Isolation and Culture of Primary Cochlear Hair Cells from Neonatal Mice

Summary

Herein, we present a detailed protocol for isolating and culturing primary cochlear hair cells from mice. Initially, the organ of Corti was dissected from neonatal (aged 3-5 days) murine cochleae under a microscope. Subsequently, cells were enzymatically digested into a single-cell suspension and identified using immunofluorescence after several days in culture.

Abstract

Cochlear hair cells are the sensory receptors of the auditory system. These cells are located in the organ of Corti, the sensory organ responsible for hearing, within the osseous labyrinth of the inner ear. Cochlear hair cells consist of two anatomically and functionally distinct types: outer and inner hair cells. Damage to either of them results in hearing loss. Notably, as inner hair cells cannot regenerate, and damage to them is permanent. Hence, in vitro cultivation of primary hair cells is indispensable for investigating the protective or regenerative effects of cochlear hair cells. This study aimed to discover a method for isolating and cultivating mouse hair cells.

After manual removal of the cochlear lateral wall, the auditory epithelium was meticulously dissected from the cochlear modiolus under a microscope, incubated in a mixture consisting of 0.25% trypsin-EDTA for 10 min at 37 °C, and gently suspended in culture medium using a 200 µL pipette tip. The cell suspension was passed through a cell filter, the filtrate was centrifuged, and cells were cultured in 24-well plates. Hair cells were identified based on their capacity to express a mechanotransduction complex, myosin-VIIa, which is involved in motor tensions, and via selective labeling of F-actin using phalloidin. Cells reached >90% confluence after 4 d in culture. This method can enhance our understanding of the biological characteristics of in vitro cultured hair cells and demonstrate the efficiency of cochlear hair cell cultures, establishing a solid methodological foundation for further auditory research.

Introduction

Cochlear hair cells play important roles in sound detection and signal transmission to the auditory nerve. Hair cells are mechanistic cells that function as primary sensory receptors and convert sound vibrations into electrical signals in vertebrates. The sensory epithelium of the mammalian inner ear comprises a single row of inner hair cells and three rows of outer hair cells. In different basic membrane areas, hair cells perceive sounds at different frequencies (between 20 and 2,000 Hz)¹. The function of outer hair cells is an active mechanical amplification process that helps fine-tune the mammalian inner ear, conferring high sensitivity to sound. Inner hair cells are responsible for detecting sounds. After graded depolarization, acoustic information is transmitted to the brain through the auditory nerve fibers².

Hearing loss may be caused by genetic defects, aging, noise trauma, or the excessive use of ototoxic drugs, which constitute a major health concern worldwide^3,4. Hearing loss mainly results from irreversible damage to hair cells⁵. Regarding noise-induced hearing loss, although researchers have reached a consensus on several details of its etiology, a comprehensive understanding of the numerous underlying mechanisms is lacking. Outer hair cells are particularly vulnerable to acoustic overexposure⁶. Mechanosensitive cochlear hair cells are involved in age-related hearing loss; however, the molecular and cellular mechanisms underlying hair cell degeneration remain unknown. Several changes in the molecular processes lead to hair cell aging, oxidative stress, DNA damage response, autophagy, and dysregulation of the expression and transcription of genes related to hair cell specialization⁷.

As the inner ear is encased in the temporal bone, deep in the hardest bone of the body, it is experimentally inaccessible, posing a challenge to investigations into the mechanisms of hair cell repair and regeneration. Hence, establishing in vitro cultures for investigating the function of hair cells has become an ideal method for research on the regeneration and injury mechanisms of the inner ear. The procedures for preparing cochlear organotypic cultures have been described in earlier studies^8,9,10. Investigators worldwide have employed various cochlear microdissection and surface preparation techniques. Despite the persistent challenges, various primary hair cell culture systems have been successfully established in vitro. Cochlear organ cultures contain various cell types, including hair cells, Deiters cells, Hensen's cells, pillar cells, and auditory nerve fibers. An in-depth understanding of the changes in hair cells at the cellular and molecular levels after injury will enable the development of more powerful research tools. This study aimed to demonstrate the steps for isolating cochlear organs from neonatal mice and enzymatically detaching the abundant hair cells for in vitro studies. The nature of the cultured cells was confirmed using immunofluorescence staining.

Protocol

All animal experiments were approved (No. 2021-847) by the Xi'an Jiaotong University Committee on the Use and Care of Animals.

1. Sterilization and material preparation

1. Sterilize the dissection tools using high-temperature and high-pressure steam disinfection and dry them in a 50 °C incubator overnight.
2. Prepare 100 mL of the culture medium containing 10% fetal bovine serum (FBS) and 10 mg/mL penicillin/streptomycin (add 10 mL of FBS and 10 µL of penicillin stock solution to 90 mL of Dulbecco's Modified Eagle Medium (EMEM))in advance and store at 4 °C.

2. Dissection and removal of the temporal bone for collection of auditory epithelia

1. Euthanize a total number of 10 newborn mice (aged between P3 and P5) by decapitation on ice. Use an alternative, ethically approved methodology, if required.
2. Hold the head in place, open the scalp along the sagittal suture using micro-operating scissors, and separate and fix the scalp bilaterally with the fingers, as shown in Figure 1A.
3. Remove the brain using a small periosteal elevator and bisect the basal skull. Cut the cranium with scissors and flip forward to open the skull; use the tip of the scissors to scrape the brain and expose the base of the skull.
4. Observe the bilateral temporal bones at the base of the skull (Figure 1C). Use scissors to cut the base of the skull along the midline, scrape away the skin, and remove any unnecessary bone (Figure 1D,D').
5. Retain and transfer the temporal bones into 35 mm sterile Petri dishes containing fresh Hank's balanced salt solution (HBSS) (Figure 1E).
6. Use two #5-pointed forceps to remove the bulla and surrounding tissue from the petrous portion of the temporal bone (Figure 1E).
   NOTE: At this stage, the bony labyrinth in the temporal bone of the mice would not be completely calcified and would be easily dissected using forceps.
7. Hold the forceps with one hand to fix the semicircular part of the temporal bone and stick the lower foot of the forceps into the round window niche with the other hand to separate the lateral bone of the cochlea from the scala vestibule. Carefully remove the petrous portion of the temporal bone without touching the OC epithelium. Subsequently, carefully separate and remove the bony labyrinth of the cochlea from the basal end to the apical end (Figure 1F).
8. Carefully micro-isolate the organ of the Corti sensory epithelium from the modiolus using #5-pointed forceps (Figure 1G).
9. Hold the spiral ligament, carefully separate it from the stria vascularis with micro-operating forceps, and transfer the clean auditory epithelium to a 3 mm sterile culture dish containing HBSS using a 200 µL pipette (Figure 1H).
10. Collect 20 specimens from each animal and quickly transfer them to a 100 mm sterile Petri dish containing HBSS for the next preparation step (Figure 1).

3. Enzymatic disaggregation for obtaining auditory hair cells

1. Transfer the auditory epithelium to 10 mL of fresh DMEM containing 0.25% trypsin and incubate at 37 °C for 12 min.
2. Using a 200 µL pipette tip, gently separate the hair cells from the basal lamina and other cells under an operating microscope.
3. Add another 10 mL of culture medium to inhibit the disaggregation.
4. Filter the suspended cells in the culture medium through a 70 µm filter, collect the filtrate in a clean 50 mL tube, and centrifuge it at 300 × g for 5 min.
5. Resuspend the hair cells in at least 5 mL of culture medium by gently pipetting them up and down using a 1,000 µL pipette tip. Avoid introducing bubbles.
6. Place a coverslip at the bottom of a six-well plate in advance. Count the cells and culture them at a density of 10⁶ cells/mL in six-well plates.
7. Grow the adherent cells in 2 mL of DMEM (containing 10% of FBS, 100 units/mL of penicillin, and 100 µg/mL of streptomycin) at 37 °C and 5% CO₂. Change the culture medium every day.
   ​NOTE: It should be mentioned that using this protocol does not result in obtaining pure hair cells. Based on this primary cell culture method, we recommend using d2 or d3 cells for further studies, as hair cells might constitute approximately 70% of cells in culture and be in a good state.

4. Immunofluorescent staining

1. Harvest the cultured cells from d1 to d6 (one well per day). Aspirate the culture medium and rinse cells 2x with phosphate-buffered saline (PBS).
2. Fix cells with 4% paraformaldehyde for 15 min at room temperature (RT).
3. Remove the fixative and rinse cells for 3 x 3 min with PBS.
4. Permeabilize the cells with PBS containing 0.2% Triton X-100 for 10 min at RT.
5. Incubate the permeabilized cells with a blocking solution consisting of 10% FBS in PBS for 20 min at RT.
6. Stain the cells with anti-myosin monoclonal antibody (diluted at 1:200 in PBS) at 4 °C overnight.
7. Wash the cells 3x with sterile PBS and incubate them with secondary antibody (Alexa Fluor 594 goat anti-rabbit MYO7A, diluted 1:500 in PBS) and fluorescein-labeled phalloidin (Alexa Fluor 488, to identify cell structure) for 2 h at RT.
8. Rinse the cells 3x with PBS to remove the secondary antibodies.
9. Add 1-2 drops of mounting medium with DAPI onto the slides, mount the coverslips, and place them under a laser scanning confocal microscope to capture photos of the cells.

5. Statistical analysis

1. Perform two-way analysis of variance (ANOVA) followed by Tukey's post hoc tests to analyze the changes in the grey values of myosin-VIIa and phalloidin over time. Use the letter marking method to mark statistical differences.
2. Perform additional one-way ANOVAs followed by Tukey's post-hoc tests to compare the phalloidin-positive cell ratio between cell samples from day 1 to day 6 and the myosin-VII-positive cell ratio on the same days.

Results

Following this protocol, we seeded the isolated cells. Primary cochlear hair cell seeds were considered successful if the cells did not float in the culture medium and spread within 24 h. We determined the number of hair cells after they adhered and spread into flat aggregates at the bottom of the dish. After 1 day, live hair cells were tightly adhered to the bottom of the culture dish and non-adherent cells were removed by rinsing with PBS. Typically, the number of cells doubled after 3 d of culture (

Discussion

Compared with the HEI-OC1 cell line, primary cultures of hair cells more accurately replicated the physiological state of cells in vivo. Therefore, the auditory primary culture method established by isolating living cells from cochlear organs and immediately culturing them appears to be a valuable tool for extensive research on auditory systems. Certain techniques are crucial for a successful culture. First, minimizing the duration of separation of the organ of Corti from the temporal bone enhances the likelihood of sust...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
100 mm BioLite cell culture dish	Thermo Fisher Scientific	130182	using for culture
35 mm Nunc cell culture dish	Thermo Fisher Scientific	150318	using for culture
6-well palate	Thermo Fisher Scientific	310109005	using for culture
70 µm cell strainers	BD Company	352350	using for filter
Alexa Fluor 488 Phalloidin	Thermo Fisher Scientific	A12379	immunofluorescent staining
Anti-rabbit IgG Alexa Fluor 488	Thermo Fisher Scientifc	A11008	immunofluorescent staining
day 3-5 neonatal murine			provided by Xi'an Jiaotong University
Dulbecco’s Modified Eagle Medium	Thermo Fisher Scientific	11965092	using for culture
Fetal Bovine Serum	Thermo Fisher Scientific	12483020	using for culture
Forceps	Dumont	5#	using for dissection
Leica anatomy microscope	Germany	S9i	using for dissection
Penicillin/streptomycin	Thermo Fisher Scientific	15140-122	using for culture
Rabbit plyclonal to Myosin VIIa	Abcam company	ab92996	immunofluorescent staining
Scissor	Belevor	10cm/04.0524.10	using for dissection
Triton X-100	Sigma Aldrich	9036-19-5	immunofluorescent staining
Trypsin	Thermo Fisher Scientific	25200072	using for culture