Abstract
The auxiliary subunit α2δ2 modulates the abundance and function of voltage-gated calcium channels. Here we show that α2δ2 mRNA is expressed in neonatal and mature hair cells. A functional α2δ2-null mouse, the ducky mouse (du), showed elevated auditory brainstem response click and frequency-dependent hearing thresholds. Otoacoustic emissions were not impaired pointing to normal outer hair cell function. Peak Ca2+ and Ba2+ currents of mature du/du inner hair cells (IHCs) were reduced by 30–40%, respectively, and gating properties, such as the voltage of half-maximum activation and voltage sensitivity, were altered, indicating that Cav1.3 channels normally coassemble with α2δ2 at IHC presynapses. The reduction of depolarization-evoked exocytosis in du/du IHCs reflected their reduced Ca2+ currents. Ca2+- and voltage-dependent K+ (BK) currents and the expression of the pore-forming BKα protein were normal. Cav1.3 and Cavβ2 protein expression was unchanged in du/du IHCs, forming clusters at presynaptic ribbons. However, the close apposition of presynaptic Cav1.3 clusters with postsynaptic glutamate receptor GluA4 and PSD-95 clusters was significantly impaired in du/du mice. This implies that, in addition to controlling the expression and gating properties of Cav1.3 channels, the largely extracellularly localized α2δ2 subunit moreover plays a so far unknown role in mediating trans-synaptic alignment of presynaptic Ca2+ channels and postsynaptic AMPA receptors.
SIGNIFICANCE STATEMENT Inner hair cells possess calcium channels that are essential for transmitting sound information into synaptic transmitter release. Voltage-gated calcium channels can coassemble with auxiliary subunit α2δ isoforms 1–4. We found that hair cells of the mouse express the auxiliary subunit α2δ2, which is needed for normal hearing thresholds. Using a mouse model with a mutant, nonfunctional α2δ2 protein, we showed that the α2δ2 protein is necessary for normal calcium currents and exocytosis in inner hair cells. Unexpectedly, the α2δ2 protein is moreover required for the optimal spatial alignment of presynaptic calcium channels and postsynaptic glutamate receptor proteins across the synaptic cleft. This suggests that α2δ2 plays a novel role in organizing the synapse.
Introduction
Voltage-gated calcium channels (VGCCs) are protein complexes composed of a main, pore-forming α1 subunit and auxiliary α2δ and β subunits. The traditional view is that auxiliary subunits modulate biophysical properties of VGCCs, which are mostly determined by the α1 subunit, and assist in the trafficking and proper surface expression of the channel complex (Catterall, 2000; Dolphin, 2012a, 2013). Recent evidence suggests an additional role of α2δ subunits in synaptogenesis (Eroglu et al., 2009; Kurshan et al., 2009; Dolphin, 2012a; Geisler et al., 2015) and for synaptic morphology (Pirone et al., 2014). α2δ subunits, encoded by one of the four genes CACNA2D1–4, consist of a large extracellular glycosylated α2 peptide linked to a small membrane-anchored δ peptide (Davies et al., 2007; Dolphin, 2012a, 2013). Coexpressing any of the isoforms α2δ1–3 with various α1 and β subunits results in functional channels with increased current amplitudes and altered channel gating (Davies et al., 2007; Dolphin, 2012a, 2013). In the brain, the isoforms α2δ1, α2δ2, and α2δ3 are widely expressed (Cole et al., 2005; Schlick et al., 2010), whereas α2δ4 is mainly found in the retina (De Sevilla Müller et al., 2013; Knoflach et al., 2013). Genetic deletion of α2δ1 or α2δ3 yielded relatively mild phenotypes (Fuller-Bicer et al., 2009; Neely et al., 2010; Pirone et al., 2014). In contrast, the “ducky” (du) mutation of Cacna2d2 encoding α2δ2 results in a functional α2δ2-null mouse (du/du mouse) with a severe phenotype, including cerebellar ataxia, epilepsy, reduced body weight, and premature death (Barclay et al., 2001; Brodbeck et al., 2002). Cerebellar Purkinje cells of ducky mice exhibit 35% smaller whole-cell Ca2+ currents mediated by P-type (Cav2.1) Ca2+ channels and abnormal morphology of their dendritic trees (Barclay et al., 2001; Brodbeck et al., 2002). The ducky mutation is a genomic rearrangement involving the Cacna2d2 gene, which is disrupted after exon 3 (of 39 exons; Barclay et al., 2001). A short truncated protein is still produced but not targeted correctly (Brodbeck et al., 2002). The characterization of other α2δ2-deficient mouse models revealed phenotypes very similar to that of du/du mice (Brill et al., 2004; Ivanov et al., 2004; Donato et al., 2006), altogether indicating that α2δ2 is indispensable for the Ca2+ channel complex consisting of Cav2.1 and Cavβ4 at inhibitory Purkinje cell presynapses. Recently in humans, a missense and a nonsense mutation in CACNA2D2 were identified, both of which cause early infantile epileptic encephalopathy with severe developmental impairment and intellectual disability (Edvardson et al., 2013; Pippucci et al., 2013).
Inner hair cells (IHCs) transform sound-evoked depolarization into graded release of the transmitter glutamate. They express voltage-gated Ca2+ channels, which almost exclusively consist of the L-type Cav1.3 subunit (Platzer et al., 2000; Brandt et al., 2003; Dou et al., 2004) and are localized predominantly in clusters underneath the synaptic ribbons (Vincent et al., 2014; Wong et al., 2014). The Cav1.3 subunits of IHCs predominantly coassemble with the β subunit Cavβ2 (Neef et al., 2009), with some contribution of Cavβ3 and Cavβ4 (Kuhn et al., 2009). The nature of the third partner of the Ca2+ channel complex, the α2δ subunit, was unknown so far.
Materials and Methods
Animals.
Mice carrying a mutation of the Cacna2d2 gene coding for α2δ2 with genomic rearrangement (Cacna2d2du or ducky allele; Barclay et al., 2001) were obtained from the The Jackson Laboratory (http://jaxmice.jax.org/strain/012889.html). They were backcrossed into C57BL/6N (obtained from Charles River) for five or more generations. Cochleae were dissected after mice had been killed by decapitation with isoflurane anesthesia or after cervical dislocation for exocytosis experiments. Animals were housed with free access to food and water at an average temperature of 22°C and a 12 h light/dark cycle. All experiments were performed in accordance with the European Communities Council Directive (86/609/EEC) and approved by regional board for scientific animal experiments of the Saarland, Germany. Genotyping of du/du mice was adapted from Brodbeck et al. (2002). Mice of either sex were used. Hearing measurements and all electrophysiological recordings were performed on du/du mice and wild-type (wt) littermates. For immunofluorescence experiments, mostly wt mice, but in some cases heterozygous (du/wt) littermates, were used as control (ctrl) animals. Heterozygous animals have a normal life expectancy and do not show any of the phenotypes of homozygous du/du mice (Barclay et al., 2001).
Quantitative real-time PCR.
For quantitative PCR (qPCR) analysis, IHCs and outer hair cells (OHCs) from postnatal day 6 (P6) or P20–P25 mice were selectively harvested with micropipettes under microscope control as described previously (Baig et al., 2011). For reverse transcription (RT), 9 μl of cell lysate was mixed with 2 μl of random primers pd(N)6 (50 μm; Applied Biosystems) and 1 μl of dNTP (deoxynucleotidetriphosphate) mix (10 mm; New England Biolabs), incubated at 65°C for 5 min and stored on ice for 1 min. An RT mix (8 μl; Life Technologies) consisting of 5× reverse transcription buffer, 0.1 m dithiothreitol, 2 U/μl RNAseOUT and 2.5 U/μl reverse transcriptase Superscript III was added, and each tube was incubated at 50°C for 150 min followed by 70°C for 30 min. cDNA was stored at −20°C. The abundance of different α2δ subunit transcripts in IHC and OHC cDNA was assessed by TaqMan qPCR using a standard curve method, as previously described (Schlick et al., 2010). cDNA of pooled hair cells (30–630) was split into 12 samples to simultaneously run qPCR-TaqMan assays for all α2δ subunit genes and the endogenous control gene Hprt1 in duplicate. TaqMan gene expression assays specific for the four α2δ isoforms were designed to span exon–exon boundaries and were purchased from Applied Biosystems. The following assays were used [identified as name (gene symbol), assay ID; Applied Biosystems]: α2δ1 (Cacna2d1), Mm00486607_m1; α2δ2 (Cacna2d2), Mm00457825_m1; α2δ3 (Cacna2d3), Mm00486613_m1; and α2δ4 (Cacna2d4), Mm01190105_m1. The expression of hypoxanthine phosphoribosyl-transferase 1 (Hprt1; Mm00446968_m1) was used as the endogenous control. The qPCR (50 cycles) was performed in duplicate using total cDNA (see above) and the specific TaqMan gene expression assay for each 20 μl reaction in TaqMan Universal PCR Master Mix (Applied Biosystems). Samples without cDNA were used as controls. Analyses were performed using the 7500 Fast System (Applied Biosystems). The Ct values for each gene expression assay were recorded for each individual preparation and the molecule numbers were calculated for each α2δ subunit from their respective standard curve (Schlick et al., 2010). Expression of Hprt1 was used to evaluate the total mRNA abundance and to allow a direct comparison between the expression levels in different preparations.
Hearing measurements.
Auditory brainstem response (ABR) and distortion product otoacoustic emissions (DPOAEs) were recorded in anesthetized mice aged 3–4 weeks as described previously (Rüttiger et al., 2004; Engel et al., 2006). For anesthesia, a mixture of ketamine-hydrochloride [75 mg/kg body weight (b.w.); Ketavet 100, Pharmacia] and xylazine-hydrochloride (5 mg/kg b.w.; Rompun 290, Bayer) was injected intraperitoneally with an injection volume of 5 ml/kg b.w. Anesthesia was maintained by subcutaneous application of one-third of the initial dose, typically in 30 min intervals. Body temperature was maintained with a temperature-controlled heating pad. ABR thresholds were determined with click (100 μs) and pure tone stimuli (3 ms plus 1 ms rise/fall time, 2–45 kHz) with electrodes placed at the ear (positive) and vertex (reference). ABR waveforms for click stimuli were determined 40 dB above threshold for each individual animal. Cubic 2 · f1 − f2 (where f is frequency) DPOAE amplitudes for the two stimulus primaries with frequencies f1 and f2, and f2 = 1.2 · f1, and sound pressure level L1 = 55 dB SPL and L2 = 45 dB SPL for the first and the second primary, respectively, were measured at either 16 kHz as well as in the range between 10 and 18 kHz using 0.5 kHz steps followed by averaging (Schimmang et al., 2003; Hecker et al., 2011). Latencies and peak-to-peak amplitudes of the ABR waveforms were extracted from each individual waveform. Click ABR thresholds, wave I latencies of ABR waveforms, and DPOAE amplitudes were analyzed using the Mann–Whitney U test; and frequency-dependent ABR thresholds were analyzed with a two-way ANOVA with Bonferroni post hoc test.
Electrophysiological recordings.
Acutely dissected organs of Corti of 3-week-old mice were used to record Ca2+/Ba2+ and K+ currents. The bath solution for Ca2+ and K+ currents (B1) contained the following (in mm): 70 lactobionate·NaOH, 83 NaCl, 10 HEPES, 5.8 KCl, 5.6 glucose, 1.3 CaCl2, 0.95 MgCl2, and 0.7 NaH2PO4. The bath solution for Ba2+ currents (B2) contained the following: 72 lactobionate·NaOH, 40 NaCl, 35 TEA, 15 4-AP, 10 BaCl2, 10 HEPES, 5.6 KCl, 5.3 glucose, and 1 MgCl2. Both solutions were adjusted to pH 7.35 and 320 mosmol/kg. For Ca2+ current recordings, the specimen was locally superfused with B3 (in mm), as follows: 72.5 lactobionate·NaOH, 40 NaCl, 35 TEA, 15 4-AP, 10 CaCl2, 10 HEPES, 5.6 KCl, 5.6 glucose, 0.9 MgCl2, 0.1 linopirdine, and 0.0005 apamin, pH 7.35 and 320 mosmol/kg. The pipette solution for Ca2+/Ba2+ currents contained the following (in mm): 110 Cs+ methane sulfonate, 20 CsCl, 10 Na+ phosphocreatine, 5 HEPES, 5 EGTA, 4 MgCl2, 4 Na2ATP, 0.3 GTP, 0.1 CaCl2; for exocytosis measurements EGTA concentration was 1 mm. The pipette solution for K+ currents consisted of the following (in mm): 110 K+ gluconate, 20 KCl, 10 Na+ phosphocreatine, 5 HEPES, 5 EGTA, 4 MgCl2, 4 Na2ATP, 0.1 GTP, and 0.1 CaCl2; and all pipette solutions were adjusted to pH 7.35 and 305 mosmol/kg.
Uncompensated series resistance was corrected by 70–80% for K+ current and membrane capacitance recordings. Linear leak subtraction was performed off-line, and voltages were corrected by subtracting liquid junction potentials (LJPs) of 6 mV (Ca2+ currents), 8 mV (Ba2+ currents), and 10 mV (K+ currents). I–V curves of Ca2+/Ba2+ currents were fitted to a second-order Boltzmann function times Goldman–Hodgkin–Katz driving force to determine parameters of the activation curve, the voltage of half-maximum activation, Vh, and the voltage sensitivity of activation, the slope factor k, according to the following equation:
where I is the current at the time the I–V was calculated (averaged over 7–8 ms after depolarization), Pmax is the maximum permeability, and ν = zFV/(RT), with z being 2, F the Faraday constant, R the universal gas constant, T the absolute temperature, and V the membrane potential. [Ca]i (set at 50 nm) and [Ca]o denote the intracellular and extracellular Ca2+ concentrations, respectively. Fits for Ba2+ as charge carrier were performed accordingly.
The degree of ICa/IBa inactivation 300 ms after peak was determined as follows:
where Ipeak is the maximum of the peak current trace at t = tpeak, and Itest is the current determined at t = tpeak + 300 ms. The inactivation time course of ICa/IBa in the 300 ms following the peak was fitted with a monoexponential function, as follows:
with A0 and A1 being constants, and τ the time constant of inactivation.
I–V curves of K+ currents were fitted to a first-order Boltzmann function times driving force according to Nernst's equation:
where Vrev is the reversal potential for K+ (−73 mV), and Gmax is the maximum conductance of the IHC K+ currents (Kros and Crawford, 1990). Activation time constants of IK over 700 μs after start of the depolarization were determined as follows:
where I(t) is the current at time t, Imax is the steady-state current, and τ is the time constant of activation (Marcotti et al., 2003).
Parameters of Ca2+, Ba2+, and K+ currents were statistically analyzed using the Mann–Whitney U test. I–V curves were analyzed with a two-way ANOVA with Bonferroni post hoc test.
Exocytosis measurements.
Voltage-evoked capacitance changes were recorded from IHCs of P18–P20 mice. The bath was perfused with B1; IHCs were locally superfused with B4 as follows (in mm): 72.5 lactobionate·NaOH, 40 NaCl, 35 TEA, 15 4-AP, 5 CaCl2, 10 HEPES, 5.6 KCl, 5.6 glucose, 1 MgCl2, and 0.1 linopirdine, at pH 7.35 and 320 mosmol/kg. An LJP of 5 mV between the bath and pipette solution was subtracted; the small LJP change due to superfusion of B4 was neglected. Membrane capacitance measurements were performed using an Optopatch amplifier (Cairn Research Ltd) at room temperature, as described previously (Brandt et al., 2007). IHCs were depolarized for 100 ms from Vh (−85 mV) up to 45 mV in 10 mV increments. The change in membrane capacitance (ΔCm) was calculated as the capacitance averaged over 300 ms after the voltage step (after a delay of 150 ms from the end of the step) minus the capacitance averaged over 300 ms before onset of the voltage step. For statistics of the capacitance–voltage (C–V) curve, a two-way ANOVA with Bonferroni post hoc test was used. To assess the Ca2+ efficiency of exocytosis, QCa was calculated by integrating the absolute value of ICa over the time of depolarization plus tail currents for voltages less than or equal to Vmax. Fits to the individual ΔCm–QCa curves were made according to the following:
where y0 is the offset and c is a scaling factor, which yielded the power N for each cell and were analyzed using Student's t test. Data are reported as the mean ± SEM.
Immunohistochemistry.
Immunolabeling was performed on whole-mounts of the organ of Corti of 3-week-old mice. The scalae of each cochlea were injected with either 2% PFA in 100 mm PBS (Pirone et al., 2014), Zamboni's fixative (Stefanini et al., 1967), or −20°C cold ethanol and immersed in the fixative for 10–20 min on ice. After replacing the fixative with PBS, the cochlear spiral was dissected into an apical part of up to 22% of the length of the basilar membrane, corresponding to 2–8 kHz, which was designated as an apical turn, and a medial part of 22% to 45% (sometimes 60%) of the length of the basilar membrane, covering a region from 9 to 17 kHz (sometimes 26 kHz), which was designated as the medial turn. Specimens were labeled with antibodies against Cav1.3 (rabbit polyclonal, 1:500; Alomone Labs), Cavβ2 (rabbit polyclonal, custom-made; 1:500), CtPB2/Ribeye (mouse monoclonal, 1:100; BD Transduction Laboratories), PSD-95 (mouse monoclonal, 1:1000; NeuroMab), BKα (rabbit polyclonal, 1:500; Alomone Labs; mouse monoclonal, 1:50; Antibodies-Online), SK2 (rabbit polyclonal, 1:200; Sigma-Aldrich), glutamate receptor 4/GRIA4 (goat polyclonal, 1:500; Biorbyt, which required ethanol fixation), and Acti-stain 670 Fluorescent Phalloidin (200 nm; Cytoskeleton, Inc.). Primary antibodies were detected with Cy3-conjugated (1:1500; Jackson ImmunoResearch) or Alexa Fluor 488-conjugated secondary antibodies (1:500; Invitrogen). For all immunolabeling experiments, at least three specimens of three or more animals were analyzed. z-stacks of fluorescence images were acquired using a confocal laser scanning microscope (model LSM700 or LSM710, Carl Zeiss Microscopy). Images (70 × 70 nm2 pixel size) were obtained using a 63× oil objective (Planapochromat, Zeiss), 1.4 numerical aperture, with a pinhole of 1 airy unit. Stacks of 0.32-μm-thick optical slices, maximum intensity projections (MIPs) and single images were analyzed with Fiji software (Schindelin et al., 2012).
For quantifying the overlap of Cav1.3 with PSD-95 clusters, a line scan analysis was used on MIPs of double-labeled images each containing four adjacent IHCs. Ten lines measuring 10 × 0.7 μm2 were placed on each MIP image, thereby covering ∼90% of all clusters, without covering clusters twice. Corresponding pixel intensity profiles, for which a threshold of 500 was applied, yielded overlapping and single peaks of the two color channels. For analysis of cluster sizes, the channel of interest of an MIP image was subjected to background subtraction, and a thresholded binary image was created. The cluster area was analyzed automatically with Fiji (Schindelin et al., 2012). For quantification of the overlap of Cav1.3 clusters with GluA4 spots, images of 67.5 × 38.9 μm2 size covering eight IHCs were acquired at equal laser and gain settings, and MIPs were calculated. The channel of interest of an MIP image was background subtracted. A thresholded binary image was created (0 below threshold; 1 above threshold) with thresholds of 10% of the maximum intensity of the green color channel (GluA4) and 15% of the red color channel (Cav1.3). After discarding GluA4 clusters <0.15 μm2 and Cav1.3 clusters <0.05 μm2, the GluA4-thresholded binary image was used as a mask for the Cav1.3-thresholded binary image to yield overlapping clusters, which were analyzed using the particle count routine in Fiji. Parameters of overlapping/segregated Cav1.3, PSD-95, and GluA4 clusters were statistically analyzed using the Mann–Whitney U test.
Results
Expression of α2δ mRNA in IHCs
To assess the contribution of α2δ subunits to Ca2+ channel complexes of hair cells, quantitative real-time PCR for α2δ1–4 was performed, and transcript numbers for all α2δ subunits were obtained from standard curves, as previously established (Schlick et al., 2010). IHCs and OHCs were selectively harvested with micropipettes under microscope control, and cDNA was synthetized from mRNA by reverse transcription. At P6, transcripts for α2δ2 and α2δ3 were reliably detected, with α2δ2 showing the highest transcript numbers (Table 1). In eight independent IHC samples of the mature cochlea (P20–P25), only α2δ2 transcripts could be detected at low numbers. The same was true for OHCs at both P6 and P20–P25. α2δ transcript numbers were very low in mature hair cells, even when several hundred hair cells were pooled (Table 1), probably because of low Cav1.3/α2δ expression levels (Knirsch et al., 2007) and the limited number of cells. Despite the low number of detected α2δ2 transcripts (1–122; Table 1) the log-transformed transcript numbers strongly and highly significantly correlated indirectly with the Ct values of the endogenous control gene Hprt1 (Pearson correlation: r(17) = −0.79, p < 0.001; mean ± SEM Ct value, 33.42 ± 0.40; range, 31.36–36.94). This shows that the number of detected α2δ2 transcripts reliably increased with the quantity of cDNA, while transcripts of the other α2δ isoforms were either very low (at P6) or not detectable (P20–P25). Due to the fact that reliable antibodies against α2δ3 and α2δ2 are not available (Dolphin, 2013), the expression of α2δ subunits in hair cells could not be tested at the protein level. Nevertheless, our qPCR revealed the consistent presence of α2δ2 transcripts in hair cells before and after hearing onset, and thus we set out to study hearing and the cellular and molecular phenotypes of a functional α2δ2-null mouse, the ducky mouse.
Transcript numbers of α2δ subunits in samples of pooled IHCs and OHCs of the apical turn in mice before (P6) and after (P20–P25) hearing onset determined by quantitative RT-PCR
Impaired auditory function in du/du mice
Next, we assessed hearing performance of α2δ2 (du/du) mutants (Fig. 1). Click ABR thresholds of du/du mice were significantly elevated (33.0 ± 12.9 dB SPL; n = 8/15 animals/ears) compared with wt mice (14.7 ± 2.9 dB SPL; n = 8/16 animals/ears; Mann–Whitney U test, p < 0.001; Fig. 1A, left). Average frequency-specific ABR thresholds were always larger for du/du mice than for wt mice between 2 and 45 kHz. This difference was significant in the best hearing range of mice, specifically at 11.3, 16, and 22.3 kHz (two-way ANOVA, p < 0.01; Fig. 1A). Hearing function was further tested by measuring DPOAEs, an objective indicator of the cochlear amplifier including the electromotility of OHCs. Unexpectedly, 2f1-f2 DPOAE maximum amplitudes for f2 averaged over 10–18 kHz in 0.5 kHz steps were significantly larger in du/du mice (Fig. 1B; wt mice: 21.9 ± 4.0 dB SPL, n = 3/6 animals/ears; du/du mice: 28.9 ± 3.0 dB SPL, n = 4/8 animals/ears; p < 0.01, Mann–Whitney U test). This suggests that OHC function was not corrupted by the lack of α2δ2 and raises the question of why DPOAE amplitudes were not reduced but were even larger despite increased ABR thresholds in du/du mice. Averaged ABR waveforms for click stimuli 40 dB above threshold revealed differences between the genotypes (Fig. 1C). Notably, wave I, the negative peak of which appears at 1.4–1.9 ms after the start of the click stimulus in wt mice and represents the discharge of auditory nerve fibers (but see Pirone et al., 2014), was delayed by 0.3 ms (p < 0.01) in du/du mice. The peak-to-peak amplitude of wave I showed a tendency toward reduced values, but, due to the high variability especially in du/du mice, this reduction was not significant. Interestingly, wave III was completely distorted, suggesting that du/du mice also exhibit a central auditory processing phenotype.
Hearing function and integrity of OHCs of ducky mice. A, ABR thresholds (mean ± SD) of du/du mice were significantly increased for click stimuli (leftmost values: wt, n = 8/16 animals/ears; du/du, n = 8/15 animals/ears, ***p < 0.001, Mann–Whitney U test) and as a function of stimulus tone frequency (wt, n = 8/16 animals/ears; du/du, n = 7/14 animals/ears; **p < 0.01, two-way ANOVA with Bonferroni post hoc test, effect of genotype at 11.3, 16, and 22.3 kHz). B, Mean DPOAE maximum amplitudes ± SD with f1 starting at 9.1 kHz, L1 = 55 dB SPL, f2 = 1.2 · f1 averaged over 10–18 kHz, and L2 = 45 dB SPL were larger in du/du (n = 4/8 animals/ears) compared with wt (n = 3/6 animals/ears, **p < 0.01, Mann–Whitney U test). C, Averaged ABR waveforms to click stimuli at 40 dB above threshold (mean ± SEM) for 8 of 16 wt animals/ears (black) and 7 of 14 du/du animals/ears (gray). D, E, Normal number of OHC rows (arrowheads) and normal OHC hair bundle morphology as revealed by phalloidin staining of the apical turn of wt (D) and du/du (E) cochleae. F, Sketch of a mouse cochlear whole-mount that illustrates the position of the apical turn representing 2–8 kHz and the medial turn representing 8–17 kHz, which were used in immunolabeling and electrophysiological experiments. The medial part partially reached frequencies up to 24 kHz. G, Immunolabeling for the basolateral K+ channels BK (left column) and SK2 (middle column; merged BK and SK2 immunolabeling in the right column) at the base of du/du OHCs of all three rows of the medial cochlear turn (bottom row) did not differ from that of the wt OHCs (top row). This indicates the normal expression of OHC K+ channels that mediate inhibition by medial olivocochlear fibers.
Because OHCs of the apical cochlear turn of Cav1.3-deficient mice undergo early degeneration (Platzer et al., 2000), we tested for morphological integrity of OHCs in whole-mount preparations of 3-week-old mice. Apical and medial cochlear turns stained for actin with fluorescent phalloidin showed normal expression of three rows of OHCs (arrows) and normal hair bundle morphology in du/du mice compared with wt mice (Fig. 1D,E, shown for apical turns). To test whether medial efferent fibers from the brainstem, which can inhibit OHC electromotility at low to medium sound pressure levels, project normally to du/du OHCs, we labeled for the inhibitory effector K+ channels SK2 and BK (Dulon et al., 1998; Oliver et al., 2000; Rüttiger et al., 2004). The location of apical and medial turn OHCs in mouse cochlear whole mounts and their corresponding frequencies (Müller et al., 2005; Engel et al., 2006) are illustrated in Figure 1F. In both du/du and wt mice, BK channels colocalized with SK2 channels in two to six patches in all three rows at the base of the OHCs, showing normal expression of the effector channels of efferent inhibitory terminals in du/du mice (Fig. 1G, shown for medial turns).
Ca2+ and Ba2+ currents are reduced and altered in mature IHCs of du/du mice
Increased hearing thresholds without a reduction of DPOAEs pointed to a defect of IHCs. Therefore, whole-cell Ca2+ and Ba2+ currents were recorded in 3-week-old du/du and wt apical turn IHCs. Selected current traces in response to 8 ms step depolarizations for the voltages indicated, with 10 mm Ca2+ as a charge carrier are shown for a typical wt IHC (Fig. 2A) and a typical du/du IHC (Fig. 2B). The peak Ca2+ current (ICa) was larger in the wt IHC compared with the du/du IHC. I–V curves averaged for 20 wt and 12 du/du IHCs showed a shift toward positive voltages and a reduction in ICa in du/du IHCs (Fig. 2C). Average ICa amplitudes, cell capacitance, current density, and biophysical parameters of ICa activation, such as Vh and slope factor of the Boltzmann function obtained from fits to the individual I–V curves (see Materials and Methods), are given in Table 2. IHCs of du/du mice showed a highly significant reduction of peak ICa of 37.2% and a reduction of ICa density (ICa divided by cell capacitance) of 29.1% (Table 2). Their capacitance was smaller by 10% compared with wt IHCs with Ca2+ as a charge carrier, but not with Ba2+ as a charge carrier (Table 2). This discrepancy might result from reduced exocytosis in du/du IHCs in the Ca2+ current recordings (see below). Exocytosis, however, should not occur with Ba2+ as a charge carrier. Moreover, the Vh value was shifted by 5.2 mV to depolarized voltages in du/du IHCs, and the slope factor of ICa activation was increased by 0.9 mV, which indicates changed voltage-dependent gating of Cav1.3 channels lacking functional α2δ2 subunits. Reduced peak ICa in du/du IHCs may result from (1) fewer Cav1.3 channels in du/du IHCs, (2) altered single-channel conductance, and (3) altered gating properties of ICa or a combination of items 1–3. The reason for item 3 is that a shift of the ICa activation curve toward positive voltage values not only increases Vh but also lowers peak ICa due to a reduced driving force. Therefore, the permeability density (permeability normalized to the cell capacitance; see Eq. 1 in Materials and Methods) was determined from fitting activation curves. On average, this parameter was significantly reduced by 22%, indicating a reduction of the product of the number of Cav1.3 channels × single-channel conductance of 22%.
Whole-cell Ca2+ and Ba2+ currents in 3-week-old IHCs of ducky mice. A, B, Representative Ca2+ currents (ICa) of a wt IHC (A) and a du/du IHC (B) of apical cochlear turns in response to 8 ms step depolarizations to the voltages indicated. C, Averaged Ca2+ I–V curves ± SD for 20 apical turn wt and 12 apical turn du/du IHCs. For clarity, the SD is plotted in one direction only (−SD or +SD). Two-way ANOVA for the effect of genotype, ***p < 0.001. D, E, Representative Ba2+ currents (IBa) of a wt (D) and a du/du IHC (E) of apical cochlear turns in response to 8 ms step depolarizations to the voltages indicated. F, Averaged Ba2+ I–V curves ± SD for 12 apical turn wt and 11 apical turn du/du IHCs. Two-way ANOVA for the effect of genotype, ***p < 0.001. G, Averaged Ba2+ I–V curves ± SD for 13 wt and 9 du/du IHCs of medial/midbasal turns. Two-way ANOVA for the effect of genotype, ***p < 0.001. H, ICa responses to step depolarizations to Vmax lasting 400 ms and monoexponential fits (red) in a wt and a du/du IHC of apical cochlear turns. I, IBa responses to step depolarizations to a Vmax lasting 400 ms and monoexponential fits (red) in an apical wt and an apical du/du IHC.
Ca2+/Ba2+ current amplitudes, membrane capacitances, and activation and inactivation parameters of ICa and IBa in mature IHCs of du/du mice
Using Ba2+ as a charge carrier yielded larger currents (IBa) compared with Ca2+, which is shown for a typical wt IHC and a typical du/du IHC from the apical turn (Fig. 2D,E). Averaged I–V curves demonstrated reduced peak IBa in apical (Fig. 2F) and medial du/du IHCs (Fig. 2G) compared with the respective wt IHCs. The average IHC IBa was significantly reduced by 29.1% (apical turn) and by 47.3% (medial turn); reduction of IBa density amounted to 29.1% for apical IHCs and to 39.0% for medial IHCs compared with wt IHCs of the respective cochlear location (Table 2). Fitting the I–V curves yielded a highly significant shift of Vh by 7.2 mV versus wt IHCs in apical du/du IHCs and by 7.8 mV in medial du/du IHCs, and the slope factor showed a tendency to larger values, which was significant for apical IHCs (Table 2). In sum, the effects of the ducky mutation on IHC Ca2+ channels were more severe in the medial turn compared with the apical turn, in accordance with the larger ABR threshold elevation in the middle frequencies (Fig. 1A).
Using depolarization steps lasting 400 ms, we tested Ca2+- and voltage-dependent inactivation in apical turn IHCs. Peak current traces for wt and du/du IHCs are shown for ICa and IBa (Fig. 2H,I, respectively). The percentage of ICa inactivation after 300 ms was not different between wt and du/du IHCs, and the same was true for IBa (Table 2). Fitting ICa (IBa) traces with monoexponential functions yielded no difference in the inactivation time constants between wt and du/du apical IHCs for both ICa and IBa (Table 2).
Together, du/du IHCs showed significantly smaller ICa and IBa amplitudes and a positive shift in Vh. The rightward shift of the I–V curves by 5–8 mV in the absence of functional α2δ2 demonstrates that α2δ2 is part of the Cav1.3 channel complex and has a clear impact on channel gating in wt IHCs.
Reduced exocytosis with normal Ca2+ efficiency in IHCs of du/du mice
Next, we asked how the altered Ca2+ currents affected exocytosis in IHCs of du/du mice. Apical IHCs of mice aged P18–P20 were held at −85 mV, stimulated with 100 ms steps to different voltages, and changes in membrane capacitance (ΔCm) reflecting exocytosis were determined with 5 mm Ca2+ as a charge carrier (Fig. 3). Figure 3A depicts typical ΔCm increases (middle) and corresponding ICa traces (bottom) upon stepping to the individual Vmax for a wt and a du/du IHC. Both the capacitance increase and the underlying ICa were smaller in the du/du IHC. Capacitance changes were also determined as a function of voltage between −85 and 45 mV in 10 mV steps. Averaged ΔCm (top) and corresponding peak ICa values (bottom) as a function of voltage are shown for 10 wt and 11 du/du IHCs (Fig. 3B), yielding C–V and I–V curves. The C–V curves largely reflected the voltage dependence of the corresponding inverted I–V curves for both wt and du/du IHCs, with significant reductions of ΔCm at −15 and −5 mV (Fig. 3B, top; two-way ANOVA, effect of genotype, p < 0.05), the range of the largest differences in ICa between genotypes (Fig. 3B, bottom). To assess the Ca2+ efficiency of exocytosis, the Ca2+ charge QCa was calculated by integrating the absolute value of ICa over the time of depolarization, including tail currents in the ascending part of the C–V (up to 5 mV), and averaged ΔCm values were plotted against QCa (Fig. 3C). Fits corresponding to Equation 6 yielded an averaged power N of 1.58 ± 0.20 (mean ± SEM) for 11 wt IHCs, which was not different from the power of 1.45 ± 0.13 (mean ± SEM; p = 0.62, Student's t test) for 10 du/du IHCs. Thus, the Ca2+ efficiency of exocytosis was unchanged in du/du IHCs, and the reduction of ΔCm between −15 and −5 mV in du/du IHCs (Fig. 3B, top) was a result of their correspondingly reduced Ca2+ currents (between −25 and +5 mV; Fig. 3B, bottom).
Reduced exocytosis with normal Ca2+ efficiency in IHCs of du/du mice. A, Example traces of a P18 wt IHC and a P18 du/du IHC of apical turns. Cells were depolarized from Vh (−85 mV) for 100 ms to Vmax (wt, −5 mV; du/du, +6 mV). Top, Stimulus protocol; middle, resulting ΔCm; bottom, corresponding ICa. B, Averaged ΔCm ± SEM (top) and corresponding ICa ± SEM (bottom) as a function of voltage of 10 wt and 11 du/du IHCs. For clarity, the SEM is plotted in one direction only (−SEM or +SEM). Two-way ANOVA for the effect of genotype, *p < 0.05. C, Averaged ΔCm ± SEM plotted against the corresponding Ca2+ charge QCa ± SEM. QCa was calculated by integrating the absolute value of ICa over the time of depolarization, including the tail currents. Values for ΔCm and ICa were taken from B between −75 mV (sometimes −55 mV) and 5 mV. Fits corresponding to Equation 6 to the individual curves yielded averaged powers N of 1.58 (wt, n = 11) and 1.45 (du/du, n = 10).
Expression of BK channels in du/du mice
The presence of Ca2+- and voltage-dependent BK channels is a hallmark of mature IHCs. Their expression starts at the onset of hearing (Kros et al., 1998; Hafidi et al., 2005) and requires the presence of substantial Cav1.3 currents in neonatal development, because BK currents are absent from Cav1.3−/− IHCs (Brandt et al., 2003) and altered in IHCs lacking Cavβ2 (Neef et al., 2009). Because mature du/du IHCs from the apical turn revealed a reduction of peak ICa by 37%, we analyzed BK currents and protein expression of the pore-forming subunit BKα (Fig. 4). Typical whole-cell outward currents are shown for a wt (Fig. 4A) and a du/du apical IHC (Fig. 4B). I–V curves were extracted from current families by averaging the current amplitudes between 1.2 and 1.3 ms after the start of the depolarization (Ifast; Fig. 4A,B), at a time when delayed rectifier K+ currents are not yet activated (Brandt et al., 2007), and plotting them as a function of voltage (Fig. 4C). I–V curves were fitted by a first-order Boltzmann function, yielding the activation parameters Vh and k. Neither the average current amplitude nor the current density (both determined at 0 mV) and the activation parameters (Vh, k) were altered in du/du mice (Table 3). Activation kinetics of Ifast was fitted by monoexponential functions from 0.3 to 0.5 ms after the start of depolarization between −25 and 10 mV (Fig. 4A,B, red curves). The resulting averaged activation time constants of du/du IHCs were slightly larger over the voltage range analyzed than those of wt IHCs, indicating that BK currents activated slightly slower in du/du IHCs at a given voltage (Fig. 4D; two-way ANOVA, p < 0.01). A similar subtle increase in activation time constants has been observed in a subset of hypothyroid IHCs, which expressed BK currents with a developmental delay of 2 weeks (Brandt et al., 2007). In du/du IHCs, BKα protein showed a normal localization at the neck of the IHCs (red dot-like labeling; Fig. 4E,F). Together, BK current amplitudes, I–V relations, and localization of BKα were normal in du/du IHCs.
BK currents and the expression of BKα protein in du/du mice. A, B, Typical outward currents in IHCs from the apical turn of a wt mouse (A, P21) and a du/du mouse (B, P20) in response to voltage steps of 8 ms duration. Red traces are monoexponential fits to the current traces. C, Corresponding I–V curves for the wt and du/du IHCs in A and B were obtained by averaging the current amplitudes from 1.2 to 1.3 ms after the start of depolarization (A, B, gray lines), which represent K+ currents through BK channels. D, Average activation time constants ± SD of BK currents obtained from monoexponential fits from 0.3 to 0.5 ms as a function of voltage for 10 wt and 8 du/du mice; **p < 0.01, two-way ANOVA for the effect of genotype. E, F, Double immunolabeling for BKα (red) and the ribbon marker CtBP2/RIBEYE (green) in whole-mount preparations of organs of Corti from apical cochlear turns. Maximum intensity projections of confocal stacks of a stretch of four IHCs are shown for wt (E) and du/du (F). The dashed line shows the outline of an IHC, respectively. Nuclei are stained in blue with DAPI, and stereocilia and cuticular plates are stained with phalloidin (purple). Scale bar, 10 μm.
Amplitudes and biophysical properties of BK currents in mature IHCs of du/du mice
Presynaptic localization of Cav1.3 and Cavβ2 protein clusters is normal in du/du IHCs
Because of reduced Ca2+ currents in du/du IHCs, we asked whether the spatial distribution of the pore-forming Cav1.3 subunit and the dominant Cavβ subunit Cavβ2 of IHC Ca2+ channel complexes were altered in du/du IHCs. In control IHCs, Cav1.3 showed some extrasynaptic and cytosolic immunolabeling, but the most obvious labeling was observed in clusters with partial overlap with the ribbons (Fig. 5A,C–E, top). The same labeling pattern was found in IHCs of du/du mice (Fig. 5B–E, bottom). Similarly to Cav1.3, Cavβ2 immunolabeling clustered at the ribbon synapses of control IHCs with partial overlap of Cavβ2 and RIBEYE fluorescence signals (Fig. 5F,H–J, top). Again, the same labeling pattern was detected in IHCs of du/du mice (Fig. 5G–J, bottom). Ribbon numbers were counted in MIPs of anti-RIBEYE immunolabeling (Fig. 5, compare A–C, F–H). The average number of ribbons, which amounted to 16.3 ± 3.1 per control IHC and 15.3 ± 3.4 per du/du IHC, was not different between the two groups (p = 0.55; ribbons counted in three to six IHCs each of seven apical turn specimens of six control and du/du mice, respectively). Together, the ducky mutation neither affected the cluster formation of both Cav1.3 and Cavβ2 at the ribbon synapses nor altered the number of IHC ribbons.
Expression of Cav1.3 and Cavβ2 protein in IHCs of ducky mice. A–J, MIP of confocal stacks of whole-mount preparations of organs of Corti from apical cochlear turns coimmunolabeled for Cav1.3 (A–E) or Cavβ2 (F–J) with the ribbon marker CtBP2/RIBEYE at P18-P25. A, B, Stretches with four IHCs of ctrl (A) and du/du mice (B) labeled with anti-Cav1.3 and anti-RIBEYE are shown, with one IHC indicated by the dashed outline. The synaptic poles of the IHCs boxed in A and B are shown enlarged and with split color channels in C and D. C, Synaptic ribbons labeled with anti-RIBEYE at the basolateral pole of a ctrl IHC (top) and a du/du IHC (bottom). D, Corresponding immunolabeling for Cav1.3. E, Overlay of the signals of C and D indicates the close apposition of Cav1.3 clusters and ribbons at the basolateral pole in both ctrl and du/du IHCs. F, G, Localization of Cavβ2 (red) and RIBEYE immunolabeling (green) in stretches of four IHCs of ctrl (F) and du/du mice (G), with one IHC indicated by the dashed outline. Stereocilia are stained in purple with phalloidin. Boxed synaptic poles from F and G are shown enlarged and with split color channels in H (RIBEYE, green) and I (Cavβ2, red) for a ctrl (top) and a du/du IHC (bottom). J, Overlay of the signals of H and I indicates close apposition of Cavβ2 clusters and ribbons at the basolateral pole in both the ctrl and du/du IHCs. Nuclei are stained in blue with DAPI. Scale bars: A, B, F, G, 10 μm; C–E, H–J, 5 μm.
The spatial coupling of presynaptic Cav1.3 channels with postsynaptic PSD-95 and AMPA receptor clusters is impaired in du/du mice
Next we analyzed the expression and localization of the postsynaptic marker PSD-95, which acts as the main scaffold for postsynaptic receptor complexes (for review, see Opazo et al., 2012). The sizes of PSD-95 clusters showed some variability in both genotypes (Fig. 6A–C). Quantification of PSD-95 cluster areas did not reveal a significant difference between control IHCs (0.43 ± 0.24 μm2, n = 572) and du/du IHCs (0.47 ± 0.43 μm2, n = 747; p = 0.228, Mann–Whitney U test). Double immunolabeling for Cav1.3 and PSD-95 revealed largely overlapping fluorescence signals in control mice, with the smaller Cav1.3 signals fully covered by the larger PSD-95 signals (Fig. 6A,C–E, top). Occasionally, a Cav1.3-labeled cluster was not accompanied by a PSD-95-positive spot. However, at du/du synapses, significantly more separated Cav1.3- and PSD-95-positive clusters were observed (Fig. 6B–E, bottom), which was further quantified using a line scan analysis (see Materials and Methods; Fig. 6E,F) applied on MIPs covering four to five IHCs, as shown in Figure 6, A and B. The average total number of PSD-95 and Cav1.3 clusters was not different between apical turn of control and du/du mice, respectively (Fig. 6G). In contrast, the number of both separated PSD-95 and separated Cav1.3 clusters was significantly increased at apical du/du IHC synapses (Fig. 6H). The same was true for IHCs of the medial turn (Fig. 6I,J). About 21% (18.5%) of all PSD-95 clusters and 11% (19.8%) of all Cav1.3 clusters were separated from the respective counterpart in apical (respectively medial) wt IHCs, whereas those numbers increased to 39% (49.5%) and 30% (50.9%) for du/du IHCs of the apical (respectively medial) cochlear turn. In other words, lack of functional α2δ2 increased the number of unpaired PSD-95 clusters by a factor of 1.9 for the apical and 2.8 for the medial turn, and that of unpaired Cav1.3 clusters by a factor of 2.7 for both apical and medial turns. These findings indicate that the α2δ2 subunit is required for an optimal apposition of IHC presynaptic Cav1.3 channels and the postsynaptic scaffold protein PSD-95.
Altered spatial coupling of presynaptic Cav1.3 and postsynaptic PSD-95 clusters at the IHC synapse in du/du mice. A, B, MIP of confocal stacks of whole-mount preparations of organs of Corti from apical cochlear turns coimmunolabeled for Cav1.3 and PSD-95 at P19–P24. Stretches of four IHCs of a ctrl (A) and du/du mouse (B) labeled with anti-Cav1.3 (red) and anti-PSD-95 (green) are shown, with a synaptic pole of an IHC indicated by the dashed outline. C, D, Boxed synaptic poles from A and B are shown enlarged and with split color channels in C (PSD-95, green) and D (Cav1.3, red) for ctrl (top) and du/du IHC (bottom). E, Merged images contain a bar tool (10 μm long, 0.7 μm wide) that is used for colocalization analysis. Scale bars: A, B, 10 μm; C–E, 5 μm. F, Fluorescence intensity profiles for Cav1.3 (red) and PSD-95 immunolabeling (green) for the corresponding bar tools in E as a function of length. G, Analysis of the total number of immunopositive spots in regions each containing four apical turn IHCs covered by 10 bar tools, respectively, for PSD-95 (p = 0.57) and Cav1.3 labeling (p = 0.81) in ctrl IHCs (n = 6/4 ears/animals) and du/du IHCs (n = 6/4 ears/animals) revealed no difference between the genotypes. H, Both the numbers of separated PSD-95 clusters and those of separated Cav1.3 clusters were significantly increased in apical du/du versus ctrl IHCs (*p < 0.05; **p < 0.01, Mann–Whitney U test). I, Similarly, the total number of immunopositive spots for PSD-95 (p = 0.24) and Cav1.3 labeling (p = 0.46) in wt (specimen of 4 IHCs each, n = 6/4 ears/animals) and du/du medial turn IHCs (n = 6/3 ears/animals) showed no differences between genotypes. J, Numbers of separated PSD-95 clusters and Cav1.3 clusters were significantly increased in medial turn du/du vs wt IHCs (**p < 0.01; ***p < 0.001, Mann–Whitney U test).
Postsynaptic AMPA receptors face the IHC synaptic cleft and could directly or indirectly interact with α2δ2 at the Cav1.3 channel complex. AMPA receptors are (hetero-)tetramers consisting of four types of GluA1–4 subunits and auxiliary subunits (Traynelis et al., 2010); and GluA2/3 and GluA4 have been shown at the IHC synapse (Matsubara et al., 1996; Khimich et al., 2005; Engel et al., 2006). For double-labeling AMPA receptors and Cav1.3 channels, a goat polyclonal GluA4 antibody was chosen, which required ethanol fixation. Cav1.3 clusters partially overlapped with the larger GluA4 clusters or were completely enclosed by them in MIPs of the wt IHCs, but much less so in du/du IHCs, shown for the medial turn IHCs (Fig. 7). Figure 7, A and B, shows an overlay of red Cav1.3 and green GluA4 immunofluorescence in a region containing four wt and du/du IHCs, respectively. GluA4 clusters looked less ordered and somewhat fragmented (Fig. 7B,D,E) in du/du IHCs. Further, the number and size of Cav1.3 clusters was more variable, with a tendency to smaller sizes in du/du IHCs compared with wt IHCs, as shown in two examples of du/du IHCs with split color channels (Fig. 7B,D,E). A quantitative analysis demonstrated that the total number of GluA4-positive clusters was not different in du/du versus wt IHCs (Fig. 7F). Quantification of GluA4 clusters overlapping with Cav1.3 clusters yielded relatively small numbers when normalized to one IHC for the wt IHC (6.4 ± 3.7 per apical turn and 10.0 ± 4.7 per medial turn IHC; Fig. 7G). In our experience, using ethanol as a fixative generally results in smaller clusters of ion channels than using the cross-linking fixative PFA. Using a minimum size threshold in the subsequent analysis to discriminate signal from background may have contributed to an underestimation of the number of Cav1.3 clusters. However, in du/du IHCs the number of GluA4 clusters overlapping Cav1.3 clusters was further and significantly reduced (p < 0.001) to 3.6 ± 2.9 (per apical turn IHC) and to 4.3 ± 3.2 (per medial turn IHC) or to 56% of wt IHCs for the apical turn and 43% of wt IHCs for the medial turn (Fig. 7G).
Impaired spatial coupling of presynaptic Cav1.3 clusters and postsynaptic GluA4 AMPA receptor clusters at the IHC synapse in du/du mice. A, B, MIP of confocal stacks of whole-mount preparations of organs of Corti from medial cochlear turns coimmunolabeled for Cav1.3 and GluA4 at P22–P23. Stretches with three to four IHCs of a wt (A) and du/du mouse (B) labeled with anti-Cav1.3 (red) and anti-GluA4 (green) are shown, with the synaptic pole of one IHC indicated by the dashed outline. C–E, Boxed synaptic poles of IHCs from A and B are shown enlarged with color channels split in red (Cav1.3, left column) and green (GluA4, middle column), and with merged colors (left column) for wt (C) and du/du mice (D, E). F, G, Quantitative analysis of the spatial overlap of Cav1.3 and GluA4 immunolabeling in specimen, each containing a stretch of eight IHCs. F, The total number of GluA4-positive clusters was unchanged at IHC postsynapses of both the apical (p = 0.084) and the medial cochlear turn in du/du mice (p = 0.29). G, The numbers of overlapping Cav1.3/GluA4 clusters were significantly decreased in du/du vs wt IHCs from both apical and medial cochlear turns. ***p < 0.001, Mann–Whitney U test. Number of ears/animals/specimens analyzed (with eight IHCs per specimen), respectively: wt: apical, 10/6/53; medial, 8/6/40; du/du: apical, 7/4/42; medial, 7/4/36. Scale bars: A, B, 10 μm; C–E, 5 μm.
In summary, the lack of functional α2δ2 not only reduced the amplitude and shifted the voltage dependence of Ca2+ currents and exocytosis in IHCs, but, moreover, impaired the trans-synaptic apposition with GluA4 glutamate receptors and the postsynaptic scaffold PSD-95, which altogether led to hearing impairment.
Discussion
Using the ducky mouse lacking functional α2δ2, we show here with voltage-clamp recordings that α2δ2 coassembles with Cav1.3 and Cavβ2 in wt IHCs. It regulates the Ca2+ current amplitude and has an impact on the gating of Cav1.3 channels.
α2δ subunits in hair cells
Real-time qPCR analysis of all α2δ subunit isoforms indicated that neonatal IHCs express mainly α2δ2, whereas mature IHCs exclusively express α2δ2 mRNA. The low expression of α2δ3 mRNA detected in IHCs before hearing onset is in accordance with a small reduction of the Ba2+ current density in immature IHCs of α2δ3−/− mice (Pirone et al., 2014). α2δ2 mRNA was clearly present in both immature and mature IHCs and OHCs. Its low level at P20–P25 can be explained by the downregulation of Cav1.3 currents during development to only one-third of the level present at P6 in IHCs (Beutner and Moser, 2001; Johnson et al., 2005) and similarly also in OHCs (Knirsch et al., 2007). The fact that α2δ1, α2δ3, and α2δ4 mRNA were not detected indicates that α2δ2 is the dominant α2δ isoform that forms Ca2+ channels with Cav1.3 and Cavβ2 in mature IHCs. Interestingly, this L-type channel complex (CaV1.3/β2/α2δ2) is distinct from the P/Q-type channel complex in cerebellar Purkinje cells, which also coassembles with α2δ2 (Cav2.1/β4/α2δ2; Barclay et al., 2001). Given that in hair cells Cav1.3 currents are indispensable for generating Ca2+ action potentials, changes in immature Ca2+ currents may alter aspects of IHC development, leading, for example, to altered BK current properties in Cavβ2−/− IHCs (Neef et al., 2009) and du/du IHCs (see below).
Cav1.3 channels and Cavβ2 subunits still reached the IHC membrane and clustered at ribbon synapses of du/du IHCs. From our whole-cell recordings, we cannot determine whether the number of Cav1.3 channels, the single-channel conductance, or both were reduced. It is more likely, however, that the number of Cav1.3 channels present at the IHC surface rather than the single-channel conductance was affected. Immunolabeling showed smaller Cav1.3 clusters at du/du IHC presynapses, which were visible with the non-cross-linking ethanol fixation (Fig. 7), suggesting fewer Cav1.3 channels in the IHC membrane. Second, the single-channel conductance of Cav2.1 channels was unchanged both in cultured neonatal du/du Purkinje cells and in COS cells transfected with Cav2.1/Cavβ4 and ducky-α2δ2 compared with the respective wt (Barclay et al., 2001; Brodbeck et al., 2002). Based on a reduced surface expression of Ca2+ channels, du/du Purkinje neurons showed a 35% reduction of the whole-cell Ba2+ current (Barclay et al., 2001), which was similar to the reduction of the du/du IHC whole-cell Ca2+/Ba2+ current amplitude by 30–40% reported here. Impaired trans-synaptic coupling due to a lack of functional α2δ2 in the du/du IHC may have led to a reduced stability and increased turnover of Cav1.3 in the membrane (Dolphin, 2013) and, hence, a reduction of the Ca2+/Ba2+ current (see below).
Coassembly of Cav1.3 with α2δ2 at IHC ribbon synapses is surprising because α2δ4 together with Cav1.4 and Cavβ2 forms presynaptic L-type Ca2+ channel complexes at ribbon synapses of retinal photoreceptor and bipolar cells (De Sevilla Müller et al., 2013). Mice with a mutation of Cacna2d4 exhibit cone–rod dysfunction (Wycisk et al., 2006a), and humans with mutations in CACNA2D4 have autosomal-recessive cone dystrophy (Wycisk et al., 2006b). A potential contribution of α2δ2 to L-type Ca2+ channels at retinal ribbon synapses has yet to be established.
α2δ2 and hearing
Ducky mice exhibited elevated ABR thresholds, which is in line with reduced Ca2+ currents and reduced exocytosis in the IHCs. Unexpectedly, their elevated ABR thresholds were accompanied by DPOAEs with amplitudes that are larger than normal (Fig. 1A,B). There are cases of severe auditory neuropathy where deafness coexists with partially normal DPOAEs, such as in humans with mutations in otoferlin (Rodríguez-Ballesteros et al., 2003; Tekin et al., 2005) or in mice deficient in Cav1.3 (Engel et al., 2006). Given that the reduction of ICa in du/du IHCs at any given voltage was ∼50% compared with wt IHCs, the increase in click or frequency ABR thresholds of du/du mice was unexpectedly mild. Two scenarios, both of which are based on the systemic ablation of functional α2δ2 in du/du mice, may have reduced the hearing loss. First, the stapedius reflex, a middle ear reflex, attenuates loud sound entering the ear (Mukerji et al., 2010). If α2δ2 was playing an important role in transmitter release at the neuromuscular junction of the stapedius muscle in wt mice, that reflex may have been attenuated in du/du mice, increasing the sound amplitude entering the cochlea and the DPOAE signal leaving the cochlea. Alternatively, inhibitory synapses at OHCs, which are activated at medium to high sound pressure levels (for review, see Guinan, 2006), could require α2δ2-containing presynaptic Ca2+ channels for transmitter release, which would result in impaired inhibition of OHCs in du/du mice. Disinhibition of the cochlear amplifier would alleviate an otherwise substantial hearing loss, inferred from altered function of the du/du IHCs alone. To assess the isolated, IHC-dependent contribution of α2δ2 to hearing function will require the analysis of a conditional, IHC-specific α2δ2 knock-out mouse, in which systemic effects of α2δ2 deletion on other synapses (e.g., of brainstem medial efferent olivocochlear or stapedius nerve fibers) can be excluded.
The drugs gabapentin and pregabalin targeting α2δ1 und α2δ2 are used for the treatment of neuropathic pain and epilepsy in humans (Dolphin, 2012b). They likely reduce the number of functional Ca2+ channels by interfering with their trafficking from the ER to the plasma membrane (Hendrich et al., 2008; Tran-Van-Minh and Dolphin, 2010). Reversible hearing loss has been reported as a side effect of gabapentin treatment under conditions of elevated plasma concentrations (Pierce et al., 2008), which may have been caused by a severe reduction of Cav1.3 channels/currents in IHC membranes.
A novel function of α2δ2 at the IHC synapse
IHCs of du/du mice exhibited a largely normal phenotype with respect to cell capacitance (i.e., cell size), amplitude of whole-cell BK currents, localization of BK channels, number of ribbons, and clustering of Cav1.3 and Cavβ2 protein at the ribbons. The shift in Vh of the Ca2+ currents in du/du IHCs, being equivalent to altered gating of Cav1.3 channels, strongly suggests that in wt IHCs the extracellularly localized α2δ2 subunit physically interacts with the extracellular loops of the Cav1.3 protein (Dolphin, 2013). Synapses of the afferent auditory pathway require ultrafast synaptic transmission, and optimal positioning of presynaptic release sites and postsynaptic receptor complexes is an important factor in keeping reaction times short. Here we have shown that trans-synaptic alignment of Cav1.3 clusters with GluA4 AMPA receptors and with the postsynaptic scaffold PSD-95 was impaired at IHC synapses lacking functional α2δ2. In general, presynaptic and postsynaptic neuronal compartments in the CNS are coupled by the cell adhesion molecules neuroligin and neurexin, which form an interaction layer in the synaptic cleft (Südhof, 2008). Presynaptic neurexins are anchored at the active zone, whereas neuroligins are anchored at the postsynaptic density (Südhof, 2008). Ablation of all three neurexin isoforms led to a marked reduction of presynaptic Cav2.2 currents in neocortical neurons (Missler et al., 2003), but the mechanism for this link is unclear. If neurexins/neuroligins are expressed at the IHC synapse, which is unknown so far, they may be responsible for the principal organization of the synapse, whereas α2δ2 might contribute to fine-tuning the apposition of clustered Cav1.3 channels and postsynaptic elements, such as GluA4 and PSD-95, for reducing reaction times (Figs. 6, 7, 8). α2δ subunits bear protein–protein interaction domains, including a von Willebrand factor A domain and a Cache domain (Dolphin, 2012a, 2013), which makes them good candidates for coupling Cav subunits (with which they interact noncovalently) with proteins of the AMPA receptor complex directly or with proteins of the extracellular matrix (Eroglu et al., 2009; Fig. 8). Recently, the extracellular matrix proteins thrombospondin 1 and 2 have been shown to be required for proper IHC synaptogenesis (Mendus et al., 2014). α2δ1 has been shown to act as a receptor for thrombospondins independently of its contribution to Ca2+ channel function (Eroglu et al., 2009), and α2δ2 might have a similar function.
Model of the role of α2δ2 in trans-synaptic coupling of IHC Cav1.3 channels with the postsynaptic AMPA receptor complex. A, In the wt IHC, the extracellularly localized α2δ2 subunit (orange) couples the Cav1.3 channel (red) with the AMPA receptor complex (blue), possibly through an unknown linker protein (black). PSD-95 (green) acts as a scaffold for AMPA receptors. The α2δ2-mediated trans-synaptic link perfectly aligns the presynaptic Cav1.3 channel with the postsynaptic AMPA receptor–PSD-95 complex. B, In du/du mice lacking functional α2δ2, the coupling between Cav1.3 channels and the AMPA receptor complex is abolished. As a result, the AMPA receptor–PSD-95 complex is no longer perfectly aligned with Cav1.3 across the synaptic cleft, leading to an increase in the number of separated Cav1.3 and GluA4/PSD-95 clusters, respectively.
From our findings, the question arises as to whether α2δ subunits, specifically α2δ2, play a role in coupling presynaptic VGCCs and postsynaptic receptors, not only in IHCs, but also at CNS synapses. Because of the uniform pore-forming subunit Cav1.3 and the unusual clustering of ∼80 Cav1.3 channels at each IHC presynapse (Brandt et al., 2005), the highly specialized IHC may enable us to dissect functions of α2δ subunits and possibly other presynaptic and postsynaptic partners, which at conventional synapses—due to the much smaller active zones—so far have remained elusive. Here we show that α2δ2 can form Ca2+ channel complexes not only with Cav2.1 and Cavβ4, as in cerebellar Purkinje cells (Barclay et al., 2001), but also with the L-type channel Cav1.3 and auxiliary Cavβ2 (Neef et al., 2009). In conclusion, the specific combination of a particular pore-forming Cavα1 subunit with a distinct α2δ isoform in vivo may be determined by the requirements of the synapse rather than by the α1 subunit itself.
Footnotes
This work was supported by Deutsche Forschungsgemeinschaft (DFG) Sonderforschungsbereich (SFB) 894 (Grants A8 to J.E. and A3 to V.F.), DFG SFB 1027 (Grant A4 to J.E.), the Austrian Science Fund (Grants P24079 and SFB F4415), and Saarland University. We thank Jennifer Ihl, Angela Di Turi, Roman Egger, and Stefanie Geisler for excellent technical assistance; and Yvonne Schwarz for help with data analysis.
The authors declare no competing financial interests.
- Correspondence should be addressed to Dr. Jutta Engel, Department of Biophysics, Center for Integrative Physiology and Molecular Medicine (CIPMM), Saarland University, School of Medicine, Building 48, 66421 Homburg, Germany. jutta.engel{at}uni-saarland.de
