Evaluation of Planar-Cell-Polarity Phenotypes in Ciliopathy Mouse Mutant Cochlea

Summary

Primary cilia influence various signaling pathways. The mammalian cochlea is ideal for examining planar cell polarity (PCP) signaling. Cilia dysfunction affects cochlear outgrowth, cellular patterning and hair cell orientation, readouts of PCP. Our goal is to analyze PCP signaling in mouse cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy.

Abstract

In recent years, primary cilia have emerged as key regulators in development and disease by influencing numerous signaling pathways. One of the earliest signaling pathways shown to be associated with ciliary function was the non-canonical Wnt signaling pathway, also referred to as planar cell polarity (PCP) signaling. One of the best places in which to study the effects of planar cell polarity (PCP) signaling during vertebrate development is the mammalian cochlea. PCP signaling disruption in the mouse cochlea disrupts cochlear outgrowth, cellular patterning and hair cell orientation, all of which are affected by cilia dysfunction. The goal of this protocol is to describe the analysis of PCP signaling in the developing mammalian cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy. Defects in convergence and extension are manifested as a shortening of the cochlear duct and/or changes in cellular patterning, which can be quantified following dissection from developing mouse mutants. Changes in stereociliary bundle orientation and kinocilia length or positioning can be observed and quantitated using either immunofluorescence or scanning electron microscopy (SEM). A deeper insight into the role of ciliary proteins in cellular signaling pathways and other biological phenomena is crucial for our understanding of cellular and developmental biology, as well as for the development of targeted treatment strategies.

Introduction

Primary cilia are long microtubule-based appendages that extend from the surface of most mammalian cells. Primary cilia are often confused with motile cilia, of which there are always multiple per cell, and whose purpose is to move fluid across membrane surfaces. Primary cilia, in contrast, adopt sensory roles and are consequently also referred to as sensory cilia. Once long forgotten, this organelle has recently been 'rediscovered' as a result of its association with a multitude of human genetic diseases¹. Ideally positioned as a signaling organelle, the primary cilium has been shown to regulate numerous signaling pathways, many of which are important not only in tissue homeostasis and disease, but also during development².

One of the first signaling pathways shown to be associated with cilia dysfunction was the non-canonical Wnt signaling pathway, known also as the planar cell polarity (PCP) pathway³. This signaling cascade initially identified in Drosophila, is critical for embryogenesis; in particular for convergence and extension processes and for the correct orientation of cells in the plane of epithelia⁴. The sequential signaling of a core set of regulatory proteins translates directional cues which ultimately lead to cytoskeletal rearrangements and result in the coordinated polarization of epithelial cells in a plane⁵. The process of convergence and extension is absolutely required for the cochlear duct to elongate and for correct cellular patterning⁶. As this is regulated via activation of the PCP pathway, one of the most striking phenotypes of cochlea PCP mutants is a shortened cochlear duct with disorganized sensory epithelia⁷. Similarly, mouse mutants, which lack cilia, also exhibit such a convergence and extension phenotype^8,9, though precisely how this is regulated remains to be elucidated.

Because convergence and extension processes are critical for the outgrowth of the cochlear duct, and cellular patterning of the sensory epithelia within the cochlear duct, the developing cochlea is an ideal organ in which to examine PCP signaling during vertebrate development. The organ of Corti, the term given to the specialized sensory epithelium that lines the cochlear duct, is comprised of non-sensory supporting cells and mechanosensory hair cells which must be uniformly oriented for the cochlea to function¹⁰. The mechanosensory hair cells are so called because of the stereociliary bundles that extend from the cuticular plate (apical surface) of each sensory hair cell¹¹. These act as primary transducers of mechanosensation and despite their nomenclature as stereocilia, are actually comprised of modified actin filament-based microvilli. Within each chevron-shaped hair bundle, three rows of stereocilia are organized in a highly ordered and regular pattern in a stair case-like manner. Real microtubule-based cilia, termed kinocilia, are required for the development and orientation of the stereociliary bundles¹². Upon each hair cell, one single kinocilium is physically attached to the stereocilia bundle, located centrally adjacent to the tallest row of stereocilia. The precise function of the kinocilium is unclear, and one hypothesis is that the kinocilium 'pulls' the stereocilia into shape as they mature from microvilli¹². In vertebrates, kinocilia in the cochlea are only present transiently and retract from the hair cells in mice prior to the onset of hearing^11,13,14.

Complete loss of cilia in the developing cochlea results in severely shortened cochlear ducts, mis-formed and mis-oriented stereociliary bundles, as well as mis-positioned basal bodies^8,9. A functional cilium is not just comprised of the ciliary axoneme. Many proteins associated with cilia function occur in complexes localized to cilia-related subdomains such as the basal body, transition zone, or ciliary axoneme¹⁵. The basal body, derived from the mother centriole of the centrosome, is also a microtubule-organizing center for microtubules extending away from the cilium into the cell body and can regulate intracellular trafficking as well as ciliary trafficking. The ciliary transition zone is another region where ciliary function is regulated in terms of organizing import and export of ciliary compounds¹⁶.

Multiple studies have identified a connection between cilia and non-canonical Wnt (PCP signaling), though the precise mechanism is unclear¹⁷. Redundancy of ciliary and PCP genes and the sensitivity of cell polarity to generalized cellular abnormalities, make it difficult to directly link a mutation to PCP-specific deficits. One of the read outs of PCP signaling is the positioning of the basal body and primary cilium, therefore segregating the primary from the secondary defects is challenging. Some studies in zebrafish and mouse mutants have suggested no connection between cilia and Wnt signaling^18-20. Discrepancies in the data may reflect species, tissue, or temporal-dependent differences in ciliary contributions towards Wnt signaling. Furthermore, normal Wnt responsiveness might be retained if basal bodies remain functional. A deeper insight into the role of ciliary proteins in cellular signaling pathways and other biological phenomena is crucial for our understanding of cellular and developmental biology, as well as for the development of targeted treatment strategies.

Protocol

Use and euthanize all animals in accordance with institutional and governmental guidelines and regulations, most commonly via CO₂ inhalation and cervical dislocation.

1. Preparation of Reagents

NOTE: Prior to beginning, prepare all reagents using analytical grade chemicals. Make solutions using molecular grade distilled and deionized water unless otherwise specified.

1. Phosphate buffered saline (1x PBS): Make 1 L of 1x PBS by dissolving 8g NaCl, 0.2g KCl, 1.44g Na₂HPO₄ and 0.24g KH₂PO₄ in 1 L of H₂O. Adjust the pH to 7.4 with HCl. PBS does not need to be sterile and can be stored at RT. Note that this buffer is made without CaCl₂ or MgCl₂.
2. Paraformaldehyde (4% PFA): Prepare a 4% solution of PFA in 1x PBS in a ventilated fume hood. Add 40g of paraformaldehyde powder to 1 L of 1x PBS. To dissolve, stir and heat to approximately 60 °C, and slowly add NaOH drop wise to raise the pH. Once the powder has dissolved, readjust the pH to 7.4 with HCl. Filter through a 0.45 µM filter and freeze aliquots. Defrost a fresh aliquot for each experiment.
3. Triton buffer: Prepare a 0.1 M Tris-HCl (pH 7.5), 0.15 M NaCl, 0.1% Triton X-100 solution by dissolving 8.77 g NaCl in 1 L of 0.1 M Tris-HCl (pH 7.5). Add 1 ml Triton X-100. Stir to dissolve. Store Triton buffer at RT.
4. Triton block: Prepare 10% goat serum in Triton buffer by adding 1 ml of goat serum to 9 ml of Triton buffer. Store at 4 °C. If using primary antibodies raised in mouse, add goat anti-mouse IgG Fab fragments at a dilution of 1 in 200 to the Triton block for the blocking step.
   NOTE: If planning to use samples for scanning electron microscopy (SEM), prepare additional reagents as listed below.
5. Hanks' Balanced Salt Solution (HBSS) with calcium and magnesium: Make a 1x solution of HBSS by diluting from a stock solution in sterile distilled H₂O. As HBSS buffer is complicated to make and there are risks involved, it is advisable to order a premade 10x stock solution. The final concentration of HBBS buffer routinely used is the following: 1.26 mM CaCl₂, 0.49 mM MgCl₂-6H2O, 0.41 mM MgSO₄-7H2O, 5.33 mM KCl, 0.44 mM KH₂PO₄, 4.17 mM NaHCO₃, 137.93 mM NaCl, 0.34 mM Na₂HPO₄, 5.56 mM Dextrose.
6. Hepes buffer: Prepare a 0.1 M Hepes solution in 1x HBSS, by dissolving 23.8 g HEPES in 1 L 1x HBSS buffer.
7. Scanning electron microscopy fixative (SEM Fix): Prepare fixative for SEM by diluting electron microscopy grade glutaraldehyde (2.5%) and paraformaldehyde (4%) in Hepes buffer. Add CaCl₂ to a final concentration of 10 mM.
8. 1% Osmium tetroxide (OsO₄) in Hepes buffer; 1% Osmium tetroxide in water: Dilute a stock solution of osmium tetroxide in Hepes buffer, and separately in sterile distilled H₂O, to a final concentration of 1%. Osmium tetroxide is highly toxic and is most commonly sold as a 4% solution.
9. 1% (w/v) Tannic acid: Dissolve 0.5 g tannic acid in 50 ml sterile distilled H₂O. Filter sterilize by filtering through a 0.45 pore size filter.
10. Graded ethanol solution series: Dilute 200 proof ethanol with sterile distilled H₂O to make 30, 50, 70, 90 and 95% ethanol solutions. For the final ethanol rinses, 100% ethanol 200 proof is required. Use a freshly opened bottle of 200 proof ethanol for the final rinses.

2. Choice of Tissue

1. Ideally, examine embryos or young pups between the ages of embryonic day 16.5 (E16.5) and postnatal day 3 (P3).
   NOTE: Ossification of the bony labyrinth and temporal bones makes dissection progressively more challenging as mice mature. Also kinocilia, the only true microtubule-based cilium found on cochlear hair cells, retracts during development and is no longer present in adult mice.
2. After fixation, decalcify adult cochleae. Place dissected (see section 3) bony labyrinths, into 2 ml EDTA (4.13% in PBS), pH 7.3, in a microcentrifuge tube for 3 - 4 days with rotation. Replenish EDTA daily. After the tissues have softened, rinse in 1.5 ml or more PBS 3 times for 5 min on a nutator with increased agitation.

3. Cochlea Dissection

1. Post euthanization via cervical dislocation or CO₂ inhalation (100% CO₂ ), remove the head. Use a scalpel blade or small pair of scissors, depending on the size of the animal, to dissect the head along the sagittal mid-line beginning at the nose and extending caudally. Remove the brain from each half of the skull with a pair of forceps. Identify the temporal bones, which contain the developing bony labyrinths of the inner ear (see arrow in Figure 1A).
2. Use a pair of forceps to carefully isolate the bony labyrinths from the skull (temporal bones). Do this under a dissecting microscope. Prise away the bony tissue from the skull by running the forceps gently underneath the bony labyrinths. In animals up to P4 the bony labyrinths are still cartilaginous and can be easily removed without breaking.
3. Following dissection, use the tip of the dissecting forceps to clear the oval and round windows and make a small hole in the apex of the cochlear spiral. Fix the bony labyrinths before further dissection of the cochlear duct. To allow for easy removal of the tectorial membrane (see 3.7), fix the bony labyrinths in 1.5 ml 4% PFA for 5 min on ice.
   NOTE: If removal of the tectorial membrane is not required, longer fixation is advised (see 4.1 - 4.3). Brief fixation of the temporal bones prior to dissection of the cochlear duct aids the dissection process and removal of the tectorial membrane. Post dissection and exposure of the organ of Corti, further fixation is required.
4. Further dissect the bony labyrinths to expose the cochlear sensory epithelium as follows. Place the bony labyrinths in a black silicone elastomere-coated dissecting dish containing PBS to facilitate dissection. Use minutien pins to immobilize the tissue if required. To use minutien pins, place them through the vestibular portion of the bony labyrinths with the ventral aspect of the cochlear spiral facing upwards.
   NOTE: Black silicone elastomere dish: Mix silicone elastomere base component with powdered charcoal to obtain an opaque black color. Add the curing agent and pour into one or more petri dishes (glass or plastic). Dry under vacuum to remove trapped air bubbles. Dry dishes completely prior to use.
5. Use fine forceps (#5) to remove the outer cartilage exposing the cochlear duct. Start at the oval window; insert the bottom tip of the forceps into the oval window and gently pry open the cartilage, slowly moving upwards towards the apex.
6. After exposure of the cochlear duct, remove the ventral surface, the Reissner's membrane. Use a fine pair of forceps to pinch the Reissner's membrane at the base of the cochlear duct and peel it off in an upward motion. Visualize the dorsal aspect of the cochlear duct, including the sensory epithelium.
7. Remove the tectorial membrane as described below (optional). For SEM or immunohistochemistry of the kinocilium or stereociliary bundles remove the tectorial membrane.
8. Use a very fine pair of forceps (#55 or finer) to pinch the tectorial membrane at the base of the cochlea and peel upwards towards the apex.
   NOTE: The tectorial membrane is optically clear which makes it difficult to identify. More often than not the membrane comes off in one piece and although it is not easily visualized, one can feel the resistance as it is pulled off from the exposed organ of Corti.
9. After exposing the organ of Corti, further fix the tissue as described below. Retain the vestibular region to aid further preparation of the tissues.

4. Fixation

1. For regular immunohistochemistry fix the dissected bony labyrinths in 1.5 ml 4% PFA at 4 °C on a nutator for 2 hr.
   NOTE: Because of the sensitivity of differing antibodies and antigens, variations in the length and composition of fixation may be required and should be optimized for each antibody.
2. After fixation, wash the samples by rinsing in 1.5 ml or more PBS 3 times for 5 min on a nutator with increased agitation. Samples can be stored in PBS at 4 °C for several weeks before processing.
3. If preparing samples for SEM, fix dissected temporal bones in SEM fix (see 1.7) for 2 hr at RT. Wash by rinsing in 1.5 ml or more Hepes buffer 3 times for 5 min on a nutator with increased agitation. Store in Hepes buffer at 4 °C until further processing. Storage for longer than a couple of weeks is not recommended.

5. Immunohistochemistry

1. Perform immunohistochemistry on dissected samples in one well of a flat-bottomed 96 well plate. Alternatively, use a PCR or microcentrifuge tube. It is advisable to retain the vestibular portion of the bony labyrinth to facilitate processing.
2. Permeabilize the tissue by incubating in 1.5 ml Triton buffer for 1 hr at RT on a nutator with gentle agitation.
3. Block the tissue by incubating in 0.5 ml Triton block for a minimum of 1 hr at RT. If using primary antibodies raised in mouse, add anti-mouse IgG Fab fragments to the block at a dilution of 1 in 200 (see section 1.4). This greatly decreases non-specific binding of anti-mouse IgG to mouse tissue.
4. Incubate the tissue with primary antibodies diluted in Triton block O/N at 4 °C with gentle agitation. Antibody dilution depends on antibodies being used (see section 6.1). Alternatively incubate primary antibodies for 2 hr at RT. Ideally use a minimum volume of 0.5 ml primary antibody dilution. If antibody is limited, use the smallest volume of antibody dilution that completely envelops the tissue.
5. Wash the tissue by rinsing in 1.5 ml or more Triton buffer, a minimum of 3 times for 15 min, on a nutator with increased agitation.
6. Incubate the tissue with fluorescent dye-conjugated secondary antibodies diluted at the manufacturers recommended concentration (typically 1:200-1:1,000) in Triton block for 1 hr at RT on a nutator. Use a minimum volume of 0.5 ml secondary antibody dilution.
   1. Centrifuge the diluted secondary antibody at 13,000 x g for 3 min prior to use to minimize non-specific binding of secondary antibody aggregates. Shield the tissue from light from this step onwards to avoid photobleaching the fluorescent dyes.
7. As in step 5.5, wash the tissue by rinsing in 1.5 ml or more Triton buffer, a minimum of 3 times for 15 min, on a nutator with increased agitation. Store samples in PBS at 4 °C until ready to mount.

6. Antibody Selection 

NOTE: The source of recommended antibodies and their dilutions are listed in the Table of Specific Materials and Equipment. Perform antibody incubations as described in section 5.

1. To visualize the microtubule-based kinocilium, use a ciliary axoneme marker such as anti-Arl13b (1:1,000) or anti-acetylated-α-tubulin (1:800). Visualize the basal body using an antibody against γ-tubulin (1:200) or any other centrosomal marker.
2. To visualize the actin filament-rich stereociliary bundles, add phalloidin (1:300 - 1,000) conjugated to one of several different fluorophores to the secondary antibody incubation. Phalloidin also labels other filamentous actin structures including the cortical actin filaments surrounding the periphery of each epithelial hair cell. This is helpful in determining the outline of each hair cell. Use an alternative membrane marker such as anti-Zo-1 (1:500) if necessary.
3. Label cochlear hair cells using antibodies against Myosin VI (1:1,000) or Myosin VIIa (1:1,000). These can be used if assessing cochlea length.

NOTE: Use appropriate fluorescent dye-conjugated secondary antibodies optimized to the microscope requirements.

7. Mounting and Imaging

1. Place the sample in a black silicone elastomere dish containing PBS. Using fine forceps, remove the vestibular region from the cochlear spiral.
2. Very carefully remove the underlying cartilage and mesenchyme from the cochlear spiral. The remaining cochlear spiral contains the inner sulcus, organ of Corti and outer sulcus.
3. Transfer the cochlear spiral into a drop of PBS placed on a microscope slide. If desired, separate the cochlear spiral into pieces equivalent to one turn. This step is not necessary however, and can lead to loss of the tissue. The apical surface (stereocilia side) of the cochlear hair cells should be facing upward towards the coverslip.
4. Wick away the PBS with an adsorbent tissue or piece of filter paper and gently reposition the cochlear spiral if the epithelia has shifted and overlaps onto itself.
5. Add a drop of mounting media directly onto the cochlea sample and gently place a coverslip on top, taking care to avoid air bubbles. No spacers are required; place the coverslip directly onto the sample. A water-soluble, non-fluorescing, semi-permanent mounting medium which does not require additional steps to hold the coverslip in place is recommended. Mounting media and coverslip thickness should be matched to microscope specifications.
6. Use an epifluorescent or laser-scanning confocal microscope equipped with high magnification (63 - 100X), high numerical aperture (1.2 - 1.4) objectives to capture images of the apical surface of the cochlear hair cell. Use lower objectives (5 - 20X) to take overlapping images of the cochlear spiral to quantify the length of the cochlear duct. Match excitation lasers and emission filters to fluorophores used for secondary antibodies.

8. Scanning Electron Microscopy

1. Continue from section 4.3. After washing SEM fixed samples in Hepes buffer, perform the following steps under ventilation. The following protocol works well for cochlea tissue and avoids the use of sputter coating with a conductive metal.
2. Post-fix in 1% OsO₄ in Hepes buffer for 1 hr, followed by 3 washes with 1 ml distilled water for 5 min each. The smallest volume of OsO₄ that completely envelops the tissue should be used to minimize toxic waste. No agitation is required.
3. Incubate samples at RT in each of the following solutions for 1 hr with 3 washes with 1 ml distilled water for 5 min each in between steps: 1% Tannic acid freshly made in water and filtered prior to use; 1% OsO₄ in water,1% Tannic acid in water,1% OsO₄ in water.
4. Dehydrate samples through a graded ethanol series by incubation for 10 min in each of the following dilutions: 30, 50, 70, 90, and 95%. No agitation is required.
5. Transfer samples through three changes, 5 min each, of 100% ethanol (from a newly opened bottle of 200 proof ethanol).
6. Run the samples though a critical point drier following manufacturer's instructions.
7. Mount the samples using conductive carbon adhesive on top of an SEM stub prior to imaging on an electron microscope. Under a stereomicroscope, use a pair of fine forceps and a paintbrush to gently position the sample with the cochlear epithelia facing up. Use the vestibular portion of the sample to 'anchor' the tissue to the stub. Store samples in a desiccator until imaging. Perform SEM imaging as described in Jones (2012)²¹.

9. Quantification

1. After immunohistochemistry preparation, acquire images using an epifluorescent or laser-scanning confocal microscope. After preparation for SEM, use a scanning electron microscope to image the specimens. Image the apical surface of the organ of Corti with the exposed cochlear hair cells and intercalated support cells.
2. Define specific positions along the cochlear duct, such as 25, 50, and 75% from the base and use these to compare cochlea regions between mutant and control samples. Analyze and compare cilia mutants with their littermate controls, as differences in genetic background can modify the cochlear phenotype.
   NOTE: The organ of Corti develops in a gradient that extends from the mid-base towards both the apex and the base. In the mouse, development is not complete until P14, so it is important to compare regions at similar developmental stages and positions.
3. Take pictures using software accompanying the microscope that allow for quantification of the particular phenotype as required (see below). Use image analysis software such as Image J or software accompanying the microscope to measure lengths, count cells and determine protein localization as described below. Graph these data as a bar chart, scatter plot, box blot or histogram, comparing mutant vs. control.
4. Total length of the cochlear duct (Figure 3A):
   1. Using a hair cell marker or phalloidin, (which marks the stereociliary bundles), determine and measure where the sensory epithelium begins and ends. The length of the cochlear duct is often shortened in cilia mutants.
5. Stereociliary bundle abnormalities (Figure 3B,D,E,G):
   1. Measure bundle convexity (height) as the shortest distance between the vertex of the bundle and the line that extends through both ends of the bundle 'arms' as shown in Figure 3G, left. Alternatively, quantify the area encompassed by the arms of outer hair cell stereociliary bundle as shown in Figure 3G, right. If hair cells with circular stereociliary bundles can be identified count these as a percentage of total hair cells.
      NOTE: In many ciliary mutants bundle abnormalities can be observed irrespective of bundle orientation. It is difficult to accurately quantify stereociliary bundle abnormalities.
6. Orientation of the stereociliary bundles (Figure 3B,C,G):
   1. Assess the orientation of each individual bundle relative to a line extending perpendicular to the row of pillar cells that separates the inner and outer hair cells. Cells with a normal orientation are aligned along this perpendicular axis and so have a rotation of 0°. Determine angles of rotation using either a 360° notation or as absolute deviations from 0°.
7. Kinocilia positioning or stereociliary bundle positioning (Figure 3B,F,H):
   1. Count the percentage of cells in which kinocilia are missing. Measure the distance from the kinocilium to the 'vertex' or center of the bundle. The 'vertex' of the bundle may not be easy to define if bundles are abnormal, and so the center of the bundle can be used instead.
8. Quantify the position of kinocilia and stereociliary bundles on the apical hair cell surface, by overlaying a positional grid onto each hair cell and marking the location of either the base of the kinocilium or the vertex/center of the stereociliary bundle. Display the data as a percentage of each category that lies within a specific region of the grid, and by overlaying the locations of each marking onto one positional grid.
   NOTE: In control tissues, kinocilia are always attached to the vertex of the stereociliary bundle. In cilia mutants, kinocilia are often missing, mislocalized yet still attached, or even completely detached from the stereociliary bundle.
9. Kinocilia length (Figure 3I):
   1. Measure the length of the kinocilium by labeling with a ciliary axoneme marker. Only measure kinocilia that lie flat within a 1.5 µm confocal plane. Confirm that cilia are lying flat by observing a cross-sectional view of the scanned tissue. Compare aged matched samples from the same region of the cochlea since the kinocilium retracts during development.
10. Mislocalization of polarity proteins:
    1. Following immunohistochemistry using antibodies against polarity proteins such as Vangl2 and Gαi3, determine the localization via imaging.

NOTE: In some cilia mutants, localization of polarity proteins is disrupted. These include Vangl2 and Gαi3, which have been shown to be disrupted in Ift20, Bbs8 and Bbs6 mutant cochlea.

NOTE: Further examples for how to quantify and display hair cell polarity can be found in the following papers. Curtin, et al. (2003) Current Biology²², Montcouquiol, et al. (2003) Nature⁷, Wang, et al (2006) The Journal of Neuroscience²³, Montcouquiol, et al. (2008) Methods in Molecular Biology²⁴, May-Simera, et al. (2012) Methods in Molecular Biology²⁵, Yin et al (2012) PloS one ²⁶.

Results

Cochlea Dissection and Tissue Preparation

After removal of the brain, post mid-line sagittal dissection of a P0 mouse head, the bony labyrinth, seen from behind, can be visualized (Figure 1A, white arrow) and removed. Figure 1B shows the isolated bony labyrinths with the cochlear hair facing upwards, ventral, (left) and from behind, dorsal (right). The white arrow points to the ...

Discussion

When preparing cochlear tissue for analysis, there are a few key points to bear in mind. Firstly, differences in genetic background can modify the cochlear phenotype, making it necessary to analyze and compare only littermate controls. Secondly, complete removal of the tectorial membrane is required to obtain the best images with immunohistochemistry and is essential for SEM. The tectorial membrane is an opaque structure and can obscure the cells of the sensory epithelium directly beneath it, making imaging more challeng...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Tools/Equipment
Silicone elastomere - Sylgard 184	Sigma-Aldrich	761028-5EA	See Note 2
Micro dissecting scissors-straight blade	Various
Fine forceps (no. 5 and 55) and blunt forceps	Various
Dissecting microscope.	Various
Uncoated glass microscope slides	Various
Microscope cover slips (22 mm × 40 mm × 0.15 mm)	Various
Transfer pipettes	Various
Minutien pins	Fine Science Tools	26002-10
SEM sample holder	tousimis	8762
Scanning electron microscopy studs	TED PELLA	16111
PELCO Tabs: Carbon adhesive	TED PELLA	16084-3
Fluorescent Microscope	Various
Critical Point Dryer	Various
Scanning Electron Microscope	Various
Glass microscope slides	Various
Glass coverslips	Various
Kimwipe Tissue	Various
Fine Paint Brush
Reagents
1× Phosphate buffered saline (PBS)	Gibco/Life Technologies	10010023
Paraformaldehyde  (PFA) (EM Grade Required for EM)	Various		Prepare a 4% solution in 1× PBS made fresh each time. EM Grade Required for EM.
2.5% Glutaraldehyde Grade1	Sigma-Aldrich	G5882
Tris-HCl (pH 7.5)	Various
NaCl	Various
CaCl 2	Various
Triton X-100	Various
Normal Goat Serum	Various
AffiniPure Fab Fragment Donkey Anti-Mouse IgG (H+L)	Jackson ImmunoResearch	715-007-003
Fluoromount-G Mounting media	SouthernBiotech	0100-01
10× Hanks’ Balanced Salt Solution (HBSS)	Gibco/Life Technologies	14065
Hepes	Gibco/Life Technologies	15630-080
Osmium tetroxide (OsO4 )	Sigma-Aldrich/Fluka Analytical	75632
Tannic acid	Sigma-Aldrich	403040
Ethanol 200 proof	Various
Antibodies
anti Arl13b	Protein Tech	17711-1-AP	Suggested concentration 1:1,000
anti acetylated tubulin (611-B1)	Sigma-Aldrich	T6793	Suggested concentration 1:800
anti gamma tubulin (GTU-88)	Sigma-Aldrich	T6557	Suggested concentration 1:200
anti Zo_1	Invitrogen	40-2300	Suggested concentration 1:500
Myosin VI	Proteus Biosciences	25-6791	Suggested concentration 1:1000
Myosin VIIa	Proteus Biosciences	25-6790	Suggested concentration 1:1,000
anti Vangl2	Merk Millipore	ABN373	Suggested concentration 1:250
anti Gαi3	Sigma-Aldrich	G4040	Suggested concentration 1:250
Alexa Fluor® 488 Phalloidin	Invitrogen/Life Technologies	A12379	Suggested concentration 1:300 - 1,000
Alexa Fluor® 568 Phalloidin	Invitrogen/Life Technologies	A12380	Suggested concentration 1:300 - 1,000