J Neurophysiol 113: 3531–3542, 2015. First published March 25, 2015; doi:10.1152/jn.00869.2014.

Activity-dependent regulation of expression in mouse outer hair cells

Yohan Song, Anping Xia, Hee Yoon Lee, Rosalie Wang, Anthony J. Ricci, and John S. Oghalai Department of Otolaryngology-Head and Neck Surgery, Stanford University, Stanford, California

Submitted 4 November 2014; accepted in final form 19 March 2015

Song Y, Xia A, Lee HY, Wang R, Ricci AJ, Oghalai JS. patch-clamp recording conditions (Kakehata and Santos-Sac- Activity-dependent regulation of prestin expression in mouse outer chi, 1995 and 1996; Santos-Sacchi 1991; Santos-Sacchi et al. hair cells. J Neurophysiol 113: 3531–3542, 2015. First published 1998a). Thus, measuring the NLC is a common way of assess- March 25, 2015; doi:10.1152/jn.00869.2014.—Prestin is a membrane necessary for outer (OHC) electromotility and normal ing the amount of functional prestin within an OHC (Abe et al. 2007; Oliver and Fakler 1999;). hearing. Its regulatory mechanisms are unknown. Several mouse Downloaded from models of hearing loss demonstrate increased prestin, inspiring us to There are many ways to modulate the voltage dependence of investigate how hearing loss might feedback onto OHCs. To test force production by prestin protein within the plasma mem- whether centrally mediated feedback regulates prestin, we developed brane, such as changing the surrounding lipid environment, a novel model of inner hair cell loss. Injection of diphtheria toxin (DT) intracellular Ca2ϩ level, membrane tension, membrane poten- into adult CBA mice produced significant loss of inner hair cells tial, and ionic composition of the intracellular and extracellular without affecting OHCs. Thus, DT-injected mice were deaf because they had no afferent auditory input despite OHCs continuing to compartments (Kakehata and Santos-Sacchi 1995; Keller et al. receive normal auditory mechanical stimulation and having normal 2014; Rajagopalan et al. 2007; Ratnanather et al. 1996; Santos- http://jn.physiology.org/ function. Patch-clamp experiments demonstrated no change in OHC Sacchi et al., 1998b and 2001; Wu and Santos-Sacchi 1998). prestin, indicating that loss of information transfer centrally did not However, the basic mechanisms of how prestin levels within alter prestin expression. To test whether local mechanical feedback the plasma membrane are regulated remain unclear. We know C1509G regulates prestin, we used Tecta mice, where the tectorial that prestin has an extremely high density within the lateral membrane is malformed and only some OHCs are stimulated. OHCs wall plasma membrane. It has a charge density of ϳ6,000– connected to the tectorial membrane had normal prestin levels, Ϫ ␮ 2 Ϫ whereas OHCs not connected to the tectorial membrane had elevated 10,000 e / m (where e is the electron charge) (Belyantseva prestin levels, supporting an activity-dependent model. To test et al. 2000; Mahendrasingam et al. 2010), and an estimated whether the endocochlear potential was necessary for prestin regula- 60–75% of the lateral wall surface area is composed of by 10.220.33.4 on September 23, 2016 tion, we studied TectaC1509G mice at different developmental ages. intramembranous particles, thought to be composed of or OHCs not connected to the tectorial membrane had lower than normal include prestin (Forge 1991; Kumano et al. 2010; Wang et al. prestin levels before the onset of the endocochlear potential and 2010). In addition, prestinϪ/Ϫ OHCs transfected with prestin higher than normal prestin levels after the onset of the endocochlear driven by the constitutively active cytomegalovirus promoter potential. Taken together, these data indicate that OHC prestin levels have a similar prestin density to untransfected wild-type OHCs are regulated through local feedback that requires mechanoelectrical (Xia et al. 2008). These data suggest that prestin production is transduction currents. This adaptation may serve to compensate for variations in the local mechanical environment. not limited at the level of transcription but argue instead that mechanical considerations physically limit the amount of pres- ; hair cell; hearing; prestin; electromotility; mechanics tin within the plasma membrane. However, there is growing evidence that prestin levels can increase in states of hearing loss. For example, prestin mRNA SOUND-INDUCED VIBRATIONS of the deflect the expression is increased after salicylate administration (Yu et al. sensory hair cell stereociliary bundle, creating a receptor po- 2008), after noise exposure (Chen 2006; Mazurek et al. 2007; tential within the soma. In response, outer hair cells (OHCs) Xia et al. 2013), and in the presence of a malformed tectorial generate force, a process termed electromotility (Brownell et membrane (Liu et al. 2011; Xia et al. 2010). These observa- al. 1985). These forces amplify the cochlear traveling wave and tions suggest the possibility that hearing loss stimulates OHCs increase organ of Corti vibration, producing the exquisite to upregulate prestin, perhaps in an effort to compensate for auditory sensitivity and frequency selectivity characteristic of decreased hearing sensitivity. However, all of these previously normal mammalian hearing. Prestin (SLC26A5), a tetrameric published reports created hearing loss in animals by affecting protein present within the lateral wall plasma membrane of OHCs in some way. Thus, it remains unclear whether higher OHCs, is necessary for electromotility (Cheatham et al. 2004; brain centers recognize a state of hearing loss and provide Dallos et al. 2008; Hallworth and Nichols 2012; Liberman et efferent feedback to residual OHCs via the medial olivoco- al. 2002; Wang et al. 2010). Prestin is often modeled as having chlear bundle (Fig. 1, A and B). Alternatively, signaling could two states so that voltage-dependent stochastic transitions fit a occur locally within the epithelium to regulate prestin levels Boltzmann function and produce electromotility (Dallos et al. through a direct reduction in the OHC stereociliary bundle 1993; Iwasa 1994). These transitions are associated with stimulation and/or reciprocal synapses that have been found in charge movement across the membrane that can be measured type II nerve fibers that innervate multiple OHCs within a row as a voltage-dependent non-linear capacitance (NLC) under (Francis and Nadol Jr. 1993; Spoendlin 1969). In the present Address for reprint requests and other correspondence: J. S. Oghalai, Dept. study, we sought to clearly differentiate between efferent of Otolaryngology-Head and Neck Surgery, Stanford Univ., 300 Pasteur Drive, feedback and local feedback as regulators of prestin level using Edwards R113, Stanford, CA 94305-5739 (e-mail: [email protected]). two different mouse models that allowed us to isolate the www.jn.org 0022-3077/15 Copyright © 2015 the American Physiological Society 3531 3532 ACTIVITY-DEPENDENT REGULATION OF PRESTIN Downloaded from http://jn.physiology.org/ by 10.220.33.4 on September 23, 2016

Fig. 1. Experimental concept and mouse models. A: illustration of a normal organ of Corti. The tectorial membrane (TM), inner hair cell (IHC), outer hair cell (OHC) rows (1, 2, and 3), (BM), afferent auditory nerves (blue arrow), medial olivocochlear efferent nerves (green arrow) are shown. The TM stimulates OHC stereociliary bundles in all three rows of OHCs in control mice. B: potential mechanisms of prestin regulation. OHCs amplify the vibration of the organ of Corti in synchrony with its stereociliary bundle stimulation. This drives IHC stimulation and an afferent auditory response. Higher brain centers could then modulate efferent neural activity, which effects a change in OHC prestin levels. We broke this feedback loop using mice with IHC loss. Alternatively, OHC bundle stimulation could directly regulate prestin within individual OHCs or along a row via reciprocal synapses in type II nerve fibers. We selectively blocked OHC bundle stimulation using TectaC1509G mice. C: mice injected with diphtheria toxin (DT inj) demonstrated severe IHC loss and thus lacked an afferent neural response. OHC stimulation was unaffected. D: TectaC1509G mice had altered TM anatomy. The TM only stimulates first-row OHCs in TectaC1509G/ϩ mice (top). None of the OHCs are stimulated in TectaC1509/C1509 mice (bottom). source of the feedback. Our findings indicate that prestin is F2 mice were crossed with wild-type CBA mice for three generations regulated locally within the epithelium in an activity-dependent to reduce the 129/Sv/Ev background to 12.5%. All mice studied were manner. F6–F7 generation littermates in this background. Auditory brain stem responses and distortion product otoacoustic emissions. Mice were anesthetized using ketamine (100 mg/kg) and METHODS xylazine (10 mg/kg), and supplemental doses were given until paw areflexia was achieved. Auditory brain stem responses (ABRs) and The Institutional Animal Care and Use Committee of Stanford distortion product otoacoustic emissions (DPOAEs) were measured as University approved the study protocol. previously described (Cho et al. 2013; Oghalai 2004a; Xia et al. Diphteria toxin mouse model. This mouse model has not been 2007). Briefly, the ABR signal was measured with a bioamplifier previously published. We used 5-wk-old wild-type CBA/CaJ mice (DP-311, Warner Instruments, Hamden, CT) from needle elec- (stock no. 000654, The Jackson Laboratory, Bar Harbor, ME). These trodes placed at the ventral surface of the tympanic bulla and the mice have stable auditory thresholds from postnatal day (P)12 to P21 skull vertex with a ground electrode placed in the hindleg. The (Ohlemiller et al. 2010). Intraperitoneal injections of diphteria toxin sound stimulus frequency ranged from 4 to 70 kHz. At each (DT) (no. 15042B1, List Biological Laboratories, San Jose, CA), each frequency, the sound intensity was raised in 10-dB steps from 10 at a dosage of 50 ng/g, were administered for 3 days in a row. to 80 dB sound pressure level (SPL). After 260 repetitions were Uninjected mice of the same age were used as controls. averaged, ABR responses for each frequency were interpolated, TectaC1509G mouse model. The TectaC1509G mutant mouse was and the threshold was determined to be when the peak to peak previously created in our laboratory (Xia et al. 2010). Heterozygous voltage was Ͼ5 SD above the noise floor. If an ABR response was

J Neurophysiol • doi:10.1152/jn.00869.2014 • www.jn.org ACTIVITY-DEPENDENT REGULATION OF PRESTIN 3533 not detectable at 80 dB SPL, we then arbitrarily defined the fit to the first derivative of a two-state Boltzmann function using Igor threshold at that frequency to be 80 dB SPL. Pro 6.22A software (WaveMetrics, Lake Owego, OR) to calculate DPOAEs were measured by placing a probe tip microphone (type total elementary charge movement (Qmax), voltage at peak capaci- ␣ 4182, Bruel & Kjaer) in the external auditory canal. The frequency tance (V1/2), and the slope factor of voltage dependence ( ) (Oliver response of this microphone was measured using a free-field micro- and Fakler 1999) as follows: phone (type 4939, Bruel & Kjaer) that has a flat frequency response Qmax out to 100 kHz and compensated for in the software. The stimuli used C͑V͒ ϭ C ϩ lin ␣ Ϫ ␣ ϩ Ϫ Ϫ␣ 2 to elicit DPOAEs were two sine wave tones 1-s in duration with exp͓͑V V1⁄2͒ ⁄ ͔͕1 exp͓͑V V1⁄2͒ ⁄ ͔͖ different frequencies, where frequency 2 (F2) ϭ 1.22 ϫ frequency 1 (1) (F1). F2 was varied from 4 to 70 kHz, and the two tones were given where C is capacitance and V is voltage. Linear capacitance (Clin) was at identical intensities ranging from 20 to 80 dB in 10-dB steps. The calculated as the average capacitance at Ϫ150 and ϩ120 mV. NLC magnitude of the acoustic signal detected from the microphones was ϫ Ϫ was calculated as the difference between peak capacitance and linear measured at the 2 F1 F2 frequency. DPOAE responses for each capacitance. Charge density was then calculated using the specific frequency were interpolated, and the threshold was determined to be Ͼ Ͼ membrane capacitance previously determined for mouse OHCs (Abe when the DPOAE magnitude was both 5 dB SPL and 2 SD from et al. 2007). The following equation was used:

the noise floor. If a DPAOE signal was not detectable at 80 dB SPL, Downloaded from Ϫ we then arbitrarily defined the threshold at that frequency to be 80 dB Qmax ⁄ e d ϭ (2) SPL. C ⁄ C Basilar membrane tuning curves. Basilar membrane vibrations lin conv were measured using a custom-built optical coherence tomography where d is charge density or the number of electrons moved per square system using previously described techniques (Gao et al., 2013 and micrometer of cell surface area, eϪ is the electron charge (1.602 ϫ Ϫ19 2014; Lee et al. 2015). Briefly, mice were anesthetized with a 10 C), and Cconv is the specific membrane capacitance of 0.008 ketamine-xylazine mixture and secured in a head holder, and the bulla pF/␮m2. was surgically opened. The mouse was oriented to view the region of Quantitative real-time PCR. As previously described (Xia et al. http://jn.physiology.org/ the cochlea one-half turn down from the helicotrema. By imaging 2013), two cochleae from each mouse were dissected in cold PBS and noninvasively through the otic capsule bone, vibratory data were placed in lysis buffer (RNAqueous Micro-Kit, AM1931, Invitrogen). collected from the midportion of the basilar membrane over the Cochleae were then homogenized in a 1.5-ml microcentrifuge tube for frequency range from 2 to 13 kHz. Stimulus intensities ranged from 1 min at 0°C using a pestle and mortar (no. 47747-370, VWR Pellet 10 to 80 dB SPL. Vibratory magnitude responses below the noise floor Mixer). RNA was then isolated using a RNA extraction kit (AM1931, of 0.11 nm were not analyzed. Q10 dB values were calculated using the RNAqueous Micro-Kit, Ambion). RNA concentration and purity were vibratory data collected with the 10 dB SPL stimulus as the charac- determined using a Nanodrop ND1000 spectrophotometer. Samples teristic frequency (the frequency of maximum vibratory amplitude) not meeting concentration and purity thresholds were discarded. divided by the bandwidth 10 dB down from the peak. Cochlear gain cDNA was then generated using iScript Reverse Transcription Super- by 10.220.33.4 on September 23, 2016 was calculated by dividing the maximum vibratory amplitude in the mix for quanitative real-time PCR (Bio-Rad). A control reaction with active cochlea (living) by that of the passive cochlea (dead) using 10 no reverse transcriptase verified that samples were free of genomic dB SPL stimuli. Since we could not measure the response of the DNA. For targeted amplification, we selected prestin, myosin VIIa, passive cochlea with a 10 dB SPL stimulus because it was below the and GAPDH primers (Xia et al. 2010). Quantification of targets noise floor of our system, we used the vibratory response of the basilar was then determined using SsoFast EvaGreen Supermix (Bio-Rad) in membrane to the 80 dB SPL stimulus and scaled it down linearly by a thermal cycler (CFX96 Real-Time System, Bio-Rad). Mean PCR Ϫ ⌬ ( CT) ϭ 70 dB to derive the 10-dB response of the passive cochlea. threshold cycles (CT) were compared using the 2 Ϫ Ϫ Measurement of NLC. Mice were killed by rapid cervical disloca- 2 (CT,target CT,control) relative quantification method. tion, and their bullas were removed for further dissection in external Whole mount preparations. Whole mount preparations of the co- solution containing (in mM) 150 NaCl, 2 KCl, 2.5 MgCl2, 4 CaCl2,10 chlear epithelium were prepared as previously described (Xia et al. HEPES, 5 glucose, 2 creatine, 2 ascorbate, and 2 pyruvate. The 2013). Briefly, cochleae were excised, fixed in 4% paraformaldehyde external solution had an osmolality of 305 mosM/kg and a pH of 7.4. at room temperature for 1 h, and immersed in 0.5 M EDTA overnight. After removal of some of the otic capsule bone, a small strip of the Cochleae were then rinsed with PBS containing 0.1% Triton X-100 organ of Corti from the apical region about a half turn down from the three times for 5 min each. The entire organ of Corti was dissected out helicotrema (9- to 12-kHz region) measuring 2–4 mm in length was of the cochlea under a microscope. Immunolabeling was performed by dissected out, and the tectorial membrane was gently brushed off. The first blocking the cochleae with 4% donkey serum (017-000-121, epithelium was then secured by two strands of dental floss in a Jackson ImmunoResearch Laboratories, West Grove, PA) in PBS in recording chamber filled with external solution. The experimental Tween 20 (PBST) for 1 h at room temperature before incubation with chamber holding the preparation was transferred to an upright micro- the primary antibody overnight at 4°C. Cochleae were washed four scope (Axioskop 2, Zeiss) and viewed through a ϫ63 water-immer- times with PBST, incubated with secondary antibody at room tem- sion objective. perature for 1 h, and rinsed with PBS. The primary antibody was Patch pipettes were made with a pipette puller (P1000, Sutter rabbit anti-myosin VIIa (1:200, Proteus Biosciences, Ramona, CA) Instruments, Novato, CA) with initial resistances of 3–4 M⍀. The diluted in PBST. Secondary antibody was Alexa fluor 563 donkey standard internal solution contained (in mM) 5 EGTA, 135 KCl, 3 anti-rabbit (1:500, Invitrogen, Grand Island, NY) diluted in PBST.

MgCl2, 3 ATP, 5 phosphocreatine, 10 HEPES, and 2 ascorbate. The After being labeled, cochleae were washed with PBS and mounted in internal solution had an osmolality of 290 mosM/kg and a pH of 7.25. antifade fluorescence mounting medium (DAKO, Carpinteria, CA) OHCs were whole cell voltage clamped within the epithelium with an and coverslipped. A confocal Zeiss LSM5 Pascal microscope system Axon 200B amplifier using jCLAMP software (SciSoft). Data from with a ϫ20/0.5 EC Plan-NEOFLUAR objective was used to image cells that appeared unhealthy (i.e., those that appeared swollen or hair cells. contained intracellular particulates in Brownian motion) were ex- Statistical analysis. All data were analyzed with Microsoft Excel cluded and not analyzed. (Microsoft Office 2007). Plots and figures were created with Sigma- Cell capacitance was measured using a continuous high-resolution Plot (11.0, Systat Software, San Jose, CA), Adobe Photoshop CS6 two-sine wave stimulus protocol superimposed onto a voltage ramp (Adobe Systems, San Jose, CA), and Adobe Illustrator CS6 (Adobe from Ϫ150 to ϩ120 mV (Santos-Sacchi 2004). Capacitance data were Systems). Statistical significance was determined using either a non-

J Neurophysiol • doi:10.1152/jn.00869.2014 • www.jn.org 3534 ACTIVITY-DEPENDENT REGULATION OF PRESTIN paired two-tailed Student’s t-test (for comparisons of two groups), DT-injected mice have normal prestin levels. To directly one-way ANOVA followed by a nonpaired two-tailed Student’s t-test assess whether prestin levels were affected in DT-injected for significant findings (for comparison of more than two groups), or mice, we patch clamped 17 first-row OHCs (8 DT-injected two-way ANOVA (for multiple comparisons among two or more Ͻ OHCs and 9 control OHCs) from 14 mice (8 DT-injected mice groups). P values of 0.05 were considered significant. Baseline and 6 control mice). We studied the experimental mice 7 days currents are reported as means Ϯ SD; all other data are presented as means Ϯ SE. after DT injection. This time delay was selected to give adequate time to ensure that feedback to the OHCs could happen and was based on our knowledge that IHCs were gone RESULTS by 3 days after DT injection and the fact that 7 days after noise DT causes inner hair cell death but leaves OHCs intact. We exposure, OHC prestin is upregulated (Xia et al. 2013). first studied a mouse that has hearing loss due to inner hair cell Baseline currents measured immediately after achieving the (IHC) loss (Fig. 1C) to investigate whether hearing loss, as whole cell configuration using a holding potential of Ϫ80 mV detected by higher brain centers, modulates efferent feedback were similar between cohorts (DT-injected cohort: 0.06 Ϯ 0.07 to regulate prestin. Thus, any efferent feedback pathways that nA and control cohort: 0.2 Ϯ 0.2 nA, P ϭ 0.14 by t-test). We regulate prestin should be activated if hearing loss upregulates then measured membrane capacitance and found the standard Downloaded from prestin. This new mouse model was discovered serendipitously bell-shaped curve characteristic of prestin function, with no while working on another research project using a transgenic obvious difference between DT-injected mice and control mice mouse where the DT receptor gene was targeted to a different (Fig. 3A). To quantify prestin function, capacitance-versus- cell population. As a control, we injected adult wild-type CBA voltage curves were fit to the first derivative of a Boltzmann mice with DT (referred to henceforth as “DT-injected mice”). function. We then calculated linear capacitance, NLC, Qmax, ␣ In these DT-injected mice, we found a dramatic loss of IHCs 3 charge density, , and V1/2. None of these parameters were days after the injection (Fig. 2, A and B). OHCs were only different between the two cohorts of mice (P Ͼ 0.1 for each http://jn.physiology.org/ minimally affected. We counted IHCs and OHCs from three to comparison; Fig. 3B), indicating that both cohorts of mice had five whole mount preparations from the middle turn of the similar functional prestin levels. The lack of a difference in cochleae from three control and three DT-injected mice. Each linear capacitance between the two groups indicates that the preparation contained 16–22 IHCs and 45–67 OHCs in control OHCs we studied were the same size, since this serves as a mice. There was a 69% loss of IHCs (P Ͻ 0.001 by t-test) and surrogate measure of cell surface area (Huang and Santos- a 2% loss of OHCs (P ϭ 0.03 by t-test) 3 days after DT Sacchi 1993). injection (Fig. 2C). Finally, we performed quantitative real-time PCR to deter-

We then studied cochlear function in a cohort of CBA mice mine whether prestin mRNA levels were affected in DT- by 10.220.33.4 on September 23, 2016 by measuring ABRs and DPAOEs from the left ears of 12 mice injected mice. For each cohort, we wanted to normalize prestin before DT injection and again 3 days after the injection (Fig. mRNA to myosin VIIa mRNA, an internal hair cell reference 2D). After DT injection, mice had no measureable ABRs to gene (Duncan et al. 2006; Xia et al., 2010 and 2013). Since the equipment limits (P Ͻ 0.01 by two-way ANOVA). In contrast, direct effect of DT-induced loss of IHCs will be to reduce DPOAE thresholds after injection were no different from myosin VIIa levels without affecting prestin levels, this ratio before injection (P ϭ 0.15 by two-way ANOVA). Further- would be expected to increase even if prestin is not upregu- more, we measured basilar membrane tuning curves in cohorts lated. Therefore, we measured this impact by also normalizing of six control mice and in six mice 3–5 days after DT injection. prestin and myosin VIIa to GAPDH, a general housekeeping These in vivo measurements were made from the midportion of gene found in all cells. Furthermore, we calculated the pre- the basilar membrane about a half turn down from the helico- dicted changes for these ratios using the cell counts shown in trema. Measurements were made in both live and dead mice. Fig. 2C (Xia et al. 2013). There were no significant differences Living DT-injected mice demonstrated nonlinear displacement in the prestin-to-GAPDH ratio, myosin VIIa-to-GAPDH ratio, responses, and the responses became linear postmortem (Fig. or prestin-to-myosin VIIa ratio between DT-injected mice and 2E). Qualitatively, these curves looked similar to those of predicted values (P Ͼ 0.15 for each comparison; Fig. 3C). In control mice. We then calculated Q10 dB and cochlear gain in addition, there were no significant changes in these ratios in both cohorts to compare these features of cochlear amplifica- DT-injected mice compared with control mice (P Ͼ 0.08 for tion. We found no differences between DT-injected and control each comparison). Taken together, these results indicate that ϭ ϭ mice for either measure (Q10 dB: P 0.51; gain: P 0.52; Fig. prestin transcription, prestin protein function, and the nonlin- 2F). This indicates that cochlear amplification, a process within earities amplified by prestin in vivo that lead to DPOAEs are the mammalian cochlea that is produced by OHCs, is normal in unaffected when hearing loss is caused by blocking afferent mice after DT injection. auditory input. Thus, the concept that centrally mediated feed- Taken together, these data indicate that DT kills IHCs back increases prestin levels in states of hearing loss was without affecting OHC survival and function in adult CBA proven false. This indicates that local factors within the epi- mice. We do not know why this occurs, as this does not happen thelium likely provide the feedback necessary to regulate OHC in neonatal mice (Cox et al. 2014); however, we assume that prestin levels. the IHC must later develop sensitivity to DT. Nevertheless, for Prestin levels in TectaC1509G mice. To address how local the purposes of this research, we used the DT-injected mouse feedback could regulate prestin, we tested whether the direct model to investigate if centrally mediated feedback is involved effect of stimulating the stereociliary bundle of the OHC in the regulation of prestin. We reasoned that if hearing loss correlated with prestin levels using the TectaC1509G mutant modulates centrally mediated feedback pathways, DT-injected mouse (Xia et al. 2010). In this mouse strain, the anatomy of mice should have increased prestin in their OHCs. the tectorial membrane varies with the genotype of the mouse

J Neurophysiol • doi:10.1152/jn.00869.2014 • www.jn.org ACTIVITY-DEPENDENT REGULATION OF PRESTIN 3535 Downloaded from http://jn.physiology.org/ by 10.220.33.4 on September 23, 2016

Fig. 2. Adult CBA mice injected with DT have IHC loss but not OHC loss. A: photomicrographs of representative hair cell epithelia in CBA mice not injected with DT (control; top) and CBA mice injected with DT (bottom). The three rows of OHCs (1, 2, and 3) and the one row of IHCs (arrow) are labeled. In both images, a pipette (P) is patch clamped to an OHC in row 1. The green bar highlights missing IHCs in the DT-injected mouse. Scale bar ϭ 10 ␮m. B: similar view of a fixed whole mount preparation immunolabeled for myosin VIIa to label hair cells. Scale bar ϭ 10 ␮m. C: loss of IHCs was noted after DT injection, whereas there was no effect on OHCs. D: auditory brain stem response (ABR) thresholds increased after DT injection, whereas distortion product otoacoustic emissions (DPOAE) thresholds were unchanged. E: representative basilar membrane tuning curves measured in a live DT-injected mouse demonstrated the expected nonlinear responses. Postmortem, the responses became linear. Stimulus intensity levels are indicated by the numbers next to each tracing. F: average

Q10 dB and cochlear gain calculated from basilar membrane tuning curves were similar between cohorts of DT-injected mice and control mice. (Fig. 1D). Tectaϩ/ϩ mice have a tectorial membrane that is closest to the pillar cells), and TectaC1509G/C1509G mice have a attached to the stereociliary bundle of every OHC within the tectorial membrane that is not attached to any OHCs. In epithelium, TectaC1509G/ϩ mice have a tectorial membrane that heterozygous mice, the altered tectorial membrane anatomy, in is only connected to OHCs that sit within the first row (the row which it has a functional connection to OHCs in the first row,

J Neurophysiol • doi:10.1152/jn.00869.2014 • www.jn.org 3536 ACTIVITY-DEPENDENT REGULATION OF PRESTIN

previous studies, based on whole organ physiology and whole cochlea mRNA analysis, suggested that that there was more prestin in TectaC1509G/ϩ and TectaC1509G/C1509G mice com- pared with Tectaϩ/ϩ mice (Liu et al. 2011; Xia et al. 2010). However, no single cell analyses or row-specific functional assays for prestin levels have been done to test whether this represents functional prestin protein within the plasma membrane. Here, we patch-clamped OHCs from adult mice at the age of P30 so that their prestin levels would be mature (Abe et al. 2007; Belyantseva et al. 2000; He et al. 1994). All OHCs studied were within the first row, so if OHC stimulation relates to prestin level, TectaC1509G/ϩ OHCs should have the same prestin level as Tectaϩ/ϩ OHCs, whereas TectaC1509G/C1509G

OHCs should have an elevated prestin level. We recorded from Downloaded from nine OHCs from six Tectaϩ/ϩ mice, eight OHCs from five TectaC1509G/ϩ mice, and nine OHCs from six TectaC1509G/C1509G mice. After the whole cell configuration had been achieved, average baseline currents using a holding potential of Ϫ80 mV were similar between the cohorts (Tectaϩ/ϩ mice: 0.2 Ϯ 0.2 nA, TectaC1509G/ϩ mice: 0.07 Ϯ 0.08 nA, and TectaC1509G/C1509G mice: 0.08 Ϯ 0.2 nA, P ϭ 0.3 by ANOVA). http://jn.physiology.org/ We then measured membrane capacitance over the voltage range (Fig. 4). NLC, Qmax, and charge density were signifi- cantly elevated in first-row OHCs from TectaC1509G/C1509G mice compared with first-row OHCs from Tectaϩ/ϩ mice (P Ͻ 0.007 for all comparisons by t-test). In contrast, first-row OHCs from TectaC1509G/ϩ mice were no different than first- row OHCs from Tectaϩ/ϩ mice (P Ͼ 0.7 for all comparisons

␣ by 10.220.33.4 on September 23, 2016 by t-test). Linear capacitance, , and V1/2 values were not significantly different between the three genotypes (P Ͼ 0.2 for all comparisons by one-way ANOVA). These data indicate that OHCs within the first row that are not attached to the tectorial membrane have increased functional prestin. In addition, the lack of a difference in linear capacitance indicates that the OHC is able to upregulate functional prestin levels without a change in surface area. To further test this concept, we proceeded to patch addi- tional OHCs from the second and third rows in each genotype and compared all three rows among all three genotypes (Fig. 5A). From nine Tectaϩ/ϩ mice, we recorded from eight cells in the second row and seven cells in the third row. In 10 Fig. 3. OHCs from DT-injected mice have normal levels of functional prestin. C1509G/ϩ A: average capacitance as a function of voltage was measured in OHCs from Tecta mice, we recorded from 8 cells in the second C1509G/C1509G control and DT-injected mice. B: linear capacitance, nonlinear capacitance, row and 10 cells in the third row. In eight Tecta total charge moved (Qmax), charge density, slope factor of voltage dependence mice, we recorded from eight cells in the second row and eight (␣), and voltage at peak capacitance (V ) were not significantly different 1/2 cells in the third row. Average baseline currents at a holding between control and DT-injected mice. C: levels of prestin mRNA and myosin Ϫ VIIa mRNA normalized to the level of GAPDH mRNA were similar between potential of 80 mV were similar to previous data, and there DT-injected mice and predicted values based on the cell counts (left and were no differences between the three rows or three genotypes middle, respectively). In addition, the level of prestin mRNA normalized to the (P Ͼ 0.3 by two-way ANOVA). Consistent with the observa- level of myosin VIIa mRNA was similar to the predicted value (right). None tions in first-row OHCs, membrane capacitance measures re- of the changes after DT injection were significantly different than control. P30, vealed increased functional prestin levels in OHCs that were postnatal day 30. not attached to the tectorial membrane (Fig. 5, B and C). This but not the second or third rows, has been confirmed by was evidenced by increased NLC, Q , and charge density in max ϩ multiple methods including histological sectioning, in situ OHCs from the second and third rows in TectaC1509G/ mice visualization of the tectorial membrane free edge and the and from all three rows in TectaC1509G/C1509G mice (P Ͻ 0.001 underlying OHC rows, in situ imaging of dynamic Ca2ϩ for all comparisons by ANOVA followed by t-tests). In con- ϩ ϩ changes within OHCs in each row during sound stimulation, in trast, OHCs from all three rows in Tecta / mice had similar Ͼ vivo cochlear microphonic recordings (Xia et al. 2010), and in NLC, Qmax, and charge density values (P 0.6 by ANOVA). situ optical imaging using optical coherence tomography (Gao Linear capacitance and ␣ values were not significantly differ- et al. 2011). In addition, noise exposure produces loss of only ent between the three genotypes or rows (P Ͼ 0.1 by two-way first row OHCs in heterozygous mice (Liu et al. 2011). Our ANOVA). There was a slight difference in V1/2 values between

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Fig. 4. OHCs in row 1 in TectaC1509/C1509 mice have ele- vated functional prestin levels compared with those in Tec- taϩ/ϩ and TectaC1509G/ϩ mice. A: average capacitance as a function of voltage in row 1 of OHCs in the three Tecta genotypes. B: linear capacitance was not significantly different among all three genotypes, whereas nonlinear capacitance,

C1509/C1509 http://jn.physiology.org/ Qmax, and charge density were higher in Tecta mice ϩ/ϩ C1509G/ϩ ␣ compared with Tecta and Tecta mice. and V1/2 were not significantly different between the groups. *P Ͻ 0.05 compared with Tectaϩ/ϩ mice (by unpaired t-test). by 10.220.33.4 on September 23, 2016

the rows (P ϭ 0.02 by two-way ANOVA), with a gradient period of postnatal cochlear development. The tectorial C1509G toward a more negative V1/2 value moving from the first row to membrane malformation in Tecta mice is present by P5 the third row; however, there was no difference in V1/2 values (Xia et al. 2010), prestin expression begins at P4–P5, prestin between the genotypes (P ϭ 0.07 by two-way ANOVA). mRNA expression is maximal by P10 (Abe et al. 2007), and Taken together, these data confirm that OHCs attached to the the stiffness of the OHC correlates strongly with the onset of tectorial membrane have normal prestin levels, whereas OHCs prestin expression (Jensen-Smith and Hallworth 2007). There- detached from the tectorial membrane have increased prestin fore, if mechanical forces stemming from tectorial membrane- levels. Thus, decreased stereociliary bundle stimulation corre- induced deflections of the OHC regulate prestin, lates with increased prestin. The size of the OHCs, as assessed differences in prestin expression should be detectable before by linear capacitance, was unaffected. P10. In contrast, if transduction currents and/or receptor po- Prestin upregulation occurs after the endocochlear potential tentials regulate prestin, the effect should not be seen until later develops. We acknowledge two possible mechanisms by which since the endocochlear potential begins to rise at P10 and bundle stimulation might feedback onto prestin levels. One reaches adult levels at ϳP15 (Schmidt and Fernandez 1963). option is that there is a direct mechanical effect by which Thus, we addressed this question by measuring the develop- forces applied to the bundle in some way alter the mechanics mental time course of prestin expression levels in TectaC1509G of the cell and modulate prestin expression. Another option is mice. that the transduction currents and receptor potential that We patch clamped first-row OHCs in all genotypes at P10, develop within the soma of the OHC in response to the P15, and P20 (Fig. 6, A and B). Linear capacitance and ␣ stimulation modulate prestin expression. We tested these values were not significantly different between the three geno- two possibilities by measuring prestin levels during the types at any age (P Ͼ 0.15 for each measure by two-way

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Fig. 5. TectaC1509G/ϩ OHCs attached to the TM have functional prestin levels that are similar to those of Tectaϩ/ϩ OHCs, whereas TectaC1509G/ϩ and TectaC1509/C1509 OHCs detached from the TM have elevated functional prestin levels. A: representative examples of the whole cell voltage-clamp technique on rows 1, 2, and 3 of OHCs. P, patch pipette. Scale bar ϭ 10 ␮m. B: average capacitance as a function of voltage for OHC in rows 1–3 in Tectaϩ/ϩ, TectaC1509G/ϩ, and TectaC1509/C1509 mice at P30. C: linear capacitance was similar among all rows of each genotype and similar between genotypes. In the TectaC1509G/ϩ mouse,

OHC in rows 2 and 3 (detached from the TM) had significantly elevated nonlinear capacitance, Qmax, and charge density compared with OHCs in row 1 (attached C1509G/C1509G to the TM). Nonlinear capacitance, Qmax, and charge density in rows 2 and 3 were similar to those in Tecta OHCs, and row 1 was similar to those ϩ/ϩ ␣ in Tecta . and V1/2 were similar among all rows of each genotype and between genotypes.

ϩ/ϩ ANOVA), but NLC, V1/2, Qmax, and charge density demon- measure). V1/2 demonstrated age-related changes in Tecta mice strated both age and genotype effects (P Ͻ 0.01 for each (P Ͻ 0.01 by one-way ANOVA) but not in the other two measure by two-way ANOVA). Further analyses revealed that genotypes (P Ͼ 0.2 by one-way ANOVA). This suggests that C1509G/C1509G NLC, Qmax, and charge density were all lower in Tecta there may be an activity-dependent component to development mice compared with Tectaϩ/ϩ and TectaC1509G/ϩ mice at P10 that is slowed in the mutants because of altered stimulation (one-way ANOVA followed by t-tests for each measure). mechanics. However, at every older age (P15, P20, and P30), these values Quantitative real-time PCR was also performed to measure were higher (one-way ANOVA followed by t-tests for each the prestin-to-myosin VIIa ratio (Fig. 6C). Prestin gene expres-

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Fig. 6. Prestin increases after the onset of hearing in OHCs not attached to the TM. A: average capacitance as a function of voltage for row 1 OHCs in Tecta genotypes at different developmental ages. B: whereas linear capacitance was similar among the Tecta genotypes at the different developmental ages, nonlinear capacitance, Qmax, and charge density were lower before the onset of hearing (P10) and increased after the onset of hearing (P15, P20, and P30) in TectaC1509G/C1509G mice. C: prestin mRNA normalized to myosin VIIa mRNA confirmed elevated prestin mRNA at P15 and P30 in TectaC1509G/C1509G mice. *P Ͻ 0.05 (by unpaired t-test). sion was higher in TectaC1509G/C1509G mice compared with potential. Thus, while we cannot rule out a role for signaling ϩ ϩ Tecta / mice at P15 (P ϭ 0.01 by one-way ANOVA fol- between OHCs via reciprocal synapses in type II nerve fibers, lowed by t-tests) and higher in both TectaC1509G/C1509G and our data are consistent with the concept that prestin expression ϩ TectaC1509G/ mice at P30 (P ϭ 0.002 and P ϭ 0.034, within the OHC is locally regulated by the steady-state trans- respectively). Taken together, these data indicate that func- ducer bias current. Blocking the bias current [as occurs when tional prestin expression increases in OHCs not attached to the the tectorial membrane is detached (Xia et al. 2010; Yuan et al. tectorial membrane only after P10 and, thus, after the endoco- 2010)] or not having a bias current (as occurs before the chlear potential develops. development of the endocochlear potential) increases prestin expression. Since both prestin protein and mRNA are potenti- DISCUSSION ated, the regulatory feedback mechanism appears to occur at Given the central role of prestin protein in producing OHC the level of transcription. electromotility, it is reasonable to assume that regulating the The concept that centrally mediated feedback does not amount of prestin within the OHC is important to normal regulate prestin is consistent with previously published data ␣ Ϫ/Ϫ hearing. In the present study, we show that functional prestin demonstrating that 9 mice, in which the receptor that protein can be increased above typically measured levels and plays a dominant role in the efferent olivocochlear signal that this process does not involve centrally mediated efferent transmission (Eybalin 1993; Guinan Jr. and Guinan 2006), feedback. Furthermore, we show that prestin regulation is have normal auditory thresholds and OHC electromotility stimulation dependent, in that OHCs stimulated by the tectorial (Elgoyhen et al. 1994; He et al. 2004; Vetter et al. 1999). membrane have a functional prestin level that is ϳ20% lower However, other efferent neurotransmitters have been identified than those not stimulated by the tectorial membrane. This in the efferent fibers (including GABA, CGRP, and various regulatory feedback mechanism depends on the cochlea being opioids) whose function remains unknown (Altschuler et al., matured to a point where it has developed an endocochlear 1984; Vetter et al. 1991; Fex and Altschuler, 1981 and 1984;

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Maison et al. 2003; Wersinger and Fuchs 2011). Thus, the lack et al. 1982). Interestingly, among mouse cells, neural cells are of an obvious difference in OHC function in ␣9Ϫ/Ϫ mice does 200 times more sensitive to DT than other somatic cells (Allt not completely rule out a role for efferent feedback in regulat- and Cavanagh 1969; Pappenheimer et al. 1982). In addition, ing prestin levels in response to hearing loss regulated by the mRNA that codes for the protein that DT damages in higher brain centers. Unfortunately, the functional role of the humans, eukaryotic elongation factor 2 (Honjo et al. 1971), is efferent system in the sense of hearing remains poorly under- expressed a higher level in IHCs than in OHCs (606 vs. 432) stood. However, our finding that severe hearing loss detected (Liu et al. 2014; National Center for Biotechnology Informa- by higher brain centers does not lead to changes in OHC tion 2014). This may in part underlie the differential suscepti- function, as measured by prestin levels, NLC, and DPOAE bility of adult hair cells to DT. In addition, DT may have other thresholds, is useful in that it suggests that the efferent system effects that were undetected by the studies to assess cochlear has little to no effect in a state of hearing loss. function that we performed. While transgenic mouse models In support of this idea, otoferlinϪ/Ϫ mice have no afferent have been used in which adult utricular hair cells selectively response detectable by ABRs but have normal DPOAEs (Roux express the DT receptor to produce hair cell death in studies of et al. 2006), which argues that OHC electromotility is un- regeneration (Golub et al. 2012), we are unaware of any DT changed in the presence of deafness and presumably a func- studies involving nonhair cell targets in adult mice where Downloaded from tioning efferent pathway. Similarly, centrally mediated feed- hearing has been assessed. To date, all other inner ear studies back via thyroid hormone (TH) may be a critical regulator of involving the administration of exogenous DT have used neo- prestin mRNA production and protein function since absence natal mice (Burns et al. 2012; Cox et al. 2014; Mellado of TH causes deafness and significantly reduces prestin mRNA Lagarde et al. 2013). Thus, as far as we are aware, this is the (Weber et al. 2002). However, electromotility is unaffected in first report of this mouse model of IHC loss. OHCs deprived of TH, indicating that hearing impairment seen Why might a mechanism to modulate prestin levels based on in TH-deficient models may actually be due to morphological stereociliary bundle stimulation have evolved? The primary http://jn.physiology.org/ or physiological changes in the cochlea rather than a problem function of OHCs is to produce mechanical force to amplify regulating prestin (He et al. 2003). the vibration of the cochlear partition (Gao et al. 2014; Oghalai The upregulation of prestin expression shown in previous 2004b). Their stereociliary bundle is fixed to the undersurface studies have been in response to transient changes, such as of the tectorial membrane, unlike the stereociliary bundle of noise exposure or salicylate injection. In TectaC1509G mice, IHCs, which is untethered. To place the OHC stereociliary however, the reduced stimulation of OHCs is inherited. How bundles at the bias point of maximum sensitivity, the center of do they know to upregulate prestin? The simplest explanation the transduction curve, it would be beneficial for the length mechanism of how mechanical stimulation regulates prestin and/or stiffness of the OHC to be adjustable. Both of these by 10.220.33.4 on September 23, 2016 transcription is that regulation occurs individually within each factors are regulated by prestin. For example, the prestinϪ/Ϫ OHC. Reduced steady-state bias currents from lack of tectorial mouse has OHCs that are shorter and less stiff than OHCs from membrane attachment would hyperpolarize the OHC in vivo, wild-type mice (Dallos et al. 2008; Liberman et al. 2002). Over which may increase in prestin levels. There are many examples time, aging, noise trauma, ototoxic exposure, or other factors of activity-dependent protein synthesis, for example, this type often lead to progressive OHC loss, which may alter the static of process is the underlying basis for some forms of long-term position of the stereociliary bundles of residual OHCs. This potentiation (for reviews, see Heise et al. 2014 and Schuman et feedback might then upregulate prestin expression so as to al. 2006). In hair cells, this concept has been noted in organo- return to the proper bias point. Similarly, during development, typic cultures whereby OHC prestin levels decrease in re- such a mechanism may help to increase prestin expression sponse to depolarizing the membrane potential by increasing initially and then turn down prestin expression when the Kϩ concentration in the external solution (Mazurek et al. correct bias point is reached. Thus, an evolutionary mechanism 2011). While the transcription factors Gata-3 and Carf are also for prestin regulation within individual OHCs may have de- modulated by this manipulation, further work is needed to veloped to maintain optimal auditory sensitivity in the face of identify the signal transduction pathway whereby transmem- slow changes over time. brane potential changes modulate prestin transcription. One additional feedback mechanism that should not be overlooked ACKNOWLEDGMENTS is the possibility of regional signaling. Type II nerve fibers Artwork was created by Chris Gralapp. innervate multiple OHCs along a single row, often with recip- rocal synapses (Francis and Nadol Jr. 1993; Spoendlin 1969). Thus, bidirectional communication could create a regional GRANTS feedback mechanism that might play a role in regulating OHC This work was funded by Department of Defense Grant W81XWH-11-2- function and prestin levels. Unfortunately, we cannot test for 0004, National Institute on Deafness and Other Communication Disorders Grants DC-014450, DC-013774, DC-003896, and DC-010363, the Howard this possibility as we are not aware of an animal model where Hughes Medical Institute, and the Stanford Society for Physician Scholars. type II neurons can be eliminated. Finally, the reason why the DT injection preferentially kills DISCLOSURES IHCs in adult CBA mice is unclear. Infection by Corynebac- terium diphtheriae, the bacterium that naturally produces DT, No conflicts of interest, financial or otherwise, are declared by the author(s). is a known cause of sensorineural hearing loss, but the patho- physiology is not known (Schubert et al. 2001). Mouse cells AUTHOR CONTRIBUTIONS are generally highly resistant to DT and can survive 4,000 Author contributions: Y.S., A.X., H.Y.L., A.J.R., and J.S.O. conception and times the DT dose required to kill a human cell (Pappenheimer design of research; Y.S., A.X., H.Y.L., R.W., and J.S.O. performed experi-

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