Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea

Summary

This manuscript describes an experimental protocol for evaluating the morphological characteristics and functional status of ribbon synapses in normal mice. The present model is also suitable for noise-induced and age-related cochlear synaptopathy-restricted models. The correlative results of previous mouse studies are also discussed.

Abstract

Cochlear inner hair cells (IHCs) transmit acoustic signals to spiral ganglion neurons (SGNs) through ribbon synapses. Several experimental studies have indicated that hair cell synapses may be the initial targets in sensorineural hearing loss (SNHL). Such studies have proposed the concept of cochlear "synaptopathy", which refers to alterations in ribbon synapse number, structure, or function that result in abnormal synaptic transmission between IHCs and SGNs. While cochlear synaptopathy is irreversible, it does not affect the hearing threshold. In noise-induced experimental models, restricted damage to IHC synapses in select frequency regions is employed to identify the environmental factors that specifically cause synaptopathy, as well as the physiological consequences of disturbing this inner ear circuit. Here, we present a protocol for analyzing cochlear synaptic morphology and function at a specific frequency region in adult mice. In this protocol, cochlear localization of specific frequency regions is performed using place-frequency maps in conjunction with cochleogram data, following which the morphological characteristics of ribbon synapses are evaluated via synaptic immunostaining. The functional status of ribbon synapses is then determined based on the amplitudes of auditory brainstem response (ABR) wave I. The present report demonstrates that this approach can be used to deepen our understanding of the pathogenesis and mechanisms of synaptic dysfunction in the cochlea, which may aid in the development of novel therapeutic interventions.

Introduction

Frequencies in the range of approximately 20‒20,000 Hz can be perceived as auditory stimuli by humans. Human hearing is normally most sensitive near 1,000 Hz, where average sound pressure level is 20 μPa in young adults (i.e., 0 decibels of sound pressure level [dB SPL]). In some pathological conditions, hearing loss is restricted to specific frequencies. For example, in the early stages of noise-induced hearing loss (NIHL), a “notch” (i.e., hearing threshold elevation) can be observed in the audiogram at 4 kHz¹. Along the mammalian cochlear partition, its gradations of stiffness and mass produce an exponential frequency map, with high-frequency sound detection at the base of the cochlea and low-frequency detection at the apex². Indeed, there is a cochlear place-frequency map along the basilar membrane, leading to what is known as tonotopic organization^2,3. Each given place on the basilar membrane has the highest sensitivity to only one particular sound frequency, which is usually termed the characteristic frequency^3,4, although responses to other frequencies can also be observed.

To date, various mouse models have been employed to investigate normal function, pathological processes, and therapeutic efficacy in the auditory system. Precise knowledge of physiological parameters in the mouse cochlea is a prerequisite for such studies of hearing loss. The mouse cochlea is anatomically divided into apical, middle, and basal turns, which correspond to different frequency regions. By labeling auditory nerve afferents at the cochlear nucleus to analyze their corresponding peripheral innervation sites in the cochlea, Müller et al. succeeded in establishing the cochlear place-frequency map in the normal mouse in vivo⁵. In the interval of 7.2–61.8 kHz, which corresponds to positions between 90% and 10% of the full length of the basilar membrane, the mouse cochlear place-frequency map can be described by a simple linear regression function, suggesting a relation between the normalized distance from the cochlear base and the logarithm of the characteristic frequency⁵. In laboratory mice, the place-frequency map can be used to explore the relationship between hearing thresholds within specific frequency ranges and cochleograms showing the numbers of missing hair cells in relative regions along the basilar membrane⁶. Importantly, the place-frequency map provides a positioning system for the investigation of minimal structural damage, such as damage to the ribbon synapses of hair cells at specific cochlear frequency locations in mice with peripheral auditory trauma^7,8.

In the mammalian cochlea, ribbon synapses are comprised of a presynaptic ribbon, an electron-dense projection that tethers a halo of release-ready synaptic vesicles containing glutamate within the IHC, and a postsynaptic density on the nerve terminal of the SGN with glutamate receptors⁹. During cochlear sound transduction, deflection of the hair cell bundle results in IHC depolarization, which leads to glutamate release from IHCs onto the postsynaptic afferent terminals, thereby activating the auditory pathway. Activation of this pathway leads to the transformation of sound-induced mechanical signals into a rate code in the SGN¹⁰. Indeed, the IHC ribbon synapse is highly specialized for indefatigable sound transmission at rates of hundreds of Hertz with high temporal precision, and is of critical importance for presynaptic mechanisms of sound encoding. Previous studies have revealed that ribbon synapses vary greatly in size and number at different frequency regions in the adult mouse cochlea^11,12, likely reflecting structural adaptation to the particular sound coding for survival needs. Recently, experimental animal studies have demonstrated that cochlear synaptopathy contributes to multiple forms of hearing impairments, including noise-induced hearing loss, age-related hearing loss, and hereditary hearing loss^13,14. Thus, methods for identifying correlated changes in synaptic number, structure, and function at specific frequency regions have been increasingly employed in studies of auditory development and inner ear disease, using models generated via experimental manipulation of genetic or environmental variables^15,16,17.

In the current report, we present a protocol for analyzing the synaptic number, structure, and function at a specific frequency region of the basilar membrane in adult mice. Cochlear frequency localization is performed using a given place-frequency map in combination with a cochleogram. The normal morphological characteristics of cochlear ribbon synapses are evaluated via presynaptic and postsynaptic immunostaining. The functional status of cochlear ribbon synapses is determined based on the suprathreshold amplitudes of ABR wave I. With minor alterations, this protocol can be used to examine physiological or pathological conditions in other animal models, including rats, guinea pigs, and gerbils.

Protocol

All procedures were carried out in accordance with the NRC/ILAR Guide for the Care and Use of Laboratory Animals (8th Edition). The study protocol was approved by the Institutional Animal Care and Use Committee of Capital Medical University, Beijing, China.

1. Animal Selection

1. For all experiments, use adult C57BL/6J male mice (8 weeks old) as the animal model.
   NOTE: C57BL/6J mice carrying a splice variant of the Cdh23 exhibit accelerated senescence in the auditory system, reflected as a 40% loss of ribbon synapses at the basal turn of the cochlea and a 10% loss at the middle turn by 6 months of age, followed by a rapid increase of this loss in the whole cochlea with age^18,19. Thus, we advise caution when using C57BL/6J mice older than 6 months for auditory research. Other strains of mice can be used depending on specific experimental aims.
2. Inspect the mice using a professional diagnostic pocket otoscope to rule out outer or middle ear pathologies prior to hearing assessments. Potential signs may include fluid or pus in the external auditory canal, redness and swelling in local tissue, and tympanic membrane perforation.
   NOTE: Although this is rarely encountered, once identified, mice with outer or middle ear diseases should be excluded.

2. Hearing Assessment

1. Anesthetize mice using an intraperitoneal injection of a mixture of ketamine hydrochloride (100 mg/kg) and xylazine hydrochloride (10 mg/kg). Judge the depth of anesthesia via painful stimuli (e.g., toe-pinch reflex).
   NOTE: When the toe-pinch reflex is completely absent, the animal has reached an adequate depth of anesthesia for auditory testing. If more time is required for bilateral ABR recordings, administer a lower dosage (one-fifth the original dosage) of anesthetics to restore the original anesthetic plane. Take care to avoid anesthetic overdose, as this may lead to death in mice.
2. Maintain the anesthetized animal’s body temperature at 37.5 °C using a thermoregulating heating pad. Place the anesthetized animal in an electrically and acoustically shielded room to avoid interference throughout the hearing test.
   NOTE: Maintain the physiological temperature during the entire procedure until the animal is totally awake, in order to prevent death caused by post-anesthesia hypothermia.
3. Place subdermal needle electrodes (20 mm, 28 G) at the vertex of the skull (recording electrode), in the ipsilateral parotid region below the pinna of measured ear (reference electrode), and in the contralateral parotid region (ground electrode), with a depth of 3 mm under the skin of the mouse head, respectively²⁰.
4. Use a closed-field speaker to perform acoustic stimulation via a 2 cm plastic tube with a cone-shaped tip. Fit the tip into the external ear canal²¹.
   NOTE: Ensure that the electrical impedance in the recording and reference electrodes is less than 3 kOhm (usually 1 kOhm). If the impedance is high, change the insertion site of the electrode, clean the electrode with alcohol, or replace the electrode to avoid alterations in ABR wave amplitude.
5. For ABR recording, generate tone pips (3 ms duration, 1 ms rise/fall times, at a rate of 21.1/s, frequency: 4–48 kHz) and present them at decreasing SPLs from 90 to 10 dB in 5‒10 dB SPL steps²⁰. During this step, the responses are amplified (10,000 times), filtered (0.1–3 kHz), and averaged (1,024 samples/stimulus level).
   NOTE: ABRs are collected for each stimulus level in 10 dB steps, with additional 5 dB steps near the threshold.
6. At each frequency, determine the ABR threshold, which refers to the minimal SPL resulting in a reliable ABR recording with one or more distinguishable waves that can be clearly identified by visual inspection (Figure 1).
   NOTE: It is usually necessary to repeat the process for low SPLs around the threshold to ensure the consistency of the waveforms. The response threshold is the lowest stimulus level at which the waveform can be observed, when a decrease of 5 dB would lead to the disappearance of the waveform.

3. Cochlear Tissue Processing

1. After ABR recording, euthanize the anesthetized mice via cervical dislocation, decapitate them, expose the bulla from the ventral side, and open with sharp scissors to gain access to the cochlea.
2. Using a fine forceps, remove the temporal bones, sever the stapes artery, remove the stapes from the oval window, and rupture the round window membrane. Make a small hole at the apex of the cochlea by gently rotating the tip of a needle (13 mm, 27 G).
3. Fix the isolated bones with 4% (wt/vol) paraformaldehyde in 0.1 M phosphate-buffered saline (PBS, pH 7.4) overnight at 4 °C. Using a fine-tipped pipette, gently flush fixative through the perilymphatic spaces via application to the oval or round windows (as an inlet) and the opening at the apex (as an outlet).
   NOTE: Some proteins require a short duration of fixation to avoid destruction of their epitopes for immunolabeling. In such cases, incubate bones in 4% paraformaldehyde at room temperature (RT) for 2 h, depending on the manufacturer’s instructions for immunohistochemistry. Fixation can also be performed via cardiac perfusion to remove the cochlear blood, avoiding background noise due to non-specific staining at later stages, particularly in mouse models of cochlear synaptopathy.
4. Rinse the bones three times for 5 min with 0.1 M cold PBS to remove residual paraformaldehyde. Decalcify the bones with 10% ethylenediaminetetraacetic acid (EDTA) either at RT for 4 h or at 4 °C for 24 h via gentle shaking in a horizontal shaker at 20 rpm. EDTA can be refreshed midway.
   NOTE: Decalcification times are subject to the concentration of EDTA and users’ preferences. Decalcified tissue should maintain a certain degree of toughness, which facilitates the manipulation of isolating cochlear whole-mounts at later steps. Temporal bones can be decalcified in 10% EDTA with rotation, allowing researchers to leave the laboratory following ABR tests and fixation experiments. The decalcification time is flexible within the range of 20 to 30 h at 4 °C.
5. Transfer one decalcified temporal bone from EDTA to 0.1 M PBS. Use #3, #5 Dumont forceps and the 27 G needle to dissect the apical, middle and basal cochlear regions in turn and then dissect the cochlea out of the bone under a stereo dissection microscope (as previously described²²). Make a series of small cuts along the spiral ligament using a razor blade, and remove the tectorial membrane and Reissner’s membrane (Figure 2).
   NOTE: As long as the dissected cochlea is intact, this process can be modified according to the individual operator's usual protocol.
6. Further dissect the remaining auditory epithelium including the spiral limbus into individual cochlear turns (apex, middle, and base with hook region) for whole-mount preparations.
7. Under a 40x oil objective of a light microscope, measure the basilar membrane length with a 250 μm scale placed in the eyepiece, which can be adjusted along the stereocilia of the IHCs.
8. Calculate the length of each cochlear turn by adding all segment lengths (250 μm per segment), and concomitantly obtain the total length of the basilar membrane by summing the lengths of each turn.
9. Convert the total length of the basilar membrane including the hook region into a percentage based on distance from the cochlear apex (0% refers to the cochlear apex, 100% to the cochlear base).
10. Convert this distance into the cochlear characteristic frequency using a logarithmic function (d(%) = 1 - 156.5 + 82.5 × log(f), with a slope of 1.25 mm/octave of frequency, where d is the normalized distance from the cochlear apex in percent, f is the frequency in kHz), as previously described^5,6. Thus frequency range in a corresponding area of the basilar membrane on each cochlear turn can be acquired.

4. Immunofluorescence Staining

1. After dissection, place each cochlear turn in a separate 2.5 mL centrifuge tube and incubate cochlear turns in 10% goat serum/PBS/0.1% Triton X-100 for 1 h at RT on a rotator.
2. Remove the above blocking/permeabilization solution from each tube using a 200 μL pipette tip under a dissection microscope and incubate the specimens with primary antibodies diluted in 5% goat serum/PBS/0.1% Triton X-100 overnight at 4 °C on a rotator.
   NOTE: For immunolabeling of cochlear synaptic ribbons, use the presynaptic marker mouse anti-carboxyl-terminal binding protein 2 IgG1 (CtBP2, labeling the B domain of the RIBEYE scaffolding protein, 1:400) and the postsynaptic marker mouse anti-glutamate receptor 2 IgG2a (GluR2, labeling a subunit of the AMPA receptor, 1:200)²³.
3. Rinse three times for 5 min with 0.1 M cold PBS to remove residual primary antibodies and incubate the specimens with secondary antibodies diluted in 5% goat serum/PBS/0.1% Triton X-100 at RT for 2‒3 h in darkness on a rotator.
   NOTE: Prepare appropriate secondary antibody mixtures using goat anti-mouse Alexa Fluor 568 (IgG1, 1:500) and goat anti-mouse Alexa Fluor 488 (IgG2a, 1:500), which are complementary to the primary antibodies used in step 4.2. To improve labeling efficiency for synaptic ribbons, we recommend selecting specific secondary antibodies. Some labs extend incubation with secondary antibodies to increase GluR2 immunolabeling²⁴.
4. Rinse three times for 5 min with 0.1 M PBS to remove residual secondary antibodies and transfer the specimens from 2.5 mL centrifuge tubes to 35 mm plates containing 0.1 M PBS.
5. Place a drop of mounting medium containing 4’,6-diamidino-2-phenylindole (DAPI) onto the slide and transfer the specimens from PBS to the mounting medium. Place one edge of a coverslip on the slide and release to let the coverslip fall gently.
   NOTE: To ensure that the hair cells face upward and that no folding or twisting of cochlear specimens occurs during the procedure, mount cochlear specimens under a stereo dissection microscope.
6. Place the slides in a slide box at 4 °C overnight to let slides dry and then image under a laser confocal microscope.

5. Morphological Evaluation of Cochlear Ribbon Synapses

1. Image slides using a confocal microscope with three lasers—a 405 nm UV diode, a 488 nm argon laser, and a 561 nm diode-pumped solid-state (DPSS) laser to excite DAPI (Excitation spectrum 409–464 nm), Alexa Fluor 488 (Excitation spectrum 496–549 nm) and Alexa Fluor 568 (Excitation spectrum 573–631 nm), respectively.
2. Acquire confocal z-stacks over a distance of 8 μm from each cochlear turn using a 63x high-resolution oil immersion lens.
   ​NOTE: Once defined, all parameters for digitizing photomicrographs should be saved and applied uniformly to all slides.
3. For synaptic punctum counts, set the z-stacks (0.3 μm step size) to span the entire length of IHCs, thereby ensuring that all synaptic puncta can be imaged.
4. Merge the images containing puncta in a z-stack to obtain the z-axis projection, and import to the image-processing software.
5. Divide synaptic total counts in each z-stack at specific frequency regions by the number of IHCs (equal to the DAPI nuclear manual counts) to calculate the number of synaptic puncta for each IHC. At each specific frequency region, average all synaptic puncta in three images of different microscopic fields containing 9–11 IHCs.
   1. Outline the region of interests (ROIs) including the basolateral regions of each IHC using freehand selections button. Use the measure function for automatic quantification of puncta, and the watershed function to distinguish between closely adjacent spots.
   2. After each automated counting, perform visual inspections with manual corrections to ensure puncta counting reliable.
      NOTE: Experimenters should remain blinded as to whether the slide is from the apex, middle, or basal turn of the cochlea.
6. Visually assess synaptic structure and distribution, to manually isolate individual IHCs from their neighbors by the Pencil Tool (M) to better visualize the cytoskeletal architecture and synaptic localization.
7. To inspect the juxtaposition of presynaptic ribbons (CtBP2) and postsynaptic receptor patches (GluR2), extract the voxel space around ribbon by the Rectangular Marquee Tool and isolate individual ribbon by Crop. Through clicking Image > Image Size, acquire a thumbnail array of these miniature projections, which can then be used to identify paired synapses (appeared as closely juxtaposed pairs of CtBP2-positive and GluR2-positive puncta) versus orphan ribbons (lacking postsynaptic glutamate receptor patches) (Figure 3).
   NOTE: Normal cochlear synapses appear as combined immunolabeling of the presynaptic ribbon within the hair cell (anti-CtBP2) and the postsynaptic glutamate receptor patch on the auditory nerve terminal (anti-GluR2)²⁵. Some labs use confocal projections in conjunction with 3D modeling to quantify synaptic patch size or volume^26,27. Prior to significant loss of ribbon synapses, ribbons exhibiting changes in size or without paired glutamate receptor patches are likely indicative of synaptic dysfunction^27,28.

6. Functional Evaluation of Cochlear Ribbon Synapses

1. Collect all ABR waves for each frequency stimulus presented at an SPL of 90 dB for the analysis of suprathreshold ABR wave I amplitudes.
   NOTE: Neurophysiological and morphological studies have demonstrated that low spontaneous-rate, high-threshold fibers are especially vulnerable to aging and noise exposure^29,30. Although the simple loss of ribbon synapses cannot affect ABR thresholds, it commonly results in significant reductions in ABR wave I amplitudes, because these afferents including low spontaneous-rate, high-threshold fibers and high spontaneous-rate, low-threshold fibers contribute heavily to the summed activities of cochlear nerve fibers^28,29,31. A suprathreshold intensity of 90 dB SPL is selected here.
2. Determine peak-to-peak wave I amplitude using an offline analysis program (Figure 4). Each wave I in the ABR test consists of a starting positive (p) deflection and the subsequent negative (n) deflection. ABR wave I amplitude is defined as the difference in voltage between Ip (the positive peak of wave I) and In (the negative peak of wave I)²⁹.
   NOTE: In pathological conditions, cochlear synaptopathy can be determined based on the suprathreshold amplitudes of ABR wave I, which reflect the summed onset responses of SGNs evoked by sound. However, cochlear sensitivity that is not compromised due to OHC dysfunction is a prerequisite for this method.

Results

ABR hearing tests were performed for 10 C57BL/6J mice (8 weeks of age) under anesthesia. ABRs were elicited using tone burst stimuli at 4, 8, 16, 32, and 48 kHz. The hearing threshold of each animal was visually detected by distinguishing at least one clear waveform in the ABR. All mice exhibited ABR thresholds in response to tone bursts, ranging between 25 and 70 dB SPL depending on the frequency of the stimulus. Our results indicated that the hearing threshold was lowest at 16 kHz (Figure 1

Discussion

Since cochlear synaptopathy was first characterized in adult mice with a temporary threshold shift (TTS) induced by 8‒16 kHz octave band noise at 100 dB SPL for 2 h³¹, researchers have increasingly investigated the effects of synaptopathy in various mammals, including monkeys and humans^32,33. In addition to noise exposure, several other conditions have been associated with cochlear synaptopathy (e.g., aging, the use of ototoxic drugs...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Ketamine hydrochloride	Gutian Pharmaceutical Co., Ltd., Fujian, China	H35020148	100 mg/kg
Xylazine hydrochloride	Sigma-Aldrich, St. Louis, MO, USA	X-1251	10 mg/kg
TDT physiology apparatus	Tucker-Davis Technologies, Alachua, FL, USA	Auditory Physiology System III
SigGen/BioSig software	Tucker-Davis Technologies, Alachua, FL, USA	Auditory Physiology System III
Electric Pad	Pet Fun	11072931136
Dumont forceps 3#	Fine Science Tools, North Vancouver, B.C., Canada	0203-3-PO
Dumont forceps 5#	Fine Science Tools, North Vancouver, B.C., Canada	0209-5-PO
Stereo dissection microscope	Nikon Corp., Tokyo, Japan	SMZ1270
Goat serum	ZSGB-BIO, Beijing,China	ZLI-9021
Anti-glutamate receptor 2, extracellular, clone 6C4	Millipore Corp., Billerica, MA, USA	MAB397	mouse
Purified Mouse Anti-CtBP2	BD Biosciences, Billerica, MA, USA	612044	mouse
Alexa Fluor 568 goat anti-mouse IgG1antibody	Thermo Fisher Scientific Inc., Waltham, MA, USA	A21124	goat
Alexa Fluor 488 goat anti-mouse IgG2a antibody	Thermo Fisher Scientific Inc., Waltham, MA, USA	A21131	goat
Mounting medium containing DAPI	ZSGB-BIO, Beijing,China	ZLI-9557
Confocal fluorescent microscopy	Leica Microsystems, Wetzlar, Germany	TCS SP8 II
Image Pro Plus software	Media Cybernetics, Bethesda, MD, USA	version 6.0
Professional diagnostic pocket otoscope	Lude Medical Apparatus and Instruments Trade Co., Ltd., Shanghai,China	HS-OT10
Needle electrode	Friendship Medical Electronics Co., Ltd., Xi'an,China	1029	20 mm, 28 G
Closed-field speaker	Tucker-Davis Technologies, Alachua, FL, USA	CF1