Null Mutation of α1D Ca2+ Channel Gene Results in Deafness but No Vestibular Defect in Mice
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- Dou, H., Vazquez, A.E., Namkung, Y. et al. JARO (2004) 5: 215. doi:10.1007/s10162-003-4020-3
Multiple Ca2+ channels confer diverse functions to hair cells of the auditory and vestibular organs in the mammalian inner ear. We used gene-targeting technology to generate α1D Ca2+ channel-deficient mice to determine the physiological role of these Ca2+ channels in hearing and balance. Analyses of auditory-evoked brainstem recordings confirmed that α1D−/− mice were deaf and revealed that heterozygous (α1D+/−) mice have increased hearing thresholds. However, hearing deficits in α1D+/− mice were manifested mainly by the increase in threshold of low-frequency sounds. In contrast to impaired hearing, α1D−/− mice have balance performances equivalent to their wild-type littermates. Light and electron microscope analyses of the inner ear revealed outer hair cell loss at the apical cochlea, but no apparent abnormality at the basal cochlea and the vestibule. We determined the mechanisms underlying the auditory function defects and the normal vestibular functions by examining the Ba2+ currents in cochlear inner and outer hair cells versus utricular hair cells in α1D+/− mice. Whereas the whole-cell Ba2+ currents in inner hair cells consist mainly of the nimodipine-sensitive current (~85%), the utricular hair cells express only ~50% of this channel subtype. Thus, differential expression of α1D channels in the cochlear and utricular hair cells confers the phenotype of the α1D null mutant mice. Because vestibular and cochlear hair cells share common features and null deletion of several genes have yielded both deafness and imbalance in mice, α1D null mutant mice may serve as a model to disentangle vestibular from auditory-specific functions.
Keywordshearingbalancinginner earhair cellsCa2+ channelsvoltage clamp
Despite the general agreement that distinct voltage-gated Ca2+ channels serve as the principal conduits for Ca2+ influx into cells, thus conferring specific Ca2+-dependent functions, the exact role(s) of individual Ca2+ channel subtypes remains largely unknown. Even for specific functions such as neurotransmitter release, neurons employ more than one type of Ca2+ channel to execute this function, suggesting substantial redundancy in the nervous system (Smith and Cunnane 1997; Wu et al. 1999). There are three families of voltage-gated Ca2+ channels (VGCCs) with intrafamily sequence identities above 80% (Ertel et al. 2000) and additional diversity is generated by alternative splicing of exons encoding the cytoplasmic loop of different repeats of the α subunits (Solatov et al. 1995; Lin et al. 1997; Kollmar et al. 1997a, b). Genes for the voltage-gated channel CaV1.3 (α1D) are expressed throughout the nervous system and play a minor role in neurotransmitter release (Wu et al. 1999; Ertel et al. 2000). In hair cells, however, the channel is the most abundant isoform (Zidanic and Fuchs 1995; Rodriguez–Contreras and Yamoah 2001, 2002), suggesting distinct physiological roles in the inner ear.
Hair cells form a tonic synapse with afferent nerve terminals of the eighth cranial nerve and are poised to respond rapidly to sustained stimuli of different intensities and to transmit amplitude and frequency modalities of the stimuli (Hudspeth 1989). The fast neurotransmitter release in hair cells relies on the voltage- and time-dependent properties of clusters of α1D Ca2+ channels at the release sites (Rodriguez–Contreras and Yamoah 2001; Rispoli et al. 2001). Moreover, the whole-cell current activates at low membrane voltages (~−50 mV), has a fast activation time constant (~0.5 ms), and has half-activation voltages ranging from −40 to +20 mV (Hudspeth and Lewis 1988; Zidanic and Fuchs 1995; Martini et al. 2000; Platzer et al. 2000; Rodriguez–Contreras and Yamoah 2001). Furthermore, hair cells may express not only α1D Ca2+ channels alone, but several other distinct Ca2+ channels as well, ensuring multiple Ca2+-dependent processes. The differential roles of distinct Ca2+ channel subtypes in hair cells have been proposed using evidence from immunocytochemical, polymerase chain reaction (PCR), and electrophysiological techniques. Variants of CaV1.2 (α1C), CaV2.2 (α1B), and CaV2.3 (α1E) have been localized in cochlear and other hair cells (Green et al. 1996; Lopez et al. 1999; Rodriguez–Contreras and Yamoah 2001).
To analyze the physiological role of α1D Ca2+ channels in hair cells, we used gene targeting to produce α1D-deficient mice. The results of our analyses demonstrate that α1D is crucial for hearing, but less so for maintaining balance.
Generation of α1D-deficient mice
Conventional gene-targeting technology was used to generate Ca2+ channel α1D null mutant mice (Namkung et al. 2001). A murine CaV1.3 genomic DNA clone having the first two exons of the gene was isolated from a 129/SVJ mouse genomic library. The region extending from half of the 3′ part of the first exon to half of the 5′ part of the second exon was deleted and replaced by the IRES β-gal expression cassette and the NEO cassette. The negative selection marker, TK cassette, was inserted into the end of the 3′ homology region of the targeting vector. The targeting vector was transfected into J1 embryonic stem (ES) cells. Chimeras were backcrossed to C57BL/6J mice. Germline transmission was determined by DNA typing of tail DNA. α1D+/− mice were then intercrossed to obtain α1D−/− mice. All mice analyzed were from F2 and F3 generations. Because the α1D−/− mice were fertile, they were further intercrossed to increase the sample number of null mutants.
Gross evaluation of auditory and vestibular functions
We evaluated the ear twitch response of ten wild-type α1D+/+, α1D+/− and α1D−/− mice with a hand clap (Preyer’s reflex) to grossly assess the hearing status of the three genotypes. To obtain a gross assessment of the vestibular (utricular) function of the mice, we performed a swim test. Mice were placed in a water bath at 37°C and allowed to swim and climb onto a dry platform. The swimming performance and the time taken to swim to the target were scored in a blind fashion. Seven α1D+/+ and α1D−/− mice each were tested. Balance was further tested using a custom-made setup as described by Xiang et al. (1997). Mice were placed on a soft fabric-covered horizontal cylinder (~7 cm in diameter) and positioned 10 cm above a foam pad. The cylinder was connected to a variable speed motor that runs from 0 to 15 rpm. Each animal’s ability to balance on both the stationary cylinder and the rotating cylinder was scored. Seven α1D+/+ and α1D−/− mice each were tested.
Auditory brainstem responses
Twenty α1D+/+, 28 α1D+/−, and 19 α1D−/− mice (age range: 5–8 weeks old) were anesthetized with avertin and auditory brainstem response (ABR) measurements were recorded as previously described (Kozel et al. 1998; Flagella et al. 1999). Briefly, a ground needle electrode and recording needle were placed subcutaneously in the scalp, and then a calibrated electrostatic speaker coupled to a hollow ear bar was placed inside the pinna. Broadband clicks and pure tones (8, 16, and 32 kHz) were presented in the animal’s ear in 10 dB increments, starting from 0 dB SPL and ending at 100 dB SPL. The ABR sweeps were computer-averaged (time-locked with onset of 128–1024 stimuli, at 20/s) out of the continuous electroencephalographic activity. The threshold of hearing was determined as the lowest intensity of sound required to elicit a characteristic waveform.
Light microscopy of the inner ear
Light microscope analyses of the cochlea and vestibular organs of the inner ear from 5–12-week-old mice of all genotypes were studied using procedures described in Kozel et al. (1998). The mice were deeply anesthetized with sodium pentobarbital and transcardially perfused with a solution containing 2% glutaraldehyde and 2% paraformaldehyde in 0.1 M cacodylate buffer, pH 7.3. The temporal bones were harvested and the inner ears were perfused with fixative through the oval and round windows and left in fixative overnight. The temporal bones were decalcified in a 0.1 M EDTA solution for 1 week. The specimens were postfixed in buffered 1% osmium tetroxide (OsO4) for 1–2 h at room temperature, dehydrated in graded ethanol solutions followed by propylene oxide, and embedded in Spurr’s resin. Serial sections (1–2 μm) parallel to the modiolus of the inner ears were cut and stained with toluidine blue and then subsequently cover-slipped.
Scanning electron microscopy of the inner ear
Five mice from each genotype were selected for scanning electron microscope analyses. Similar to the procedures for light microscopy, the temporal bones were harvested and fixed in 2% glutaraldehyde in 0.1 M phosphate buffer saline (PBS) for 36 h at room temperature (~21°C) and decalcified in 0.1 M EDTA for 5 days. The cochlea, utricle, saccule, and the semicircular canals were dissected out of the specimen and postfixed in 1% OsO4 for 2 min. Dehydration of the specimen was carried out in a graded ethanol series. The specimens were critical-point dried from liquid CO2, sputter-coated with gold–palladium, and examined in a scanning electron microscope (Philips XL30). Digital images were captured and stored in a personal computer interfaced with the microscope.
Distortion product otoacoustic emissions (DPOAE)
Mice were anesthetized with ketamine (95 mg/kg) and xylazine (4 mg/kg). The f1 and f2 primary tones were generated by a 2-channel frequency synthesizer (Hewlett Packard 3326A), presented over two tweeters (Realistic), and delivered through a small soft rubber probe tip. Ear-canal sound pressure was measured with a commercial acoustic probe (Etymotic Research 10B+). The ear-canal sound pressure was sampled and synchronously averaged (n = 8) by a digital signal processor for frequencies <20.1 kHz, and by a dynamic signal analyzer (Hewlett Packard 3561A) for frequencies >20.1 kHz. DF-grams were collected over a range of geometric mean frequencies between 5.6 and 48.5 kHz (f2 = 6.3–54.2 kHz), in 0.5-octave intervals at stimulus levels of L1 = L2 = 65 dB SPL, with f2/f1 = 1.25.
Whole-cell recording of Ba2+ currents
Standard patch-clamp recording techniques were used to record whole-cell currents to study the Ba2+ currents in cochlear and utricular hair cells in intact sensory epithelia. Utricles and cochleae were excised from young wild-type (+/+) and homozygous mutant (−/−) mice between postnatal days 1–10, as described in previous reports (Rüsch and Eatock 1996; Holt et al. 1997; Moser and Beutner 2000). All chemicals were obtained from Sigma Chemical Co. (St Louis, MO), unless indicated. Mice were sacrificed by cervical dislocation and decapitation. The sensory epithelium was removed from each inner ear while immersed in MEM solution (GIBCO, pH 7.4 with 10 mM HEPES: Gaithersburg, MD). The bony labyrinth was opened, and once the epithelium of interest was exposed, the tissue was treated with protease XXVII (100 μg/ml, 15 min). The surrounding extraneous tissue and nerve were trimmed away and the sensory epithelium was then mounted on a recording chamber with Cell-Tak (Collaborative Biomedical Products, Bedford, MA). The mounted epithelia were placed under an upright Olympus compound microscope (IX50). Solutions used in the experiments were as follows with mM quantities of each component given: Modified Tyrode, consisting of 130 Na+, 3 K+, 4 Ca2+, 5 glucose, and 5 HEPES, pH 7.4 (NaOH). The external solution for recording Ba2+ current contained 105 Na+, 25 TEA, 5 4-AP, 1 Ba2+, 5 HEPES, and 5 glucose. The patch-pipette filling solution contained 80 NMG, 40 Cs+, 1 Mg2+, 5 EGTA, 10 HEPES, 3 ATP, 1.5 GTP, pH 7.4 (CsOH). Thus, outward K+ currents were suppressed with 4-AP, TEA, Cs+, and the nonpermeant cation NMG. Stock solutions of Bay K 8644 (Bay K: Calbiochem, La Jolla, CA) and nimodipine (Calbiochem) were dissolved in 100% dimethyl sulfoxide (DMSO). The final concentrations of Bay K and nimodipine were 10 and 20 μM, respectively.
Patch-pipettes (1.5 mm o.d. and 1 mm i.d.) were pulled from borosilicate glass using the horizontal puller, P87 (Sutter Instrument, Novato, CA). Patch electrodes had 2–5-MΩ tip resistances when filled with the pipette solution. Recordings were done using an Axopatch 200B patch-clamp amplifier interfaced with a personal computer equipped with pClamp software (Axon Instruments, Foster City, CA). Currents were filtered at 2 kHz and sampled at 10 kHz. Leakage and capacity currents were subtracted using the p/4 method. Data analyses were performed using the pClamp and Origin software (MicroCal Inc., Northampton, MA). Where appropriate, pooled data were presented as means ± SD. Significant differences between groups were tested using Student’s t-test [data with p > 0.05 were considered not significant (NS)].
Hearing defect and normal balance in α1D−/− mice
Histology of the inner ear
Whole-cell Ba2+ currents in cochlear and utricular hair cells
α1D−/− mice were generated to better understand the physiological functions of this particular Ca2+ channel isoform in hearing and balancing. The most robust feature of the α1D−/− mice was deafness and increased thresholds for audiological response to sound in the heterozygote littermates, which was restricted mainly to low-frequency sounds. A completely unexpected finding was that the hearing loss was not associated with gross structural changes in IHCs, suggesting that α1D Ca2+ channels play a minimal role in maintaining IHC morphology. The findings that the OHCs at the apical turn of the cochlea expressed mostly the nimodipine-sensitive Ca2+ currents (>95%: α1D Ca2+ channels) and that hair cells in this region of the cochlea degenerated in α1D-deficient mice may indicate that Ca2+ is required for the maintenance of OHC morphology. Other Ca2+ channel subtypes are known to be present in hair cells and they carry a residual current following block by nimodipine. Although this nimodipine-insensitive current in IHCs in α1D−/− mice may sustain the cellular architecture, the current may not be sufficient to mediate adequate neurotransmitter release to confer hearing. In contrast, the results for hair cells in the vestibule indicate that the non-α1D Ca2+ channels may be sufficient in maintaining both the structure and functions of the cells. Although gross assessment of the vestibular function suggested that the α1D−/− mice have no vestibular defect, a more direct diagnosis of vestibular function may be required. Nonetheless, we infer that differential expression of diverse Ca2+ channel subtypes in hair cells of the inner ear helps explain the balance and auditory phenotypes of the α1D Ca2+ channel-deficient mice.
Hearing in vertebrates requires the precise synchronization of cochlear IHC activity and the auditory nerve. Previous studies have shown that the kinetics and voltage-dependent activation properties of the L-type channels in mammalian cochlear IHCs and hair cells from other vertebrates may suffice to confer both the tonic and phasic Ca2+-dependent release of neurotransmitters (Hudspeth 1989; Zidanic and Fuchs 1995; Martinez–Dunst et al. 1997; Moser and Beutner 2000; Beutner et al. 2001). However, recent reports have demonstrated that hair cells in lower vertebrates and mammals do express nimodipine-insensitive Ca2+ currents (Su et al. 1995; Martini et al. 2000; Platzer et al. 2000; Rodriguez–Contreras and Yamoah 2001). Although the functions of the non-L-type current remain unclear, their voltage-dependent properties suggest a possible function in tonic release of neurotransmitters (Martini et al. 2000; Rodriguez–Contreras and Yamoah 2001). Measurements of hair cell capacitance as an index for exocytotic release of neurotransmitters from hair cells have shown that the nimodipine-sensitive component of the whole-cell Ca2+ (L-type) current plays a marked role in phasic neurotransmitter release. Moreover, the functions of non-L-type current may be obscured by its baseline activity and minimal contribution toward the hair cell Ca2+ current (Moser and Beutner 2000; Spassova et al. 2001). A more sensitive method for resolving neurotransmitter release, such as postsynaptic recordings, may be required to determine the contribution of the non-L-type current (von Gersdorff et al. 1998; Spassova et al. 2001). Nonetheless, the in vivo and in vitro results presented strongly indicate that the α1D Ca2+ channel is critically required for hair cell functioning and hearing.
Our findings on the properties and expression patterns of the L-type current in IHCs and OHCs are consistent with previous reports using hair cells from frog (Hudspeth and Lewis 1988; Su et al. 1995; Smotherman and Narrins 1999; Rodriguez–Contreras and Yamoah 2001), chick (Kimitsuki et al. 1994; Zidanic and Fuchs 1995), and guinea pig (Rennie and Ashmore 1991). The activation threshold (~50 mV), rapid onset (time-to-peak = ~1 ms), and slow inactivation of the nimodipine-sensitive current are reminiscent of those previously described in cochlear and vestibular hair cells. Similar to whole-cell Ca2+ currents in other hair cells, the expression pattern is heterogenous. Our observations add significantly new findings, which demonstrate that for OHCs, the contribution of the non-L-type current toward the total Ca2+ current increases along the axis of the cochlea with cells at the apical turn expressing little non-α1D channel current and those at the basal turn expressing ~15% of the non-L-type current. Because OHCs at the basal turn of the cochlea have sufficient non-L-type current and retain their gross cellular morphology and because the cells at the apical turn virtually lack the non-L-type Ca2+ current in the α1D−/− mice, hence having little Ca2+ influx leading to OHC degeneration, the possibility that the non-L-type Ca2+ channel in hair cells may play housekeeping roles in addition to other cell-specific functions is raised. Furthermore, synaptic connections between OHCs and afferent nerve terminals are physiologically silent (Robertson and Gummer 1985). Thus, the L-type current is not expected to play an obvious role in neurotransmitter release. Moreover, there are a plethora of physiological activities in cells which require Ca2+ as a second messenger (Fuchs 1996).
At the macroscopic level, the results of the present studies are similar to recent data from α1D null mutant mice published by Platzer et al. (2000). However, our extension of the previous studies revealed important differences at the microscopic level. Serial sectioning of the entire cochlea at the light microscope level and scanning electron microscopy clearly showed that only the OHCs at the apical turn of the cochlea degenerate in 5–7-week-old α1D−/− mice. This is in sharp contrast to an earlier report that both IHCs and OHCs degenerated after P14 (Platzer et al. 2000). Although the previous report may reflect genuine differences between the two models, it is likely that evaluation of different planes of sections without complete serial reconstruction led to the earlier results (Platzer et al. 2000). Thus, our findings confirm the earlier report that α1D Ca2+ channel knockout mice have hearing impairment despite apparent normal IHC morphology. We have extended the scope of these studies to demonstrate that with the exception of OHCs at the apical turn of the cochlea which degenerate, cochlear hair cells do remain intact in the α1D−/− mice.
Finally, neurotransmitter release by hair cells onto afferent nerve terminals of the vestibular nerve is mediated by Ca2+ entry via presynaptic VGCCs. Although the nimodipine-sensitive Ca2+ current contributes substantially to the VGCC in vestibular hair cells (Hudspeth and Lewis 1988; Prigioni et al. 1992; Martini et al. 2000; Rispoli et al. 2001), our behavioral and electrophysiological analyses of the vestibular system of the α1D−/− mice show that in addition, there is a non-nimodipine-sensitive current. In contrast to the auditory system, the vestibular hair cells employ multiple VGCCs that mediate neurotransmitter release to ensure proper balance. Hair cells in both the cochlea and the vestibule share common morphological and functional traits that are usually difficult to disentangle. Thus, the differential expression and functions of Ca2+ channel isoforms in vestibular and cochlear hair cells will ultimately constitute an invaluable functional and molecular biological tool to study the two systems in isolation.
We thank Dr N. Chiamvimonvat and members of our laboratory for their constructive comments on the manuscript. This work was supported by grants to ENY from the NIH (R01 DC03828, DC04512).