This work was supported by the National Institute of Environmental Health Sciences [K22ES012264 to A.J.R.B.] and an American Cancer Society InstitutionalResearch Grant [ACS-IRG-00-173-04]pilot projectaward [to A.J.R.B.]. We also thank members of the Bishop lab for critical reading of the manuscript and comments on the video and Adam Brown in particular, for the example of what not to do. We thank Dr. Donald McEwen of Greehey Children's Cancer Research Institute for allowing us the use of his dissecting scope/ video camera set-up for filming of the dissection video. We thank Daron Brown at Corte Instruments for sharpening and repair of our microdissection tools.

1. Remove Extraneous tissue from the outside of the eye
<ol>
	<li>Pour 1X PBS into the lid of a 35mm dish. The level should be just below the lip of the lid.</li>
	<li>Using straight forceps, transfer eye(s) from the storage tube to the 35mm dish. Eyes are submerged in PBS for two reasons: 1. The extraneous tissues will "float" away from the eye so you can see and remove them easily, and 2. During dissection, the eye will naturally maintain its spherical shape while in suspension, allowing you to work with this shape rather than against it.</li>
	<li>Using both pairs of angled forceps (45&deg; and 15&deg;), gently remove as much muscle and connective tissue as possible, pulling the tissue against the grain (i.e. toward the cornea). Some connective tissue will remain. Effort should be made to squeeze the eyeball as little as possible.</li>
	<li>Trade your 15&deg; forceps for spring scissors. While holding the eye steady with 45&deg; forceps, use the spring scissors to cut away remaining muscle and connective tissue, as well as the optic nerve. *Tip: Use the outside edge of the scissors to push the tissue against the grain (toward the corneo-scleral divide) and then trim. Continue until all extraneous tissue has been removed from the eye.</li>
</ol>
2. Remove the cornea and lens
<ol>
	<li>With the cornea facing up, use the 45&deg; forceps to pinch up a small fold in the center of the cornea.</li>
	<li>Using the spring scissors, make a small incision at the base of the flap. Let go of the flap.</li>
	<li>Holding the eyes still with your forceps, insert the lower scissor blade into and perpendicular to the incision and make a cut toward the corneo-scleral divide. Try to keep your lower blade under the iris so both the cornea and iris are cut. Do not cut all the way to the corneo-scleral divide. Keep scissors inserted in the eye.</li>
	<li>Using your 45&deg; forceps, carefully rotate the eye so that your scissors are parallel to the corneo-scleral divide and make a small cut. Slowly rotate the eye 360&deg;, making small incisions all the way around. Again, try to keep the lower blade of your spring scissors under the iris so both the iris and cornea are cut. Lift the iris and cornea away from the rest of the eye and discard them.</li>
	<li>Tilt the eye on its side and press down gently on the posterior half of the eye, forcing the lens out. Discard the lens. The inside of the remaining eyecup is lined by the neural retina, which should look smooth and opaque.</li>
</ol>
3. Quarter the resulting eyecup, resulting in a 4-"petaled", flower-like structure
<ol>
	<li>The eye should be positioned such that the opening of the eyecup is perpendicular to your body. Use your 45&deg; forceps to steady and position the eyecup.</li>
	<li>Holding your spring scissors in front of you so they are perpendicular to your body, insert the lower blade of your spring scissors into the top edge of the open eye cup. Using your 45&deg; forceps, carefully align the eye cup so that a single, straight cut can be made from the corneo-scleral divide toward the optic nerve head. All cuts should be perpendicular to the corneo-scleral divide and unless specified, should go as close to the optic nerve head as possible without cutting into it.</li>
	<li>Rotate the eyecup 180&deg;, maintaining its perpendicularity with your body. Use technique in step 3.2 to make a second perpendicular cut.</li>
	<li>Rotate the eyecup and repeat the technique described in 3.2 to make two more cuts. All four cuts should be as close to 90&deg; apart as possible. Keep in mind that the eye is not perfectly spherical so there will be some variability.</li>
</ol>
4. Cut each of the four "petals" in half, resulting in an eight-petal flower-like structure
<ol>
	<li>Transfer the quartered eyecup from the 35mm dish to a clean slide. With the eyecup facing up, scoop under it with the 45&deg; forceps. Carefully lift it out of the liquid and place it on a clean slide.</li>
	<li>Using both pairs of angled forceps, gently open out the eye, with the neural retina facing up. DO NOT REMOVE THE NEURAL RETINA- it helps the eye maintain some of its spherical shape so you are not working against it. It also protects the RPE from being accidently damaged in the following procedures.</li>
	<li>Rotate the slide until one of the petals is perpendicular to your body.</li>
	<li>Use your 15&deg; forceps, gently grab the corner of the petal. The corner of the petal should be cornea, not neural retina or RPE. Lift up on the corner gently, allowing it to retain its curvature.</li>
	<li>Insert the bottom blade of your spring scissors between the petal and the slide. Line up your scissors such that a single, straight, perpendicular cut can be made from the corneo-scleral divide toward the optic nerve head. Cut as close to the optic nerve head as possible without cutting into it. Let go of the petal, allowing it to lie on the slide.</li>
	<li>Rotate the slide 90&deg;, so that the next petal is perpendicular to your body. Repeat step 4.5.</li>
	<li>Repeat steps 4.5 and 4.6 twice more. The resulting structure should look like an eight-petal flower.</li>
</ol>
5. Remove the neural retina
<ol>
	<li>Pipette 100-200 &mu;L of 1X PBS onto the specimen. This will prevent the neural retina from sticking as you peel it off of the RPE.</li>
	<li>Typically the neural retina is loosely attached to the RPE. However, it does take practice and skill to remove only the neural retina and not the RPE. Gently grasp a petal of the neural retina with one pair of angled forceps while stabilizing (holding down) the rest of the eye with the other pair of angled forceps. Pull the neural retina away from the rest of the eye. Continue this procedure until the entire neural retina has been removed. *Tip: stick the tip of your 15&deg; forceps into the optic nerve head to anchor the eye. Then use the 45&deg; forceps to sweep up underneath the neural retina and lift it off and away. It is usually best to lift the neural retina from the optic nerve head toward the corneo-scleral divide. Be sure to pull away any loose remnants of the iris. The iris is quite sticky and can otherwise get stuck to the RPE.</li>
	<li>Rinse the specimen with 1X PBS. Pipette 100-200 &mu;L of 1X PBS into the specimen and suck it back up. Discard the PBS. Repeat as necessary to remove dust and debris.</li>
</ol>
6. Mount you specimen(s) on a slide
<ol>
	<li>Pipette 80-90 &mu;L of 90% glycerol onto a new slide in a rectangular shape. Use the side of the pipette tip to make an even, rectangular smear of glycerol. (This is the procedure for mounting two RPE on one slide.)</li>
	<li>Insert the lower arm of 15&deg; forceps under the specimen. Gently lift the specimen off the dissecting slide.</li>
	<li>Slowly lower the specimen into the glycerol. Gently wiggle the forceps as you pull them out from under the specimen.</li>
	<li>Repeat steps 6.1-6.3 for the other specimen.</li>
	<li>Rotate the slide so that the long side is perpendicular to your body. Hold your cover glass at a 45&deg; angle to the slide so that it is touching the slide. Move it toward the glycerol smear until they touch. Maintaining contact between the cover glass and slide, slowly move the cover glass away from the smear, toward the edge of the slide. Line up the long sides of the slide and the cover glass so that they are parallel. Hold the corners of the cover glass between your thumb and forefinger.</li>
	<li>Pick up the 15&deg; forceps in your other hand. Cradle the upper (free) edge of the cover glass in the crook of the arm. Very slowly, lower the cover glass. When contact is made between the cover glass and the glycerol, stop and wait for the glycerol to migrate. Repeat this lowering and waiting until the RPE whole mounts are fully covered. This is to prevent the highly viscous glycerol from capturing air bubbles. DO NOT PUSH DOWN ON THE COVERGLASS - the glycerol will squeeze out from under it, making adhesion very poor.</li>
	<li>With clear nail polish, paint around the perimeter where the cover glass and slide meet.</li>
	<li>Place the slide on top of an empty rack to dry.</li>
</ol>
7. Representative Results:
The result of this procedure should be a structure that looks like a flower and should be fairly symmetrical.
<img src="/files/ftp_upload/2563/2563fig1.jpg" alt="Figure 1" /> 
Figure 1. Whole mount RPE from a wildtype C67Bl/6J mouse. RPE from black or agouti animals should be dark brown in color and should have a smooth surface. It is normal to notice undulations in the topography of the petals. The pigmentation on any give specimen may be somewhat variable, due to variable density of pigmentation of both the RPE and the underlying choroid. White arrows point to hypopigmented "channels"- this is normal and is due to the underlying vasculature of the eye. Blue arrow points to physical damage to both the RPE and the underlying choroid.
<img src="/files/ftp_upload/2563/2563fig2.jpg" alt="Figure 2" /> 
Figure 2. Whole mount RPE from a dilute mouse. Specimens harvested from dilute animals can range in color from nearly transparent to caf&eacute; au lait and any one specimen is like to have variability within it, indicated in black circles here. In general, RPE harvested from younger animals are lighter and those harvested from older animals are darker. Black arrows indicate some of the underlying vasculature, which appears hypopigmented.
<img src="/files/ftp_upload/2563/2563fig3.jpg" alt="Figure 3" /> 
Figure 3. Phalloidin staining can be used to detect physical damage to the RPE. Phalloidin staining clearly outlines the cell membranes, showing the hexagonal shape of the epithelium. (A) Example from a black mouse. (B) Example from a dilute mouse.
<img src="/files/ftp_upload/2563/2563fig4.jpg" alt="Figure 4" /> 
Figure 4. A poorly dissected RPE from a dilute mouse. (A) Brackets indicate margins of cornea that are too wide, which can cause buckling and/ or folding. Within the black circle you can see excessive amounts of extraneous tissue that have not been removed from the back of the eye. They are particularly obvious because the sample is from a dilute animal. The petal on the bottom left is partially folded over. (B) This whole mount has the appearance of a pinwheel. Black arrows indicate some of the cuts that were made. Many of the cuts from the corneo-scleral divide toward the optic nerve head are not in line with the diameter. (C) Black arrows indicate how the cuts should have been made, in line with the diameter and perpendicular to the tangent.

<ol>
	<li>Bishop, A. J., Kosaras, B., Sidman, R. L., Scheistl, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Benzo(a)pyrene+andX-rays+induce+reversions+of+the+pink-eyed+unstable+mutation+in+the+retinal+pigmentepithelium+of+mice.">Benzo(a)pyrene andX-rays induce reversions of the pink-eyed unstable mutation in the retinal pigmentepithelium of mice.</a> Mutat. Res.. 457, 31-31 (2000).</li><li>Reliene, R. H. l. a. v. a. c. o. v. e., Mahadevan, A., Baird, B., M, W., Schiestl, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diesel+exhaust+particles+cause+increased+levels+of+DNA+deletions+after+transplacental+exposure+in+mice.">Diesel exhaust particles cause increased levels of DNA deletions after transplacental exposure in mice.</a> Mutat. Res. 570, 245-2452 (2005).</li><li>Bishop, A. J., Barlow, C., Wynshaw-Boris, A. J., Scheistl, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atm+deficiency+causes+increased+frequency+of+intrachromosomal+homologous+recombination+in+mice.">Atm deficiency causes increased frequency of intrachromosomal homologous recombination in mice.</a> Cancer Res. 60, 395-399 (2000).</li><li>Brown, A. D., Claybon, A. B., Bishop, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+WRN+helicase+domain+is+not+required+for+spontaneous+homologous+recombination-mediated+DNA+deletion.">Mouse WRN helicase domain is not required for spontaneous homologous recombination-mediated DNA deletion.</a> J. Nucleic Acids. , (2010).</li><li>Claybon, A., Karia, B., Bruce, C., Bishop, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PARP1+suppresses+homologous+recombination+events+in+mice+in+vivo.">PARP1 suppresses homologous recombination events in mice in vivo.</a> Nucleic Acids Res. , (2010).</li><li>Bishop, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=p53-,+and+Gadd45a-deficient+mice+show+an+increased+frequency+ofhomologous+recombination+at+different+stages+during+development.">p53-, and Gadd45a-deficient mice show an increased frequency ofhomologous recombination at different stages during development.</a> Cancer Res. 63, 5335-5343 (2003).</li><li>Bodenstein, L., Sidman, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+and+development+of+the+mouse+retinalpigment+epithelium.+Part+I.+Cell+and+tissue+morphometrics+and+topography+of+mitoticactivity.">Growth and development of the mouse retinalpigment epithelium. Part I. Cell and tissue morphometrics and topography of mitoticactivity.</a> Dev Biol. 121, 192-204 (1987).</li><li>Burke, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epithelial+phenotype+and+the+RPE:+is+the+answer+blowing+in+the+Wnt?.">Epithelial phenotype and the RPE: is the answer blowing in the Wnt?.</a> Prog Retin. Eye Res. 27, 579-595 (2008).</li></ol>

No conflicts of interest declared.

The RPE is the site of the pun assay, an in vivo assay of homology-directed repair. The pun assay has been used to study the effects of different DNA damages1,2 and DNA damage signaling and repair genes3,4,5 on the frequency of homology-directed repair. This assay is highly sensitive, detecting single-cell events on the RPE1 . It can also detect differences in the timing of homology-directed repair events during development6. The...

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		<div>
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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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<tr>
<td>straight forceps</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-4903</td>
<td>tip: .08 x .04 mm material: INOX</td>
</tr>
<tr>
<td>45&deg; forceps</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5005</td>
<td>tip: .05 x.01 mm material: INOX</td>
</tr>
<tr>
<td>15&deg; "up" forceps</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5045A</td>
<td>tip: .1 x.06 mm material: INOX</td>
</tr>
<tr>
<td>spring scissors</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5604</td>
<td>comb. tip width 0.3mm cutting edge length 3mm material: stainless steel</td>
</tr>
<tr>
<td>binocular dissecting microscope</td>
<td>&nbsp;</td>
<td>Ziess</td>
<td>Discovery V.8</td>
<td>use reflected light source</td>
</tr>
<tr>
<td>phalloidin</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>A22283</td>
<td>Alexa Fluor 546</td>
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dissection of a mouse eye for a whole mount of the retinal pigment epithelium

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and cones). The RPE is a monolayer of pigmented cuboidal cells and associates closely with the neural retina just above it. This association makes the RPE of great interest to researchers studying retinal diseases. The RPE is also the site of an in vivo assay of homology-directed DNA repair, the pun assay. The mouse eye is particularly difficult to dissect due to its small size (about 3.5mm in diameter) and its spherical shape. This article demonstrates in detail a procedure for dissection of the eye resulting in a whole mount of the RPE. In this procedure, we show how to work with, rather than against, the spherical structure of the eye. Briefly, the connective tissue, muscle, and optic nerve are removed from the back of the eye. Then, the cornea and lens are removed. Next, strategic cuts are made that result in significant flattening of the remaining tissue. Finally, the neural retina is gently lifted off, revealing an intact RPE, which is still attached to the underlying choroid and sclera. This whole mount can be used to perform the pun assay or for immunohistochemistry or immunofluorescent assessment of the RPE tissue.

Watch this Scientific Journal Video about Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium at JoVE.com

Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium

A formal demonstration of the dissection of a mouse eye, resulting in a whole mount of the retinal pigment epithelium.

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and ...

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49.2K Views.  Greehey Children's Cancer Research Institute and Department of Cellular and Structural Biology. A formal demonstration of the dissection of a mouse eye, resulting in a whole mount of the retinal pigment epithelium.

Video: Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium

Experimental Models for Study of Retinal Pigment Epithelial Physiology and Pathophysiology

We have developed a cell culture procedure that can produce large quantities of confluent monolayers of primary human fetal retinal pigment epithelium (hfRPE) cultures with morphological, physiological and genetic characteristics of native human RPE. These hfRPE cell cultures exhibit heavy pigmentation, and electron microscopy show extensive apical membrane microvilli. The junctional complexes were identified with immunofluorescence labeling of various tight junction proteins. Epithelial polarity and function of these easily reproducible primary cultures closely resemble previously studied mammalian models of native RPE, including human. These results were extended by the development of therapeutic interventions in several animal models of human eye disease. We have focused on strategies for the removal of abnormal fluid accumulation in the retina or subretinal space. The extracellular subretinal space separates the photoreceptor outer segments and the apical membrane of the RPE and is critical for maintenance of retinal attachments and a whole host of RPE/retina interactions.

We provide a reproducible method for culturing confluent monolayers of human fetal retinal pigment epithelial cells (hfRPE) cells that exhibit morphology, physiology, polarity, and protein and gene expression patterns of adult native tissue. This work has been extended to an animal model of several eye diseases.

We have developed a cell culture procedure that can produce large quantities of confluent monolayers of primary human fetal retinal pigment epithelium ...

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled samples with little to no tissue damage enables us to analyze cell morphology and to compare individual samples to each other, hence allowing the identification of mutant anomalies.
In the demonstrated dissection method the nervous system remains mostly inside the adult fly. Through a dorsal incision, the abdomen and thorax are opened and most of the internal organs are removed. Only the dorsal side of the ventral nerve cord (VNC) and the cervical connective 
(CvC) containing the big axons of the giant fibers (GFs)1 are exposed, while the brain containing the GF cell body and dendrites remains2 in the 
intact head. In this preparation most nerves of the VNC should remain attached to their muscles. 
Following the dissection, the intracellular filling of the giant fiber (GF) with a fluorescent dye is demonstrated. In the CvC the GF axons are located at the dorsal surface and thus can be easily visualized under a microscope with differential interference contrast (DIC) optics. This allows the injection of the GF axons with dye at this site to label the entire GF including the axons and their terminals in the VNC. This method results in reliable and strong staining of the GFs allowing the neurons to be imaged immediately after filling with an epifluorescent microscope. Alternatively, the fluorescent signal can be enhanced using standard immunohistochemistry procedures3 suitable for high resolution confocal microscopy.

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

An in vivo dissection of the adult Drosophila ventral nerve cord (VNC) is demonstrated. This particular dissection method causes little damage to the VNC allowing the subsequent labeling of the giant fiber neurons with fluorescent dye for high resolution imaging.

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

The process by which the anterior region of the neural plate gives rise to the vertebrate retina continues to be a major focus of both clinical and basic research. In addition to the obvious medical relevance for understanding and treating retinal disease, the development of the vertebrate retina continues to serve as an important and elegant model system for understanding neuronal cell type determination and differentiation1-16. The neural retina consists of six discrete cell types (ganglion, amacrine, horizontal, photoreceptors, bipolar cells, and M&uuml;ller glial cells) arranged in stereotypical layers, a pattern that is largely conserved among all vertebrates 12,14-18.
While studying the retina in the intact developing embryo is clearly required for understanding how this complex organ develops from a protrusion of the forebrain into a layered structure, there are many questions that benefit from employing approaches using primary cell culture of presumptive retinal cells 7,19-23. For example, analyzing cells from tissues removed and dissociated at different stages allows one to discern the state of specification of individual cells at different developmental stages, that is, the fate of the cells in the absence of interactions with neighboring tissues 8,19-22,24-33. Primary cell culture also allows the investigator to treat the culture with specific reagents and analyze the results on a single cell level 5,8,21,24,27-30,33-39. Xenopus laevis, a classic model system for the study of early neural development 19,27,29,31-32,40-42, serves as a particularly suitable system for retinal primary cell culture 10,38,43-45.
Presumptive retinal tissue is accessible from the earliest stages of development, immediately following neural induction 25,38,43. In addition, given that each cell in the embryo contains a supply of yolk, retinal cells can be cultured in a very simple defined media consisting of a buffered salt solution, thus removing the confounding effects of incubation or other sera-based products 10,24,44-45.
However, the isolation of the retinal tissue from surrounding tissues and the subsequent processing is challenging. Here, we present a method for the dissection and dissociation of retinal cells in Xenopus laevis that will be used to prepare primary cell cultures that will, in turn, be analyzed for calcium activity and gene expression at the resolution of single cells. While the topic presented in this paper is the analysis of spontaneous calcium transients, the technique is broadly applicable to a wide array of research questions and approaches (Figure 1).

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

Xenopus laevis provides an ideal model system for studying cell fate specification and physiological function of individual retinal cells in primary cell culture. Here we present a technique for dissecting retinal tissues and generating primary cell cultures that are imaged for calcium activity and analyzed by in situ hybridization.

The process by which  the anterior region of the neural plate gives rise to the vertebrate retina  continues to be a major focus of both clinical and ...

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 ...

The Corneal Micropocket Assay: A Model of Angiogenesis in the Mouse Eye

The mouse corneal micropocket assay is a robust and quantitative in vivo assay for evaluating angiogenesis. By using standardized slow-release pellets containing specific growth factors that trigger blood vessel growth throughout the naturally avascular cornea, angiogenesis can be measured and quantified. In this assay the angiogenic response is generated over the course of several days, depending on the type and dose of growth factor used. The induction of neovascularization is commonly triggered by either basic fibroblast growth factor (bFGF) or vascular endothelial growth factor (VEGF). By combining these growth factors with sucralfate and hydron (poly-HEMA (poly(2-hydroxyethyl methacrylate))) and casting the mixture into pellets, they can be surgically implanted in the mouse eye. These uniform pellets slowly-release the growth factors over five or six days (bFGF or VEGF respectively) enabling sufficient angiogenic response required for vessel area quantification using a slit lamp. This assay can be used for different applications, including the evaluation of angiogenic modulator drugs or treatments as well as comparison between different genetic backgrounds affecting angiogenesis. A skilled investigator after practicing this assay can implant a pellet in less than 5 min per eye.

The protocol describes the corneal micropocket assay as developed in mice.

The mouse corneal micropocket assay is a robust and quantitative in vivo assay for evaluating angiogenesis. By using standardized slow-release pellets ...

Limbal Approach-Subretinal Injection of Viral Vectors for Gene Therapy in Mice Retinal Pigment Epithelium

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in retinopathies using non-viral and viral gene delivery to the retina and retinal pigment epithelium (RPE). Subretinal injection is the best approach to deliver viral vectors directly to RPE cells. Before the clinical trial of a gene therapy, it is inevitable to validate the efficacy of the therapy in animal models of various retinopathies. Thus, subretinal injection in mice becomes a fundamental technique for an ocular gene therapy. In this protocol, we provide the easy and replicable technique for subretinal injection of viral vectors to experimental mice. This technique is modified from the intravitreal injection, which is widely used technique in ophthalmology clinics. The representative results of RPE/choroid/scleral complex flat-mount will help to understand the efficacy of this technique and adjust the volume and titer of viral vectors for the extent of gene transduction.

Subretinal injection is a surgical technique for effective gene delivery to retinal pigment epithelium in the mouse eye. Here we describe an easy and replicable method for subretinal injection of viral vectors to retinal pigment epithelium in experimental mice.

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in ...

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 ...

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 ...

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for determining the mechanisms underlying retinal diseases. Maintaining structural integrity throughout the tissue preparation is vital for obtaining reproducible retinal IHC data but can be challenging due to the fragility and complexity of retinal cytoarchitecture. Strong fixatives like 10% formalin or Bouin's solution optimally preserve the retinal structure, they often impede IHC analysis by enhancing the background fluorescence and/or diminishing antibody-epitope interactions, a process known as epitope masking. Milder fixatives, on the other hand, like 4% paraformaldehyde, reduces background fluorescence and epitope-masking, meticulous dissection techniques must be utilized to preserve the retinal structure. In this article, we present a comprehensive method to prepare mouse ocular posterior cups for IHC that is sufficient to preserve most antibody-epitope interactions without loss of retinal structural integrity. We include representative IHC with antibodies to various retinal cell type markers to illustrate tissue preservation and orientation under optimal and sub-optimal conditions. Our goal is to optimize IHC studies of the retina by providing a complete protocol from ocular posterior cup dissection to IHC.

This report describes comprehensive methods for preparing frozen mouse retina sections for immunohistochemistry (IHC). Methods described include dissection of the ocular posterior cup, paraformaldehyde fixation, embedding in Optimal Cutting Temperature (OCT) media and tissue orientation, sectioning and immunostaining.