Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

NOTE: All protocols using live animals must be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved methods for the care and use of the laboratory animals. All the dissections should be performed using sterile technique on a clean laminar flow bench. Gloves and a mask, if desired, should be worn during this procedure.

1. Dissection of the Embryonic Mouse Inner Ear

1. Setting up for organ of Corti explant cultures:
   1. Sterilize the laminar flow tissue culture hood by turning on UV light for about 30 min and disinfect the surface with 70% ethanol prior to use. In addition, sterilize all dissection instruments including fine dissection tools and Sylgard-coated dishes for embryo dissection via autoclave or by soaking in 70% ethanol for at least 20 min before use. Pour off 70% ethanol and allow the dishes and instruments to air dry in a clean laminar flow clean bench prior to use.
   2. Prepare sterile Hank’s Balanced Salt Solution (HBSS)/(4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) (HEPES) mixture by adding 10% of 10x HBSS and 0.5% HEPES and adjust the pH to approximately 7.2 and filter sterilize the final solution and store at 4 °C. Carry out all dissections in chilled HBSS/HEPES solution to better preserve the tissue throughout the procedure.
   3. Coat glass bottom dishes with basement membrane matrix by adding 5 ml of cold (4 °C) sterile Dulbecco’s Modified Eagle Medium (DMEM) to a 300 µl aliquot of basement membrane matrix in a sterile 15 ml polypropylene tube. Mix by vortexing the contents and add 150 µl of the basement membrane matrix-DMEM mixture to the center of each culture dish so that it covers the entire culture well present in the center of the dish. Cover the dishes and store in a 37 °C incubator for at least 45 min before use.
   4. Pour 4-5 ml chilled HBSS into three sterile polystyrene petri dishes and leave closed in the hood.
   5. Prepare culture medium in a sterile 15 ml polypropylene tube by adding 9 ml of DMEM, 100 µl of N2 supplements, 10 g/ml Cipro and 1 ml of FBS.
2. Isolation of embryos:
   NOTE: In this procedure for culturing cochlear explants from embryonic mice, harvest inner ears from embryonic day (E) 13 temporal bones with the day after fertilization being considered as E1⁹. The protocol can also be used with embryonic mouse inner ears beginning from E12 to E18.
   1. Euthanize a timed-pregnant mouse with CO₂ and sacrifice animal by cervical dislocation or appropriate approved protocols. Carry out euthanization away from laminar-flow tissue culture hood to maintain sterility in the tissue culture room.
   2. Place euthanized mouse on paper towel with abdomen facing up and disinfect abdomen with 70% ethanol.
   3. Open the abdominal cavity by grabbing skin with curved forceps with one hand and cut epidermis and muscle along the midline with scissors using another hand. Make two perpendicular cuts on both sides and pull out the uterus carefully by lifting up bilateral uterine horns with forceps and detach it from underneath connective tissue using scissors.
   4. Place the embryonic chain in a petri dish containing chilled dissection solution (HBSS/HEPES mixture) and transfer to the laminar-flow clean bench.
   5. Remove embryos carefully from placenta and place them into one of the sterile Petri dishes containing dissection solution. If the solution turns bloody, move them to a new Petri dish.
      NOTE: At this point, embryos can be staged using a Theiler staging guide.
   6. Decapitate embryos by pinching at the neck region with a pair of clean forceps or scissors and place the heads in a fresh Petri dish containing chilled dissection solution.
3. Dissection of the inner ears:
   1. Place one embryo head in a sterile sylgard dish containing cold dissecting
   2. solution. Working under a dissecting microscope at a magnification of ~1.6X, immobilize the embryo head by placing Minutien pins around or closer to eye region (Figure 1A).
   3. Using two pairs of sterile forceps carefully remove the skin and open up the skull on the dorsal side along the midline (Figure 1B). Remove brain from cranial cavity and inner ears located within temporal bones can be identified at this stage (Figure 1C, D). The lining of the blood vessel, which is usually present around the inner ears, can be used as a landmark for identifying the inner ear at this stage (Figure 1D).
   4. Dissect the inner ear from the temporal bones by placing the forceps underneath the tissue and isolating the inner ear from base of the skull and transfer to a new dish containing cold dissecting solution (Figure 1E, F).

2. Generation of Organ of Corti Explant Cultures

NOTE: At this stage of development, the tissue is cartilaginous and can be easily dissected using forceps.

1. Orient the inner ear so that ventral side is facing up (Figure 2A) and stabilize the inner ear by gently inserting the sharp end of Minutien pins through the vestibular portion of the inner ear (Figure 2B).
2. Using ultrafine forceps cut open the overlying cartilage by making an incision using one end of forceps near the oval window and carefully remove the cartilage from the cochlea (Figure 2C).
   NOTE: It is important to make sure that forceps are not inserted too deeply into the cartilage as overlying cartilage is sometimes fused with the cochlear duct in which case it is helpful to use closed forceps to release cartilage from the underlying cochlear duct by scraping below the surface of the cartilage. At this stage of development, the cochlear spiral is only three quarters of a turn in length.
3. Next, expose the sensory epithelium by placing the forceps at the preferred region of the cochlear duct, either the base or at the very apex, and gently pulling out the roof of the cochlea (Figure 2D).
   NOTE: The roof of the cochlea must be removed completely as it can mask the sensory epithelium hindering the analysis.
   NOTE: This procedure must be done gently as it is easy to tear the sensory epithelium.
4. As a last step, carefully remove the underlying connective tissue from the exposed sensory epithelium so that the base of the cochlear explant is uniformly flat.
5. Isolate the dissected cochlea from the vestibular portion of the inner ear using forceps. This can be performed before or after step 2.4 (Figure 2E).
6. Transfer the dissected cochlear sensory epithelium into a basement membrane matrix-coated culture well using a 1.5 mm sterile scooper (Figure 2F).
7. Orient the cochlear explant with the lumenal surface of the epithelium facing up and aspirate the basement membrane matrix-DMEM solution to carefully flatten the tissue and replace it with 150 µl of fresh culture medium by gently adding to the dish. Care should be taken to make sure that each explant adheres well to the coated glass coverslip and is not floating in the culture medium. Ensure that the tip of the forceps is pointed up while affixing to the glass well to avoid damaging the cochlear explant.
8. Gently transfer the culture dish into a sterile 150 mm culture dish and place in a tissue culture incubator at 37 °C with 5% CO₂ for desired period of time, usually 3-6 days in vitro (DIV). After 1 DIV, examine the cultures under a dissecting microscope to make sure that the cochlear explant has adhered well to the culture dish.
9. After incubation in vitro for desired lengths of time, cultures are processed for immunohistochemistry.
   NOTE: The explant cultures usually are incubated for 6 DIV which is equivalent to developmental stage, P0. After incubation in vitro, cultures can be processed for further downstream applications like immunohistochemistry (Figure 3B), RT-PCR, in situ hybridization, and/or western blot.

3. Electroporation-mediated Gene Transfer

1. Setting up for electroporation:
   1. Sterilize one 100 mm Sylgard-coated glass dish by autoclaving or soaking in 70% ethanol for about 20 min and allow to air dry in a clean bench prior to use.
   2. Prepare DNA from an expression vector of choice using appropriate maxi- or midi-prep kits. The expression vectors used in this protocol are pIRES2-Atoh1.EGFP and pCLIG-NeuroD1.EGFP. The final concentration of DNA should be at least 1 µg/µl in sterile DNA/RNAse-free water.
2. Electroporating DNA into embryonic cochlear explants:
   1. Add 10 µl of plasmid DNA solution to a fresh 100 mm Sylgard coated dish and transfer a fully dissected cochlear explant (obtained from step 2.5) into the DNA solution.
   2. With lumenal surface of the epithelium facing up, tilt the cochlea slightly so that it’s perpendicular to the plane of the dish.
   3. Place the electrode paddles on either side of the cochlea with negative paddle (cathode) towards the sensory epithelium and the positive paddle (anode) located adjacent to the base of the cochlea.
      NOTE: Since DNA is negatively charged, it migrates from negative to positive electrode and by placing the cathode end of the probe adjacent to the sensory epithelium; the DNA will mostly go through the surface of the sensory epithelium.
   4. Using electroporator, deliver 9 to 10 pulses of 24 mV, 30 msec pulse duration with the foot pedal switch on the electroporator, and then add 100 µl of warm culture medium to the electroporated cochlea.
   5. Repeat steps 3.2.1-4 for all the cochlear explants and for DNA of all genes of interest and transfer the electroporated cochleae into the basement membrane matrix-coated dish for plating (step 2.7). Culture all the electroporated cochleae in a 37 °C, 5% CO₂ humidified incubator for desired lengths of time which will then be processed for immunocytochemistry.
      NOTE: The cochlea from an entire litter of CD1 mouse (approximately 8-12 pups) can be isolated, microdissected, electroporated and plated in <4 hr.

4. Analysis of Cochlear Explant Cultures

NOTE: The cultures are usually incubated for 6 DIV which is equivalent to developmental stage, P0. After incubation in vitro, the cultures are fixed and processed for immunocytochemistry.

1. After desired length of incubation, remove culture medium and quickly rinse the cochlear explant in phosphate-buffered saline (PBS) and incubate in 4% paraformaldehyde (prepared in PBS) for 15 min at room temperature.
2. Wash the fixative by adding PBS and rinse in PBS three times for 15 min.
3. Permeabilize using PBS-T (PBS + 0.5% Tween) for 30 min before blocking with PBS-T containing 10% goat serum.
4. Incubate in primary antibody solution containing primary antibody in PBS-T with 1% goat serum overnight at 4 °C.
5. Wash the cochlea three times at room temperature using PBS-T 15 min each before adding appropriate Alexa-conjugated secondary antibodies.
6. Since the cochlear explant is cultured on glass coverslip it can be handled individually by isolating the coverslip from the dish and mounting on a slide. This can be achieved by soaking the culture dish in OS30 solvent for at least 1 hr at room temperature. Using the sterile razor blade, detach the coverslip from the dish and mount on a slide and visualize under a fluorescent microscope.