Harvest of Vestibular End-Organs under Physiologic Conditions during Labyrinthectomy

Here, we describe a technique for harvesting human vestibular end-organs under physiologic conditions during labyrinthectomy and their analysis using immunostaining.

The living human inner ear is challenging to study because it is encased within dense otic capsule bone that limits access to biological tissue. Traditional temporal bone histopathology methods rely on lengthy, expensive decalcification protocols that take 9-10 months and reduce the types of tissue analysis possible due to RNA degradation. There is a critical need to develop methods to access fresh human inner ear tissue to better understand otologic diseases, such as Ménière's disease, at the cellular and molecular level. This paper describes a technique for the harvest of human vestibular end organs from a living donor under physiologic conditions. An individual with Ménière's disease and 'drops attacks' that were refractory to intratympanic gentamicin injection underwent labyrinthectomy. A traditional mastoidectomy was first performed, and the horizontal and superior semicircular canals (SCC) were identified. The mastoid cavity was filled with a balanced salt solution so that the labyrinth could be opened under more physiologic conditions to preserve cellular integrity. A zero-degree endoscope fit with a lens-cleaning sheath irrigation system was used to visualize the submerged mastoid cavity, and a 2 mm diamond burr was used to skeletonize and open the horizontal and superior SCCs, followed by the vestibule. The ampullae and portion of the canal ducts for the superior and lateral SCCs were harvested. The utricle was similarly harvested. Harvested tissue was immediately placed in an ice-cold buffer and then fixed for one hour in 4% paraformaldehyde in phosphate-buffered saline (PBS). The tissue was rinsed several times in 1x PBS and stored for 48 h at 4 °C. The tissue samples underwent immunostaining with a combination of primary antibodies against tenascin-C (Calyx), oncomodulin (streolar hair cells), calretinin (Calyx and Type II hair cells), synaptic vesicle protein 2 (efferent fibers and boutons), β-tubulin 1 (Calyx and afferent boutons), followed by incubation with fluorophore-conjugated secondary antibodies. The tissue samples were then rinsed and mounted for confocal microscopy examination. Images revealed the presence of ampullar and macular hair cells and neural structures. This protocol demonstrates that it is possible to harvest intact, high-quality human inner ear tissue from living donors and may provide an important tool for the study of otologic disease.

The living human inner ear is challenging to study due to its location within the dense otic capsule bone of the temporal bone. Consequently, access to human inner tissue has been limited, and researchers have mainly relied upon post-mortem tissue harvest. Post-mortem temporal bone histopathology (TBH) has been a critical tool for understanding human otologic disease for over 100 years^1,2,3. Tissue for TBH is prepared by the post-mortem harvest of the temporal bone, a lengthy (9-10 month) decalcification and tissue preparation process, followed by hematoxylin and eosin staining. While TBH will remain an essential tool for revealing new information about the healthy and diseased human inner ear, lengthy post-mortem times and long and harsh tissue processing methods limit its utility for certain purposes, necessitating adjunct methods to study human inner ear tissue. High-resolution magnetic resonance imaging can visualize inner ear organs but lacks sufficient resolution to view structures at the cellular or molecular level^4,5. Due to these challenges, many human inner ear diseases remain poorly understood.

An alternative approach is to harvest inner ear tissue during surgery. During labyrinthectomy or translabyrinthine vestibular schwannoma resection, the inner ear tissues are intentionally sacrificed. Utricles harvested from patients during translabyrinthine vestibular schwannoma resection have been used to characterize vestibular hair cell morphology^6,7,8 and study hair cell regeneration^9,10. More recently, techniques have been developed to harvest inner ear organs from organ donors using a transcanal approach that can be used to remove the utricle and potentially other vestibular end organs through a widened oval window with minimal tissue trauma^11,12. Using this technique, it has been possible to characterize single-cell transcriptomic profiles for the human utricle¹³. However, these techniques expose the inner ear organs to non-physiologic conditions during harvest. Specifically, inner ear organs may be exposed to the absence of perilymphatic fluid and submersion in normal saline drill irrigation, which has a substantially different ion composition than perilymphatic fluid. Further, the dehydrated membranous labyrinth is difficult to visualize, even with maximal magnification of the operating microscope, which makes atraumatic surgical dissection challenging. Mechanical trauma may further damage tissue, and our anecdotal experience suggests that surgical tissue is often of insufficient quality of immunostaining due to mechanical damage and cellular degeneration. There is a need for new techniques to atraumatically harvest human inner ear tissue for biological studies that may elucidate poorly understood human inner ear diseases. Here we describe an underwater technique for harvesting human vestibular end-organs under more physiologic conditions during labyrinthectomy and their analysis using immunostaining.

This protocol was developed with the approval of the institutional review board (IRB) of Johns Hopkins University School of Medicine (IRB00203441) and per institutional policies for using human tissue and potentially infectious material. Tissue collection was performed during labyrinthectomy, which is part of standard clinical care for recalcitrant Ménière's disease with drop attacks.

1. Labyrinthectomy and tissue harvest

1. Obtain local institutional review board (IRB) board approval, review institutional policies for the use of human and infectious material, and obtain consent for tissue donation before utilizing this protocol. Perform tissue harvest during surgical labyrinthectomy or translabyrinthine approach to the internal auditory canal, which is part of standard clinical care.
2. Prepare the patient.
   1. Before the induction of anesthesia, discuss with the anesthesia team the avoidance of long-acting paralysis so that the facial nerve may be monitored.
   2. Once general anesthesia is induced, intubate the patient and rotate the operating table 180°. Place facial nerve monitoring electrodes and ensure that pre-operative antibiotic prophylaxis is administered.
   3. Inject 1% lidocaine with 1:100,000 epinephrine into the external auditory canal and postauricular area. Prepare the ear in the standard fashion using betadine.
3. Use a #15 blade to create a standard (5-6 cm) postauricular curvilinear incision 1 cm posterior to the postauricular fold.
4. Lift the postauricular soft tissue.
   1. Identify the temporalis fascia superiorly and elevate the postauricular skin flap anteriorly towards the ear canal.
   2. Create a standard 7-shaped periosteal incision and elevate the periosteum anteriorly until the spine of Henle and the bony ear canal are identified. Place self-retraining retractors.
5. Perform mastoidectomy.
   1. Perform a complete mastoidectomy, identifying the tegmen superiorly, sigmoid sinus posteriorly, and thinning the external auditory canal anteriorly.
   2. Identify the horizontal semicircular canal and the short process of the incus.
      NOTE: The mastoid should not be saucerized to ensure it remains a vessel capable of holding liquid for underwater tissue collection.
6. Identify the semicircular canals (SC).
   1. Under the operating microscope (2-4x magnification), use a 3 mm coarse diamond burr to remove the perilabyrinthine air cells lateral to the superior SC, the subarcuate air cells, and retro-labyrinthine air cells behind the posterior SC, extending superiorly to the common crus.
7. Submerge the labyrinth.
   1. Submerge the mastoid cavity in a balanced salt solution (BSS) and visualize the labyrinth using a zero-degree endoscope with a lens-cleaning sheath irrigation system.
   2. Use the lens-cleaning sheath irrigation system to irrigate the mastoid cavity with BSS, washing away blood and allowing improved visualization of the labyrinth.
8. Blue-line' SCs.
   1. While maintaining an adequate level of BSS in the mastoid cavity and using the endoscope for visualization, use a 3 mm diamond burr to 'blue line' all three semicircular canals by carefully drilling away the otic capsule bone until the semicircular canal appears as a bluish line when viewed with the endoscope.
   2. Irrigate intermittently with BSS solution using the irrigation system to wash away blood and achieve adequate visualization, as illustrated in Figure 1.
9. Harvest SC ampullae.
   1. Under BSS, enter the dome of the lateral semicircular canal and follow this anteriorly until its ampullae are identified. Enter the superior SC and follow this medially towards its ampullae, which may be harvested with the ampullae of the superior SC. Cut the lateral SC duct sharply to facilitate removal.
   2. Elevate the horizontal and superior SC ampullae off the crista using a Rosen needle and separate the afferent fibers from the epithelia and membranous labyrinth. Then, remove these structures.
   3. Transect the dome of the posterior SC and follow this inferiorly and anteriorly to its ampullae. Similarly, harvest the posterior SC ampullae and place all tissue in BSS on ice.
10. Harvest macula.
    1. Remove the bone between the horizontal and posterior SC ampullae to expose the vestibule. The membranous walls of the utricle will have been torn with the removal of the ampullae. So long as a fluid level is maintained, the maculae will remain attached anteriorly; elevate and remove it.
    2. The saccule sits within the spherical recess; sharply elevate and remove it from the recess. Similarly, place the tissue samples in BSS on ice.
11. Transport tissue samples on ice from the operating room to the laboratory for fixation within 1 h of harvesting.

2. Immunohistochemistry and imaging

1. Fix the sample.
   1. Place the tissue sample in 4% paraformaldehyde (PFA) in 1x phosphate-buffered saline (1x PBS) at room temperature (RT) with gentle rocking or orbital shaking (100 rpm) for 1 h.
      ​NOTE: 4% PFA solution (for 5 mL total): 625 mL of 32% PFA stock solution + 500 mL of 10xPBS + 3875 mL of ddH₂O.
2. Transfer tissue samples to 1x PBS solution and rinse 3x 15 min (or longer if necessary) at RT with gentle rocking or orbital shaking.
3. Remove excess connective tissue under the dissecting microscope in 1x PBS at RT.
4. Decalcify the sample.
   1. Transfer tissue samples to 5% ethylenediaminetetraacetic acid (EDTA) in 1x PBS solution to clear the sensory epithelia from fine bone debris and otoconia. Run incubation for 15-30 min at RT, with gentle rocking or orbital shaking.
5. Again, transfer tissue samples to 1x PBS solution and rinse 3x 15 min (or longer if necessary) at RT with gentle rocking or orbital shaking.
6. Apply 125-200 mL of 2x blocking buffer (2x BB) to each sample and incubate for 1.5-5 h at RT or overnight (ON) at 4 °C. Run the incubation under constant orbital shaking.
   ​NOTE: 2x Blocking buffer (2x BB): 1 g of bovine serum albumin (BSA) dissolved in up to 5 mL of 1x PBS supplemented with 0.5% Triton X-100. The BSA serves as the blocking agent for non-specific antigen binding sites.
7. Stain using primary antibodies.
   1. Apply 120-150 mL of a combination of primary antibodies dissolved in 1x BB to the desired dilution. Apply rabbit anti-tenascin-C antibodies at 1:200 dilution (2.5 μg/mL), goat anti-oncomodulin antibodies at 1:200 dilution (2.5 μg/mL), mouse anti-calretinin at 1:300 dilution (2.2 μg/mL), and DAPI at 1:1000 dilution (1 μg/mL). Then, transfer the sample (covered) to 4 °C (fridge or cold room) under gentle orbital shaking for ON incubation.
      ​NOTE: 1x BB: Mix equal volume of 2x BB and 1x PBS supplemented with 0.5% Triton X-100.
8. Rinse tissue samples in 1x PBS solution, 3x 15 min (or longer if necessary), at RT with gentle rocking.
9. Stain using secondary antibody.
   1. Apply 120-150 mL of a combination of fluorophore-conjugated secondary antibodies dissolved in 1x BB at 1:2000 dilution (1 μg/mL). Apply 488 nm conjugated anti-rabbit secondary antibody, 568 nm anti-mouse secondary antibody, and 647 nm anti-goat secondary antibody.
   2. Then, incubate the sample (covered) at RT under gentle orbital shaking for 1.5-2 h (or ON at 4 °C).
10. Rinse tissue samples in 1x PBS solution, 3x 15 min (or longer if necessary), at RT with gentle rocking.
11. Perform microdissection and mounting.
    1. Under the dissecting microscope, remove excess tissue and membranes from around the sensory vestibular epithelia. Use 70% ethanol to clean the surface of a glass histology slide, air dry, and apply a 55 mL drop of the appropriate mounting medium.
    2. Transfer the sensory tissue samples to the mounting medium. Use fine forceps to gently orient the samples with the hair bundle side up in the mounting medium.
    3. Gently lay a cover glass of the appropriate size onto the mounting medium containing the samples, trying to limit air bubbles. Ensure that the cover glass is of #1.5 thickness grade required for confocal microscopy.
    4. Place the slides in a dark area(e.g., a bench drawer) to let the mounting medium set. Seal the cover glass to the slide using clear nail polish and let it dry in a dark area before examining it with the confocal microscope. Afterward, store the sealed slides in the dark (slide box of slide folder) at 4° until imaging.
12. Use a confocal microscope at 40x-100x magnification with 488 nm and 657 nm channels to image the specimen.

Using this technique the human utricle and lateral and superior canal ampullae were harvested intact with minimal trauma (Figure 2). As can be seen in Figure 2, the ampullae can be harvested with a substantial portion of the membranous duct. Immunofluorescent labeling with anti-tenascin-C (extracellular matrix protein) and anti-oncomodulin (small calcium-binding protein of the parvalbumin protein family) showed intact type 1 vestibular hair cells (Figure 3). Hair cell density was 82 per 10,000 μm². These results demonstrate the utility of this technique for harvesting human inner ear tissue with minimal trauma to the neuroepithelium.

Figure 1: Illustration of surgical set-up. Illustration of surgical set-up: The left mastoid cavity is shown with the lateral, superior, and posterior semicircular canals. The mastoid cavity is submerged in balanced salt solution and a zero-degree endoscope is used to visualize the mastoid cavity while a diamond drill is used to open the labyrinth. Please click here to view a larger version of this figure.

Figure 2: Membranous labyrinth. The harvested (A) human utricle and lateral and (B) superior canal ampullae under a light stereomicroscope at 5x magnification. Please click here to view a larger version of this figure.

Figure 3: Utricle. Maximum intensity projections of immunofluorescent labeling of the human utricle at 40x magnification taken with a confocal microscope. Calyx synaptic clefts are labeled with anti-tenascin-C antibodies (green). Type 1 vestibular cells are stained with anti-oncomodulin (red). Scale bar: 50 µm. Please click here to view a larger version of this figure.

This paper describes a new technique for the underwater harvesting of vestibular end organs in BSS using endoscopes and their analysis using immunofluorescent imaging. Here, we demonstrate the harvest of intact vestibular end-organs with intact vestibular hair cells and sufficient tissue quality for successful immunolabeling. The hair cell density in our specimen was similar to those obtained in other studies from live organ donors¹³. To our knowledge, this is the first report of an underwater technique for vestibular end-organ harvesting that allows for the maintenance of near physiologic conditions. This was an adaptation of an underwater technique for superior semicircular canal dehiscence repair that uses BSS to maintain physiologic conditions while the labyrinth is opened for plugging^14,15.

This work expands upon previous techniques developed to harvest vestibular organs during labyrinthectomy or vestibular schwannoma resection^6,7,8 and from living organ donors^11,12. The underwater technique is easily adaptable to vestibular schwannoma resection, during which the labyrinth is similarly opened following a mastoidectomy. The harvest of vestibular end organs from living organ donors has been performed on a limited basis and through a transcanal approach¹². This transcanal approach relies upon removing the tympanic membrane and ossicles and widening the oval window before vestibular end-organ harvesting. Tissue harvest from living organ donors requires working concurrently with the transplant surgery team in a limited space that is shared with anesthesia at the head of the patient's bed. It is unlikely that the technique presented here would be adaptable to the time and space constraints of vestibular end organ harvest during living organ donation. However, this technique offers the advantage of easily exposing and harvesting the ampullae of the semicircular canals, which would be difficult to perform through the oval window.

The study of human inner ear tissue will be critical for elucidating the pathophysiology of inner ear disorders, many of which remain idiopathic. For example, Ménière's disease is commonly encountered in otologic clinics, yet its etiology remains unknown, and there are no proven treatment options^16,17. Techniques such as spatial transcriptomics and immunofluorescent imaging offer an exciting opportunity to shed light on inner ear disease and, so far, have only been utilized on a limited basis. Methods to harvest preserved inner ear organs provide an opportunity to use newer molecular techniques and may offer a useful adjunct to standard TBH. While the method described here has only been utilized for immunofluorescent staining, we believe this tissue would be of sufficient quality for spatial transcriptomics.

The main limitation of this protocol is that it relies on readily available surgical material. Labyrinthectomy is currently an infrequently utilized treatment for Ménière's disease, and even busy referral centers may only perform several of these procedures a year. Translabyrinthine vestibular schwannoma resection is performed more frequently; however, this technique may add time to an already lengthy procedure. We estimate that submersion of the mastoid cavity and opening of the canals underwater add roughly 30 min to surgery. Further, a side-by-side analysis has not yet been performed comparing the quality of tissue obtained using this technique to standard vestibular organ harvest during labyrinthectomy or vestibular schwannoma resection, as it is not possible to perform such an analysis with tissue harvested from one donor. Further research is needed to determine the effect of conditions during organ harvest on tissue quality for microscopy and other techniques.

In summary, we describe a new technique for the underwater harvest of vestibular end organs during labyrinthectomy and its capability of yielding high-quality tissue samples with preserved vestibular hair cells and neural contacts. This technique allows tissue harvest in an environment as close as possible to normal physiologic conditions and may provide a useful tool for studying the human inner ear.

The authors have nothing to disclose.

We thank Mohamed Lehar for his assistance with this project. This work was supported by the National Institute on Deafness and Other Communication Disorders (U24DC020850).