than in control IHCs (Δτ= 0.11 ± 0.02 ms for control vs. 0.05 ± 0.01 ms for dfcr, P= 0.02 by Mann–Whitney rank sum test; Fig. 8D). Reduced VDF in mature dfcr IHCs was not secondary to differences in control and dfcr IHCs in terms of CDI or VDI (Supplemental Fig. 1, available online only), or voltage-dependent activation of I_Ba (Table 1). Therefore, in addition to regulating Ca_v1.3 channel density in the plasma membrane, harmonin also potentiates Ca_v1.3 VDF in mature mouse IHCs.

Figure 7. VDF is weaker in dfcr compared to control IHCs after hearing onset in mice.

A, measurement of VDF onset. Left, voltage protocol and representative current traces obtained with indicated prepulse durations. P1 and P2 currents are overlaid for comparison. Right, P2/P1 ratios were expressed as percentage of the maximal VDF obtained with a 165 ms prepulse and plotted against prepulse duration. Smooth line represents single-exponential fit. The mean time constant (±SEM) is indicated. B and C, top, voltage protocol for IHC VDF showing 20 ms test pulses (P1, P2) from −75 to −15 mV separated by a 50 ms prepulse to various voltages. Representative traces for I_Ba obtained with +50 mV prepulse for IHCs from postnatal days (p) 16–18 (B) or p6–8 (C) control (n= 25 for p16–18, n= 18 for p6–8) or dfcr (n= 26 for p16–18, n= 18 for p6–8) mice. Dashed line represents initial amplitude of P1 current. Bottom, F_ratio was calculated as P2 divided by P1 current amplitude and plotted against prepulse voltage.

Figure 8. VDF accelerates I_Ba activation to a greater extent in control than in dfcr IHCs.

A, voltage protocol and representative traces for P1 and P2 currents overlaid for comparison of activation kinetics. Results are shown from IHCs from p16–18 control or dfcr mice. B and C, time constant (τ) from exponential fit of the rising phase of P1 and P2 I_Ba traces as shown in A from control (n= 37, B) and dfcr (n= 30, C) IHCs. Symbols represent values from individual cells, bars represent mean ± SEM. D, acceleration in activation due to VDF measured as the difference in τ for P1 and P2 (Δτ) measured as in B for control and dfcr IHCs. P value determined by t test.

Table 1.

I–V parameters for I_Ca and I_Ba in control and dfcr IHCs

IBa (whole-cell)	Control (n= 43)	dfcr (n= 41)	P value
G (nS)	7.5 ± 0.5	7.0 ± 0.5	0.49
V1/2 (mV)	−18.2 ± 0.6	−19.1 ± 0.7	0.30
E (mV)	48.9 ± 1.0	49.1 ± 1.0	0.91
k	6.4 ± 0.2	6.3 ± 0.2	0.57

ICa (whole-cell)	Control (n= 11)	dfcr (n= 13)	P value
G (nS)	5.41 ± 0.19	5.86 ± 0.37	0.30
V1/2 (mV)	−21.4 ± 0.8	−26.1± 1.0	0.002
E (mV)	43.4 ± 0.8	43.3 ± 0.6	0.94
k	6.7 ± 0.3	7.3 ± 0.3	0.26

ICa (perforated- patch)	Control (n= 22)	dfcr (n= 40)	P value
G (nS)	5.91 ± 0.25	5.98 ± 0.28	0.87
V1/2 (mV)	−24.3 ± 0.7	−27.9 ± 0.5	<0.001
E (mV)	49.6 ± 1.12	47.8 ± 0.8	0.20
k	4.7 ± 0.2	4.6 ± 0.2	0.71

I_Ba and I_Ca were recorded in whole-cell (ruptured patch) or perforated-patch configuration and analysed as described in Methods. I-V relationships were fit with the following equation: where G is conductance, V is test potential, E is apparent reversal potential, V_1/2 is potential of half activation, k is the slope factor. Values represent mean ± SEM. n represents number of IHCs analysed. P values were obtained from unpaired t tests or Wilcoxon rank test.

We next evaluated modulation of VDF by harmonin using Ca²⁺ as the charge carrier. We combined whole-cell patch-clamp recordings of total I_Ca in the plasma membrane and fast confocal Ca²⁺ imaging of individual IHC active zones (Frank et al. 2009; Gregory et al. 2011). With this approach, we found a small but significant VDF of the P2 relative to the P1 peak I_Ca and charge integral (Table 2) in control and dfcr IHCs, without any significant difference between the genotypes. As shown previously (Gregory et al. 2011), the amplitude of presynaptic Ca²⁺ microdomains was significantly elevated in dfcr compared to control IHCs (baseline-normalized fluorescence change ΔF/F₀, Table 2). While the ΔF/F₀ amplitude did not increase following the conditioning pulse, the Ca²⁺ microdomains build-up tended to be faster in response to P2 compared to the P1 test pulse indicating the presence of modest VDF. The difference in kinetics of P1 and P2 signals did not reach statistical significance, perhaps related to the slower time course of the active zone Ca²⁺ signals (τ_P1∼2 ms) compared to whole-cell measurements of I_Ba (τ_P1∼0.6 ms). While multiple factors could hinder detection of differences in VDF of whole-cell I_Ca and synaptic Ca²⁺ signals in control and dfcr IHCs (see Discussion), these results demonstrate that VDF is a feature of Ca_v1.3 channels in IHCs using Ca²⁺ as the permeant ion.

Table 2.

VDF probed by whole-cell patch clamp recording and Ca²⁺ imaging at active zones

	Control (n= 41 spots, 14 IHCs)	dfcr (n= 41 spots, 16 IHCs)	P value
VDF (ICa)	2.9 ± 1.1%	3.2 ± 0.7%	0.79 (ut)
ICa (P1) vs. ICa (P2)	P= 0.003 (pt)	P= 0.0004 (pt)
VDF (QCa)	1.2 ± 0.6%	2.0 ± 0.6%	0.16 (w)
QCa (P1) vs. QCa (P2)	P= 0.045 (pt)	P= 0.002 (pt)
ΔF/F0 (P1)	0.64 ± 0.36	0.98 ± 0.84	0.02 (w)
ΔF/F0 (P2)	0.62 ± 0.34	0.95 ± 0.79	0.01 (w)
τΔF, P1	1.6 ± 0.5 ms	1.8 ± 0.6 ms	0.40 (w)
τΔF, P2	1.5 ± 0.4 ms	1.7 ± 0.5 ms	0.13 (w)

Whole-cell Ca²⁺ currents (I_Ca) were elicited by 20 ms test pulses (to −32 mV) before (P1) and after (P2) a 50 ms prepulse (to +63 mV) in control and dfcr IHCs. VDF of I_Ca was calculated as the percentage increase in the peak amplitude (VDF (I_Ca)) or the current integral (VDF (Q_Ca)) of I_Ca evoked by the P2 pulse compared to the P1 pulse. Data represent mean ± SEM. Ca²⁺ influx at individual active zones of these cells was approximated by the change in fluorescence of the low affinity Ca²⁺ indicator Fluo-5N (background-subtracted and normalized; ΔF/F₀). Time constants corresponding to the rise in Ca²⁺ derived from single exponential fitting to the depolarization-evoked ΔF (τ_{ΔF, P1} for P1; τ_{ΔF, P2} for P2)_. Data represent mean ± SD. [Ca²⁺]_i was 5 mm in these experiments to better resolve ΔF/F₀. The present analysis was performed on data extracted from the same recordings performed in a previous study in which effects of the dfcr mutation on only the P1 (control) current were reported (Gregory et al. 2011). Data were tested for randomness, normal distribution and equality of variances, then appropriate statistical tests were chosen (pt, paired t test; ut, unpaired t test; w, Wilcoxon rank test).

Impaired exocytosis in dfcr IHCs

Ca_v1.3 mediates stimulus–secretion coupling in IHCs (Platzer et al. 2000; Brandt et al. 2003), whereby exocytosis of a synaptic vesicle may be controlled by few Ca_v1.3 channels within nanometer proximity (Brandt et al. 2005). Compared to wild-type harmonin, the dfcr mutant causes abnormally high Ca_V1.3 current density (Gregory et al. 2011) and reduced VDF (Figs 7 and 8), which could have dual, and potentially complex, effects on IHC exocytosis. To gain insight into the functional consequences of the dfcr mutation on IHC exocytosis, we performed perforated-patch (Fig. 9A–D) and ruptured-patch (Fig. 9E–H) recordings of membrane capacitance changes (ΔC_m) evoked by step depolarizations. The test voltage evoking the maximal I_Ca was determined in current–voltage (I–V) relations (Fig. 9A and E). Interestingly, Boltzmann fits of the I–V curves for I_Ca revealed a small but significant negative shift in the half-maximal activation voltage for dfcr compared to control IHCs both in perforated-patch and ruptured-patch recordings, which was not observed for I_Ba (Table 1). The increased Ca_v1.3 current density in dfcr IHCs was less apparent than reported previously (Gregory et al. 2011), perhaps due to the larger age range of mice in these experiments (p13–20 vs. p16–18). With a 20 ms step to elicit maximal I_Ca in each cell, ΔC_m was significantly reduced in dfcr compared to control IHCs (Fig. 9B).

Figure 9. Presynaptic function is altered in dfcr IHCs.

I_Ca and exocytic membrane capacitance changes (ΔC_m) were measured in perforated-patch (A–D) and ruptured-patch configurations (4 mm EGTA + 2 mm Ca²⁺ in the pipette, E–H). The extracellular Ca²⁺ was 10 mm to adequately resolve ΔC_m. A, I–V relationship for I_Ca evoked by 10 ms voltage steps from −96.6 mV in perforated-patch recordings (control: n= 23, dfcr: n= 40). B, representative I_Ca and ΔC_m (C_m filtered at 50 Hz) in response to 20 ms depolarization to voltage eliciting peak I_Ca. C, ΔC_m (top) and corresponding I_Ca integral (Q_Ca, bottom) in response to step pulses for the indicated durations. Depolarizations were from −96.6 mV to the voltage eliciting peak I_Ca (between −26.6 and −16.6 mV) for each cell. Data represent grand averages (calculated from the means of the individual cells, n= 26 for dfcr and 15 for control IHCs (p13–19) ± SEM. D, reduced Ca²⁺ efficiency of synchronous exocytosis in dfcr IHCs in response to depolarizations eliciting peak I_Ca in perforated- patch recordings. ΔC_m was normalized to Q_Ca obtained with different stimulus durations in B and C and shown for control and dfcr IHCs (***P < 0.01, by Student's t test). E, I–V relationship for I_Ca evoked by 10 ms voltage steps from −96.2 mV (control: n= 11, dfcr: n= 13) in ruptured-patch recordings. F and G, same as in B and C, but for data obtained with ruptured-patch configuration with 4 mm EGTA and 2 mm Ca²⁺ in the intracellular solution. In F, n= 10 for dfcr and 10 for control IHCs. G and H, tendency towards reduced Ca²⁺ efficiency of synchronous exocytosis was also observed in EGTA-loaded dfcr IHCs. H, same as in D, but for ruptured-patch configuration (n= 10 for control, n= 10 for dfcr).

Previous work has demonstrated the existence of distinct populations of exocytic vesicles that differ in release kinetics and Ca²⁺ dependence. A readily releasable pool (RRP) of vesicles undergoes fast, synchronous exocytosis and is efficiently recruited by short (10–20 ms) stimuli. A slower phase of exocytosis is observed during sustained depolarizations (Moser & Beutner, 2000; Goutman & Glowatzki, 2007). To further characterize the exocytic defect in dfcr IHCs, we compared ΔC_m evoked by varying stimulus durations. With short (10–20 ms) depolarizing pulses, ΔC_m was significantly reduced in dfcr compared to control IHCs (∼41%, P < 0.007 for 10 ms pulses, ∼36%, P= 0.011 for 20 ms pulses; Fig. 9C), while there was no difference in ΔC_m evoked by longer (50–100 ms) depolarizations (P= 0.85 for 50 ms, P= 0.49 for 100 ms by Wilcoxon rank test; Fig. 9C). These results suggested impairment in synchronous exocytosis of the RRP in dfcr IHCs. Harmonin-dependent VDF may enhance rapid Ca²⁺ signalling at active zones in control IHCs. If so, then the difference in ΔC_m in dfcr and control IHCs should be eliminated upon normalizing ΔC_m to the charge integral of I_Ca (ΔC_m/Q_Ca). By this analysis, maximal efficiency of exocytosis was ∼3 fF pC⁻¹ (Fig. 9D), similar to that reported in frog auditory hair cells (Graydon et al. 2011). ΔC_m/Q_Ca was still significantly smaller in dfcr compared to control IHCs (Fig. 9D), which suggested a reduced efficiency of Ca²⁺ influx for driving exocytosis in dfcr compared to control IHCs.

To follow up this possibility, we performed whole-cell patch clamp measurements of ΔC_m including EGTA (4 mm+ 2 mm Ca²⁺, calculated free intracellular [Ca²⁺]= 106 nm) in the intracellular recording solution. As a slow Ca²⁺ buffer, EGTA affects sustained exocytosis more strongly than RRP exocytosis, which probably reflects a requirement for long-distance Ca²⁺ signalling involved in replenishment of depleted vesicles (Moser & Beutner, 2000). If there was looser spatial coupling of Ca_v1.3 channels and RRP vesicles in dfcr compared to control IHCs, we would expect a stronger effect of EGTA on synchronous exocytosis in dfcr IHCs than in control IHCs. However, with EGTA in ruptured-patch recordings, we found RRP exocytosis of dfcr IHCs to be more similar to control IHCs (Fig. 9G) than in perforated-patch recordings. ΔC_m/Q_Ca still tended to be smaller in dfcr than in control IHCs with EGTA (Fig. 9H), although not quite reaching statistical significance (P= 0.053 comparing dfcr vs. control ΔC_m/Q_Ca response to 20 ms depolarization in ruptured patch). There was no significant difference in dfcr IHCs in ΔC_m/Q_Ca evoked by 20 ms depolarizations in ruptured-patch or perforated-patch recordings (P= 0.12; Fig. 9D and H). These results argue against a looser spatial coupling of the RRP to Ca_v1.3 channels in dfcr IHCs.

To determine if smaller numbers of IHC synapses could contribute to the exocytic impairment in dfcr IHCs, we quantified afferent IHC synapses in stacks of confocal sections from organs of Corti immunolabelled for presynaptic ribbons (CtBP2/RIBEYE) and postsynaptic AMPA glutamate receptors (GluA2/3; Khimich et al. 2005). However, there was no significant difference between control and dfcr IHCs in the cochlear apex (8.5 ± 0.6 for control, n= 85 cells vs. 9.7 ± 0.4 for dfcr, n= 49 cells; P= 0.2 by t test). These results suggest that the exocytic defect in dfcr IHCs does not involve changes in synapse number, but could depend on presynaptic alterations in RRP dynamics at the subset of synapses characterized by harmonin/Ca_v1.3 interactions. Our results indicate a multi-faceted role of harmonin in regulating Ca_v1.3 channel function and RRP exocytosis in mature IHCs, both of which are compromised in the dfcr model of Usher syndrome.

Discussion

Our study provides multiple new insights into the regulation of Ca_v1.3 channels by harmonin. First, harmonin binding to the distal C-terminus of α₁1.3 enhances VDF of Ca_v1.3 current, the extent to which depends on the identity of the Ca_vβ subunit. Second, in mouse IHCs, VDF characterizes Ca_v1.3 currents after, but not before, hearing onset and potentiates presynaptic Ca_v1.3 currents by accelerating Ca_v1.3 activation kinetics. Third, Ca_v1.3 VDF and exocytosis are impaired in IHCs from dfcr mice expressing a mutant form of harmonin. We conclude that harmonin is an important determinant of Ca_v1.3 properties and presynaptic function in mouse IHCs.

Dual regulation of Ca_v1.3 channels by harmonin

Facilitation of Ca_v1 channels occurs by multiple mechanisms that have been studied extensively in the context of nerve and muscle (Dolphin, 1996). VDF has been reported for Ca_v1.3 channels in transformed cell lines (Safa et al. 2001; Calin-Jageman et al. 2007), neonatal mouse outer hair cells (Michna et al. 2003), and mouse sinoatrial nodal cells in the heart (Christel et al. 2012). Our results provide the first evidence that Ca_v1.3 channels in IHCs undergo VDF and that harmonin contributes to this process. The mechanism involves harmonin binding to the PDZ-binding sequence in the α₁1.3 dCT, since mutation or deletion of this motif prevents modulation of VDF (Fig. 4). In addition, the dfcr mutant, which cannot bind the α₁1.3 dCT (Gregory et al. 2011) does not enhance VDF (Fig. 5). Similar to the PDZ protein erbin, binding of harmonin to the α₁1.3 dCT may relieve an autoinhibitory regulation of VDF imposed by the α₁1.3 dCT (Calin-Jageman et al. 2007). However, harmonin binding to the α₁1.3 dCT also promotes ubiquitination of α₁1.3, which limits Ca_v1.3 current density by promoting proteosomal degradation of the channel (Gregory et al. 2011). Harmonin may dually regulate Ca_v1.3 channels by limiting channel trafficking to, or enhancing removal from, the plasma membrane through targeting to proteosomal pathways. Those channels that remain associated with harmonin at the IHC synapse exhibit enhanced VDF due to the α₁1.3 dCT interaction. Our findings that Ca_v1.3 current density is increased (Gregory et al. 2011) and VDF is decreased (Figs 7 and 8) in IHCs from dfcr compared to control mice support the bifunctional role of harmonin with respect to Ca_v1.3 regulation in vivo. Our coimmunoprecipitation experiments suggest that harmonin may interact with sites in the channel complex in addition to the dCT (Fig. 3), raising the possibility that harmonin may regulate yet other aspects of Ca_v1.3 function.

VDF in HEK293T cells cotransfected with Ca_v1.3 and harmonin (Figs 1B and 2B) was stronger and occurred at more negative potentials compared to VDF in control IHCs (Fig. 7B). A contributing factor is that VDF is somewhat limited in Ca_v1.3 channels containing the β₂ subunit (Fig. 2C and D), which is the primary Ca_vβ subunit in mouse IHCs (Neef et al. 2009). In addition, harmonin is found at only ∼50% of mature IHC synapses, whereas Ca_v1.3 is present at every synapse (Brandt et al. 2005; Gregory et al. 2011). Since our whole-cell recordings summate the activity of all IHC Ca_v1.3 channels, the impact of harmonin on VDF would be diluted by the nominal VDF exhibited by channels not associated with harmonin. Considering the proportion of harmonin-positive synapses in mature IHCs (∼50%), the ∼13% increase in I_Ba amplitude due to a +30 mV prepulse in mature IHCs VDF may be twice as large at individual synapses characterized by Ca_v1.3/harmonin interactions and thus roughly similar to what we find for Ca_vβ₂-containing channels in HEK293T cells (∼25% for a +30 mV prepulse; Fig. 2B).

Considering the above argument, our inability to measure harmonin-dependent VDF using Ca²⁺ as the charge carrier, both in electrophysiological recordings of I_Ca and Ca²⁺ imaging of presynaptic active zones is not unexpected (Table 2). Moreover, VDF of I_Ca in IHCs is likely to be further diminished by CDI. While not as prominent for Ca_v1.3 in transfected HEK293T cells as in IHCs (compare Fig. 6 and Supplemental Fig. 1), CDI is still fast enough in IHCs (τ_fast∼5–7 ms, Supplemental Fig. 1) to partially occlude VDF (τ∼11 ms, Fig. 7A) in whole-cell recordings. Given these caveats, we propose that VDF is physiologically relevant for controlling I_Ca at the subset of synapses characterized by Ca_v1.3/harmonin interactions in mature IHCs.

Physiological significance of Ca_v1.3 VDF in IHCs

VDF due to harmonin is measurable in mature IHCs but not in IHCs prior to hearing onset (Fig. 7B), which coincides with the developmental upregulation of harmonin localization at IHC synapses (Gregory et al. 2011). At very positive voltages (+50 mV), VDF causes ∼20% acceleration in the activation rate of the macroscopic Ca_v1.3 current (Fig. 8B), an effect that could be twice as large at harmonin-positive synapses. Consistent with this possibility, VDF due to a +50 mV prepulse caused a ∼36% speeding of I_Ba activation in HEK293T cells cotransfected with Ca_v1.3 and harmonin (Fig. 5B). Facilitation of I_Ca in response to prior depolarization has been reported for I_Ca at rat IHC synapses (Goutman & Glowatzki, 2011; Goutman, 2012) and at retinal bipolar synapses (Cho & von Gersdorff, 2012) and so may be a fundamental form of Ca_v1 regulation at ribbon synapses.

Exocytosis of the RRP is triggered by the opening of relatively few Ca_v1.3 channels (Brandt et al. 2005) such that changes in the microscopic properties of Ca_v1.3 may substantially affect transmitter release. In this respect, modulation of Ca_v1.3 gating by harmonin is consistent with the impairment of synchronous exocytosis in dfcr IHCs (Fig. 9). By amplifying presynaptic Ca_v1.3 influx as a function of membrane depolarization, VDF could also contribute to short-term facilitation of IHC transmission. This form of presynaptic plasticity has been documented in afferent recordings at synapses of frog auditory hair cells (Cho et al. 2011) and rat IHCs, where it manifests as a decrease in synaptic failures and delays following pre-depolarizations (Goutman & Glowatzki, 2011). Ca²⁺ current facilitation in the latter study on immature IHCs was relatively modest and not thought to significantly contribute to the synaptic facilitation, which could be due to the decreased localization of harmonin at immature IHC synapses (Gregory et al., 2011). Thus, at select harmonin-positive IHC synapses, Ca_v1.3 VDF may increase the gain and enhance the timing of release events, thus improving aspects of intensity and temporal coding of sound, respectively. Maintaining Ca_v1.3 channel availability during graded changes in the IHC membrane potential is important for the continuous encoding of sound information (Lewis & Hudspeth, 1983; Moser et al. 2006) as well as for ongoing spontaneous afferent firing (Robertson & Paki, 2002; Sueta et al. 2004). Although Ca_v1.3 channels show less inactivation (both CDI and VDI) in IHCs compared to other cell types (Platzer et al. 2000; Koschak et al. 2001), CDI and VDI are still measurable and can, along with RRP depletion (Moser & Beutner, 2000), lead to significant synaptic depression if unopposed (Cho et al. 2011).

A role for harmonin in synchronous exocytosis

Based on our findings that Ca²⁺ influx–exocytosis coupling is impaired in dfcr IHCs, we expected that intracellular dialysis with EGTA in whole-cell recordings should more strongly inhibit RRP exocytosis in dfcr IHCs compared to control IHCs, which was not the case (Fig. 9). This result could be explained by a role for harmonin in regulating Ca²⁺ coupling to only one component of the RRP, which could not be resolved in our measurements of ΔC_m. While paired pre- and postsynaptic recordings of synaptic transmitter release indicate two components of the RRP (Wu & Borst, 1999; Sakaba & Neher, 2001; Goutman & Glowatzki, 2007; but see Li et al. 2009), only one RRP was reported with ΔC_m measurements of exocytosis (Moser & Beutner, 2000). Thus, our ΔC_m recordings may have lacked the sensitivity to detect a stronger effect of EGTA on RRP exocytosis in dfcr IHCs when compared to control, which would further be challenged by an effect of harmonin at only a subset of IHC active zones (Gregory et al. 2011).

Alterations in exocytosis in dfcr IHCs could suggest additional synaptic functions of harmonin that may be independent of Ca_v1.3 modulation. Analogous to its properties in apical IHC hair bundles (Adato et al. 2005), harmonin could engage in multivalent interactions with Ca_v1.3 channels and/or other synaptic molecules that could regulate Ca²⁺ coupling to exocytosis in IHCs. For example, harmonin, either alone or in complex with cadherin-23, binds to phosphatidylinositol 4,5-bisphosphate (PIP₂) (Bahloul et al. 2010). PIP₂ is a known regulator of exocytosis in other cell types (Koch & Holt, 2012) and reductions in PIP₂ synthesis causes defects in Ca²⁺ signalling and high-frequency hearing-impairment mice (Rodriguez et al. 2012). A detailed understanding of the molecular mechanism by which harmonin regulates the Ca²⁺ efficiency of exocytosis will require more in-depth analyses, such as paired recordings of IHCs and postsynaptic afferents in genetically modified mice.

A presynaptic role for harmonin in IHCs

In apical hair bundles, the role of harmonin has been elegantly elucidated. The PDZ domains of harmonin bind to multiple proteins (sans, myosin VIIa, cadherin-23) implicated in hair bundle development (Adato et al. 2005). Mutations in harmonin or these interacting proteins cause deafness in humans with Usher syndrome and animal models (Petit, 2001), due to improper formation of hair bundles and subsequent failure of mechanotransduction. In dfcr mice, hair bundles of IHCs are grossly normal (Grillet et al. 2009), but mechanotransduction currents in dfcr outer hair cells show weaker sensitivity and slower kinetics in response to physical displacement of hair bundles, due to the inability of dfcr harmonin to interact with proteins required for gating mechanotransduction channels (Grillet et al. 2009). These results provide an intriguing parallel to our findings that Ca_v1.3 properties and exocytosis are altered in dfcr IHCs. Other Usher syndrome-associated proteins have also been found to interact with and regulate ion channels. For example, the USH2D protein whirlin associates with Ca_v1.3 in photoreceptors (Kersten et al. 2010). The Drosophila homologue of whirlin, dysc, interacts with SLO Ca²⁺-activated K⁺ channels and enhances the expression of these channels in neurons (Jepson et al. 2012). Understanding how members of the Usher interactome collectively or individually alter membrane excitability and/or synaptic transmission may provide new clues into the cellular pathology leading to deafness and blindness in human patients.

Glossary

CDI

Ca²⁺-dependent inactivation

dCT

distal C-terminus

IHC

inner hair cell

p

postnatal day

RRP

readily releasable pool

VDF

voltage-dependent facilitation

VDI

voltage-dependent inactivation

Additional information

Competing interests

None.

Author contributions

The experiments in this study were performed at the University of Iowa, Dominican University and University of Göttingen. Author contributions are as follows: conception and design of experiments, collection analysis, and interpretation of data: F.D.G., I.E.C.-J., T.P., T.M., A.L.; drafting of the article or revising it critically for important intellectual content: F.D.G., T.M., A.L. All authors approved the final version of the manuscript.

Funding

This work was supported by the NIH (DC009433, HL087120 and DC010362 to A.L.; DA015040 and K12/GM00068 to F.D.G.; and DC008417 to I.E.C.-J.), the Carver Research Program of Excellence (to A.L.), a fellowship of the Alexander von Humboldt foundation to T.P., and the German Research Foundation through the Collaborative Research Center 889 (to T.M.).

Supplementary material

Supplemental Fig. 1