We gratefully acknowledge support from NCBS, TIFR, Infosys-TIFR Leading Edge Research Grant, DST-SERB, and the Royal National Institute for the Deaf. We would like to thank Central Poultry Development Organization and Training Institute, Hesaraghatta, Bengaluru. We are grateful to CIFF and EM facility and lab support at NCBS. We thank Yoshiko Takahashi and Koichi Kawakami for the Tol2-eGFP and T2TP constructs, and Guy Richardson for HCA and G19 Pcdh15 antibody. We are grateful to Earlab members for their constant support and valuable feedback on the protocol.

Protocols involving the procurement, culture, and use of fertilized chicken eggs and unhatched embryos were approved by the Institutional Animal Ethics Committee of the National Centre for Biological Sciences, Bengaluru, Karnataka.
1. In ovo electroporation of chick auditory precursors
<ol>
	<li>sgRNA design and cloning for CRISPR/Cas9 gene knockout
	<ol>
		<li>For creating gene knockouts, design guide RNAs to disrupt the exon regions of the gene, preferably closer to the 5&#39; end of the coding region.</li>
		<li>Select the potential guide RNAs using a web tool CRISPOR23. Set the browser data to Gallus gallus and the protospacer adjacent motif (PAM) sequence to 5&#39;- NGG -3&#39;. The program determines guide RNAs from the input sequence and assigns different scores based on on-target and off-target activity. Select the top four guides for further studies.</li>
		<li>For the designing of template specific oligos for guide (g)RNA production, remove the PAM sequence (5&#8242;- NGG -3&#8242;) from the gRNA design tool output. This is not needed for targeting, but contains the Cas9 cleavage recognition sequence. Synthesize two HPLC purified complementary oligos with BsmBI restriction site at both ends, for each potential gRNA.</li>
		<li>Clone the guide sequence in frame with a tracrRNA scaffold of a vector of choice (here, pcU6_1sgRNA vector was used24).</li>
		<li>Dissolve the sgRNA oligonucleotides at a concentration of 100 &#181;M in DNase/RNase-free water. Perform annealing of the two sense and antisense oligo guides using a thermocycler with the following parameters: 95 &#176;C for 3 min ,then 37 &#176;C for 15 min, and then decrease to 4 &#176;C.</li>
		<li>Using standard molecular biology techniques as described in25, setup restriction digestion of the annealed oligos and pcU6_1sgRNA cloning vector with BsmBI enzyme overnight. Setup ligation with the gel-purified linear BsmBI-digested pcU6_1sgRNA vector and digested sgRNA oligos. Transform into a DH5-alpha competent cell and sequence confirm the obtained clone.</li>
	</ol>
	</li>
	<li>Egg handling and windowing
	<ol>
		<li>Procure freshly laid eggs and clean them by wiping them with 70% ethanol to prevent contamination. Incubate at 37-38 &#176;C, with 45% humidity for 3.5-4 days.</li>
		<li>After incubation, place the egg on its side for at least 5 min before opening. This allows the embryo to reposition to the top of the yolk. Use forceps to make small holes at the top and blunt end of the egg such that a 21G needle can pass through.</li>
		<li>To prevent damage during windowing, remove albumin to lower the embryo away from the shell. To do this, use a 5 mL syringe and a 21G needle to carefully draw 2 mL of albumin from a hole at the blunt end of the egg. Use clear tape to cover the hole at the blunt end.</li>
		<li>To make the egg window, affix clear tape to the top of the eggshell. Cut open a window, around 2 cm long and 1.5 cm wide, using spring bow scissors and expose the embryo. Use forceps to open the chorionic membranes overlying the embryo, allowing access to the embryo.</li>
	</ol>
	</li>
	<li>Microinjecting plasmids
	<ol>
		<li>For the gene knockout experiment, prepare two solutions: the knock-down mix containing SpCas9 protein and guide plasmid - pcU6_1sgRNA, and the tracer plasmid mix of T2K-eGFP (this is a chick &#223;-actin promoter driving GFP cassette surrounded by the transposon sites of Tol2) together with the T2TP (the Tol2 transposase cloned in Tol2 construct26,27). The tracer is used to track electroporation efficiency.</li>
		<li>When electroporating multiple plasmids, ensure that the final concentration of DNA is at least 4 &#181;g/&#181;L. Mix the three constructs, guide plasmid - pcU6_1sgRNA, T2K-eGFP, and T2TP, in a 1:1:1 ratio with 1 &#181;g of SpCas9 protein, 30% sucrose, and 0.1% Fast Green dye in a final volume of 10 &#181;L.</li>
		<li>Pull needles for microinjection from standard glass capillaries (length 3 in, OD 1.0 mm) using a vertical pipette puller. Use fine forceps to break off the capillary tip after pulling to obtain a tip diameter of approximately 50 &#181;m with a tapered end.</li>
		<li>Lay the embryos at E3.5 on their left side with the head facing right. This way only the right otic vesicle is accessible for micro-injection. Micro-inject the knock-down mix into the otic vesicle. Inject around a 200 nL volume of DNA solution mix to fill the otic vesicle.</li>
		<li>Determine the guide efficiency using a T7 endonuclease assay28.</li>
	</ol>
	</li>
	<li>Electroporation
	<ol>
		<li>Add a few drops of 0.719% saline on top of the embryo prior to electroporation to lower the electric resistance and prevent overheating of the embryo.</li>
		<li>Place the positive electrode through the hole made at the blunt side of the egg when the albumin was removed. Maneuver the electrode so that it is under the yolk. Place the negative electrode over the filled otocyst.</li>
		<li>Use electroporation to transfect plasmids into the cells of the embryo. Use a square pulse generator and apply five pulses of 25 V and 100 ms duration each, 50 ms apart. Determine the conditions empirically based on an individual electroporation setup.</li>
		<li>Hydrate the embryo after electroporation by adding a few drops of 0.719% saline. To clean denatured albumin from the surface of the electrodes, thoroughly rinse it with distilled water. Reseal the egg with clear tape and return to the humidified incubator at 37-38 &#176;C for further incubation. 
		&#8203;NOTE: Embryos can be cultured until hatching after electroporation, however there is a steep reduction in viability.</li>
	</ol>
	</li>
</ol>
2. Basilar papilla dissection
<ol>
	<li>Use 70% ethanol to disinfect the surgical table, microscope stage, and surrounding area. Heat or alcohol sterilize microdissection equipment which include minimally spring-bow scissors, micro-curette, and two pairs of fine forceps.</li>
	<li>Prepare the following dissection plates: a glass Petri dish with a black silicon base, a 90 mm plastic Petri dish, and a 60 mm Petri dish. Chill either phosphate-buffered saline (PBS) or Hanks&#39; Balanced Salt Solution (HBSS) for dissections.</li>
	<li>Gently crack the egg into the 90 mm Petri dish. Identify the outer ear of the chick. Decapitate the embryo by cutting the neck just below the level of the lower jaw using scissors. Transfer the head to a 60 mm Petri dish filled with ice-cold PBS.</li>
	<li>Orient the embryo with the top of the beak facing toward the experimenter and hold the beak using one of the #5 forceps. Scoop out the eyes using the second #5 forceps. Going from rostral to caudal, cut the skull along its midline. Scoop out the brain.</li>
	<li>Add more ice-cold PBS or HBSS and locate the two shiny structures, close to the level of the pinna. These are the otoliths of the lagena at the end of the cochlear duct and are found close to the midline.</li>
	<li>Cut roughly between the two lagenas, and well above and below this region to isolate two inner ears. Under oblique lighting, visualize the outline of the inner ear. Remove extraneous tissue and the vestibule.</li>
	<li>Transfer the isolated cochlea to a black silicone base plate with ice cold PBS. Using #5 forceps, peel away the cartilaginous cochlear capsule to obtain the cochlear duct. Locate the undulated layer (tegmentum) of the cochlear duct and remove using #55 forceps to expose the BP. Using #55 forceps, remove the tectorial membrane to expose HCs and SCs.</li>
</ol>
3. Culture of basilar papilla explants
<ol>
	<li>Membrane culture of the BP
	<ol>
		<li>Take a six-well tissue culture plate and arrange one culture membrane insert per well.</li>
		<li>Collect the dissected basilar papilla (explant) in a 200 &#181;L pipette with 1x HBSS buffer and transfer it onto a membrane. To prevent the tissue from sticking to the wall of the pipette, draw up some media before aspirating the tissue.</li>
		<li>Orient the explant such that the basilar papilla is facing upward, so that the hair and support cells are visible from the top29. Once the explant is positioned, aspirate the HBSS buffer slowly from the culture membrane surface. In this process, the explant will attach to the culture membrane.</li>
		<li>Add 1.2 mL of Dulbecco&#39;s modified Eagle medium (DMEM) culture media between the membrane insert and the well wall to fill up the wells of the six-well plate. Up to six explants can be cultured on a single 30 mm culture membrane.</li>
	</ol>
	</li>
	<li>Collagen culture of the cochlear duct
	<ol>
		<li>Prepare collagen mixture by adding 400 &#181;L of 3 mg/mL rat tail collagen, 50 &#181;L of 10x DMEM, 30 &#181;L of 7.5% NaHCO3, and 5 &#181;L of HEPES in a tissue culture hood.</li>
		<li>Take a four-well plate and add three drops of collagen mixture into each well. Transfer dissected cochlear duct to each collagen drop. Incubate the plate for 10 min at 37 &#176;C, 5% CO2 to cure the collagen matrix.</li>
	</ol>
	</li>
	<li>Small molecule treatment of cultures
	<ol>
		<li>Prepare culture media (DMEM, N-2 supplement, Penicillin) with either the pharmacological modulator or its solvent alone (for use as a control). Replace the culture media with 700 &#181;L of media supplemented with the inhibitor. Culture the explants in an incubator at 37 &#176;C, 5% CO2.</li>
		<li>Replace 50% of the culture media every day. After appropriate incubation time, remove the culture media and use the explants for downstream assays.</li>
	</ol>
	</li>
</ol>
4. Imaging and analysis
<ol>
	<li>Immunofluorescent analysis
	<ol>
		<li>Remove culture media from wells and wash the explants twice with 1x HBSS. Using a pipette, add 1 mL of 4% paraformaldehyde (PFA) to the well and incubate for 20 min at room temperature.</li>
		<li>Remove PFA and wash the explants three times with 1x PBS at room temperature. To collect the explants from the culture membrane, make a small cut around the membrane containing the piece of the tissue using small spring bow scissors and transfer the tissue along with the membrane to a 48-well culture plate using forceps. 
		NOTE: If the tissues are floating after the fixation, then aspirate with a 200 &#181;L pipette and transfer it to 48-well plates for further processing. Avoid forceful detachment of the tissue from the membrane.</li>
		<li>For collagen droplet culture, use forceps to transfer the entire collagen drop to a silicon plate. Add 200 &#181;L of 1x PBS and remove collagen with the help of forceps, and then transfer the tissue to a 48-well culture plate.</li>
		<li>Permeabilize the explants with 1 mL of 1x PBS supplemented with 0.3% Tween-20 (PBST) for 30 min at room temperature. Incubate the explants with 200 &#181;L of blocking buffer (10% goat serum + 1% bovine serum albumin in PBST) for 1 h at room temperature.</li>
		<li>Incubate the explants with 200 &#181;L of primary antibody solution (1:300, in blocking buffer) overnight at 4&#160;&#176;C. Remove the primary antibody solution and wash the explants thoroughly 5 x 20 min with PBST.</li>
		<li>Incubate the explants with 200 &#181;L of phalloidin and secondary antibody solution (in blocking buffer) for 1 h at room temperature in the dark. Remove the phalloidin and secondary antibody solution and wash the explants thoroughly 5 x 20 min with PBST. 
		NOTE: Secondary antibody concentration and wash length must be determined for each individual imaging application. For imaging using super resolution microscopy, we typically increase the concentration of the secondary antibody from 1:500 to 1:200 and increase the concentration of phalloidin from 1:400 to 1:200.</li>
		<li>Incubate the explants with a solution of DAPI (1:1000 in PBST) for 15 min at room temperature. Remove the DAPI solution and wash the explants 3 x 5 min with PBST.</li>
		<li>Use the mounting media to mount explants on a slide with BP facing in an upward direction (against the coverslip). For confocal imaging use antifade mounting media. Let the mounting media dry overnight at room temperature in the dark. Either image directly or store the slides at 4&#160;&#176;C until imaging.</li>
	</ol>
	</li>
	<li>OTOTO fixation for SEM analysis
	<ol>
		<li>Prepare all solutions fresh and handle according to local safety guidelines. Fix membrane-cultured explants (from step 4.1.4) in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.3) with 3 mM CaCl2. Perform fixation at 4&#160;&#176;C for 24 to 72 h with a change of fixative every 24 h.</li>
		<li>Remove the fixative and wash the tissue with 0.1 M sodium cacodylate 3 x 5 min at room temperature. Secondary fix with 1% OsO4 (diluted from a 4% OsO4 stock using 0.1 M sodium cacodylate buffer) for 1 h at room temperature. Perform this and the subsequent steps until dehydration in a fume hood.</li>
		<li>Rinse using 0.1 M sodium cacodylate buffer 3 x 5 min at room temperature. Then rinse in double distilled or ultrapure water 3 x 5 min at room temperature.</li>
		<li>Prepare a 0.5% solution of thiocarbohydrazide (TCH) in ultrapure water. Stir at 75 &#176;C for 10 min and filter the solution when it has cooled to room temperature. 
		NOTE: TCH is extremely hazardous. Usage must be approved by the local safety committee, and care must be taken when handling.</li>
		<li>Remove ultrapure water from the sample and add 0.5% TCH drop by drop. If the solution turns brown, stop and rinse the sample with ultrapure water, before carefully adding the 0.5% TCH solution. Once the solution is clear, replace it with 0.5% TCH. Incubate at room temperature for 20 min30,31.</li>
		<li>Rinse the sample with ultrapure water 3 x 5 min at room temperature. Repeat steps 4.2.2, 4.2.3, and 4.2.5 two more times ending with OsO4 incubation followed by rinsing.</li>
		<li>Dehydrate the samples in an ethanol series to 100% anhydrous ethanol (EtOH). Incubate first in 25% ETOH for 10 min, then 50% ETOH for 10 min, 75% EtOH for 10 min, 95% ETOH for 10 min, before incubating for a total of 3x for 10 min each in 100% ETOH.</li>
		<li>Perform critical point drying using liquid CO2. Immediately mount the samples on a&#160;SEM stub with double-sided carbon adhesive tape. Proceed to sputter coat to provide 5-10 nm of coating. If not imaging immediately, store the samples in a desiccator under vacuum.</li>
	</ol>
	</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organotypic+culture+of+neonatal+murine+inner+ear+explants.">Organotypic culture of neonatal murine inner ear explants.</a> Frontiers in Cellular Neuroscience. 13, 170 (2019).</li><li>Davies, S., Forge, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+the+mammalian+organ+of+Corti+for+scanning+electron+microscopy.">Preparation of the mammalian organ of Corti for scanning electron microscopy.</a> Journal of Microscopy. 147, 89-101 (1987).</li><li>Parker, A., Chessum, L., Mburu, P., Sanderson, J., Bowl, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light+and+electron+microscopy+methods+for+examination+of+cochlear+morphology+in+mouse+models+of+deafness.">Light and electron microscopy methods for examination of cochlear morphology in mouse models of deafness.</a> Current Protocols in Mouse Biology. 6 (3), 272-306 (2016).</li><li>Bermingham, N. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Math1:+an+essential+gene+for+the+generation+of+inner+ear+hair+cells.">Math1: an essential gene for the generation of inner ear hair cells.</a> Science. 284 (5421), 1837-1841 (1999).</li><li>Goodyear, R. J., Forge, A., Legan, P. K., Richardson, G. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asymmetric+distribution+of+cadherin+23+and+protocadherin+15+in+the+kinocilial+links+of+avian+sensory+hair+cells.">Asymmetric distribution of cadherin 23 and protocadherin 15 in the kinocilial links of avian sensory hair cells.</a> The Journal of Comparative Neurology. 518 (21), 4288-4297 (2010).</li><li>Bartolami, S., Goodyear, R., Richardson, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Appearance+and+distribution+of+the+275+kD+hair-cell+antigen+during+development+of+the+avian+inner+ear.">Appearance and distribution of the 275 kD hair-cell antigen during development of the avian inner ear.</a> The Journal of Comparative Neurology. 314 (4), 777-788 (1991).</li><li>Funahashi, J., Nakamura, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroporation+in+avian+embryos.">Electroporation in avian embryos.</a> Methods in Molecular Biology. 461, 377-382 (2008).</li><li>Nakamura, H., Funahashi, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroporation:+past,+present+and+future.">Electroporation: past, present and future.</a> Development, Growth &amp; Differentiation. 55 (1), 15-19 (2013).</li><li>Olaya-Sanchez, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fgf3+and+Fgf16+expression+patterns+define+spatial+and+temporal+domains+in+the+developing+chick+inner+ear.">Fgf3 and Fgf16 expression patterns define spatial and temporal domains in the developing chick inner ear.</a> Brain Structure &amp; Function. 222 (1), 131-149 (2017).</li><li>Jones, J. M., Warchol, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+the+Gata3+transcription+factor+in+the+acoustic+ganglion+of+the+developing+avian+inner+ear.">Expression of the Gata3 transcription factor in the acoustic ganglion of the developing avian inner ear.</a> The Journal of Comparative Neurology. 516 (6), 507-518 (2009).</li><li>Serralbo, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transgenesis+and+web+resources+in+quail.">Transgenesis and web resources in quail.</a> Elife. 9, 56312 (2020).</li></ol>

The authors have no competing interests to disclose.

The chick is a cost-effective and convenient addition to the model organisms that a lab may use to research the inner ear. The methods described here are routinely used in our lab and complement ongoing research in the mammalian inner ear. In ovo electroporation is used to introduce genetic manipulations into the chick genome. Electroporation can also be used to introduce constructs that encode fluorescent proteins targeted to particular organelles or subcellular structures35,<...

The inner ear of all vertebrates is derived from a simple epithelium known as the otic placode1,2. This will give rise to all the structural elements and the cell types necessary to transduce the mechanosensory information associated with hearing and balance perception. Hair cells (HCs), the ciliated sensor of the inner ear, are surrounded by supporting cells (SCs). HCs relay information to the auditory hindbrain through the neurons of the eighth cranial nerve. These are also generated from the otic placode3. The primary transduction of sound is achieved at the apical surface of the auditory HC, through a mechanically sensitive hair bundle4. This is mediated through modified actin-based protrusions called stereocilia, which are arranged in a graded, staircase pattern5. In addition, a modified primary cilium, called the kinocilium, organizes hair bundle formation and is adjacent to the tallest row of stereocilia6,7,8. The architecture of stereocilia is critical for this role in converting mechanical stimuli derived from acoustic energy to electrical neural signals9. Damage to the auditory HC through ageing, infection, otoacoustic trauma, or ototoxic shock can result in partial or complete hearing loss that, in mammals, is irreversible10.
Cellular replacement therapies have been proposed that might repair such damage11,12. The approach of this research has been to understand the normal development of the mammalian hair cell and ask if development programs can be reinitiated in progenitor-like cells that may exist within the inner ear13. A second approach has been to look outside of mammals, to non-mammalian vertebrates in which robust regeneration of auditory hair cells takes place, such as birds14,15. In birds, hair cell regeneration occurs predominantly through the de-differentiation of a supporting cell to a progenitor-like state, followed by asymmetric&#160;mitotic division to generate a hair cell and supporting cell16. In addition, direct differentiation of a supporting cell to generate a hair cell has also been observed17.
While the mechanisms of avian auditory development do show significant similarities with that of mammals, there are differences18. HC and SC differentiation in the chick BP is apparent from embryonic day (E) 7, becoming more distinct over time. By E12, a well-patterned and well-polarized basilar papilla (BP) can be visualized, and by E17 well-developed hair cells can be seen19. These time points provide windows into the mechanisms of differentiation, patterning, and polarity, as well as hair cell maturation. Understanding whether such mechanisms are conserved or divergent is important, as they provide insights into the deep homology of the origins of mechanosensory hair cells.
Here, we demonstrate an array of techniques performed at early and late embryonic stages to study cellular processes such as proliferation, fate specification, differentiation, patterning, and maintenance throughout the development of the inner ear organ. This complements other protocols on understanding inner ear development in explant culture20,21,22. We first discuss the introduction of exogenous DNA or RNA into BP precursors within the E3.5 otocyst using in ovo electroporation. Although genetic manipulations can provide valuable insights, the phenotypes thus generated can be pleiotropic and consequently confounding. This is particularly true during later inner ear development, where fundamental processes such as cytoskeletal remodeling play multiple roles in cell division, tissue morphogenesis, and cellular specialization. We present protocols for pharmacological inhibition in cultured explants, which offer advantages in controlling dosage and treatment timing and duration, offering precise spatiotemporal manipulation of developmental mechanisms.
Different organ culture methods can be utilized depending on the treatment duration of small inhibitors. Here we demonstrate two methods of organ culture that allow insights into epithelial morphogenesis and cellular specialization. A method for 3D culture using collagen as a matrix to culture the cochlear duct enables robust culturing and live visualization of the developing BP. For understanding the formation of stereocilia, we present a membrane culture method such that epithelial tissue is cultured on a stiff matrix enabling actin protrusions to grow freely. Both methods allow downstream processing such as live-cell imaging, immunohistochemistry, scanning electron microscopy (SEM), cell recording, etc. These techniques provide a roadmap for the effective use of the chick as a model system to understand and manipulate the development, maturation, and regeneration of the avian auditory epithelium.

<table><tbody><tr><td>Alexa Fluor 488 Phalloidin</td><td>Thermo Fisher Scientific</td><td>A12379</td><td></td></tr><tr><td>Alexa Fluor 647 Phalloidin</td><td>Thermo Fisher Scientific</td><td>A22287</td><td></td></tr><tr><td>Alt-R S.p. HiFi Cas9 Nuclease V3</td><td>Integrated DNA Technologies</td><td>1081061</td><td>High fidelity Cas9 protein</td></tr><tr><td>Anti-GFP antibody</td><td>Abcam</td><td>ab290</td><td>Rabbit polyclonal to GFP</td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma-Aldrich</td><td>A9647</td><td></td></tr><tr><td>Calcium Chloride Dihydrate</td><td>Thermo Fisher Scientific</td><td>Q12135</td><td></td></tr><tr><td>Collagen I, rat tail</td><td>Thermo Fisher Scientific</td><td>A1048301</td><td></td></tr><tr><td>Critical Point Dryer Leica EM CPD300</td><td>Leica</td><td></td><td></td></tr><tr><td>CUY-21 Electroporator</td><td>Nepagene</td><td></td><td></td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>Sigma-Aldrich</td><td>D8418</td><td></td></tr><tr><td>DM5000B Widefield Microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>DMEM, high glucose, GlutaMAX Supplement, pyruvate</td><td>Thermo Fisher Scientific</td><td>10569010</td><td></td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11251-20</td><td></td></tr><tr><td>Dumont #55 Forceps</td><td>Fine Science Tools</td><td>11255-20</td><td></td></tr><tr><td>Fast Green FCF</td><td>Sigma-Aldrich</td><td>F7252</td><td></td></tr><tr><td>Fluoroshield</td><td>Sigma-Aldrich</td><td>F6182</td><td></td></tr><tr><td>FLUOVIEW 3000 Laser Scanning Microscope</td><td>Olympus</td><td></td><td></td></tr><tr><td>Glutaraldehyde (25 %)</td><td>Sigma-Aldrich</td><td>340855</td><td></td></tr><tr><td>Goat anti-Mouse IgG Secondary Antibody, Alexa Fluor 488</td><td>Thermo Fisher Scientific</td><td>A-11001</td><td></td></tr><tr><td>Goat anti-Mouse IgG Secondary Antibody, Alexa Fluor 594</td><td>Thermo Fisher Scientific</td><td>A-11032</td><td></td></tr><tr><td>Goat anti-Rabbit IgG Secondary Antibody, Alexa Fluor 488</td><td>Thermo Fisher Scientific</td><td>A-11008</td><td></td></tr><tr><td>Goat Serum Sterile filtered</td><td>HiMedia</td><td>RM10701</td><td>Heat inactivated</td></tr><tr><td>Hanks&#039; Balanced Salt Solution (HBSS)</td><td>Thermo Fisher Scientific</td><td>14025092</td><td></td></tr><tr><td>LSM980 Airyscan Microscope</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Millicell Cell Culture Insert, 30 mm, hydrophilic PTFE, 0.4 &amp;micro;m</td><td>Sigma-Aldrich</td><td>PICM03050</td><td></td></tr><tr><td>MVX10 Stereo Microscope</td><td>Olympus</td><td></td><td></td></tr><tr><td>MYO7A antibody</td><td>DSHB</td><td>138-1</td><td>Mouse monoclonal to Unconventional myosin-VIIa</td></tr><tr><td>MZ16 Dissecting microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>N-2 Supplement (100X)</td><td>Thermo Fisher Scientific</td><td>17502048</td><td></td></tr><tr><td>Noyes Scissors, 14cm (5.5&#039;&#039;)</td><td>World Precision Instruments</td><td>501237</td><td></td></tr><tr><td>Osmium tetroxide (4%)</td><td>Sigma-Aldrich</td><td>75632</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>158127</td><td></td></tr><tr><td>PC-10 Puller</td><td>Narishige</td><td></td><td></td></tr><tr><td>pcU6_1sgRNA</td><td>Addgene</td><td>92395</td><td>Mini vector with modified chicken U6 promoter</td></tr><tr><td>Penicillin G sodium salt</td><td>Sigma-Aldrich</td><td>P3032</td><td></td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>Thermo Fisher Scientific</td><td>10010023</td><td></td></tr><tr><td>ProLong Gold Antifade Mountant</td><td>Thermo Fisher Scientific</td><td>P36934</td><td></td></tr><tr><td>SMZ1500 Dissecting microscope</td><td>Nikon</td><td></td><td></td></tr><tr><td>Sodium Cacodylate Buffer, 0.2M</td><td>Electron Microscopy Sciences</td><td>11652</td><td></td></tr><tr><td>Sodium chloride</td><td>HiMedia</td><td>GRM853</td><td></td></tr><tr><td>Sputtre Coater K550X</td><td>Emitech</td><td></td><td></td></tr><tr><td>Standard Glass Capillaries 3 in, OD 1.0 mm, No Filament</td><td>World Precision Instruments</td><td>1B100-3</td><td></td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>84097</td><td></td></tr><tr><td>The MERLIN Compact VP</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Thiocarbohydrazide</td><td>Alfa Aesar</td><td>L01205</td><td></td></tr><tr><td>TWEEN 20</td><td>Sigma-Aldrich</td><td>P1379</td><td></td></tr></tbody></table>

in ovo and ex ovo methods to study avian inner ear development

The inner ear perceives sound and maintains balance using the cochlea and vestibule. It does this by using a dedicated mechanosensory cell type known as the hair cell. Basic research in the inner ear has led to a deep understanding of how the hair cell functions, and how dysregulation can lead to hearing loss and vertigo. For this research, the mouse has been the pre-eminent model system. However, mice, like all mammals, have lost the ability to replace hair cells. Thus, when trying to understand cellular therapies for restoring inner ear function, complementary studies in other vertebrate species could provide further insights. The auditory epithelium of birds, the basilar papilla (BP), is a sheet of epithelium composed of mechanosensory hair cells (HCs) intercalated by supporting cells (SCs). Although the anatomical architecture of the basilar papilla and the mammalian cochlea differ, the molecular mechanisms of inner ear development and hearing are similar. This makes the basilar papilla a useful system for not only comparative studies but also to understand regeneration. Here, we describe dissection and manipulation techniques for the chicken inner ear. The technique shows genetic and small molecule inhibition methods, which offer a potent tool for studying the molecular mechanisms of inner ear development. In this paper, we discuss in ovo electroporation techniques to genetically perturb the basilar papilla using CRIPSR-Cas9 deletions, followed by dissection of the basilar papilla. We also demonstrate the BP organ culture and optimal use of culture matrices, to observe the development of the epithelium and the hair cells.

In the electroporation setup, electrode positioning can play a role in the domain of transfection. The positive electrode is placed under the yolk, and the negative above the embryo (Figure 1A). This results in higher GFP expression in much of the inner ear and both vestibular organs (Figure 1B), and auditory basilar papilla (Figure 1C,D), confirming transfection.
In assessing the phenoty...

Watch this Scientific Journal Video about In Ovo and Ex Ovo Methods to Study Avian Inner Ear Development at JoVE.com

In Ovo and Ex Ovo Methods to Study Avian Inner Ear Development

The chick is a cost-effective, accessible, and widely available model organism for a variety of studies. Here, a series of protocols is detailed to understand the molecular mechanisms underlying avian inner ear development and regeneration.

In Ovo and Ex Ovo Methods to Study Avian Inner Ear Development

The inner ear perceives sound and maintains balance using the cochlea and vestibule. It does this by using a dedicated mechanosensory cell type known as ...

in-ovo-and-ex-ovo-methods-to-study-avian-inner-ear-development

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

2.2K Views.  Tata Institute of Fundamental Research. The chick is a cost-effective, accessible, and widely available model organism for a variety of studies. Here, a series of protocols is detailed to understand the molecular mechanisms underlying avian inner ear development and regeneration. 

Video: In Ovo and Ex Ovo Methods to Study Avian Inner Ear Development

Optimized Ex-ovo Culturing of Chick Embryos to Advanced Stages of Development

Research in anatomy, embryology, and developmental biology has largely relied on the use of model organisms. In order to study development in live embryos model organisms, such as the chicken, are often used. The chicken is an excellent model organism due to its low cost and minimal maintenance, however they present observational challenges because they are enclosed in an opaque eggshell. In order to properly view the embryo as it develops, the shell must be windowed or removed. Both windowing and ex ovo techniques have been developed to assist researchers in the study of embryonic development. However, each of the methods has limitations and challenges. Here, we present a simple, optimized ex ovo culture technique for chicken embryos that enables the observation of embryonic development from stage HH 19 into late stages of development (HH 40), when many organs have developed. This technique is easy to adopt in both undergraduate classes and more advanced research laboratories where embryo manipulations are conducted.

Optimized Ex-ovo Culturing of Chick Embryos to Advanced Stages of Development

Viewing and accessing the chicken embryo during development can be challenging. We have developed an ex ovo method that is simple, cost effective, and can easily be used in a classroom or research setting. This method provides access to the embryo into late stages of embryonic development (HH 40).

Research in anatomy, embryology, and developmental biology has largely relied on the use of model organisms. In order to study development in live embryos ...

Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair cells are aligned in one row of inner hair cells and three rows of outer hair cells that extend along the basal to apical axis of the cochlea. The explant culture technique described here provides an efficient method to isolate and maintain cochlear explants from the embryonic mouse inner ear. Also, the morphology and molecular characteristics of sensory hair cells and nonsensory supporting cells within the cochlear explant cultures resemble those observed in vivo and can be studied within its intrinsic cellular environment. The cochlear explants can serve as important experimental tools for the identification and characterization of molecular and genetic pathways that are involved in cellular specification and patterning. Although transgenic mouse models provide an effective approach for gene expression studies, a considerable number of mouse mutants die during embryonic development thereby hindering the analysis and interpretation of developmental phenotypes. The organ of Corti from mutant mice that die before birth can be cultured so that their in vitro development and responses to different factors can be analyzed. Additionally, we describe a technique for electroporating embryonic cochlear explants ex vivo which can be used to downregulate or overexpress specific gene(s) and analyze their potential endogenous function and test whether specific gene product is necessary or sufficient in a given context to influence mammalian cochlear development1-8.

We present a method that describes isolation and culture of cochlear explants from embryonic mouse inner ear. We also demonstrate a method for gene transfer into cochlear explants via square-wave electroporation. The in vitro explant culture coupled with gene transfer technique enables researchers to study the effects of altering gene expression during development.

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair ...

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. The inner ear auditory sensory organ is critical for our ability to hear. It houses a highly specialized sensory epithelium that consists of mechano-transducing hair cells (HCs) and surrounding glial-like supporting cells (SCs). Despite structural differences in the auditory organs, molecular mechanisms regulating the development of the auditory organ are evolutionarily conserved between mammals and aves. In ovo electroporation is largely limited to early stages at E1 - E3. Due to the relative late development of the auditory organ at E5, manipulations of the auditory organ by in ovo electroporation past E3 are difficult due to the advanced development of the chicken embryo at later stages. The method presented here is a transient gene transfer method for targeting genes of interest at stage E4 - E4.5 in the developing chicken auditory sensory organ via in ovo micro-electroporation. This method is applicable for gain- and loss-of-functions with conventional plasmid DNA-based expression vectors and can be combined with in ovo cell proliferation assay by adding EdU (5-ethynyl-2&#180;-deoxyuridine) to the whole embryo at the time of electroporation. The use of green or red fluorescent protein (GFP or RFP) expression plasmids allows the experimenter to quickly determine whether the electroporation successfully targeted the auditory portion of the developing inner ear. In this method paper, representative examples of GFP electroporated specimens are illustrated; embryos were harvested 18 - 96 hr&#160;after electroporation and targeting of GFP to the pro-sensory area of the auditory organ was confirmed by RNA in situ hybridization. The method paper also provides an optimized protocol for the use of the thymidine analog EdU to analyze cell proliferation; an example of an EdU based cell proliferation assay that combines immuno-labeling and click EdU chemistry is provided.

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

The auditory organ mediates hearing. Here we present a modified in ovo micro-electroporation method optimized for studying auditory progenitor cell proliferation and differentiation in the developing chicken auditory organ.

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. ...

Investigating Outer Hair Cell Motility with a Combination of External Alternating Electrical Field Stimulation and High-speed Image Analysis

OHCs are cylindrical sensorimotor cells located in the Organ of Corti, the auditory organ inside the mammalian inner ear. The name "hair cells" derives from their characteristic apical bundle of stereocilia, a critical element for detection and transduction of sound energy 1. OHCs are able to change shape &#x2014;elongate, shorten and bend&#x2014; in response to electrical, mechanical and chemical stimulation, a motor response considered crucial for cochlear amplification of acoustic signals 2.
OHC stimulation induces two different motile responses: i) electromotility, a.k.a fast motility, changes in length in the microsecond range derived from electrically-driven conformational changes in motor proteins densely packed in OHC plasma membrane, and ii) slow motility, shape changes in the millisecond to seconds range involving cytoskeletal reorganization 2, 3. OHC bending is associated with electromotility, and result either from an asymmetric distribution of motor proteins in the lateral plasma membrane, or asymmetric electrical stimulation of those motor proteins (e.g., with an electrical field perpendicular to the long axis of the cells) 4. Mechanical and chemical stimuli induce essentially slow motile responses, even though changes in the ionic conditions of the cells and/or their environment can also stimulate the plasma membrane-embedded motor proteins 5, 6. Since OHC motile responses are an essential component of the cochlear amplifier, the qualitative and quantitative analysis of these motile responses at acoustic frequencies (roughly from 20 Hz to 20 kHz in humans) is a very important matter in the field of hearing research 7.
The development of new imaging technology combining high-speed videocameras, LED-based illumination systems, and sophisticated image analysis software now provides the ability to perform reliable qualitative and quantitative studies of the motile response of isolated OHCs to an external alternating electrical field (EAEF) 8. This is a simple and non-invasive technique that circumvents most of the limitations of previous approaches 9-11. Moreover, the LED-based illumination system provides extreme brightness with insignificant thermal effects on the samples and, because of the use of video microscopy, optical resolution is at least 10-fold higher than with conventional light microscopy techniques 12. For instance, with the experimental setup described here, changes in cell length of about 20 nm can be routinely and reliably detected at frequencies of 10 kHz, and this resolution can be further improved at lower frequencies. 
We are confident that this experimental approach will help to extend our understanding of the cellular and molecular mechanisms underlying OHC motility.

A reliable method to investigate outer hair cell (OHC) motile responses, including electromotility, slow motility and bending, is described. OHC motility is elicited by stimulation with an external alternating electrical field, and the method takes advantage of high-speed image recording, LED-based illumination, and last generation image analysis software.

OHCs are cylindrical sensorimotor cells located in the Organ of Corti, the auditory organ inside the mammalian inner ear. The name "hair cells" derives ...

Neuroscience

In Ovo Electroporation in the Chicken Auditory Brainstem

Electroporation is a method that introduces genes of interest into biologically relevant organisms like the chicken embryo. It is long established that the chicken embryo is an effective research model for studying basic biological functions of auditory system development. More recently, the chicken embryo has become particularly valuable in studying gene expression, regulation and function associated with hearing. In ovo electroporation can be used to target auditory brainstem regions responsible for highly specialized auditory functions. These regions include the chicken nucleus magnocellularis (NM) and nucleus laminaris (NL). NM and NL neurons arise from distinct precursors of rhombomeres 5 and 6 (R5/R6). Here, we present in ovo electroporation of plasmid-encoded genes to study gene-related properties in these regions. We show a method for spatial and temporal control of gene expression that promote either gain or loss of functional phenotypes. By targeting auditory neural progenitor regions associated with R5/R6, we show plasmid transfection in NM and NL. Temporal regulation of gene expression can be achieved by adopting a tet-on vector system. This is a drug inducible procedure that expresses the genes of interest in the presence of doxycycline (Dox). The in ovo electroporation technique &#8211; together with either biochemical, pharmacological, and or in vivo functional assays &#8211; provides an innovative approach to study auditory neuron development and associated pathophysiological phenomena.

In Ovo Electroporation in the Chicken Auditory Brainstem

Auditory brainstem neurons of avians and mammals are specialized for fast neural encoding, a fundamental process for normal hearing functions. These neurons arise from distinct precursors of embryonic hindbrain. We present techniques utilizing electroporation to express genes in the hindbrain of chicken embryos to study gene function during auditory development.

Electroporation is a method that introduces genes of interest into biologically relevant organisms like the chicken embryo. It is long established that ...

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and the organ of Corti in the coiled cochlea. The cristae and maculae contain vestibular hair cells that transduce mechanical stimuli to subserve the special sense of balance, while auditory hair cells in the organ of Corti are the primary transducers for hearing 1. Cell fate specification in these sensory epithelia and morphogenesis of the semicircular canals and cochlea take place during the second week of gestation in the mouse and are largely completed before birth 2,3. Developmental studies of the mouse inner ear are routinely conducted by harvesting transgenic embryos at different embryonic or postnatal stages to gain insight into the molecular basis of cellular and/or morphological phenotypes 4,5. We hypothesize that gene transfer to the developing mouse inner ear in utero in the context of gain- and loss-of-function studies represents a complimentary approach to traditional mouse transgenesis for the interrogation of the genetic mechanisms underlying mammalian inner ear development6.
The experimental paradigm to conduct gene misexpression studies in the developing mouse inner ear demonstrated here resolves into three general steps: 1) ventral laparotomy; 2) transuterine microinjection; and 3) in vivo electroporation. Ventral laparotomy is a mouse survival surgical technique that permits externalization of the uterus to gain experimental access to the implanted embryos7. Transuterine microinjection is the use of beveled, glass capillary micropipettes to introduce expression plasmid into the lumen of the otic vesicle or otocyst. In vivo electroporation is the application of square wave, direct current pulses to drive expression plasmid into progenitor cells8-10. 
We previously described this electroporation-based gene transfer technique and included detailed notes on each step of the protocol11. Mouse experimental embryological techniques can be difficult to learn from prose and still images alone. In the present work, we demonstrate the 3 steps in the gene transfer procedure. Most critically, we deploy digital video microscopy to show precisely how to: 1) identify embryo orientation in utero; 2) reorient embryos for targeting injections to the otocyst; 3) microinject DNA mixed with tracer dye solution into the otocyst at embryonic days 11.5 and 12.5; 4) electroporate the injected otocyst; and 5) label electroporated embryos for postnatal selection at birth. We provide representative examples of successfully transfected inner ears; a pictorial guide to the most common causes of otocyst mistargeting; discuss how to avoid common methodological errors; and present guidelines for writing an in utero gene transfer animal care protocol.

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mouse inner ear is a placode-derived sensory organ whose developmental program is elaborated during gestation. We define an in utero gene transfer technique consisting of three steps: mouse ventral laparotomy, transuterine microinjection, and in vivo electroporation. We use digital video microscopy to demonstrate the critical experimental embryological techniques.

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and ...

Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling of auditory axons within the VIIIth cranial nerve would enhance the study of central auditory circuit development. This approach is challenging because multiple sensory organs of the inner ear contribute to the VIIIth nerve 1. Moreover, markers that reliably distinguish auditory versus vestibular groups of axons within the avian VIIIth nerve have yet to be identified. Auditory and vestibular pathways cannot be distinguished functionally in early embryos, as sensory-evoked responses are not present before the circuits are formed. Centrally projecting VIIIth nerve axons have been traced in some studies, but auditory axon labeling was accompanied by labeling from other VIIIth nerve components 2,3. Here, we describe a method for anterograde tracing from the acoustic ganglion to selectively label auditory axons within the developing VIIIth nerve. First, after partial dissection of the anterior cephalic region of an 8-day chick embryo immersed in oxygenated artificial cerebrospinal fluid, the cochlear duct is identified by anatomical landmarks. Next, a fine pulled glass micropipette is positioned to inject a small amount of rhodamine dextran amine into the duct and adjacent deep region where the acoustic ganglion cells are located. Within thirty minutes following the injection, auditory axons are traced centrally into the hindbrain and can later be visualized following histologic preparation. This method provides a useful tool for developmental studies of peripheral to central auditory circuit formation.

Here we describe a microdissection technique followed by fluorescent dye injection into the acoustic ganglion of early chick embryos for selective tracing of auditory axon fibers in the nerve and hindbrain.

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for discovering new therapeutic strategies. Advancements in imaging technology have enabled quantitative investigation of the organization and maturing of the body plan, but later stage embryonic morphogenesis is less clear. Chicken embryos are an attractive vertebrate animal model system for this application because of its ease of culture and surgical manipulation. Early embryos can be cultured for a short time on filter paper rings, which enables complete optical access for cell patterning and fate studies1,2. Studying advanced developmental processes such as cardiac morphogenesis are traditionally performed through a window of the eggshell3-5, but this technique limits optical access due to window size. We previously developed a simple method to culture whole embryos ex-ovo on hexagonal weigh boats for up to 10 days, which enabled high resolution imaging via ultrasonography6,7. These cultures were difficult to transport, limiting the types of imaging tools available for live experiments. We here present an improved shell-less culture system with a cost-effective, portable environmental chamber. Eggs were cracked onto a hammock created by a polyurethane membrane (cling wrap) affixed circumferentially to a plastic cup partially filled with sterile water. The dimensions of the circumference and depth of the hammock were both critical to maintain surface tension, while the mechanics of the hammock and water beneath helped dampen vibrations induced by transportation. A small footprint circulating water bath was also developed to enable continuous temperature control during experimentation. We demonstrate the ability to culture embryos in this way for at least 14 days without morphogenic defect or delay and employ this system in several microsurgical and imaging applications.

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

In this article, we present a simple methodology to enable long-term ex-ovo avian embryo culture. This technique is ideal for longitudinal experimentation requiring complete optical accessibility and/or sterile transportation in avian embryos.

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for ...

Biology

Slicing the Embryonic Chicken Auditory Brainstem to Evaluate Tonotopic Gradients and Microcircuits

The chicken embryo is a widely accepted animal model to study the auditory brainstem, composed of highly specialized microcircuitry and neuronal topology differentially oriented along a tonotopic (i.e., frequency) axis. The tonotopic axis permits the segregated encoding of high-frequency sounds in the rostral-medial plane and low-frequency encoding in caudo-lateral regions. Traditionally, coronal brainstem slices of embryonic tissue permit the study of relative individual iso-frequency lamina. Although sufficient to investigate anatomical and physiological questions pertaining to individual iso-frequency regions, the study of tonotopic variation and its development across larger auditory brainstem areas is somewhat limited. This protocol reports brainstem slicing techniques from chicken embryos that encompass larger gradients of frequency regions in the lower auditory brainstem. The utilization of different slicing methods for chicken auditory brainstem tissue permits electrophysiological and anatomical experiments within one brainstem slice, where larger gradients of tonotopic properties and developmental trajectories are better preserved than coronal sections. Multiple slicing techniques allow for improved investigation of the diverse anatomical, biophysical, and tonotopic properties of auditory brainstem microcircuits.

Here we present a protocol for obtaining non-coronal auditory brainstem slices of the chicken embryo for the investigation of tonotopic properties and developmental trajectories within one brainstem slice. These slices include sagittal, horizontal, and horizontal/transverse sections encompassing larger tonotopic regions within an individual slice plane that the traditional coronal sections.

The chicken embryo is a widely accepted animal model to study the auditory brainstem, composed of highly specialized microcircuitry and neuronal topology ...

In Ovo and Ex Ovo Methods to Study Avian Inner Ear Development

Summary

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Chapters in this video

Optimized Ex-ovo Culturing of Chick Embryos to Advanced Stages of Development

Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

Investigating Outer Hair Cell Motility with a Combination of External Alternating Electrical Field Stimulation and High-speed Image Analysis

In Ovo Electroporation in the Chicken Auditory Brainstem

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve

Long-term Time Lapse Imaging of Mouse Cochlear Explants

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

Slicing the Embryonic Chicken Auditory Brainstem to Evaluate Tonotopic Gradients and Microcircuits