Age, infection, and ototoxic drugs can cause degeneration of hair cells of the inner ear, which in us leads to hearing loss. In non-mammalian vertebrates, such as chick, hair cells can be regenerated. The techniques described here make use of the cost effectiveness of working on chick embryos, the ease of obtaining embryos, and good exo development of tissue explants.
Understanding avian inner ear hairs development and regeneration can provide important insights into hearing loss and potential therapies, and explant culture can be important for evaluating the ototoxic effects of various drugs. Before starting the experiment, procure freshly laid eggs, and clean them with 70%ethanol to prevent contamination. Incubate the eggs for 3.5 to 4 days, at 37 to 38 degrees Celsius with 45%humidity.
After incubation, reposition the embryo to the top of the yolk, by placing an egg on its side for five minutes. Then, use forceps to make a small hole at the top and blunt end of the egg for a 21 gauge needle to pass through. Using a five milliliter syringe and a 21 gauge needle, remove two milliliters of the albumin from a hole at the blunt end of the egg.
Cover the hole at the blunt end using clear tape. To make the egg window affix the clear tape to the top of the egg shell. Hold the needles from microinjection from standard glass capillaries using a vertical pipette puller.
After pulling, break off the capillary tip using fine forceps to obtain a tip diameter of approximately 50 micrometers with a tapered end. For the gene knockout experiment, mix the three constructs, namely the guide plasmid, pcU6.1sgRNA, T2KeGFP, T2TP in a one-to-one-to-one ratio with one microgram of SP Cas9 protein, 30%sucrose, and 0.1%fast green dye, and a final volume of 10 microliters. While electroporating multiple plasmids, ensure the final concentration of DNA is at least four micrograms per microliter.
With the help of springbow scissors cut open around a two centimeter long and 1.5 centimeter wide window, and expose the embryo. Then use forceps to open the chorionic membranes overlying the embryo, allowing access to the embryo. Inject around 200 nanoliter volume of DNA solution mix to fill the otic vesicle.
To determine the guide RNA efficiency perform a T7 endonuclease assay. Add 0.719%saline drops to the embryo to lower the electric resistance, and prevent overheating. Next, place the positive electrode through the hole made at the blunt side of the egg, and maneuver the electrode so that it is under the yolk.
Place the negative electrode over the filled otocyst. To transfect the plasmid into the otic vesicle with electroporation, use a square pulse generator, and apply five pulses of 25 volts for 100 milliseconds each, 50 milliseconds apart. Determine the conditions empirically based on an individual electroporation set up.
After electroporation, hydrate the embryo by adding a few drops of 0.719%saline. Reseal the egg with clear tape, and return to the humidified incubator at 37 to 38 degrees Celsius for further incubation. Disinfect the surgical table, microscope stage, and surrounding area using 70%ethanol and heat, or alcohol sterilize the micro dissection equipment, including minimally springbow scissors, micro curet, and two pairs of fine forceps.
Prepare the dissection plates including a glass Petri dish with a black silicone base, a 90 millimeter plastic Petri dish, and a 60 millimeter Petri dish. Keep chilled PBS or Hanks'balanced salt solution also known as HBSS ready for dissections. Gently crack the egg into the 90 millimeter Petri dish.
Then identify the outer ear of the chick, and transfer the head to a 60 millimeter Petri dish filled with ice cold PBS. Next, orient the embryo with the top of the beak facing inside, and hold the beak using one of the number five forceps. Scoop out the eyes using the second number five forceps.
Then, from rostral to caudal, cut along the skull along its midline, and scoop out the brain. Add more ice cold PBS, or HBSS, and locate the two otoliths of the lagena as shiny structures at the end of the cochlear duct, and close to the midline. To isolate two inner ears, cut between the two lagenas, and well above and below the region.
Then under oblique lighting visualize the outline of the inner ear, and remove extraneous tissue and the vestibule. Transfer the isolated cochlea to a black silicone base plate with ice cold PBS. Peel away the cartilaginous cochlear capsule using number five forceps to obtain the cochlear duct.
Then locate the undulated layer or tegmentum of the cochlear duct, and remove it using number 55 forceps to expose the basilar papilla or BP.Next, remove the tectorial membrane to expose hair cells and supporting cells using number 55 forceps. For membrane culture of the basilar papilla, take a six well tissue culture plate, and arrange one culture membrane insert per well. Draw up some media in a 200 microliter pipette, and then aspirate the dissected basilar papilla explant with one X HBSS buffer.
Transfer the explant onto a membrane, and orient it so that the basilar papilla faces upward, and the hair and support cells are visible from the top. 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.
Fill up the well of the six well plate by adding 1.2 milliliters of dulbecco's modified eagle medium, or DMEM culture media, between the membrane insert and the well wall. To prepare the collagen culture of the cochlear duct add three drops of freshly prepared collagen mixture into each well of a four well plate, and transfer dissected cochlear duct to each collagen drop. Incubate the plate for 10 minutes at 37 degrees Celsius, and 5%carbon dioxide to cure the collagen matrix.
For a small molecule treatment of the cultures, replace the culture media with 700 microliters of media supplemented with the pharmacological modulator, such as penicillin, and incubate the plates as demonstrated before. Replace 50%of the culture media every day. After appropriate incubation time, remove the culture media, and use the explants for downstream assays.
In the electroporation setup the correct electrode positioning resulted in higher GFP expression in the inner ear in both vestibular organs, and auditory basilar papilla confirming transfection. CRISPR Cas9 mediated Atoh1 gene knockout in inner ear, resulted in loss of hair cells. Atoh1 gene knockout via an ovo-electroporation followed by incubation until E10 showed reduced hair cell development compared to empty plasmid control.
Although electroporation is mosaic, control electroporated cells were able to form hair cells. In Atoh1 gRNA electroporated samples, green fluorescent protein positive cells never showed the markers of hair cell development. The organ culture of the basilar papilla in a 3D matrix, such as, collagen and stain with antibodies against hair cell antigen provided the excellent preservation of tissue morphology for up to five days.
The organization of hair cells and supporting cells were maintained in these culture conditions. The organ cultures on a membrane can be maintained for up to five days while maintaining the hair bundle integrity. This can be seen in the representative image by the localization of the tip link protein, protocadherin 15.
For investigating the development of the hair bundle, higher resolution imaging, such as, super resolution microscopy, or scanning electron microscopy can provide more information. Sterile conditions should be maintained throughout the procedure. While electroporating, ensure that the embryos do not dry out.
When titrating pharmacological inhibitors, one may reduce the concentration to overcome any lethality. Both embryos and explants can be taken for imaging experiments. Either by performing realtime video microscoping, or static light confocal electron microscoping, as shown in the protocol.
Given the ease and accessibility of using chick inner ear explants, we can use medium through proof screening to identify new molecules essential for hair cell regeneration.
