The authors wish to thank Dr. Candace Hsieh for suggestions and assistance with imaging techniques and Dr. Doris Wu for expertise on chick inner ear anatomy during early embryogenesis. This work was supported by NSF IOS-0642346, NIH T32-DC010775, NIH T32-GM008620, NIH R01-DC010796, and DOE GAANN P200A120165.

1. Prepare the Following Dissection Tools and Reagents
<ul>
	<li>Artificial cerebrospinal fluid (aCSF; 130 mM NaCl, 3 mM KCl, 1.2 mM KH2PO4, 20 mM NaHCO3, 3 mM HEPES, 10 mM Glucose, 2 mM CaCl2, 1.3 mM MgSO4) continuously infused with 95% O2 / 5% CO2 at room temperature. For infusion, fill to 2/3 a 500 ml wide-mouth Nalgene jar with a hole drilled in the lid. Tank will be attached by tubing to a glass stem bubbler, which penetrates the aCSF through the hole in the jar lid. Adjust pressure to obtain a constant stream of bubbles that reach about 1/3 of the liquid but are not forceful enough to cause splashing over the rim. Submerge 5 ml glass vials (up to 6) into the Nalgene jar to separate individual samples from each other and from the gas flow turbulence. ACSF should be oxygenated for at least 20 min prior to any tissue incubation.</li>
	<li>Small silicone dissection dish (35 mm x 10 mm Petri coated using Sylgard Elastomere Kit) with two dissection pins placed into small hexagonal polystyrene (47 mm bottom) weigh boat.</li>
	<li>Two fine-tip forceps, one curved-tip forceps, a 50 ml beaker, and a container for tissue waste.</li>
	<li>Pulled glass micropipette (1.2 OD / 0.9 ID) broken to approximately 20-50 &mu;m opening at the tip and filled with rhodamine dextran amine (RDA, MW = 3,000, Invitrogen) in a 6.25% solution with 0.4% Triton X-100 in PBS. Attach by fine tubing to picospritzer and stabilize to micromanipulator.</li>
</ul>
2. Micro-dissection to Reveal Basilar Papilla at E8 4,5
<ol>
	<li>With curved forceps, cut E8 embryo just below the brainstem, placing head directly onto silicone-coated dissection dish within weigh boat.</li>
	<li>Using dissection pins, position the head with a ventral view so the ear on the side of interest is slightly visible, the head tilted slightly dorsally and pointing 10-30 degrees away from center (Figure 2A). Immediately secure the head with pins. If bilateral tracing is desired, center the head at 0 degrees, or a full ventral view so both sides can be accessed. Similarly, for a left-sided injection, secure the head at -10 to -30 degrees from center so the left side is accessible.</li>
	<li>Using the 50 ml beaker, transfer 30 ml of oxygenated aCSF into the dish, fully submerging the tissue.</li>
	<li>Gently grasp and peel down the lower jaw using fine-tip forceps, then remove the lower eyelid on the side of interest.</li>
	<li>Remove the overlying vascular tissue, including any remaining palate and gullet. These soft tissues should peel away revealing the smooth surface of developing chondrocranium with the distinctive vertebral arteries at its surface. By this point, the characteristic white otoconial mass should be visible.</li>
	<li>Carefully dissect the skin around the external meatus, cartilage of the jaw joint, and the middle ear while leaving the inner ear intact. Use the forceps to gently cut away these cartilaginous structures.</li>
	<li>Remove vertebral arteries and pooling blood using a gentle sweeping motion with the fine forceps tips. Blood can obscure view and coagulation may plug the injection pipette opening.</li>
	<li>The cochlear duct with adjacent sensory epithelium (basilar papilla) and acoustic ganglion lies directly beneath the chondrocranium 6 and can be visualized as an elongated finger-like projection extending from the inner ear ventrally, with the distal-most tip (apex) near the anterior midline 5. There may be a small pooling of blood (red) from the dissection and/or otolithic calcifications (white) at the apex of the cochlear duct (Figure 2B) that can aid in visualization. At the apex is the lagenar macula, a sensory patch thought to be of vestibular function 7,8.</li>
</ol>
3. Selective Labeling of CN VIII Auditory Fibers
<ol>
	<li>Place the injection pipette near the embryo surface at the cochlear duct and basilar papilla region.</li>
	<li>Slowly lower the micropipette to first puncture the cranium. This cranial region is thinner than surrounding areas and requires less force to penetrate. The micropipette should now be inside the cochlear duct. Optional: a small injection of tracing dye can be made here to aid in visualizing the basilar papilla.</li>
	<li>Without repositioning the micropipette, continue lowering until it just breaches the deep border of the cochlear duct (Figure 1). Perform an injection here and repeat in multiple regions along the length of the basilar papilla (Figures 2C, 2D), excluding the most proximal tip where the vestibular lagena is located. With the picospritzer set between 10-30 psi, several injections are made. The volume of injected dye fluid varies, and depends on the extent of labeling desired. Each injection should result in a fluorescent dye-labeled spot of approximately 200 - 400 &mu;m diameter (Figure 2D).</li>
	<li>Once the injection is complete, use forceps to transect the sample above the brainstem and use a transfer pipette to place the caudal portion into a small jar within the container of aCSF continually being perfused with 95% O2 / 5% CO2.</li>
	<li>For each embryo, allow 20-30 min for dye transfer, depending on the distance of central tracing desired.</li>
	<li>Remove tissue and prepare for histological sectioning. In the example shown here, tissue is immersed in 4% paraformaldehyde (in 1X PBS) overnight, 30% sucrose (in 1X PBS) overnight. </li>
</ol>
4. Counterstaining and Analysis
<ol>
	<li>Counterstaining is useful to help orient the observer to anatomical location as well as identify particular regions of interest. An effective counterstain in this preparation is neurofilament immunolabeling (Figures 1, 3B, 3E), which strongly labels axons and allows for demarcation of axonal tracts in the CNS.</li>
	<li>Analysis should be performed throughout the labeled region, as the projections can extend several hundred microns or more along the rostrocaudal axis. Some examples of basic analyses are: targeting error rates, timing and/or changes in divergence of projection patterns. RDA-labeled axons are visualized in the peripheral and central nervous system with high resolution (Figures 3A, 3D).</li>
</ol>

<ol>
	<li>Groves, A. K., Fekete, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shaping+sound+in+space:+the+regulation+of+inner+ear+patterning.">Shaping sound in space: the regulation of inner ear patterning.</a> Development. 139, 245-257 (2012).</li><li>Pflieger, J. F., Cabana, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+vestibular+primary+afferents+and+the+vestibulospinal+projections+in+the+developing+and+adult+opossum,+Monodelphis+domestica.">The vestibular primary afferents and the vestibulospinal projections in the developing and adult opossum, Monodelphis domestica.</a> Anatomy and Embryology. 194, 75-88 (1996).</li><li>Molea, D., 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=Timing+and+topography+of+nucleus+magnocellularis+innervation+by+the+cochlear+ganglion.">Timing and topography of nucleus magnocellularis innervation by the cochlear ganglion.</a> The Journal of Comparative Neurology. 466, 577-591 (2003).</li><li>Bissonnette, J. P., Fekete, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Standard+atlas+of+the+gross+anatomy+of+the+developing+inner+ear+of+the+chicken.">Standard atlas of the gross anatomy of the developing inner ear of the chicken.</a> The Journal of Comparative Neurology. 368, 620-630 (1996).</li><li>Brigande, J. V., Kiernan, A. E., Gao, X., Iten, L. E., Fekete, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+genetics+of+pattern+formation+in+the+inner+ear:+do+compartment+boundaries+play+a+role.">Molecular genetics of pattern formation in the inner ear: do compartment boundaries play a role.</a> Proceedings of the National Academy of Sciences of the United States of America. 97, 11700-11706 (1073).</li><li>Bellairs, R., Osmond, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The atlas of chick development. , (2005).</li><li>Manley, G. A., Haeseler, C., Brix, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Innervation+patterns+and+spontaneous+activity+of+afferent+fibres+to+the+lagenar+macula+and+apical+basilar+papilla+of+the+chick's+cochlea.">Innervation patterns and spontaneous activity of afferent fibres to the lagenar macula and apical basilar papilla of the chick's cochlea.</a> Hearing Research. 56, 211-226 (1991).</li><li>Code, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efferent+neurons+to+the+macular+lagena+in+the+embryonic+chick.">Efferent neurons to the macular lagena in the embryonic chick.</a> Hearing Research. 82, 26-30 (1995).</li><li>Maklad, A., 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=Development+of+vestibular+afferent+projections+into+the+hindbrain+and+their+central+targets.">Development of vestibular afferent projections into the hindbrain and their central targets.</a> Brain Research Bulletin. 60, 497-510 (2003).</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>Satoh, T., Fekete, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lineage+analysis+of+inner+ear+cells+using+genomic+tags+for+clonal+identification.">Lineage analysis of inner ear cells using genomic tags for clonal identification.</a> Methods Mol. Biol. 493, 47-63 (2009).</li><li>Bok, J., Chang, W., 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=Patterning+and+morphogenesis+of+the+vertebrate+inner+ear.">Patterning and morphogenesis of the vertebrate inner ear.</a> The International Journal of Developmental Biology. 51, 521-533 (2007).</li><li>Appler, J. M., Goodrich, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Connecting+the+ear+to+the+brain:+Molecular+mechanisms+of+auditory+circuit+assembly.">Connecting the ear to the brain: Molecular mechanisms of auditory circuit assembly.</a> Progress in Neurobiology. 93, 488-508 (2011).</li><li>Bulankina, A. V., Moser, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+circuit+development+in+the+mammalian+cochlea.">Neural circuit development in the mammalian cochlea.</a> Physiology (Bethesda). 27, 100-112 (2012).</li><li>Fekete, D. M., Campero, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axon+guidance+in+the+inner+ear.">Axon guidance in the inner ear.</a> The International Journal of Developmental Biology. 51, 549-556 (2007).</li><li>Momose-Sato, Y., Glover, J. C., Sato, K. <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+functional+synaptic+connections+in+the+auditory+system+visualized+with+optical+recording:+afferent-evoked+activity+is+present+from+early+stages.">Development of functional synaptic connections in the auditory system visualized with optical recording: afferent-evoked activity is present from early stages.</a> Journal of Neurophysiology. 96, 1949-1962 (2006).</li><li>Marrs, G. S., Spirou, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryonic+assembly+of+auditory+circuits:+spiral+ganglion+and+brainstem.">Embryonic assembly of auditory circuits: spiral ganglion and brainstem.</a> The Journal of Physiology. 590, 2391-2408 (2012).</li><li>Milo, M., 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=Genomic+analysis+of+the+function+of+the+transcription+factor+gata3+during+development+of+the+mammalian+inner+ear.">Genomic analysis of the function of the transcription factor gata3 during development of the mammalian inner ear.</a> PloS One. 4, e7144 (2009).</li><li>Fritzsch, B., Eberl, D. F., Beisel, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+bHLH+genes+in+ear+development+and+evolution:+revisiting+a+10-year-old+hypothesis.">The role of bHLH genes in ear development and evolution: revisiting a 10-year-old hypothesis.</a> Cellular and Molecular Life Sciences : CMLS. 67, 3089-3099 (2010).</li><li>Jahan, I., Kersigo, J., Pan, N., 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=Neurod1+regulates+survival+and+formation+of+connections+in+mouse+ear+and+brain.">Neurod1 regulates survival and formation of connections in mouse ear and brain.</a> Cell and Tissue Research. 341, 95-110 (2010).</li><li>Huang, E. J., 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=Brn3a+is+a+transcriptional+regulator+of+soma+size,+target+field+innervation+and+axon+pathfinding+of+inner+ear+sensory+neurons.">Brn3a is a transcriptional regulator of soma size, target field innervation and axon pathfinding of inner ear sensory neurons.</a> Development. 128, 2421-2432 (2001).</li><li>Jones, J. M., Warchol, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+the+Gata3+transcription+factor+in+the+acoustic+ganglion+of+the+developing+avian+inner+ear.">Expression of the Gata3 transcription factor in the acoustic ganglion of the developing avian inner ear.</a> The Journal of Comparative Neurology. 516, 507-518 (2009).</li><li>Lu, C. C., Appler, J. M., Houseman, E. A., Goodrich, L. 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No conflicts of interest are declared.

Studies of the early development of the VIIIth nerve have been limited in part because of the difficulty in identifying embryonic axons arising from multiple distinct ganglia. Several studies have explored the molecular signals guiding auditory and vestibular sensory cell and ganglion cell fates during early development, 5,11,12 but the processes regulating central innervation have yet to be determined. Reports of acoustic ganglion cell projections typically describe peripheral processes to sensory epit...

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Polystyrene Weigh Dish</td>
 <td>Fisher Scientific</td>
 <td>02-202-101</td>
 <td></td>
 </tr>
 <tr>
 <td>Petri Dish, 35 X 10 mm</td>
 <td>Fisher Scientific</td>
 <td>50820644</td>
 <td>Use to make silicone dissection dish</td>
 </tr>
 <tr>
 <td>Sylgard Silicone Elastomer Kit</td>
 <td>World Precision Instruments</td>
 <td>SYLG184</td>
 <td>Coat Petri to make dissection dish</td>
 </tr>
 <tr>
 <td>Dissection Pins</td>
 <td>Various</td>
 <td></td>
 <td>Holds embryo in place during dissection</td>
 </tr>
 <tr>
 <td>NaCL</td>
 <td>Various</td>
 <td></td>
 <td>part of aCSF recipe</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>Various</td>
 <td></td>
 <td>part of aCSF recipe</td>
 </tr>
 <tr>
 <td>KH2PO4</td>
 <td>Various</td>
 <td></td>
 <td>part of aCSF recipe</td>
 </tr>
 <tr>
 <td>NaHCO3</td>
 <td>Various</td>
 <td></td>
 <td>part of aCSF recipe</td>
 </tr>
 <tr>
 <td>Glucose</td>
 <td>Various</td>
 <td></td>
 <td>part of aCSF recipe</td>
 </tr>
 <tr>
 <td>CaCl2</td>
 <td>Various</td>
 <td></td>
 <td>part of aCSF recipe</td>
 </tr>
 <tr>
 <td>MgSO4</td>
 <td>Various</td>
 <td></td>
 <td>part of aCSF recipe</td>
 </tr>
 <tr>
 <td>Container for aCSF. Suggest translucent wide-mouth Nalgene jar, 500 ml (16 oz) with lid.</td>
 <td>CPLabSafety</td>
 <td>QP-PLC-03717</td>
 <td>Drill hole opening in top of lid for glass bubling stem to penetrate liquid</td>
 </tr>
 <tr>
 <td>Empty 5 ml glass vial or comparable transparent vial</td>
 <td>American Pharmaceutical Partners, Inc</td>
 <td>6332300105</td>
 <td>Use during aCSF incubation to keep samples separate from each other and from the bubbling stream</td>
 </tr>
 <tr>
 <td>Tank of carbogen (95%O2 / 5%CO2) connected by tubing to bubbler</td>
 <td>Various</td>
 <td></td>
 <td>Attach by tubing to glass stem bubbler for infusion into aCSF</td>
 </tr>
 <tr>
 <td>Glass stem bubbler</td>
 <td>Various</td>
 <td></td>
 <td>To infuse carbogen into aCSF</td>
 </tr>
 <tr>
 <td>Curved-tip forceps</td>
 <td>World Precision Instruments</td>
 <td>501008</td>
 <td>To remove embryo head from egg</td>
 </tr>
 <tr>
 <td>Two fine-tip forceps</td>
 <td>World Precision Instruments</td>
 <td>501985</td>
 <td>For micro-dissection</td>
 </tr>
 <tr>
 <td>50 ml Beaker</td>
 <td>various</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Rhodamine Dextran Amine (RDA)</td>
 <td>Invitrogen</td>
 <td>various</td>
 <td>Fluorescent axon tracer</td>
 </tr>
 <tr>
 <td>Triton X-100</td>
 <td>ICN Biomedicals</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Phosphate Buffered Saline, (1X PBS)</td>
 <td>Various</td>
 <td></td>
 <td>Standard lab reagent</td>
 </tr>
 <tr>
 <td>Thin Wall Glass Capillaries, 1.2 OD, .9 ID 4" (100 mm) length</td>
 <td>World Precision Instruments</td>
 <td>TW120F-4</td>
 <td>Load with RDA. Each capillary makes two glass micropipettes</td>
 </tr>
 <tr>
 <td>Needle / Pipette puller</td>
 <td>David Kopf Instruments</td>
 <td>Model 720</td>
 <td>Settings used: Heat 16.4, Solenoid 2.2</td>
 </tr>
 <tr>
 <td>Picospritzer</td>
 <td>Parker Instrumentation</td>
 <td>various</td>
 <td>Attach by fine tubing to glass micropipette</td>
 </tr>
 <tr>
 <td>Micromanipulator</td>
 <td>Narishige</td>
 <td>various</td>
 <td></td>
 </tr>
 <tr>
 <td>Dissection microscope with fluorescence</td>
 <td>Various</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>4% Paraformaldehyde</td>
 <td>Various</td>
 <td></td>
 <td>Standard lab reagent</td>
 </tr>
 <tr>
 <td>anti-Neurofilament antibody, optional</td>
 <td>Millipore</td>
 <td>AB1991</td>
 <td>Follow histological protocol recommended by manufacturer</td>
 </tr>
 <tr>
 <td>Cryostat and associated materials for sectioning</td>
 <td>Leica</td>
 <td>various</td>
 <td></td>
 </tr>
 <tr>
 <td>Epifluorescent microscope for imaging</td>
 <td>Zeiss, various</td>
 <td></td>
 <td></td>
 </tr>
 </tbody>
</table>

selective tracing of auditory fibers in the avian embryonic vestibulocochlear nerve

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

The components of the VIIIth nerve and the anatomy of the nerve itself are complex and convoluted (Figures 1, 3). By selectively tracing fibers arising from acoustic ganglion cells, segments of the VIIIth nerve as well as primary auditory afferents within the brainstem can be cleanly traced and distinguished from their vestibular counterparts (Figures 2, 3). Likewise, this technique could be used to study peripheral projections of the acoustic ganglion cells (Figure 3G),...

Watch this Scientific Journal Video about Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve at JoVE.com

Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve

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

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

selective-tracing-auditory-fibers-avian-embryonic-vestibulocochlear

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JoVE Journal

Neuroscience

9.1K Views.  University of California, Irvine. Here we describe a microdissection technique followed by fluorescent dye injection into the acoustic ganglion of early chick embryos for selective tracing of auditory axon fibers in the nerve and hindbrain.

Video: Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

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 ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Conditional Genetic Transsynaptic Tracing in the Embryonic Mouse Brain

Anatomical path tracing is of pivotal importance to decipher the relationship between brain and behavior. Unraveling the formation of neural circuits during embryonic maturation of the brain however is technically challenging because most transsynaptic tracing methods developed to date depend on stereotaxic tracer injection. To overcome this problem, we developed a binary genetic strategy for conditional genetic transsynaptic tracing in the mouse brain. Towards this end we generated two complementary knock-in mouse strains to selectively express the bidirectional transsynaptic tracer barley lectin (BL) and the retrograde transsynaptic tracer Tetanus Toxin fragment C from the ROSA26 locus after Cre-mediated recombination. Cell-specific tracer production in these mice is genetically encoded and does not depend on mechanical tracer injection. Therefore our experimental approach is suitable to study neural circuit formation in the embryonic murine brain. Furthermore, because tracer transfer across synapses depends on synaptic activity, these mouse strains can be used to analyze the communication between genetically defined neuronal populations during brain development at a single cell resolution. Here we provide a detailed protocol for transsynaptic tracing in mouse embryos using the novel recombinant ROSA26 alleles. We have utilized this experimental technique in order to delineate the neural circuitry underlying maturation of the reproductive axis in the developing female mouse brain.

Capitalizing on a binary genetic strategy we provide a detailed protocol for neural circuit tracing in mice that express complementary transsynaptic tracers after Cre-mediated recombination. Because cell-specific tracer production is genetically encoded, our experimental approach is suitable to study the formation and maturation of neural circuitry during murine embryonic brain development at a single cell resolution.

Anatomical path tracing is of pivotal importance to decipher the relationship between brain and behavior. Unraveling the formation of neural circuits ...

Laser-guided Neuronal Tracing In Brain Explants

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake through electroporation in brain explants with targeted laser illumination and matching filter goggles during the procedure. The described technique of in vitro electroporation by itself yields relatively good visual control for targetting certain areas of the brain. By combining it with laser excitation of fluorescent genetic markers and their read-out through band-passing filter goggles, which can pick up the emissions of the genetically labeled cells/nuclei and the fluorescent tracing dye, a researcher can substantially increase the accuracy of injections by finding the area of interest and controlling for the dye-spread/uptake in the injection area much more efficiently. In addition, the laser illumination technique allows to study the functionality of a given neurocircuit by providing information about the type of neurons projecting to a certain area in cases where the GFP expression is linked to the type of transmitter expressed by a subpopulation of neurons.

We describe a technique to label neurons and their processes via anterograde or retrograde tracer injections into brain nuclei using an in vitro preparation. We modified an existing method of in vitro tracer electroporation by taking advantage of fluorescently labeled mouse mutants and basic optical equipment in order to increase labeling accuracy.

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake ...

Three-dimensional Imaging and Analysis of Mitochondria within Human Intraepidermal Nerve Fibers

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to isolate nerve-specific mitochondria and evaluate disease-induced alterations of mitochondria in the distal tip of sensory nerves. The protocol combines fluorescence immunohistochemistry, confocal microscopy and 3D image analysis techniques to visualize and quantify nerve-specific mitochondria. Detailed parameters are defined throughout the procedures in order to provide a concrete example of how to use these techniques to isolate nerve-specific mitochondria. Antibodies were used to label nerve and mitochondrial signals within tissue sections of skin punch biopsies, which was followed by indirect immunofluorescence to visualize nerves and mitochondria with a green and red fluorescent signal respectively. Z-series images were acquired with confocal microscopy and 3D analysis software was used to process and analyze the signals. It is not necessary to follow the exact parameters described within, but it is important to be consistent with the ones chosen throughout the staining, acquisition and analysis steps. The strength of this protocol is that it is applicable to a wide variety of circumstances where one fluorescent signal is used to isolate other signals that would otherwise be impossible to study alone.

This protocol uses three-dimensional (3D) imaging and analysis techniques to visualize and quantify nerve-specific mitochondria. The techniques are applicable to other situations where one fluorescent signal is used to isolate a subset of data from another fluorescent signal.

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

In Vitro Characterization of the Electrophysiological Properties of Colonic Afferent Fibers in Rats

Dysfunction of the colonic sensory nerves has been implicated in the pathophysiology of several common conditions, including functional and inflammatory bowel diseases and diabetes. Here, we describe a protocol for the in vitro characterization of the electrophysiological properties of colonic afferents in rats. The colorectum, with the intact pelvic ganglion (PG) attached, is removed from the rat; superfused with carbogenated Krebs solution in the recording chamber; and cannulated at the oral and anal ends to allow for distension. A fine nerve bundle emanating from the PG is identified, and the multiunit afferent nerve activity is recorded using a suction electrode. Distension of the colonic segment elicits gradual increases in multiunit discharge. A principal component analysis is conducted to differentiate the low-threshold, the high-threshold, and the wide-dynamic range afferent fibers. Chemical sensitivity of colonic afferents can be studied through the bath or intraluminal administration of test compounds. This protocol can be modified for application to other species, such as mice and guinea pigs, and to study the differences in the electrophysiological properties of thoracolumbar/hypogastric and lumbosacral/pelvic afferents of the descending colon in normal and pathological conditions.

In Vitro Characterization of the Electrophysiological Properties of Colonic Afferent Fibers in Rats

Abnormal sensory function underlies visceral pain and other symptoms of functional and inflammatory bowel diseases. A protocol for the electrophysiological recording of the colonic afferent nerves in an ex vivo rat colorectum preparation is presented here.

Dysfunction of the colonic sensory nerves has been implicated in the pathophysiology of several common conditions, including functional and inflammatory ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...

Targeting Alpha Synuclein Aggregates in Cutaneous Peripheral Nerve Fibers by Free-floating Immunofluorescence Assay

To date, for most neurodegenerative diseases only a post-mortem histopathological definitive diagnosis is available. For Parkinson&#39;s disease (PD), the diagnosis still relies only on clinical signs of motor involvement that appear later on in the disease course, when most of the dopaminergic neurons are already lost. Hence, there is a strong need for a biomarker that can identify patients at the beginning of disease or at the risk of developing it. Over the last few years, skin biopsy has proved to be an excellent research and diagnostic tool for peripheral nerve diseases such as small fiber neuropathy. Interestingly, a small fiber neuropathy and alpha synuclein (&#945;Syn) neural deposits have been shown by skin biopsy in PD patients. Indeed, skin biopsy has the great advantage of being an easily accessible, minimally invasive and painless procedure that allows the analysis of peripheral nervous tissue prone to the pathology. Moreover, the possibility of repeating the skin biopsy in the course of the follow-up of the same patient allows studying the longitudinal correlation with the disease progression. We set up a standardized reliable protocol to investigate the presence of &#945;Syn aggregates in skin nerve fibers of the PD patient. This protocol involves few short fixation steps, a cryotome sectioning and then a free-floating immunofluorescence double-staining with two specific antibodies: anti Protein Gene Product 9.5 (PGP9.5) to mark the cutaneous nerve fibers and anti 5G4 for detecting &#945;Syn aggregates. It is a versatile, sensitive and easy to perform protocol that can also be applied for targeting other proteins of interest in skin nerves. The ability to mark &#945;Syn aggregates is another step forward to the use of skin biopsy as a tool for establishing a pre-mortem histopathological diagnosis of PD.

Here, we present a protocol for a free-floating indirect immunofluorescence assay on skin biopsy sections that allows for the identification of disease specific conformation variants of alpha synuclein involved in Parkinson disease and multiple proteins of the peripheral nervous system.

To date, for most neurodegenerative diseases only a post-mortem histopathological definitive diagnosis is available. For Parkinson&#39;s disease (PD), the ...