This study was supported by: a National Natural Science Foundation of China grant (No. 32000697); the Science and Technology Program of Guangzhou (202102080139); the Guangdong Natural Science Foundation (2019A1515110625, 2021A1515010619); the Fundamental Research Funds for the Central Universities (11620324); a Research Grant of Key Laboratory of Regenerative Medicine, Ministry of Education, Jinan University (No. ZSYXM202107); the Fundamental Research Funds for the Central Universities of China (21621054); and the Medical Scientific Research Foundation of Guangdong Province of China (20191118142729581). We thank the medical experimental center of Jinan University. We thank Dr. Terra Bradley for careful editing of the manuscript.

Eggs and chicken embryos were handled with care and respect in accordance with the animal protocols approved by the Jinan University Animal Care and Use Committee.
1. Egg and plasmid preparation
<ol>
	<li>Egg preparation
	<ol>
		<li>Obtain fresh fertilized chicken eggs (Gallus gallus) from the South China Agricultural University and store at 16 &#176;C before incubation. For optimal viability, set all eggs for incubation within a week of arrival.</li>
		<li>Place eggs horizontally and incubate at 38 &#176;C for 46-48 h until Hamburger and Hamilton (HH) stage 1225 for neural tube transfection, or for 54-56 h until HH13 for otocyst transfection. Because the animal pole of the egg is lighter than the vegetal pole, horizontal placement positions the embryo on the top of the egg for easy manipulation. Use caution to keep the egg horizontal in this and all following steps.</li>
		<li>The location of the embryo appears as a dark area on the eggshell when casting light from the bottom of the egg using a flashlight. At the desired HH stages, use this flashlight method to mark the location of the embryo on the eggshell with pencil.</li>
		<li>Wipe the eggs with gauze containing 75% ethanol. Drill a hole in the pointy end of the eggs using the tip of scissors and remove 2 mL of albumin with an 18 G needle syringe. Ensure the hole is only large enough to allow needle insertion. Wipe away any leaking albumin with gauze and seal the hole with clear tape.</li>
		<li>To minimize cracks and to prevent falling shell debris during windowing, cover the top of eggs with clear tape, centering on the pencil-marked dark area.</li>
	</ol>
	</li>
	<li>Plasmid DNA extraction
	<ol>
		<li>Clone chicken Fmr1 shRNA using a transposon-based Tet-on system as described previously20. This system contains three plasmids: pCAGGS-T2TP, pT2K-CAGGS-rtTA-M2, and pT2K-BI-TRE-EGFP-Fmr1 shRNA, providing a drug-inducible vector system by which the expression of Fmr1 shRNA and EGFP is silenced after transfection and can be turned on by doxycycline (Dox) induction. 
		&#8203;NOTE: These plasmids are not currently available in a repository. The authors agree to provide the plasmids upon reasonable request.</li>
		<li>Extract plasmid DNAs using an endotoxin-free preparation kit according to manufacturer&#39;s instruction and precipitate by adding 1/15 volume of 7.5 M sodium acetate and 1 volume of 100% isopropanol.</li>
		<li>Precipitate plasmids at -20 &#176;C overnight and centrifuge at 13,000 x g for 10 min. Dissolve the pellet with the Tris-EDTA buffer provided in the kit to a final concentration of 4-5 &#181;g/&#181;L. Plasmids can be stored at -20 &#176;C for 12 months without reduced transfection efficiency.</li>
		<li>On the day of electroporation, thoroughly mix 2 &#181;L of each plasmid in a sterile centrifuge tube using a pipette tip. The resultant 6 &#181;L volume of the plasmid mixture is enough to electroporate three dozen eggs. To help visualize the plasmid during electroporation, add 1 &#181;L of 0.1% fast green (prepared in sterile ddH2O) for every 6 &#181;L of DNA mixture26, which turns the plasmid mixture blue.</li>
	</ol>
	</li>
</ol>
2. In ovo electroporation
<ol>
	<li>Egg windowing and embryo identification
	<ol>
		<li>On the day of electroporation, remove the eggs from the incubator, one dozen eggs at a time. Keeping eggs out of their incubator for more than 1 h introduces developmental variations and reduces viability.</li>
		<li>Wipe all surgery tools with gauze containing 75% ethanol.</li>
		<li>Place each egg in a custom-made foam holder. Wipe the tape-covered area with 75% ethanol and cut a window (1-2 cm2) around the circumference of the pencil mark using a small pair of scissors (Figure 1A). When cutting, hold the scissors flat to avoid damaging the embryo underneath.</li>
		<li>Place the windowed egg under a stereomicroscope with a 10x eyepiece and 2x zoom and identify the embryo. Adjust the angle and brightness of the light source for better visualization. 
		NOTE: It takes practice and experience to visualize the embryo and identify the neural tube at this stage.
		<ol>
			<li>For beginners, use the following optional ink injection to facilitate embryo identification. Prepare 10% Indian ink solution in 0.01 M phosphate-buffered saline (PBS) and autoclave in advance. Fill a 1 mL syringe with the ink solution and then fit a 27G needle. Bend the needle to a 45&#176; angle with forceps.</li>
			<li>Under the microscope, carefully poke from the edge of the area opaca and insert the needle beneath the embryo. Inject ~50 &#181;L of ink, which will diffuse below the area pellucida for embryo visualization. The ink will form a dark background for a clear visualization of the embryo. 
			NOTE: Expel the air from the syringe before insertion and injection. When skilled at visualizing, avoid the ink injection step as it reduces the survival rate.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Neural tube injection and electroporation 
	NOTE: This procedure is performed at HH12 for transfecting NM neurons in the brainstem.
	<ol>
		<li>Pull glass capillaries (~1 mm in diameter and 100 mm long) into pipettes using a pipette puller. Under a dissecting microscope, carefully open the tip of the capillary needle to 10-20 &#181;m in diameter with forceps. Store the pipettes (usually 5-10 at one time) in a storage box until use.</li>
		<li>Fill a capillary pipette with 0.5-1 &#181;L of the plasmid mixture by applying negative pressure through a rubber tube at the end of the pipette. This can be achieved by a syringe or air pump.</li>
		<li>Place the egg under the microscope so that the embryo is vertically oriented with the tail near to you (or horizontally for injecting capillary pipettes using a three-axis manipulator). Hold the capillary pipette with one hand or with the three-axis manipulator and drive the tip of the pipette to the r5/6 in a tail-to-head direction. 
		NOTE: The most anterior hindbrain area is r1 and each subsequent bulge along the neural tube is an individual rhombomere. R5/6 is at the same anteroposterior level as the otocyst, which is a readily recognizable cup-like structure (Figure 1B-C).</li>
		<li>Gently poke the tip through the vitelline membrane and into the dorsal neural tube and then withdraw the pipette a bit so the tip is in the lumen of the neural tube. Inject the plasmid mixture by applying air pressure until the tinted plasmid diffuses fully into r5/6 and extends into r3 and r4. 
		NOTE: It is not necessary to make a slot on the vitelline membrane for injection.</li>
		<li>Check for a successful injection, which is achieved when the blue plasmid solution rapidly diffuses down the neural tube without leaking (Figure 1B-C). When leaking occurs, the blue quickly fades.</li>
		<li>Immediately after the injection, place a platinum bipolar electrode on either side of the neural tube (Figure 1D). Deliver two pulses of 12 V and 50 ms duration with an electroporator. Observe air bubbles at the ends of the bipolar electrode, with more on the negative side.</li>
		<li>Check for successful electroporation, which is achieved when the tinted plasmid mixture enters the neural tube tissue near the positive side of the electrode. After electroporation, carefully remove the bipolar electrode.</li>
		<li>Cover the window on the eggshell with a piece of transparent film that is pre-cut into 2 in squares and sprayed with 75% ethanol. Place the egg back into its incubator.</li>
		<li>Clean the bipolar electrode by delivering 10-20 pulses of 12 V and 50 ms duration in saline before proceeding to the next egg.</li>
	</ol>
	</li>
	<li>Otocyst injection and electroporation 
	NOTE: This procedure is performed at HH13 for transfecting hair cells and AG neurons in the inner ear.
	<ol>
		<li>At HH13 (which is ~embryonic day 2.5), the embryo has turned so the right side of the head faces up. Under the microscope, place the egg so that the embryo is vertical, with the tail near to you. Hold the capillary pipette and gently poke the right otocyst in a dorsolateral direction (Figure 2A). 
		NOTE: At this stage, the otocyst appears as a small circular structure on the top of the body at the same anteroposterior level as r5/6.</li>
		<li>Inject the plasmid mixture with air pressure until the otocyst is filled with blue solution27. Check for a successful injection, which is achieved when the blue plasmid mixture is confined within the otocyst and doesn&#39;t leak.</li>
		<li>Immediately after the injection, place the bipolar electrode on the otocyst as depicted in Figure 2B. Position the positive and negative sides anterior and posterior to the otocyst, respectively. Deliver two pulses of 12 V and 50 ms duration with the electroporator.</li>
		<li>Check for successful electroporation, which is achieved when the blue plasmid mixture enters the tissue of the otocyst near the positive side of the electrode. After electroporation, carefully remove the bipolar electrode. Cover the window on the eggshell with a transparent film and return the egg to the incubator.</li>
	</ol>
	</li>
</ol>
3. Administration of Dox to initiate and maintain plasmid transcription
<ol>
	<li>Preparation of the Dox solution
	<ol>
		<li>Measure 100 mg of Dox powder under a chemical hood and dissolve it in 100 mL of sterile PBS to make a 1 mg/mL of working solution. Filter the solution with a 0.22 &#181;m filter and store 1 mL aliquots at -20 &#176;C. Protect from light20,28.</li>
	</ol>
	</li>
	<li>Administration of Dox
	<ol>
		<li>For full-strength gene editing, at 24 h before the desired age, thaw a Dox aliquot on ice. Drop 50 &#181;L of Dox directly on the chorioallantoic membrane of the egg using a syringe penetrating the transparent film. Seal the needle hole with transparent film or tape after injection.</li>
		<li>To maintain the knockdown effect, administer Dox every other day until tissue harvest at the desired developmental stage.</li>
	</ol>
	</li>
</ol>
4. Tissue dissection and sectioning
<ol>
	<li>Brainstem
	<ol>
		<li>For embryos at embryonic stage 3 (E3) and E6, open the eggshell and cut the associated membrane with scissors. Spoon the embryo out and immerse it in 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer.</li>
		<li>For E9 embryos, open the eggshell, decapitate the embryo with scissors, and transfer the head to a silicon-bottomed plate filled with PBS. Pin the head down through the orbits and cut the top of the skull apart. Carefully place the forceps beneath the brain from both sides and elevate the entire brain from the skull with the shoulders of the forceps. Immerse the brain in 4% PFA.</li>
		<li>For E15 and E19 embryos, open the eggshell, decapitate the embryo with scissors, and place the head on a silicon-bottomed plate filled with PBS. Cut open the skin on top of the head to expose the skull.</li>
		<li>Incise through the skull vertically with a razor to separate the forebrain and the caudal brain. Immerse the caudal block in PBS, remove the top of the skull, and then take the brain off the skull.</li>
		<li>Pin the brain down through the optic lobe and separate the cerebellum and brainstem. Take care not to damage the auditory brainstem, which is located right below the cerebellum. Remove as much dura as possible from the brainstem and then immerse it in 4% PFA.</li>
		<li>Keep embryos and brain tissues in 4% PFA at 4 &#176;C overnight except for E19 brainstems, which should be kept under the same condition for 24 h.</li>
		<li>Following fixation, dehydrate the tissue in PBS containing 30% sucrose until it settles, which usually takes 1-3 days, depending on the age.</li>
		<li>Section the brainstem (E15 and E19) at 30 &#181;m at the coronal plane using a sliding microtome. Collect sections in PBS and store at 4 &#176;C before immunostaining.</li>
		<li>For E3 and E6 embryos and E9 brainstems, section at 50 &#181;m transversely. Collect sections in PBS and store at 4 &#176;C. Mount the sections on gelatin-coated microscope slides before immunostaining. Collecting thicker sections for these ages facilitates tissue handling.</li>
	</ol>
	</li>
	<li>Auditory duct
	<ol>
		<li>After removing the brainstems from E9, E15, and E19 embryos, the auditory duct, which is embedded in the temporal bone, is readily identifiable lying beneath the skull, and the temporal bone is easily separatable from the surrounding skull. For E9 embryos, fix the whole temporal bone in 4% PFA. For older embryos, remove the bony structure of the temporal bone to isolate the auditory duct and fix the auditory duct in 4% PFA.</li>
		<li>Keep all temporal bones and auditory ducts in 4% PFA at 4 &#176;C overnight before tissue embedding.</li>
		<li>Perform tissue embedding as described. Prepare the embedding solution as follows: Soak 10% gelatin (from bovine skin) in cold water until the gelatin granules swell and settle (about 30 min). Heat the mixture to ~50 &#176;C to dissolve the gelatin and add 20% sucrose into the gelatin solution and stir to dissolve. Store the gelatin-sucrose solution at 4 &#176;C for up to a month.</li>
		<li>For use, warm the gelatin-sucrose solution at 37 &#176;C. Soak the temporal bones/auditory ducts in the warm gelatin-sucrose solution on a 96-well plate at 37 &#176;C until they settle, usually 30-60 min.</li>
		<li>Line the bottom of the wells of a 12-well plate with a strip of transparent film. Add a layer of warm gelatin-sucrose solution and wait until it solidifies. Transfer a temporal bone/auditory duct to each well.</li>
		<li>Implant the tissue with a second layer of warm gelatin-sucrose solution. Adjust the tissue&#39;s position so that it is in the center of the gel and the auditory duct is oriented horizontally. Carefully move the plate into a 4 &#176;C refrigerator and wait until the gel is firm.</li>
		<li>Pull the transparent film strip to move the gel-tissue block out of the well. Trim extra gel into a square block and cut a corner of the block to identify the orientation. Wrap the gelatin block with a piece of foil, freeze on dry ice, and then store at -80 &#176;C until sectioning.</li>
		<li>Section the block at 20 &#181;m along the longitude of the auditory duct with a cryostat. Mount the sections directly on gelatin-coated microscope slides. Store the slides at -80 &#176;C before immunostaining.</li>
		<li>Prior to immunostaining, immerse the slides in pre-warmed PBS (45 &#176;C) for 5 min to dissolve the gelatin, and then wash the slides 3x with room temperature PBS to remove residual gelatin.</li>
	</ol>
	</li>
</ol>
5. Immunostaining and microscope imaging
NOTE: Two types of immunostaining are performed depending on whether sections are mounted on slides or free-floating in PBS.
<ol>
	<li>Immunostaining on slides
	<ol>
		<li>Wash the slides 3x for 10 min each with PBS. Circle the region containing the sections with an oil pen to prevent liquid leaking during staining. Cover the sections with a blocking solution containing 5% normal goat serum in PBS and incubate for 30 min at room temperature.</li>
		<li>Dilute the primary antibodies (for the final concentration used refer to Table of Materials) in PBS containing 0.3% TritonX-100 and 5% normal goat serum. Remove the blocking solution and then cover the sections with the primary antibody solution. Incubate the slides at 4 &#176;C overnight in a box containing a layer of distilled water at the bottom.</li>
		<li>Rinse the slides 3x for 10 min with PBS. Apply fluorescent secondary antibodies diluted at 1:500 in PBS containing 0.3% TritonX-100 on slides. Incubate the slides at room temperature for 4 h, shielded from light.</li>
		<li>Wash the slides 3x with PBS. Gently drop two drops of mounting medium on the slides and cover the sections with coverslips. Take care not to generate air bubbles between the coverslip and the slide.</li>
	</ol>
	</li>
	<li>Immunostaining for whole-mount embryos and free-floating sections
	<ol>
		<li>Wash the embryos/sections with PBS 3x for 10 min each in a well plate. Dilute the primary antibodies in PBS containing 0.3% TritonX-100 and 5% normal goat serum in centrifugal tubes. Transfer each embryo/section into a tube with a glass hook and incubate on a shaker at 60 rpm overnight at 4 &#176;C.</li>
		<li>Wash embryos/sections 3x with PBS for 10 min each in a well plate. Transfer each embryo/section into a dark centrifugal tube filled with fluorescent secondary antibody solution and incubate at room temperature for 4 h on the shaker.</li>
		<li>Wash the embryos/sections 3x with PBS in a well plate. Use a brush to mount the sections on gelatin-coated microscope slides in a 15 cm dish containing PBS. After the slides are dried slightly (~5 min), apply two drops of mounting medium and place a coverslip. Take care not to generate air bubbles between the coverslip and the slide. Keep the whole mount tissues in PBS for imaging.</li>
	</ol>
	</li>
	<li>Imaging
	<ol>
		<li>Image the whole mount tissues in PBS in a dish with a silicon bottom containing carbon powder, which provides a black background. Capture and process images using a commercial image processing Olympus software package, as described previously29.</li>
		<li>Image the sections on the slides with a confocal microscope as described previously20. Image the sections at a single focal plane with 10x, 20x, and 63x objectives. Capture all images from the same animal using the same parameters. Take care to adjust the laser level and imaging acquisition settings to avoid signal saturation.</li>
		<li>Perform image processing using the Fiji software. For illustration, adjust image brightness, contrast, and gamma using a professional image editing software.</li>
	</ol>
	</li>
</ol>

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R., Schecterson, L., Lu, Y., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Postsynaptic+FMRP+regulates+synaptogenesis+in+vivo+in+the+developing+cochlear+nucleus.">Postsynaptic FMRP regulates synaptogenesis in vivo in the developing cochlear nucleus.</a> Journal of Neuroscience. 38 (29), 6445-6460 (2018).</li><li>Wang, X., 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=Temporal-specific+roles+of+fragile+X+mental+retardation+protein+in+the+development+of+the+hindbrain+auditory+circuit.">Temporal-specific roles of fragile X mental retardation protein in the development of the hindbrain auditory circuit.</a> Development. 147 (21), (2020).</li><li>Rubel, E. W., Fritzsch, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Auditory+system+development:+primary+auditory+neurons+and+their+targets.">Auditory system development: primary auditory neurons and their targets.</a> Annual Review of Neuroscience. 25, 51-101 (2002).</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> Developmental Biology. 224 (2), 138-151 (2000).</li><li>Chervenak, A. P., Hakim, I. S., Barald, K. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatiotemporal+expression+of+Zic+genes+during+vertebrate+inner+ear+development.">Spatiotemporal expression of Zic genes during vertebrate inner ear development.</a> Developmental Dynamics. 242 (7), 897-908 (2013).</li><li>Hamburger, V., Hamilton, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+series+of+normal+stages+in+the+development+of+the+chick+embryo.">A series of normal stages in the development of the chick embryo.</a> Journal of Morphology. 88 (1), 49-92 (1951).</li><li>Lu, T., Cohen, A. L., Sanchez, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+ovo+electroporation+in+the+chicken+auditory+brainstem.">In ovo electroporation in the chicken auditory brainstem.</a> Journal of Visualized Experiments. (124), e56628 (2017).</li><li>Evsen, L., Doetzlhofer, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+transfer+into+the+chicken+auditory+organ+by+in+ovo+micro-electroporation.">Gene transfer into the chicken auditory organ by in ovo micro-electroporation.</a> Journal of Visualized Experiments. (110), e53864 (2016).</li><li>Schecterson, L. C., Sanchez, J. T., Rubel, E. W., Bothwell, M. <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> Journal of Neuroscience. 32 (40), 14000-14009 (2012).</li><li>Wang, X. Y., 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=High+glucose+environment+inhibits+cranial+neural+crest+survival+by+activating+excessive+autophagy+in+the+chick+embryo.">High glucose environment inhibits cranial neural crest survival by activating excessive autophagy in the chick embryo.</a> Scientific Reports. 5, 18321 (2015).</li><li>Yu, X., Wang, X., Sakano, H., Zorio, D. A. R., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+the+fragile+X+mental+retardation+protein+correlates+with+cellular+and+synaptic+properties+in+primary+auditory+neurons+following+afferent+deprivation.">Dynamics of the fragile X mental retardation protein correlates with cellular and synaptic properties in primary auditory neurons following afferent deprivation.</a> Journal of Comparative Neurology. 529 (3), 481-500 (2021).</li><li>Li, H., 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=Islet-1+expression+in+the+developing+chicken+inner+ear.">Islet-1 expression in the developing chicken inner ear.</a> Journal of Comparative Neurology. 477 (1), 1-10 (2004).</li><li>Carr, C. E., Boudreau, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Central+projections+of+auditory+nerve+fibers+in+the+barn+owl.">Central projections of auditory nerve fibers in the barn owl.</a> Journal of Comparative Neurology. 314 (2), 306-318 (1991).</li><li>Sandell, L. L., Butler Tjaden, N. E., Barlow, A. J., Trainor, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cochleovestibular+nerve+development+is+integrated+with+migratory+neural+crest+cells.">Cochleovestibular nerve development is integrated with migratory neural crest cells.</a> Developmental Biology. 385 (2), 200-210 (2014).</li><li>Cramer, K. S., Bermingham-McDonogh, O., Krull, C. 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=EphA4+signaling+promotes+axon+segregation+in+the+developing+auditory+system.">EphA4 signaling promotes axon segregation in the developing auditory system.</a> Developmental Biology. 269 (1), 26-35 (2004).</li><li>Evsen, L., Sugahara, S., Uchikawa, M., Kondoh, H., Wu, 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=Progression+of+neurogenesis+in+the+inner+ear+requires+inhibition+of+Sox2+transcription+by+neurogenin1+and+neurod1.">Progression of neurogenesis in the inner ear requires inhibition of Sox2 transcription by neurogenin1 and neurod1.</a> Journal of Neuroscience. 33 (9), 3879-3890 (2013).</li><li>Curnow, E., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+animal+models+for+understanding+FMRP+functions+and+FXS+pathology.">New animal models for understanding FMRP functions and FXS pathology.</a> Cells. 11 (10), 1628 (2022).</li></ol>

The authors have nothing to disclose.

To determine the cell-autonomous function of FMRP, manipulating its expression in individual cell groups or cell types is necessary. Since one of the major functions of FMRP is to regulate synaptic formation and plasticity, selectively manipulating each synaptic component of a certain circuit is prerequisite for a full understanding of FMRP mechanism in synaptic communication. Using in ovo electroporation of chicken embryos, we demonstrated a method to target FMRP expression in t...

Fragile X syndrome (FXS) is a neurodevelopmental disorder characterized by intellectual disability, sensory abnormalities, and autistic behaviors. In most cases, FXS is caused by a global loss of fragile X mental retardation protein (FMRP; encoded by Fmr1 gene) starting at early embryonic stages1. FMRP is an RNA-binding protein that is normally expressed in most neurons and glial cells in the brain, as well as in sensory organs2,3,4. In mammalian brains, FMRP is likely associated with hundreds of mRNAs that encode proteins that are important for various neural activities5. Studies of conventional Fmr1 knockout animals demonstrated that FMRP expression is particularly important to the assembly and plasticity of synaptic neurotransmission6. Several conditional and mosaic knockout models have further demonstrated that FMRP actions and signals vary across brain regions, cell types, and synaptic sites&#160;during several developmental events including axonal projection, dendritic patterning, and synaptic plasticity7,8,9,10,11,12,13,14. Acute function of FMRP in regulating synaptic transmission was studied by intracellular delivery of inhibitory FMRP antibodies or FMRP itself in brain slices or cultured neurons15,16,17,18. These methods, however, do not offer the ability to track FMRP misexpression-induced consequences during development. Thus, developing in vivo methods to investigate the cell-autonomous functions of FMRP is in great need, and expected to help determine whether the reported anomalies in FXS patients are direct consequences of FMRP loss in the associated neurons and circuits, or secondary consequences derived from network-wide changes during development19.
The auditory brainstem of chicken embryos offers a uniquely advantageous model for in-depth functional analyses of FMRP regulation in circuit and synapse development. The easy access to embryonic chicken brains and the well-established in ovo electroporation technique for genetic manipulation have contributed greatly to our understanding of brain development at early embryonic stages. In a recently published study, this technique was combined with advanced molecular tools that allow temporal control of FMRP misexpression20,21. Here, the methodology is advanced to induce selective manipulations of presynaptic and postsynaptic neurons separately. This method was developed in the auditory brainstem circuit. Acoustic signal is detected by hair cells in the auditory inner ear and then conveyed to the auditory ganglion (AG; also called the spiral ganglion in mammals). Bipolar neurons in the AG innervate hair cells with their peripheral processes and in turn send a central projection (the auditory nerve) to the brainstem where they terminate in two primary cochlear nuclei, the nucleus magnocellularis (NM) and the nucleus angularis (NA). Neurons in the NM are structurally and functionally comparable to the spherical bushy cells of the mammalian anteroventral cochlear nucleus. Within the NM, auditory nerve fibers (ANFs) synapse on the somata of NM neurons via the large endbulb of Held terminals22. During development, NM neurons arise from rhombomeres 5 and 6 (r5/6) in the hindbrain23, while AG neurons are derived from neuroblasts residing in the otocyst24. Here, we describe the procedure to selectively knockdown FMRP expression in the presynaptic AG neurons and in the postsynaptic NM neurons separately.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Egg incubation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>16 &#176;C refrigerator</td>
			<td>MAGAT</td>
			<td></td>
			<td>Used for fertilized egg storage.</td>
		</tr>
		<tr>
			<td>Egg incubator</td>
			<td>SHANGHAI BOXUN</td>
			<td>GZX-9240MBE</td>
			<td></td>
		</tr>
		<tr>
			<td>Fertilized eggs</td>
			<td>Farm of South China Agricultural University</td>
			<td></td>
			<td>Eggs must be used in one week for optimal viability.</td>
		</tr>
		<tr>
			<td>Plasmid preparation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Sigma</td>
			<td>10016</td>
			<td></td>
		</tr>
		<tr>
			<td>Fast green</td>
			<td>Solarbio</td>
			<td>G1661</td>
			<td>Make 0.1% working solution in distilled water and autoclave.</td>
		</tr>
		<tr>
			<td>Plasmid Maxi-prep kit</td>
			<td>QIAGEN</td>
			<td>12162</td>
			<td>Dissolve plasmid DNA in Tris-EDTA (TE) buffer; endotoxin-free preparation kit</td>
		</tr>
		<tr>
			<td>Sodium Acetate</td>
			<td>Sigma-Aldrich</td>
			<td>S2889</td>
			<td>Make 7.5M working solution in nuclase-free water.</td>
		</tr>
		<tr>
			<td>Electroporation and Doxycycline Administration</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electroporator</td>
			<td>BTX</td>
			<td>ECM399</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL / 5 mL Syringe</td>
			<td>GUANGZHOU KANGFULAI</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>CNOPTEC</td>
			<td>SZM-42</td>
			<td></td>
		</tr>
		<tr>
			<td>Doxcycline</td>
			<td>Sigma-Aldrich</td>
			<td>D9891</td>
			<td>Use fresh aliquots for each dose and store at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>Glass capillary</td>
			<td>BEIBOBOMEI</td>
			<td>RD0910</td>
			<td>0.9-1.1 mm*100 mm</td>
		</tr>
		<tr>
			<td>Laboratory parafilm</td>
			<td>PARAFILM</td>
			<td>PM996</td>
			<td>transparent film</td>
		</tr>
		<tr>
			<td>Pipette puller</td>
			<td>CHENGDU INSTRUMENT FACTORY</td>
			<td>WD-2</td>
			<td>Pulling condition: 500 &#176;C for 15 s</td>
		</tr>
		<tr>
			<td>Platinum elctrodes</td>
			<td>Home made</td>
			<td></td>
			<td>0.5 mm diameter, 1.5 mm interval.</td>
		</tr>
		<tr>
			<td>Platinum elctrodes</td>
			<td>Home made</td>
			<td></td>
			<td>0.5 mm diameter, 1.5 mm interval.</td>
		</tr>
		<tr>
			<td>Rubber tube</td>
			<td>Sigma-Aldrich</td>
			<td>A5177</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Dissection and Fixation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>RWD</td>
			<td>F11020-11</td>
			<td>Tip size: 0.05*0.01 mm</td>
		</tr>
		<tr>
			<td>Other surgery tools</td>
			<td>RWD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>158127</td>
			<td>Freshly made 4% PFA solution in phosphate-buffered saline can be stored in 4 &#176;C for up to 1 week.</td>
		</tr>
		<tr>
			<td>SYLGARD 184 Silicone Elastomer Kit</td>
			<td>DOW</td>
			<td>01673921</td>
			<td>For black background plates, food-grade carbon powder is applied.</td>
		</tr>
		<tr>
			<td>Sectioning</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cryostat</td>
			<td>LEICA</td>
			<td>CM1850</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin</td>
			<td>Sigma-Aldrich</td>
			<td>G9391</td>
			<td>From bovine skin.</td>
		</tr>
		<tr>
			<td>Sliding microtome</td>
			<td>LEICA</td>
			<td>SM2010</td>
			<td></td>
		</tr>
		<tr>
			<td>Immunostaining</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 goat anti-Mouse</td>
			<td>Abcam</td>
			<td>ab150113</td>
			<td>1:500 dilution, RRID: AB_2576208</td>
		</tr>
		<tr>
			<td>Alexa Fluor 555 goat anti-rabbit</td>
			<td>Abcam</td>
			<td>ab150078</td>
			<td>1:500 dilution, RRID: AB_2722519</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Abcam</td>
			<td>ab285390</td>
			<td>1: 1000 dilution</td>
		</tr>
		<tr>
			<td>Fluoromount-G mounting medium</td>
			<td>Southern&#160;Biotech</td>
			<td>Sb-0100-01</td>
			<td></td>
		</tr>
		<tr>
			<td>FMRP antibody</td>
			<td>Y. Wang, Florida State University</td>
			<td>#8263</td>
			<td>1:1000 dilution, RRID: AB_2861242</td>
		</tr>
		<tr>
			<td>Islet-1 antibody</td>
			<td>DSHB</td>
			<td>39.3F7</td>
			<td>1:100 dilution, RRID: AB_1157901</td>
		</tr>
		<tr>
			<td>Netwell plate</td>
			<td>Corning</td>
			<td>3478</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurofilament antibody</td>
			<td>Sigma-Aldrich</td>
			<td>N4142</td>
			<td>1:1000 dilution, RRID: AB_477272</td>
		</tr>
		<tr>
			<td>Parvalbumin antibody</td>
			<td>Sigma-Aldrich</td>
			<td>P3088</td>
			<td>1:10000 dilution, RRID: AB_477329</td>
		</tr>
		<tr>
			<td>SNAP25 antibody</td>
			<td>Abcam</td>
			<td>ab66066</td>
			<td>1:1000 dilution, RRID: AB_2192052</td>
		</tr>
		<tr>
			<td>Imaging</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Adobe photoshop</td>
			<td>ADOBE</td>
			<td></td>
			<td>image editing software</td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>LEICA</td>
			<td>SP8</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent stereomicroscope</td>
			<td>OLYMPUS</td>
			<td>MVX10</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus Image-Pro Plus 7.0</td>
			<td>OlYMPUS</td>
			<td></td>
			<td>commercial image processing software package</td>
		</tr>
	</tbody>
</table>

dissecting cell-autonomous function of fragile x mental retardation protein in an auditory circuit by in ovo electroporation

Fragile X mental retardation protein (FMRP) is an mRNA-binding protein that regulates local protein translation. FMRP loss or dysfunction leads to aberrant neuronal and synaptic activities in fragile X syndrome (FXS), which is characterized by intellectual disability, sensory abnormalities, and social communication problems. Studies of FMRP function and FXS pathogenesis have primarily been conducted with Fmr1 (the gene encoding FMRP) knockout in transgenic animals. Here we report an in vivo method for determining the cell-autonomous function of FMRP during the period of circuit assembly and synaptic formation using chicken embryos. This method employs stage-, site-, and direction-specific electroporation of a drug-inducible vector system containing Fmr1 small hairpin RNA (shRNA) and an EGFP reporter. With this method, we achieved selective FMRP knockdown in the auditory ganglion (AG) and in one of its brainstem targets, the nucleus magnocellularis (NM), thus providing a component-specific manipulation within the AG-NM circuit. Additionally, the mosaic pattern of the transfection allows within-animal controls and neighboring neuron/fiber comparisons for enhanced reliability and sensitivity in data analyzing. The inducible vector system provides temporal control of gene editing onset to minimize accumulating developmental effects. The combination of these strategies provides an innovative tool to dissect the cell-autonomous function of FMRP in synaptic and circuit development.

By performing in ovo electroporation at different sites and at different developmental stages, we achieved selective FMRP knockdown either in the auditory periphery or in the auditory brainstem.
FMRP knockdown in NM 
Small hairpin RNA (shRNA) against the chicken Fmr1 was designed and cloned into the Tet-On vector system as described previously20. The setup for in ovo electroporation is shown in Figure...

Watch this Scientific Journal Video about Dissecting Cell-Autonomous Function of Fragile X Mental Retardation Protein in an Auditory Circuit by In Ovo Electroporation at JoVE.com

Dissecting Cell-Autonomous Function of Fragile X Mental Retardation Protein in an Auditory Circuit by In Ovo Electroporation

Using in ovo electroporation, we devised a method to selectively transfect the auditory inner ear and cochlear nucleus in chicken embryos to achieve a cell-group-specific knockdown of fragile X mental retardation protein during discrete periods of circuit assembly.

Dissecting Cell-Autonomous Function of Fragile X Mental Retardation Protein in an Auditory Circuit by In Ovo Electroporation

Fragile X mental retardation protein (FMRP) is an mRNA-binding protein that regulates local protein translation. FMRP loss or dysfunction leads to ...

dissecting-cell-autonomous-function-fragile-x-mental-retardation

Research

JoVE Journal

Neuroscience

2.1K Views.  Jinan University. Using in ovo electroporation, we devised a method to selectively transfect the auditory inner ear and cochlear nucleus in chicken embryos to achieve a cell-group-specific knockdown of fragile X mental retardation protein during discrete periods of circuit assembly.

Video: Dissecting Cell-Autonomous Function of Fragile X Mental Retardation Protein in an Auditory Circuit by In Ovo Electroporation

In utero Electroporation followed by Primary Neuronal Culture for Studying Gene Function in Subset of Cortical Neurons

In vitro study of primary neuronal cultures allows for quantitative analyses of neurite outgrowth. In order to study how genetic alterations affect neuronal process outgrowth, shRNA or cDNA constructs can be introduced into primary neurons via chemical transfection or viral transduction. However, with primary cortical cells, a heterogeneous pool of cell types (glutamatergic neurons from different layers, inhibitory neurons, glial cells) are transfected using these methods. The use of in utero electroporation to introduce DNA constructs in the embryonic rodent cortex allows for certain subsets of cells to be targeted: while electroporation of early embryonic cortex targets deep layers of the cortex, electroporation at late embryonic timepoints targets more superficial layers. Further, differential placement of electrodes across the heads of individual embryos results in the targeting of dorsal-medial versus ventral-lateral regions of the cortex. Following electroporation, transfected cells can be dissected out, dissociated, and plated in vitro for quantitative analysis of neurite outgrowth. Here, we provide a step-by-step method to quantitatively measure neuronal process outgrowth in subsets of cortical cells.
The basic protocol for in utero electroporation has been described in detail in two other JoVE articles from the Kriegstein lab 1, 2. We will provide an overview of our protocol for in utero electroporation, focusing on the most important details, followed by a description of our protocol that applies in utero electroporation to the study of gene function in neuronal process outgrowth.

In utero Electroporation followed by Primary Neuronal Culture for Studying Gene Function in Subset of Cortical Neurons

In utero electroporation is a valuable method for transfecting neuronal progenitor cells in vivo. Depending upon the placement of the electrodes and the developmental timepoint of electroporation, certain subsets of cortical cells can be targeted. Targeted cells can then be analyzed in vivo or in vitro for effects of genetic alteration.

In vitro study of primary neuronal cultures allows for quantitative analyses of neurite outgrowth. In order to study how genetic alterations affect ...

TMS: Using the Theta-Burst Protocol to Explore Mechanism of Plasticity in Individuals with Fragile X Syndrome and Autism

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS display abnormalities in the expression of FMR1 - a protein required for typical, healthy neural development.3 Recent data has suggested that the loss of this protein can cause the cortex to be hyperexcitable thereby affecting overall patterns of neural plasticity.4,5
In addition, Fragile X shows a strong comorbidity with autism: in fact, 30% of children with FXS are diagnosed with autism, and 2 - 5% of autistic children suffer from FXS.6
Transcranial Magnetic Stimulation (a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability via the application of localized magnetic field pulses 7,8) represents a unique method of exploring plasticity and the manifestations of FXS within affected individuals. More specifically, Theta-Burst Stimulation (TBS), a specific stimulatory protocol shown to modulate cortical plasticity for a duration up to 30 minutes after stimulation cessation in healthy populations, has already proven an efficacious tool in the exploration of abnormal plasticity.9,10 
Recent studies have shown the effects of TBS last considerably longer in individuals on the autistic spectrum - up to 90 minutes.11 This extended effect-duration suggests an underlying abnormality in the brain's natural plasticity state in autistic individuals - similar to the hyperexcitability induced by Fragile X Syndrome.
In this experiment, utilizing single-pulse motor-evoked potentials (MEPs) as our benchmark, we will explore the effects of both intermittent and continuous TBS on cortical plasticity in individuals suffering from FXS and individuals on the Autistic Spectrum.

In this article, we examine the effects of Theta-Burst TMS stimulation on cortical plasticity in individuals suffering from Fragile X syndrome and individuals on the autistic spectrum.

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

In ovo Electroporation in Chick Midbrain for Studying Gene Function in Dopaminergic Neuron Development

Dopaminergic neurons located in the ventral midbrain control movement, emotional behavior, and reward mechanisms1-3. The dysfunction of ventral midbrain dopaminergic neurons is implicated in Parkinson's disease, Schizophrenia, depression, and dementia1-5. Thus, studying the regulation of midbrain dopaminergic neuron differentiation could not only provide important insight into mechanisms regulating midbrain development and neural progenitor fate specification, but also help develop new therapeutic strategies for treating a variety of human neurological disorders.
Dopaminergic neurons differentiate from neural progenitors lining the ventricular zone of embryonic ventral midbrain. The development of neural progenitors is controlled by gene expression programs6,7. Here we report techniques utilizing electroporation to express genes specifically in the midbrain of Hamburger Hamilton (HH) stage 11 (thirteen somites, 42 hours) chick embryos8,9. The external development of chick embryos allows for convenient experimental manipulations at specific embryonic stages, with the effects determined at later developmental time points10-13. Chick embryonic neural tubes earlier than HH stage 13 (nineteen somites, 48 hours) consist of multipotent neural progenitors that are capable of differentiating into distinct cell types of the nervous system. The pCAG vector, which contains both a CMV promoter and a chick &beta;-actin enhancer, allows for robust expression of Flag or other epitope-tagged constructs in embryonic chick neural tubes14. In this report, we emphasize special measures to achieve regionally restricted gene expression in embryonic midbrain dopaminergic neuron progenitors, including how to inject DNA constructs specifically into the embryonic midbrain region and how to pinpoint electroporation with small custom-made electrodes. Analyzing chick midbrain at later stages provides an excellent in vivo system for plasmid vector-mediated gain-of-function and loss-of-function studies of midbrain development. Modification of the experimental system may extend the assay to other parts of the nervous system for performing fate mapping analysis and for investigating the regulation of gene expression.

In ovo Electroporation in Chick Midbrain for Studying Gene Function in Dopaminergic Neuron Development

To assess the function and the regulation of genes during the development of midbrain dopaminergic neurons, we describe a method that involves in ovo electroporation of plasmid DNA constructs into embryonic chick ventral midbrain dopaminergic neuron progenitors. This technique can be used to achieve efficient expression of genes of interest to study different aspects of midbrain development and dopaminergic neuron differentiation.

Dopaminergic neurons located in the ventral midbrain control movement, emotional behavior, and reward mechanisms1-3. The dysfunction of ventral midbrain ...

In ovo Electroporation of miRNA-based Plasmids in the Developing Neural Tube and Assessment of Phenotypes by DiI Injection in Open-book Preparations

Commissural dI1 neurons have been extensively studied to elucidate the mechanisms underlying axon guidance during development1,2. These neurons are located in the dorsal spinal cord and send their axons along stereotyped trajectories. Commissural axons initially project ventrally towards and then across the floorplate. After crossing the midline, these axons make a sharp rostral turn and project longitudinally towards the brain. Each of these steps is regulated by the coordinated activities of attractive and repulsive guidance cues. The correct interpretation of these cues is crucial to the guidance of axons along their demarcated pathway. Thus, the physiological contribution of a particular molecule to commissural axon guidance is ideally investigated in the context of the living embryo. Accordingly, gene knockdown in vivo must be precisely controlled in order to carefully distinguish axon guidance activities of genes that may play multiple roles during development.
Here, we describe a method to knockdown gene expression in the chicken neural tube in a cell type-specific, traceable manner. We use novel plasmid vectors3 harboring cell type-specific promoters/enhancers that drive the expression of a fluorescent protein marker, followed directly by a miR30-RNAi transcript4 (located within the 3'-UTR of the cDNA encoding the fluorescent protein) (Figure 1). When electroporated into the developing neural tube, these vectors elicit efficient downregulation of gene expression and express bright fluorescent marker proteins to enable direct tracing of the cells experiencing knockdown3. Mixing different RNAi vectors prior to electroporation allows the simultaneous knockdown of two or more genes in independent regions of the spinal cord. This permits complex cellular and molecular interactions to be examined during development, in a manner that is fast, simple, precise and inexpensive. In combination with DiI tracing of commissural axon trajectories in open-book preparations5, this method is a useful tool for in vivo studies of the cellular and molecular mechanisms of commissural axon growth and guidance. In principle, any promoter/enhancer could be used, potentially making the technique more widely applicable for in vivo studies of gene function during development6.
This video first demonstrates how to handle and window eggs, the injection of DNA plasmids into the neural tube and the electroporation procedure. To investigate commissural axon guidance, the spinal cord is removed from the embryo as an open-book preparation, fixed, and injected with DiI to enable axon pathways to be traced. The spinal cord is mounted between coverslips and visualized using confocal microscopy.

In ovo Electroporation of miRNA-based Plasmids in the Developing Neural Tube and Assessment of Phenotypes by DiI Injection in Open-book Preparations

A method by which gene expression in the neural tube can be downregulated in a cell type-specific, traceable manner is described. We demonstrate how in ovo electroporation of microRNA-based plasmids that elicit spatiotemporally controlled RNA interference can be used to investigate commissural axon guidance in the developing neural tube.

Commissural dI1 neurons have been extensively studied to elucidate the mechanisms underlying axon guidance during development1,2. These neurons are ...

A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and sensorimotor plasticity1. Consistent sensory perturbations result in compensatory changes in motor output, reflecting changes in feedforward motor control that reduce the experienced feedback error. By quantifying how different sensory feedback errors affect human behavior, prior studies have explored how visual signals are used to recalibrate arm movements2,3 and auditory feedback is used to modify speech production4-7. The strength of this approach rests on the ability to mimic naturalistic errors in behavior, allowing the experimenter to observe how experienced errors in production are used to recalibrate motor output.
Songbirds provide an excellent animal model for investigating the neural basis of sensorimotor control and plasticity8,9. The songbird brain provides a well-defined circuit in which the areas necessary for song learning are spatially separated from those required for song production, and neural recording and lesion studies have made significant advances in understanding how different brain areas contribute to vocal behavior9-12. However, the lack of a naturalistic error-correction paradigm - in which a known acoustic parameter is perturbed by the experimenter and then corrected by the songbird - has made it difficult to understand the computations underlying vocal learning or how different elements of the neural circuit contribute to the correction of vocal errors13.
The technique described here gives the experimenter precise control over auditory feedback errors in singing birds, allowing the introduction of arbitrary sensory errors that can be used to drive vocal learning. Online sound-processing equipment is used to introduce a known perturbation to the acoustics of song, and a miniaturized headphones apparatus is used to replace a songbird's natural auditory feedback with the perturbed signal in real time. We have used this paradigm to perturb the fundamental frequency (pitch) of auditory feedback in adult songbirds, providing the first demonstration that adult birds maintain vocal performance using error correction14. The present protocol can be used to implement a wide range of sensory feedback perturbations (including but not limited to pitch shifts) to investigate the computational and neurophysiological basis of vocal learning.

We describe the design and assembly of miniaturized headphones suitable for replacing a songbird&#x2019;s natural auditory feedback with a manipulated acoustic signal. Online sound processing hardware is used to manipulate song output, introduce real-time errors in auditory feedback via the headphones, and drive vocal motor learning.

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and ...

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent improvements in methodologies and applications for the study of theta oscillations. A simple characterization of the isolated hippocampal preparation is presented whereby the relationship between internal hippocampal theta oscillators is examined together with the activity of pyramidal cells, and GABAergic interneurons, of the cornu ammonis-1 (CA1) and subiculum (SUB) areas. Overall, we show that the isolated hippocampus is capable of generating intrinsic theta oscillations in vitro and that rhythmicity generated within the hippocampus can be precisely manipulated by optogenetic stimulation of parvalbumin-positive (PV) interneurons. The in vitro isolated hippocampal preparation offers a unique opportunity to use simultaneous field and intracellular patch-clamp recordings from visually-identified neurons to better understand the mechanisms underlying theta rhythm generation.

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

Here, we present a protocol for recording rhythmic neuronal network theta and gamma oscillations from an isolated whole hippocampal preparation. We describe the experimental steps from extraction of the hippocampus to details of field, unitary and whole-cell patch clamp recordings as well as optogenetic pacing of the theta rhythm.

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent ...