We would like to thank Bing Ye, Nathan Spix, and Hao Liu for technical support. This work was supported by the National Institute of Health Grants EY022640 (D.-Q.Z.) and Oakland University Provost Undergraduate Student Research Award (E.J.A.).

Mouse care and all experimental procedures were conducted according to the National Institutes of Health guidelines for laboratory animals and were approved by the Institutional Animal Care and Use Committees at Oakland University (protocol no. 18071).
NOTE: Names of the solutions and their compositions are listed in Table 1.
1. Tissue preparation
<ol>
	<li>Euthanize the mouse with an overdose of CO2, followed by cervical dislocation.</li>
	<li>Enucleate the eyes with curved forceps and transfer them to a small petri dish with 0.1 M PBS (Table 1). Under a dissection microscope, poke a small hole along the cornea-sclera junction with a needle. Transfer to 4% paraformaldehyde (PFA) for 1 hour.</li>
	<li>Transfer the eye back to a dish with PBS. Under a dissection microscope, use dissection scissors to cut all the way around the cornea-sclera junction. Remove the cornea and lens. Cut at the base of the optic nerve and carefully peel the sclera off with forceps to isolate the retina.</li>
	<li>Make four small cuts evenly around the retina and use a fine tip brush dipped in PBS to lay it flat (GCL side down) in a clover-like shape on a small square cut from nitrocellulose filter paper to stabilize the retina.</li>
	<li>Transfer the retina using forceps to hold the corner of the nitrocellulose paper (without touching the mounted retina) and place it in a 48-well plate with 4% PFA for 1 hour.</li>
	<li>Transfer the filter paper and retina to a well with PBS and wash (3x for 5 min each).</li>
	<li>Transfer to A4P0 (Table 1) and incubate overnight at 4 &#176;C with gentle agitation.</li>
	<li>Pipette vegetable oil into the well to completely cover the A4P0 solution. Incubate in a water bath at 40 &#176;C for 3 hours with no shaking.</li>
	<li>Wash (3x for 5 min each) in PBS, making sure all the oil has been rinsed off. If necessary, use a pipet to carefully remove remaining oil from the top of the well before the last rinse.</li>
	<li>Incubate in 10% SDS at 40 &#176;C for two days with gentle shaking. Replace SDS with fresh solution on the second day.</li>
	<li>Transfer the filter paper and retina to PBS with Triton-X-100 (PBST, Table 1) and wash (5x for 1.5 h each).</li>
	<li>Store at 4 &#176;C in PBST with 0.01% sodium azide (NaN3) or move directly to immunostaining.</li>
</ol>
2. Immunostaining and refractive index matching
<ol>
	<li>Remove the retina from the filter paper by gently peeling it off with a fine tip brush in PBST.</li>
	<li>Incubate the retina in primary antibody (Table 2) diluted in blocking solution (Table 1) for 2 days at 40 &#176;C with gentle shaking.</li>
	<li>Wash (5x, 1.5 h each) in PBST.</li>
	<li>Incubate with the appropriate secondary antibodies (Table 3) diluted in blocking solution for 2 days at 40 &#176;C with gentle shaking and protect from light through the remainder of the procedure.</li>
	<li>Wash (5x, 1.5 h each) in 0.02 M phosphate buffer (see Table 1).</li>
	<li>Incubate in sorbitol-based Refractive Index Matching Solution (sRIMS, see Table 1) at 40 &#176;C overnight with gentle shaking.</li>
</ol>
3. Mounting
<ol>
	<li>Outline a 18 mm x 18 mm x 1.5 mm glass coverslip with a fine-tip permanent marker to mark a square boundary on the back of a glass microscope slide.</li>
	<li>Flip the slide over and use a syringe to trace the boundary with a thin line of silicone grease on the front of the slide, leaving a small gap in one corner for excess mounting solution to escape.</li>
	<li>Transfer the retina to the center of the bounded area and arrange with a fine-tip brush so that it lies flat with the photoreceptor side against the glass slide.</li>
	<li>Pipette approximately 60 &#181;L of sRIMS so that it covers the flattened retina and extends to one corner of the enclosure, taking care that the retina stays flat and in place.</li>
	<li>Apply the coverslip starting from the corner with the sRIMS and slowly lower it until it touches the grease on all sides, avoiding the formation of air bubbles.</li>
	<li>Place a stack of 3 coverslips on each side of the mounted retina as a spacer. Use the long edge of another slide to press down the coverslip so that the mount is flat and even.</li>
	<li>Store slides flat at 4 &#176;C until imaging.</li>
</ol>
4. Imaging
<ol>
	<li>Image samples on either a conventional fluorescence microscope or a confocal microscope (Table of Materials). Begin by placing the slide on the microscope stage and locating the sample. 
	NOTE: If an inverted objective microscope is being used, place the slide upside down on the stage, first ensuring that the exposed areas of the slide are clear of all silicone grease and mounting solution.</li>
	<li>To obtain z-stacked images of co-labeled samples, first focus on the signal in each channel individually and set the exposure time or scanning speed, for fluorescence or confocal microscopes, respectively.</li>
	<li>Set the range for the z-stack either by manually setting the focal plane at the top and bottom of the desired range, or by setting the midpoint and then specifying a range around the midpoint.</li>
	<li>Adjust the step size or number of slices as desired.</li>
	<li>Capture the image and save the original file as well as exporting it as a TIFF file or other desired format.</li>
</ol>
5. Image analysis
<ol>
	<li>Use the image analysis software of choice (Table of Materials) to adjust the brightness and contrast in each channel until optimum clarity is achieved in both the single images and the 3-dimensional rendering of the z-stack. 
	NOTE: If the selected step size is sufficiently small, 3D deconvolution can also be performed to enhance the signal.</li>
</ol>

<ol>
	<li>Witkovsky, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dopamine+and+retinal+function.">Dopamine and retinal function.</a> Documenta Ophthalmologica. 108 (1), 17-40 (2004).</li><li>McMahon, D. G., Iuvone, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circadian+organization+of+the+mammalian+retina:+from+gene+regulation+to+physiology+and+diseases.">Circadian organization of the mammalian retina: from gene regulation to physiology and diseases.</a> Progress in Retinal and Eye Research. 39, 58-76 (2014).</li><li>Prigge, C. L., 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=M1+ipRGCs+Influence+Visual+Function+through+Retrograde+Signaling+in+the+Retina.">M1 ipRGCs Influence Visual Function through Retrograde Signaling in the Retina.</a> Journal of Neuroscience. 36 (27), 7184-7197 (2016).</li><li>Zhang, D. Q., Belenky, M. A., Sollars, P. J., Pickard, G. E., McMahon, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Melanopsin+mediates+retrograde+visual+signaling+in+the+retina.">Melanopsin mediates retrograde visual signaling in the retina.</a> PLoS One. 7 (8), 42647 (2012).</li><li>Liu, L. L., Alessio, E. J., Spix, N. J., Zhang, D. Q. <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+GluA2-containing+calcium-impermeable+AMPA+receptors+on+dopaminergic+amacrine+cells+in+the+mouse+retina.">Expression of GluA2-containing calcium-impermeable AMPA receptors on dopaminergic amacrine cells in the mouse retina.</a> Molecular Vision. 25, 780-790 (2019).</li><li>Liu, L. L., Spix, N. J., Zhang, D. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NMDA+Receptors+Contribute+to+Retrograde+Synaptic+Transmission+from+Ganglion+Cell+Photoreceptors+to+Dopaminergic+Amacrine+Cells.">NMDA Receptors Contribute to Retrograde Synaptic Transmission from Ganglion Cell Photoreceptors to Dopaminergic Amacrine Cells.</a> Frontiers in Cellular Neuroscience. 11, 279 (2017).</li><li>Chung, K., 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=Structural+and+molecular+interrogation+of+intact+biological+systems.">Structural and molecular interrogation of intact biological systems.</a> Nature. 497 (7449), 332 (2013).</li><li>Poguzhelskaya, E., Artamonov, D., Bolshakova, A., Vlasova, O., Bezprozvanny, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simplified+method+to+perform+CLARITY+imaging.">Simplified method to perform CLARITY imaging.</a> Molecular Neurodegeneration. 9, 19 (2014).</li><li>Epp, J. R., 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=Optimization+of+CLARITY+for+Clearing+Whole-Brain+and+Other+Intact+Organs.">Optimization of CLARITY for Clearing Whole-Brain and Other Intact Organs.</a> eNeuro. 2 (3), (2015).</li><li>Magliaro, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clarifying+CLARITY:+Quantitative+Optimization+of+the+Diffusion+Based+Delipidation+Protocol+for+Genetically+Labeled+Tissue.">Clarifying CLARITY: Quantitative Optimization of the Diffusion Based Delipidation Protocol for Genetically Labeled Tissue.</a> Frontiers in Neuroscience. 10, 179 (2016).</li><li>Yang, B., 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=Single-cell+phenotyping+within+transparent+intact+tissue+through+whole-body+clearing.">Single-cell phenotyping within transparent intact tissue through whole-body clearing.</a> Cell. 158 (4), 945-958 (2014).</li><li>Zheng, H., Rinaman, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simplified+CLARITY+for+visualizing+immunofluorescence+labeling+in+the+developing+rat+brain.">Simplified CLARITY for visualizing immunofluorescence labeling in the developing rat brain.</a> Brain Structure and Function. 221 (4), 2375-2383 (2016).</li><li>Witkovsky, P., Arango-Gonzalez, B., Haycock, J. W., Kohler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+retinal+dopaminergic+neurons:+differential+maturation+of+somatodendritic+and+axonal+compartments.">Rat retinal dopaminergic neurons: differential maturation of somatodendritic and axonal compartments.</a> Journal of Comparative Neurology. 481 (4), 352-362 (2005).</li></ol>

The authors declare no competing financial interests.

Modification of the CLARITY protocol for whole-mount retinas. 
We have simplified the CLARITY protocol to achieve adequate polymerization without the need for a vacuum evacuation or desiccation chamber, as is used in most previous studies7,9,11. The polymerization process is inhibited by oxygen, requiring that the sample be isolated from air during the polymerization step of the protocol. However, rather th...

The vertebrate retina is perhaps the most accessible part of the central nervous system (CNS), and it serves as an excellent model for studying the development, structure, and function of the brain. Five classes of neurons in the retina are distributed in three nuclear layers separated by two plexiform layers. The outer nuclear layer (ONL) consists of classical photoreceptors (rods and cones) that convert light into electrical signals. Electrical signals are processed by neurons in the inner nuclear layer (INL), including bipolar, horizontal, and amacrine cells, and then transmitted to retinal ganglion cells (RGCs) in the ganglion cell layer (GCL). RGCs are the output neurons of the retina, with the axons projecting to the brain to contribute to image-forming and non-image-forming visual function. In addition, three types of glial cells (Muller cells, astroglia, and microglia) provide nutrients to neurons and protect neurons from harmful changes in their extracellular environment.
One specialized subpopulation of amacrine cells produces and releases dopamine, an important neuromodulator in the CNS, reconfiguring retinal neural circuits during light adaptation1,2. Dopaminergic amacrine cells (DACs) have a unique feature of morphological profiles. Their somata are located in the proximal INL with dendrites ramifying in the most distal part of the inner plexiform layer (IPL). Axon-like processes of DACs are unmyelinated, thin and long, sparsely branched, and bear varicosities (the sites of dopamine release). They form a dense plexus with dendrites in the IPL, including ring-like structures around the somata of AII amacrine cells. The axons also run through the INL toward the OPL, forming a centrifugal pathway across the retina3. We have demonstrated that DAC processes express receptors in response to glutamate release from presynaptic neurons, including bipolar cells and intrinsically photosensitive retinal ganglion cells (ipRGCs)4,5,6. However, it is unclear whether glutamate receptors express on the axons, dendrites, or both since they are cut off in vertical retinal sections and cannot be distinguished from each other5,6. Immunostaining needs to be carried out in whole-mount retinas to reveal three-dimensional branching of DACs and the presence of glutamate receptors on subcellular compartments. Although the retina is relatively transparent, the thickness of a mouse whole-mount retina is approximately 200 &#181;m, which limits the penetration of antibodies into the deep tissue as well as reduces light penetration for high-resolution imaging due to tissue light-scattering. To overcome these limitations, we adapted the immunostaining compatible tissue hydrogel delipidation method (CLARITY) developed recently for thick brain sections to whole-mount mouse retinas7.
The CLARITY method was originally developed by the Deisseroth laboratory for immunostaining and imaging of thick brain samples7. It uses a strong detergent, sodium dodecyl sulfate (SDS) and electrophoresis to remove the lipid components (that cause tissue light-scattering), leaving the proteins and nucleic acids in place. The removed lipids are replaced with a transparent scaffold made up of hydrogel monomers such as acrylamide to support the remaining protein structure. The cleared tissue can be labeled via immunohistochemistry and imaged with substantially increased light penetration depth through the tissue (up to several millimeters below the tissue surface). Since then, the CLARITY method has been optimized and simplified by several research groups8,9,10. A modified CLARITY protocol uses a passive clearing technique to avoid the possible tissue damage produced by electrophoresis for clearing the whole-brain and other intact organs11. However, this method has not yet been applied to whole-mount retinas. Here, we adapted the passive CLARITY technique for whole-mount retinas to make them more transparent for immunohistochemistry and imaging. We found that a majority of the retinal proteins tested were preserved during this process for immunohistochemistry. Using the refractive index matching solution, we were able to image retinal neurons across the approximately 200 &#181;m thickness from the ONL to the GCL in whole-mount retinas.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>16% Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710</td>
			<td>Fixative</td>
		</tr>
		<tr>
			<td>Acrylamide</td>
			<td>Fisher Biotech</td>
			<td>BP170</td>
			<td>Hydrogel monomer</td>
		</tr>
		<tr>
			<td>Axio Imager.Z2</td>
			<td>Zeiss</td>
			<td></td>
			<td>Fluorscence microscope</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Fisher Scientific</td>
			<td>BP1600</td>
			<td>Blocking agent</td>
		</tr>
		<tr>
			<td>Eclipse Ti</td>
			<td>Nikon Instruments</td>
			<td></td>
			<td>Scanning confocal microscope</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>VWR</td>
			<td>BDH0258</td>
			<td>Buffer component</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Sigma</td>
			<td>P5655</td>
			<td>Buffer component</td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>Sigma Aldrich</td>
			<td>S9763</td>
			<td>Buffer component</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma Aldrich</td>
			<td>S7653</td>
			<td>Buffer component</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Sigma Aldrich</td>
			<td>S0751</td>
			<td>Buffer component</td>
		</tr>
		<tr>
			<td>NaN3</td>
			<td>Sigma Aldrich</td>
			<td>S2002</td>
			<td>Bacteriostatic preservative</td>
		</tr>
		<tr>
			<td>NDS</td>
			<td>Aurion</td>
			<td>900.122</td>
			<td>Blocking agent</td>
		</tr>
		<tr>
			<td>NIS Elements AR</td>
			<td>Nikon</td>
			<td></td>
			<td>Image analysis software</td>
		</tr>
		<tr>
			<td>SDS</td>
			<td>BioRad</td>
			<td>1610301</td>
			<td>Delipidation agent</td>
		</tr>
		<tr>
			<td>Sorbitol</td>
			<td>Sigma Aldrich</td>
			<td>51876</td>
			<td>Buffer component</td>
		</tr>
		<tr>
			<td>Triton-X-100</td>
			<td>Sigma</td>
			<td>T8787</td>
			<td>Surfactant</td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Fisher Scientific</td>
			<td>BP337</td>
			<td>Surfactant</td>
		</tr>
		<tr>
			<td>VA-044</td>
			<td>Wako Chemicals</td>
			<td>011-19365</td>
			<td>Thermal initiator</td>
		</tr>
	</tbody>
</table>

immunostaining of whole-mount retinas with the clarity tissue clearing method

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining and imaging of thick brain samples. However, this advanced technology has not yet been used for whole-mount retinas. Although the retina is partially transparent, its thickness of approximately 200 &#181;m (in mice) still limits the penetration of antibodies into the deep tissue as well as reducing light penetration for high-resolution imaging. Here, we adapted the CLARITY method for whole-mount mouse retinas by polymerizing them with an acrylamide monomer to form a nanoporous hydrogel and then clearing them in sodium dodecyl sulfate to minimize protein loss and avoid tissue damage. CLARITY-processed retinas were immunostained with antibodies for retinal neurons, glial cells, and synaptic proteins, mounted in a refractive index matching solution, and imaged. Our data demonstrate that CLARITY&#160;can improve the quality of standard immunohistochemical staining and imaging for retinal neurons and glial cells in whole-mount preparation. For instance, 3D resolution of fine axon-like and dendritic structures of dopaminergic amacrine cells were much improved by CLARITY. Compared to non-processed whole-mount retinas, CLARITY can reveal immunostaining for synaptic proteins such as postsynaptic density protein 95. Our results show that CLARITY renders the retina more optically transparent after the removal of lipids and preserves fine structures of retinal neurons and their proteins, which can be routinely used for obtaining high-resolution imaging of retinal neurons and their subcellular structures in whole-mount preparation.

Modified CLARITY-processed retinas are optically transparent tissue. 
To formulate a tissue clearing method that is compatible with immunohistochemical applications in the retina while providing adequate delipidation and retaining the structural integrity of the cellular proteins, we adapted the CLARITY tissue clearing method to whole-mount mouse retinas. We were able to simplify the protocol and modify it for whole-mount retinas (see Protocol). After completing tissue hybridization, clearing, and r...

Watch this Scientific Journal Video about Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method at JoVE.com

Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method

Here we present a protocol to adapt the CLARITY method of the brain tissues for whole-mount retinas to improve the quality of standard immunohistochemical staining and high-resolution imaging of retinal neurons and their subcellular structures.

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining ...

immunostaining-whole-mount-retinas-with-clarity-tissue-clearing

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JoVE Journal

Neuroscience

7.2K Views.  Oakland University. Here we present a protocol to adapt the CLARITY method of the brain tissues for whole-mount retinas to improve the quality of standard immunohistochemical staining and high-resolution imaging of retinal neurons and their subcellular structures.

Video: Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

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 ...

Dissection of the Transversus Abdominis Muscle for Whole-mount Neuromuscular Junction Analysis

Analysis of neuromuscular junction morphology can give important insight into the physiological status of a given motor neuron. Analysis of thin flat muscles can offer significant advantage over traditionally used thicker muscles, such as those from the hind limb (e.g. gastrocnemius). Thin muscles allow for comprehensive overview of the entire innervation pattern for a given muscle, which in turn permits identification of selectively vulnerable pools of motor neurons. These muscles also allow analysis of parameters such as motor unit size, axonal branching, and terminal/nodal sprouting. A common obstacle in using such muscles is gaining the technical expertise to dissect them. In this video, we detail the protocol for dissecting the transversus abdominis (TVA) muscle from young mice and performing immunofluorescence to visualize axons and neuromuscular junctions (NMJs). We demonstrate that this technique gives a complete overview of the innervation pattern of the TVA muscle&#160;and can be used to investigate NMJ pathology in a mouse model of the childhood motor neuron disease, spinal muscular atrophy.

Dissection of the Transversus Abdominis Muscle for Whole-mount Neuromuscular Junction Analysis

In this video we demonstrate a protocol for dissection of the transversus abdominis muscle of the mouse and use immunofluorescence and microscopy to visualize neuromuscular junctions.

Analysis of neuromuscular junction morphology can give important insight into the physiological status of a given motor neuron. Analysis of thin flat ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

Whole Mount Labeling of Cilia in the Main Olfactory System of Mice

The mouse olfactory system comprises 6-10 million olfactory sensory neurons in the epithelium lining the nasal cavity. Olfactory neurons extend a single dendrite to the surface of the epithelium, ending in a structure called dendritic knob. Cilia emanate from this knob into the mucus covering the epithelial surface. The proteins of the olfactory signal transduction cascade are mainly localized in the ciliary membrane, being in direct contact with volatile substances in the environment. For a detailed understanding of olfactory signal transduction, one important aspect is the exact morphological analysis of signaling protein distribution. Using light microscopical approaches in conventional cryosections, protein localization in olfactory cilia is difficult to determine due to the density of ciliary structures. To overcome this problem, we optimized an approach for whole mount labeling of cilia, leading to improved visualization of their morphology and the distribution of signaling proteins. We demonstrate the power of this approach by comparing whole mount and conventional cryosection labeling of Kirrel2. This axon-guidance adhesion molecule is known to localize in a subset of sensory neurons and their axons in an activity-dependent manner. Whole mount cilia labeling revealed an additional and novel picture of the localization of this protein.

Cilia of olfactory sensory neurons contain proteins of the signal transduction cascade, but a detailed spatial analysis of their distribution is difficult in cryosections. We describe here an optimized approach for whole mount labeling and en face visualization of ciliary proteins.

The mouse olfactory system comprises 6-10 million olfactory sensory neurons in the epithelium lining the nasal cavity. Olfactory neurons extend a single ...

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered tissue samples. Traditional immunolabeling procedures require cutting the sample into thin sections, which restricts the ability to label and examine intact structures. However, if brain tissue can remain intact during processing, structures and circuits can remain intact for the analysis. Previously established clearing methods take significant time to completely clear the tissue, and the harsh chemicals can often damage sensitive antibodies. The iDISCO method quickly and completely clears tissue, is compatible with many antibodies, and requires no special lab equipment. This technique was initially validated for the use in mice tissue, but the current protocol adapts this method to image hemispheres of control and transgenic rat brains. In addition to this, the present protocol also makes several adjustments to preexisting protocol to provide clearer images with less background staining. Antibodies for Iba-1 and tyrosine hydroxylase were validated in the HIV-1 transgenic rat and in F344/N control rats using the present hydrophobic tissue clearing method. The brain is an interwoven network, where structures work together more often than separately of one another. Analyzing the brain as a whole system as opposed to a combination of individual pieces is the greatest benefit of this whole brain clearing method.

Here we present a hydrophobic tissue clearing method that allows for the viewing of target molecules as part of intact brain structures. This technique has now been validated for F344/N control and HIV-1 transgenic rats of both sexes.

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered ...

Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the morphology of processes and substructures within these neurons. As such, much of what is known about brain function has arisen alongside developments in imaging technologies that allow further insight into how neurons are organized and connected in the brain. Improvements in tissue clearing have allowed for high-resolution imaging of thick brain slices, facilitating morphological reconstruction and analyses of neuronal substructures, such as dendritic arbors and spines. In tandem, advances in image processing software provide methods of quickly analyzing large imaging datasets. This work presents a relatively rapid method of processing, visualizing, and analyzing thick slices of labeled neural tissue at high-resolution using CLARITY tissue clearing, confocal microscopy, and image analysis. This protocol will facilitate efforts toward understanding the connectivity patterns and neuronal morphologies that characterize healthy brains, and the changes in these characteristics that arise in diseased brain states.

Neuronal dendritic morphology often underlies function. Indeed, many disease processes that affect the development of neurons manifest with a morphological phenotype. This protocol describes a simple and powerful method for analyzing intact dendritic arbors and their associated spines.