We would like to thank Drs. Leslayann Schecterson, Yuan Wang, Andres Barria and Mrs. Ximena Optiz-Araya for initial assistance with protocol set-up and for providing plasmids. This work was supported by NIH/NIDCD grant DC013841 (JTS).

All procedures were approved by Northwestern University Institutional Animal Care and Use Committees, and carried out in accordance with the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals.
1. Egg Handling
<ol>
	<li>Purchase fertilized eggs from a local vendor. Store eggs at 13 &#176;C in a refrigerator for no longer than 5 days prior to incubation. Embryo viability significantly decreases after 1 week.</li>
	<li>Swab each egg with 70% ethanol before setting in the incubator.</li>
	<li>Set 1-2 dozen(s) of eggs on their sides in an incubator with the temperature set to 38 &#176;C at ~50% humidity. This ensures proper embryo positioning for electroporation and optimal viability, respectively. 
	NOTE: After 48-52 h of incubation, the eggs will be at Hamburger-Hamilton (HH) stage ~12-13 and ready for electroporation.</li>
</ol>
2. Preparation
<ol>
	<li>Plasmid Mixing
	<ol>
		<li>Add 1 &#181;L of 0.1% fast green (0.1% in 18 M&#937; deionized and distilled H2O [ddH2O]) for every 7 &#181;L of DNA mixture.</li>
		<li>For co-electroporation of multiple plasmids, mix plasmids in a 1:1 ratio. The final DNA concentration should be 5-8 &#181;g/&#181;L.</li>
	</ol>
	</li>
	<li>Fill Injection Pipette
	<ol>
		<li>Pull pipettes on a micropipette puller. Fill the pipette with 1-2 &#181;L plasmids using a 28G syringe needle.</li>
		<li>Break pipette tip to 10 - 20 &#181;m using forceps. Use a microscope to assist in determining the size of the broken pipette tip. Attach the pipette to the pipette holder of the picospritzer.</li>
	</ol>
	</li>
</ol>
3. Windowing
NOTE: The windowing procedure has been published before13. Please see reference 13 for additional visual information.
<ol>
	<li>Wipe all instruments and the work area with 70% ethanol.</li>
	<li>Take the desired number of set eggs out of the incubator (between 6-10), leave at room temperature, and individually swab eggs with 70% ethanol again. Eggs remain viable for at least 2 h after incubator removal.</li>
	<li>Place the egg on top of a light illuminator to identify embryo positioning. Circle the center of the dark area (i.e., embryo) with a pencil.</li>
	<li>Using a 19G needle, poke a hole in the blunt end of the egg.</li>
	<li>Poke another hole near the drawn circle on the pointed side of the egg. Remove a small piece of the outer eggshell (about 16 mm2) with the 19G needle and then remove a small piece of the inner eggshell (about 8 mm2) with forceps.</li>
	<li>With a 3 mL syringe fitted with a 19G needle, withdraw 1.5-2.5 mL of albumin from the egg through the blunt-end hole. Be sure to angle the needle down at 45&#176;.</li>
	<li>Cover the blunt-end hole with a small piece of tape (1 cm x 1 cm). Cover the drawn circle and the second hole with tape (2.5 cm x 4 cm).</li>
	<li>With curved scissors (cutting edge = 2.5 cm), cut a window within the circle. Scissors should be parallel to the eggshell and the diameter of the window should be less than 2 cm.</li>
</ol>
4. Plasmid Injection
<ol>
	<li>Turn on the picospritzer. Set the pressure to 18 psi and the duration to 5 &#181;s. Turn on the air tank and lamp for the dissection microscope.</li>
	<li>Make 30 mL stock solution of a 1:10 dilution of store-bought Indian ink mixed in sterile phosphate-buffered saline (PBS). Store at 4 &#176;C. Injection of Indian ink is an important step to get a better visualization of the chicken embryo.
	<ol>
		<li>Fill a 1 mL syringe with the diluted Indian ink solution. Attach a 27G needle and expel any bubbles that are present in the syringe.</li>
		<li>Insert needle just under the yolk membrane ~2 - 3 mm from the embryo and inject ~0.2 mL of Indian ink gently under the embryo. Do not inject more than 0.5 mL of the Indian ink as it may decrease embryo survival.</li>
	</ol>
	</li>
	<li>Place the windowed egg on an egg holder under the dissection microscope. Use the highest magnification for best visualization of plasmid injection site.</li>
	<li>Using forceps, remove the membrane over the injection area. Auditory brainstem regions of interest (i.e., NM and NL) arise from Rhombomeres 5 and 6 (R5/R6). R5/R6 are located adjacent to the otocysts (an embryonic structure in vertebrates that develops into the inner ear) that serve as landmarks for R5/R6 plasmid injections.</li>
	<li>Lower the DNA-filled pipette into the neural tube overlying the R5/R6 region and between otocysts using the micromanipulator. A schematic of ideal placement is shown in Figure 1A.</li>
	<li>Apply air pressure (18 psi, 5 &#181;s duration) from the picospritzer to eject DNA in the neural tube.</li>
</ol>
5. Electroporation
<ol>
	<li>Turn on the current/voltage stimulator.</li>
	<li>Fill a 3 mL syringe with PBS and attach the syringe filter. Add 1-2 drops of PBS onto the embryo.</li>
	<li>Lower the bipolar electrode to the embryo using the micromanipulator with the negative electrode above the injection site (medial) and the positive electrode lateral to the R5/R6 (Figure 1A). Avoid direct contact with the embryo. Use bipolar electrodes where the electrode material is platinum iridium. Adjust the spacing between the tips to 0.5 mm with forceps.</li>
	<li>Using a current/voltage stimulator, pulse 20 times at 50 V for 1 ms at 1 s intervals.</li>
	<li>After electroporation, add one drop of PBS over the exposed area. Gently remove the electrode and clean it with a laboratory tissue and 70% ethanol. Close the window exposing the embryo with tape.</li>
	<li>Label eggshell with the injected plasmid type and the hatch date.</li>
	<li>Place eggs back into incubator with window side up. Incubate at 38 &#176;C with 50% humidity until desired developmental stage is reached.</li>
	<li>If a tet-on vector is electroporated for temporal control of gene expression, apply Doxycycline (Dox) every 24 h to induce gene expression (see Section 6).</li>
</ol>
6. Tet-on System for Temporal Control of Gene Expression
<ol>
	<li>Use the following plasmids: 
	pCAGGS-T2TP: a transposase expressing plasmid. 
	pT2K-CAGGS-rtTA-M2: a plasmid that stably expresses the Dox-binding protein. 
	pT2K-BI-TRE-EGFP: a plasmid that contains a bidirectional tet-on promoter (TRE)-driving expression of the EGFP reporter and a second gene of interest. 
	NOTE: The above three plasmids need to be co-electroporated in a 1:1:1 ratio.</li>
	<li>Stock solution preparation:
	<ol>
		<li>Dissolve Dox in sterile PBS to produce a 1 mg/mL stock solution. Store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Dox application:
	<ol>
		<li>To induce the expression of the gene of interest during certain developmental stages, apply Dox every 24 h.</li>
		<li>When applying Dox, take the egg out, open the window, pipette 50 &#181;L Dox onto the chorioallointoic membrane, close the window and put the egg back into the incubator.</li>
	</ol>
	</li>
</ol>
7. Preparation of Dissecting Area for Brainstem Slices
NOTE: The following sections (7 - 9) and procedures have been published before14. Please see these references for additional visual information.
<ol>
	<li>Prepare an agar solution (40 mg/mL agarose, i.e. 4% in ddH2O). Pour agar solution into a Petri dish and allow it to solidify at room temperature. Store covered agar plate in the refrigerator for future use during brainstem slicing of tissue.</li>
	<li>Continually bubble artificial cerebral spinal fluid (ACSF) with 95% O2 and 5% CO2 (pH 7.2-7.4 and osmolarity: 295-310 mOsm/L).</li>
	<li>Clean the working area and vibratome with 70% EtOH and rinse the blade with distilled water.</li>
	<li>Place a clean fluid absorption pad on the working area with appropriate dissecting tools.</li>
	<li>Take the agar plate out of the refrigerator, cut a small agar cube (10 mm x 10 mm x 8 mm). Glue the agar block to the stage of a vibratome-slicing chamber using commercially available superglue.</li>
</ol>
8. Isolation of Chicken Auditory Brainstem
<ol>
	<li>Open the egg at taped window site. Puncture the membrane sac with a scalpel and remove the head of the chicken embryo.</li>
	<li>Decapitate the chicken with sharp scissors.</li>
	<li>Place the razor blade tip slightly posterior to the eyes. Incise through the skull at the rostral-to-caudal midline. Apply slight pressure depending on the age of the embryo. Older embryos require more pressure.</li>
	<li>Gently push aside skin and feathers to expose skull and verify midline cut.</li>
	<li>Applying strong pressure, slice the rostral portion of skull with a razor blade. Immediately place blade posterior to eyes and cut through entire skull and brain tissue.</li>
	<li>Make midline-to-lateral incisions with scissors in the skull&#34;s caudal region, slightly anterior to neck muscles on both sides of the head. Expose brain and cerebellum by pulling away skull and excess tissue.</li>
	<li>Cut tissue attached to the brainstem with scissors and remove the brainstem, which freely detaches from the skull.</li>
</ol>
9. Preparation of Brainstem Slices for In Vivo Electrophysiology or Imaging
<ol>
	<li>Pin down brainstem through the optic tecta.</li>
	<li>Remove the cerebellum by cutting peduncles with scissors, exposing the floor of the fourth ventricle.</li>
	<li>Using tweezers, remove any membranous tissue and blood vessels from the brainstem surface.</li>
	<li>To block the brainstem, make a horizontal cut at the most rostral end of the floor of the fourth ventricle, just caudal to the cortex. Make lateral cuts perpendicular through the optic tecta to isolate brainstem.</li>
	<li>Place a small amount of super glue on the vibratome stage directly in front of the agar block.</li>
	<li>Lift the brainstem at the spinal cord using tweezers and place it on the super glue with rostral side down and dorsal side towards the vibratome blade. Remove excess glue with a laboratory tissue or filter paper.</li>
	<li>Pour oxygenated ACSF into the vibratome stage. Set the vibratome blade at a 20-22&#176; angle. Start vibratome at maximum-amplitude oscillation. Move the stage up towards the blade so the top of the tissue is parallel with the blade.</li>
	<li>Rapidly move blade towards brainstem tissue. Slow down the blade considerably prior to the blade&#34;s contact with tissue.</li>
	<li>Slice tissue with the slowest possible forward speed. Once the blade is through the entire coronal tissue section, gently remove the slice using a transfer pipette.</li>
	<li>Lower the stage 200-300 &#181;m and slice again. Repeat until anatomical landmarks of auditory nuclei become visible, e.g., the distinct dorsal/ventral neuropil region of NL and the medial location of NM relative to NL.</li>
	<li>Carefully maintain slices in ACFS. This can be done at room temperature (22&#176; C) or near physiological conditions (~41&#176; C) using a warm water bath. Note the order of slicing. The first slice corresponds to the most caudal region of tissue and the last slice corresponds to the most rostral region. This roughly represents the tonotopic organization of the auditory brainstem, from low- to high-frequency encoding regions, respectively.</li>
</ol>

<ol>
	<li>Grothe, B., Pecka, M., McAlpine, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+sound+localization+in+mammals.">Mechanisms of sound localization in mammals.</a> Physiol Rev. 90 (3), 983-1012 (2010).</li><li>Anderson, S., 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=Neural+timing+is+linked+to+speech+perception+in+noise.">Neural timing is linked to speech perception in noise.</a> J Neurosci. 30 (14), 4922-4926 (2010).</li><li>Shannon, R. V., 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=Speech+recognition+with+primarily+temporal+cues.">Speech recognition with primarily temporal cues.</a> Science. 270 (5234), 303-304 (1995).</li><li>Carr, C. E., 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=Evolution+and+development+of+time+coding+systems.">Evolution and development of time coding systems.</a> Curr Opin Neurobiol. 11 (6), 727-733 (2001).</li><li>Carr, C. E., Soares, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+convergence+and+shared+computational+principles+in+the+auditory+system.">Evolutionary convergence and shared computational principles in the auditory system.</a> Brain Behav Evol. 59 (5-6), 294-311 (2002).</li><li>Rubel, E. W., Parks, T. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+and+development+of+brain+stem+auditory+nuclei+of+the+chicken:+tonotopic+organization+of+n.+magnocellularis+and+n.+laminaris.">Organization and development of brain stem auditory nuclei of the chicken: tonotopic organization of n. magnocellularis and n. laminaris.</a> J Comp Neurol. 164 (4), 411-433 (1975).</li><li>Rubel, E. W., Smith, D. J., Miller, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+and+development+of+brain+stem+auditory+nuclei+of+the+chicken:+ontogeny+of+n.+magnocellularis+and+n.+laminaris.">Organization and development of brain stem auditory nuclei of the chicken: ontogeny of n. magnocellularis and n. laminaris.</a> J Comp Neurol. 166 (4), 469-489 (1976).</li><li>Schecterson, L. C., 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=TrkB+downregulation+is+required+for+dendrite+retraction+in+developing+neurons+of+chicken+nucleus+magnocellularis.">TrkB downregulation is required for dendrite retraction in developing neurons of chicken nucleus magnocellularis.</a> J Neurosci. 32 (40), 14000-14009 (2012).</li><li>Chesnutt, C., Niswander, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasmid-based+short-hairpin+RNA+interference+in+the+chicken+embryo.">Plasmid-based short-hairpin RNA interference in the chicken embryo.</a> Genesis. 39 (2), 73-78 (2004).</li><li>Oertel, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Encoding+of+timing+in+the+brain+stem+auditory+nuclei+of+vertebrates.">Encoding of timing in the brain stem auditory nuclei of vertebrates.</a> Neuron. 19 (5), 959-962 (1997).</li><li>Cramer, K. S., Fraser, S. E., Rubel, E. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryonic+origins+of+auditory+brain-stem+nuclei+in+the+chick+hindbrain.">Embryonic origins of auditory brain-stem nuclei in the chick hindbrain.</a> Dev Biol. 224 (2), 138-151 (2000).</li><li>Cramer, K. S., 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=EphA4+signaling+promotes+axon+segregation+in+the+developing+auditory+system.">EphA4 signaling promotes axon segregation in the developing auditory system.</a> Dev Biol. 269 (1), 26-35 (2004).</li><li>Korn, M. J., Cramer, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Windowing+chicken+eggs+for+developmental+studies.">Windowing chicken eggs for developmental studies.</a> J Vis Exp. (8), e306 (2007).</li><li>Sanchez, J. T., 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=Preparation+and+culture+of+chicken+auditory+brainstem+slices.">Preparation and culture of chicken auditory brainstem slices.</a> J Vis Exp. (49), (2011).</li><li>Jhaveri, S., Morest, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+architecture+in+nucleus+magnocellularis+of+the+chicken+auditory+system+with+observations+on+nucleus+laminaris:+a+light+and+electron+microscope+study.">Neuronal architecture in nucleus magnocellularis of the chicken auditory system with observations on nucleus laminaris: a light and electron microscope study.</a> Neuroscience. 7 (4), 809-836 (1982).</li><li>Matsui, R., Tanabe, Y., Watanabe, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Avian+adeno-associated+virus+vector+efficiently+transduces+neurons+in+the+embryonic+and+post-embryonic+chicken+brain.">Avian adeno-associated virus vector efficiently transduces neurons in the embryonic and post-embryonic chicken brain.</a> PLoS One. 7 (11), e48730 (2012).</li><li>Koppl, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Auditory+nerve+terminals+in+the+cochlear+nucleus+magnocellularis:+differences+between+low+and+high+frequencies.">Auditory nerve terminals in the cochlear nucleus magnocellularis: differences between low and high frequencies.</a> J Comp Neurol. 339 (3), 438-446 (1994).</li><li>Hyson, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+analysis+of+interaural+time+differences+in+the+chick+brain+stem.">The analysis of interaural time differences in the chick brain stem.</a> Physiol Behav. 86 (3), 297-305 (2005).</li><li>Jones, T. A., Jones, S. M., Paggett, K. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emergence+of+hearing+in+the+chicken+embryo.">Emergence of hearing in the chicken embryo.</a> J Neurophysiol. 96 (1), 128-141 (2006).</li><li>Saunders, J. C., Coles, R. B., Gates, G. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+of+auditory+evoked+responses+in+the+cochlea+and+cochlear+nuclei+of+the+chick.">The development of auditory evoked responses in the cochlea and cochlear nuclei of the chick.</a> Brain Res. 63, 59-74 (1973).</li><li>Woolf, N. K., Ryan, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+of+auditory+function+in+the+cochlea+of+the+mongolian+gerbil.">The development of auditory function in the cochlea of the mongolian gerbil.</a> Hear Res. 13 (3), 277-283 (1984).</li><li>Walsh, E. J., McGee, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Postnatal+development+of+auditory+nerve+and+cochlear+nucleus+neuronal+responses+in+kittens.">Postnatal development of auditory nerve and cochlear nucleus neuronal responses in kittens.</a> Hear Res. 28 (1), 97-116 (1987).</li><li>Uziel, A., Romand, R., Marot, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+cochlear+potentials+in+rats.">Development of cochlear potentials in rats.</a> Audiology. 20 (2), 89-100 (1981).</li></ol>

The authors have nothing to disclose.

In ovo electroporation is a method of expressing or knocking down genes of interest in order to analyze in vivo gene function8,9. In the chicken embryo, it is an innovative method for expressing plasmid-encoded genes into different auditory brainstem regions8. To ensure optimal expression, several critical steps are required. First, only inject embryos whose otocysts are clearly visible. If otocysts are not visible the embryo is not at the correct developmental stage for electroporation. Second, smaller pipette tips make it easier to penetrate the neural tube and results in less tissue damage. However, the broken tip should be large enough to expel the DNA. Third, fill the entire neural tube region overlying R5/R6 with DNA (as visualized by spread of the fast green solution). Finally, placement of the stimulator electrode as close to the embryo as possible is important. However, make sure to avoid direct contact with the embryo, as current pulses can scorch and damage tissue.
Modifications to the technique can help improve the viability of embryos and the expression of plasmids. First, it is important to set eggs within 24 h of arrival to maintain high viability. Second, the window of the egg must be tightly sealed with tape as loss of fluid (i.e., dehydration) through an inadequate seal increases embryo death. Third, to optimize visualization of the otocyst, inject Indian ink from the rostral side of the embryo, using the smallest amount of ink as possible. Finally, stimulating electrodes can be moved slightly between R5/R6 regions to spread the electrically charged area without increasing current strength.
Despite the suggestions above, it should be noted that limitations exist. These include variable rates of transfected neurons, their location within the brainstem slice, and embryo survival. Although transfection efficiency varies amongst embryos, we previously reported with the electroporation parameter described here ~5-10% of NM/NL neurons are consistently transfected8. Even at higher rates of neuronal transfection, the preparation of brainstem tissue for in vitro patch-clamp electrophysiology poses challenges. For example, brainstem slices are between 200 - 300 &#181;m thick and occasionally, transfected neurons are too deep within the tissue to adequately patch and perform functional assays. Finally, embryo survival post successful electroporation is also variable, albeit not as much as transfection rate. Depending on the desired embryo age for experimental use, survival variability can rage between 80-20%, the latter of which corresponds to the oldest embryos (&#62;E18).
The significance of in ovo electroporation in chicken is advantageous over mammalian model systems for several reasons16. First, it is relatively inexpensive and an excellent alternative to transgenic or knockdown animals. Second, the chicken shares analogous nuclei with mammals that process temporal information of sound at the cellular, synaptic, and neural network level17. An example of similar auditory function occurs between AVCN/MSO in mammals and NM/NL of chickens, analogous auditory brainstem structures, respectively4,5. However, unlike traditional high-frequency-hearing mammalian research models, such as mice and rats, chickens utilize cues provided by both low- and high-frequency signals to encode auditory information18. Finally, chickens are auditory precocious animals; that is, their auditory system is near functional maturation at birth and the onset and refinement of hearing occurs during embryonic stages without environmental influences19,20. In contrast, the onset of hearing for typical mammalian research models begins 10-13 days after birth21,23.
In conjunction with biochemical, pharmacological, and functional assays, in ovo genetic manipulations provide an innovative approach to study neuron-specific development of anatomical and physiological specializations, permitting control of well-defined time periods in auditory brainstem development.

Fast neural encoding of sound is essential for normal auditory functions. These include sound localization abilities1, speech in noise discrimination2, and the comprehension of other behaviorally relevant communication signals3. Analogous neurons located in the auditory brainstem of both avians and mammals are highly specialized for fast neural encoding4. These include the chicken nucleus magnocellularis (NM), the nucleus laminaris (NL) and their mammalian analogs, the anteroventral cochlear nucleus (AVCN) and the medial superior olive (MSO), respectively5. However, developmental mechanisms regulating fast neural encoding are poorly understood in the auditory brainstem. Therefore, it is advantageous to study specific genes that are responsible for fast neural encoding in order to better understand their expression, regulation and function in auditory development.
The developing chicken embryo is an effective and well-established research tool to study basic biological questions of auditory system development6,7. Recent molecular advances have addressed these biological questions in the developing chicken embryo by expressing or knocking down genes of interest in order to analyze in vivo gene function8,9. Investigating the regulatory role of specific genes is a significant advancement in understanding pathologies associated with auditory deficits. Here, we present in ovo electroporation of plasmid-encoded genes into the chicken auditory brainstem where fast neural encoding of sound occurs10. By targeting auditory neural progenitor regions associated with rhombomeres 5 and 611,12 (R5/R6), we show spatial control of plasmid transfection in NM and NL. In addition, we show temporal regulation of expression 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)8.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Fertilized white leghorn chicken eggs</td>
			<td>Sunnyside Inc. (Beaver Dam, WI)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Picospritzer</td>
			<td>Parker Hannifin</td>
			<td>052-0500-900</td>
			<td>Picospritzer III, single or dual channel</td>
		</tr>
		<tr>
			<td>Current/voltage stimulator</td>
			<td>Grass Technologies</td>
			<td>SD9</td>
			<td>SD9</td>
		</tr>
		<tr>
			<td>Microfil syringe needles</td>
			<td>World Precision Instruments</td>
			<td>MF28G67-5</td>
			<td>28 Gauge, 67 mm Long, (Pack of 5)</td>
		</tr>
		<tr>
			<td>Electrode holder</td>
			<td>Warner Instruments</td>
			<td>64-1280</td>
			<td>MP Series: Non-Electrical Pressure Applications</td>
		</tr>
		<tr>
			<td>Stimulating microelectrode</td>
			<td>FHC</td>
			<td>PBSA1075</td>
			<td>PBSA1075</td>
		</tr>
		<tr>
			<td>Air tank/regulator</td>
			<td>NU Laboratory Services</td>
			<td></td>
			<td>Air dry 300 CF</td>
		</tr>
		<tr>
			<td>Fast green</td>
			<td>Sigma Aldrich</td>
			<td>F7258-25G</td>
			<td>F7258-25G</td>
		</tr>
		<tr>
			<td>Clear plastic tape</td>
			<td>Scotch</td>
			<td>191</td>
			<td></td>
		</tr>
		<tr>
			<td>Doxycycline hyclate</td>
			<td>Sigma Aldrich</td>
			<td>D9891-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Egg refrigerator</td>
			<td>Vissani Wine Refrigerator</td>
			<td></td>
			<td>13.3-16.1&#176; C (56-61&#176; F)</td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>Hova-Bator</td>
			<td></td>
			<td>37.8&#176; C (100&#176; F), ~50% humidity</td>
		</tr>
		<tr>
			<td>Dissection scope</td>
			<td>Zeiss</td>
			<td>4.35E+15</td>
			<td>SteREO Discovery, V8 Microscope, 50.4X</td>
		</tr>
		<tr>
			<td>Cold-light source</td>
			<td>Zeiss</td>
			<td>4.36E+15</td>
			<td>CL6000 LED</td>
		</tr>
		<tr>
			<td>Micromanipulators</td>
			<td>Narishige Japan</td>
			<td>Model: MM-3</td>
			<td>2 Micromanipulators</td>
		</tr>
		<tr>
			<td>Capillary tubes</td>
			<td>Sutter Instrument</td>
			<td>BF150-86-10</td>
			<td>Thick-walled borosilicate (dimensions)</td>
		</tr>
		<tr>
			<td>Syringes</td>
			<td></td>
			<td></td>
			<td>1 mL, 3 mL</td>
		</tr>
		<tr>
			<td>Needles</td>
			<td>BD Precision Glide&#160;</td>
			<td></td>
			<td>27 G x 1 1/4, 19 G x 1 1/2</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Stoelting</td>
			<td></td>
			<td>No. 5 Super Fine Dumont</td>
		</tr>
		<tr>
			<td>Egg holder</td>
			<td>Custom Made</td>
			<td></td>
			<td>Clay base works as well</td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instrument</td>
			<td></td>
			<td>Model P-97</td>
		</tr>
		<tr>
			<td>Syringe filter</td>
			<td>Ultra Cruz</td>
			<td>sc-358811</td>
			<td>PVDF 0.22 &#956;m</td>
		</tr>
	</tbody>
</table>

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.

We show here that in ovo electroporation permits gene expression in a normally developing biological system. Plasmid-encoded genes are focally injected into the neural tube overlying R5/R6. A schematic example of the electrode and pipette placements relative to important anatomical markers is shown in Figure 1A. The correct location of plasmid injection is confirmed 24 h after electroporation and shown in Figure 1B. The targeted injection of plasmids into the neural tube overlying R5/R6 permits expression in specific auditory brainstem regions, namely NM and NL(11). Figure 1C shows an in vitro brainstem slice from an E12 embryo under differential interference contrast (DIC) illumination. The auditory regions of interest are outlined. With florescent illumination (Figure 1C1), YFP expressing neurons within NM and NL are visible. Figure 1D shows DIC illumination for an E18 embryo under high magnification. The classic adendritic cell bodies of numerous NM neurons are apparent. With florescent illumination (Figure 1D1), an YFP expressing NM neuron is clearly visible while other transfected neurons are out of the focal plane. Because only a fraction of neurons in the targeted NM region are transfected, it allows other neurons within the same nucleus to serve as internal controls during dual patch-clamp electrophysiology experiments. Transgenes can also be temporally regulated by drug application, enabling gene expression and manipulation at precise developmental time periods8.
<img alt="Figure 1" src="/files/ftp_upload/55628/55628fig1.jpg" /> 
Figure 1: In Ovo Electroporation in the Chicken Auditory Brainstem. (A) Schematic of electroporation in the hindbrain of HH stage 12 chicken embryos. The injection pipette is filled with plasmid DNA and fast green dye for visualization of the injected hindbrain (rhombomeres 5/6 [R5/R6]). In this study, a plasmid that coexpressed a gene of interest and the yellow-fluorescent protein (YFP) reporter was used. Electroporated embryos are further incubated until desired developmental stage is reached. (B) Embryonic (E) day 4 chicken embryo showing site of YFP expression relative to the left eye (arrow, E) and left otocyst (arrow, O). Dashed white line represents the dorsal brain and brainstem border. Scale bar = 480 &#181;m. (C and C1) Brainstem slice (300 &#181;m thick) from an E12 chicken under differential interference contrast (DIC, C) and fluorescent (YFP, C1) illumination. Dashed white lines represent borders of nucleus magnocellularis (NM) and nucleus laminaris (NL). Note, the brainstem slice shows only one side of the tissue. White arrowheads in represents midline cleft of the brainstem slice. White arrows show YFP expressing regions of NM and NL. V = ventral, M = medial. Scale bar = 240 &#181;m. (D and D1) High magnification (80X water immersion objective) of an E18 chicken embryo brainstem slice containing NM. The classic adendritic cell bodies15 are clearly visible under DIC illumination (upward arrow, D). With fluorescent illumination for the same slice, a transfected NM neuron identified by YFP fluorescence is clearly visible (upward arrow, D1). Asterisks = YFP expressing NM neurons just below the focal plane. Scale bar = 30 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/55628/55628fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Ovo Electroporation in the Chicken Auditory Brainstem at JoVE.com

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.

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 ...

in-ovo-electroporation-in-the-chicken-auditory-brainstem

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

8.4K Views.  Northwestern University. 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. 

Video: In Ovo Electroporation in the Chicken Auditory Brainstem

Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

The cone photoreceptor-enriched cultures derived from embryonic chick retinas have become an indispensable tool for researchers around the world studying the biology of retinal neurons, particularly photoreceptors. The applications of this system go beyond basic research, as they can easily be adapted to high throughput technologies for drug development. However, genetic manipulation of retinal photoreceptors in these cultures has proven to be very challenging, posing an important limitation to the usefulness of the system. We have recently developed and validated an ex&#160;ovo plasmid electroporation technique that increases the rate of transfection of retinal cells in these cultures by five-fold compared to other currently available protocols1. In this method embryonic chick eyes are enucleated at stage 27, the RPE is removed, and the retinal cup is placed in a plasmid-containing solution and electroporated using easily constructed custom-made electrodes. The retinas are then dissociated and cultured using standard procedures. This technique can be applied to overexpression studies as well as to the downregulation of gene expression, for example via the use of plasmid-driven RNAi technology, commonly achieving transgene expression in 25% of the photoreceptor population. The video format of the present publication will make this technology easily accessible to researchers in the field, enabling the study of gene function in primary retinal cultures. We have also included detailed explanations of the critical steps of this procedure for a successful outcome and reproducibility.

Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

Embryonic chick retinal cell cultures constitute valuable tools for the study of photoreceptor biology. We have developed an efficient gene transfer technique based on ex&#160;ovo plasmid electroporation of the retinas prior to culture. This technique considerably increases transfection efficiencies over currently available protocols, making genetic manipulation feasible for this system.

The cone photoreceptor-enriched cultures derived from embryonic chick retinas have become an indispensable tool for researchers around the world studying ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Whole Retinal Explants from Chicken Embryos for Electroporation and Chemical Reagent Treatments

The retina is a good model for the developing central nervous system. The large size of the eye and most importantly the accessibility for experimental manipulations in ovo/in vivo makes the chicken embryonic retina a versatile and very efficient experimental model. Although the chicken retina is easy to target in ovo by intraocular injections or electroporation, the effective and exact concentration of the reagents within the retina may be difficult to fully control. This may be due to variations of the exact injection site, leakage from the eye or uneven diffusion of the substances. Furthermore, the frequency of malformations and mortality after invasive manipulations such as electroporation is rather high.
This protocol describes an ex ovo technique for culturing whole retinal explants from chicken embryos and provides a method for controlled exposure of the retina to reagents. The protocol describes how to dissect, experimentally manipulate, and culture whole retinal explants from chicken embryos. The explants can be cultured for approximately 24 hr and be subjected to different manipulations such as electroporation. The major advantages are that the experiment is not dependent on the survival of the embryo and that the concentration of the introduced reagent can be varied and controlled in order to determine and optimize the effective concentration. Furthermore, the technique is rapid, cheap and together with its high experimental success rate, it ensures reproducible results. It should be emphasized that it serves as an excellent complement to experiments performed in ovo.

This protocol describes a method to dissect, experimentally manipulate and culture whole retinal explants from chicken embryos. The explant cultures are useful when high success rate, efficacy and reproducibility are needed to test the effects of plasmids for electroporation and/or reagent substances, i.e., enzymatic inhibitors.

The retina is a good model for the developing central nervous system. The large size of the eye and most importantly the accessibility for experimental ...

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

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. ...

A Novel Ex Ovo Banding Technique to Alter Intracardiac Hemodynamics in an Embryonic Chicken System

The new model presented here can be used to understand the influence of hemodynamics on specific cardiac developmental processes, at the cellular and molecular level. To alter intracardiac hemodynamics, fertilized chicken eggs are incubated in a humidified chamber to obtain embryos of the desired stage (HH17). Once this developmental stage is achieved, the embryo is maintained ex ovo and hemodynamics in the embryonic heart are altered by partially constricting the outflow tract (OFT) with a surgical suture at the junction of the OFT and ventricle (OVJ). Control embryos are also cultured ex ovo but are not subjected to the surgical intervention. Banded and control embryos are then incubated in a humidified incubator for the desired period of time, after which 2D ultrasound is employed to analyze the change in blood flow velocity at the OVJ as a result of OFT banding. Once embryos are maintained ex ovo, it is important to ensure adequate hydration in the incubation chamber so as to prevent drying and eventually embryo death. Using this new banded model, it is now possible to perform analyses of changes in the expression of key players involved in valve development and to understand the role of hemodynamics on cellular responses in vivo, which could not be achieved previously.

A Novel Ex Ovo Banding Technique to Alter Intracardiac Hemodynamics in an Embryonic Chicken System

For the first time we present here a reproducible banding procedure to alter hemodynamics in the developing heart ex ovo. This is achieved by partially constricting the outflow tract (OFT).

The new model presented here can be used to understand the influence of hemodynamics on specific cardiac developmental processes, at the cellular and ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain and protect cellular identities. The conversion of germ cells into specific neurons in the nematode Caenorhabditis elegans (C. elegans) is a powerful tool for performing genetic screens in order to dissect regulatory pathways that safeguard established cell identities. Reprogramming of germ cells to a specific type of neurons termed ASE requires transgenic animals that allow broad over-expression of the Zn-finger transcription factor (TF) CHE-1. Endogenous CHE-1 is expressed exclusively in two head neurons and is required to specify the glutamatergic ASE neurons fate, which can easily be visualized by the gcy-5prom::gfp reporter. A trans gene containing the heat-shock promoter-driven che-1 gene expression construct allows broad mis-expression of CHE-1 in the entire animal upon heat-shock treatment. The combination of RNAi against the chromatin-regulating factor LIN-53 and heat-shock-induced che-1 over-expression leads to reprogramming of germ cell into ASE neuron-like cells. We describe here the specific RNAi procedure and appropriate conditions for heat-shock treatment of transgenic animals in order to successfully induce germ cell to neuron conversion.

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

This protocol describes how to study cellular processes during cell fate conversion in Caenorhabditis elegans in vivo. Using transgenic animals, allowing heat-shock promoter-driven overexpression of the neuron fate-inducing transcription factor CHE-1 and RNAi-mediated depletion of the chromatin-regulating factor LIN-53 germ cell to neuron reprogramming can be observed in vivo.

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...