The authors are grateful to Dr. Shimon P. Francis for generating the confocal micrographs.
This work was supported by the Division of Intramural Research at the National Institute on Deafness and Other Communication Disorders. Additional support was provided by NIDCD 5R01 DC007613.

1. Utricle Dissection and Culture
<ol>
 <li>Euthanize an adult (4 weeks of age or older) mouse using an approved protocol and decapitate.</li>
 <li>Snip the external auditory canal on both sides of the head and pull the skin forward towards the nose. Bisect the head from back to front and remove the brain from both sides to reveal the bony labyrinth.</li>
 <li>Trim the skull away from the bony cochlea and transfer the bony labyrinth (including the bulla) to a tissue culture hood equipped with a dissecting microscope (Note: Adenovirus work should be performed in a Class II biological safety cabinet).</li>
 <li>In the hood, transfer each cochlea to a 35mm tissue culture dish containing sterile dissecting media (M199, Gibco/Invitrogen #12350) (Figure 1A).</li>
 <li>If the auditory bulla is still intact (it is possible to remove the bulla during the gross dissection by inserting your thumbnail between the bulla and the apex of the cochlea), use two forceps (one #3 and one #5) to break the bulla (Figure 1B).</li>
 <li>Identify the landmarks of the bony cochlea preparation: apex, base, ossicles, oval and round windows, VIIIth nerve root, semicircular canals (Figure 1 C, D).</li>
 <li>Place the inner ear in the dish with the medial side facing up, such that the oval and round windows and apex are on the bottom of the dish and the VIIIth nerve root is facing up. Hold the preparation using a #3 forceps in the region of the semicircular canals (Figure 1D).</li>
 <li>Using a scalpel equipped with a #11 blade, cut off the apex of the cochlea just apical to the nerve root (Figure 1D).</li>
 <li>Turn the preparation so that the cut portion is facing up. Use the anterior semicircular canal as a handle to stabilize the preparation (Figure 1E).</li>
 <li>Using #5 forceps, remove the cochlea at the modiolus down to the base. At the point where the hook region of the cochlea turns downward, use a fine probe (half of a broken #55 forceps) to lift the last bony shelf of the osseous spiral lamina (Figure 1F), revealing the saccule. Just beneath the saccule is the stapes footplate, which is an important landmark. The utricle is immediately adjacent to the stapes footplate (Figure 1G), and it is surrounded by bone. Use the probe to chip the bone away enough to reveal about 1/3 of the utricle. Carefully remove the utricle using #55 forceps (Figure 1H). This procedure will usually result in removal of the pigmented roof epithelium as the utricle is pulled from the bony preparation. However, if the utricle is removed with the roof epithelium intact, the roof can be removed using #55 forceps.</li>
 <li>Utricles for adenovirus infection (or live imaging experiments) must have the otoconia removed during the dissection (prior to infection). In this case, use a luer-lock syringe that is filled with dissecting media and fitted with a small-bore needle (usually 26-28 gauge). Hold the utricle at the edge using #55 forceps (Figure 1I). Bring the needle tip close to the utricle with the bevel of the needle pointing down. Use a stream of dissecting media to blow off the otoconia (Figure 1J). Alternatively, otoconia can be removed by brushing them off using an eyelash tool (Ted Pella, Inc. #113). Utricles that are not going to undergo adenoviral infection are normally cultured free-floating in 24-well tissue culture plates with the otoconia intact (see below for post-fix otoconia removal, which is easier).</li>
 <li>Once all utricles are dissected, change the dissecting media to sterile culture media: DMEM F12 (Gibco #11320) supplemented with 5% FBS and 50 U/ml penicillin G (Sigma). </li>
 <li>Utricles are cultured at 37 &deg;C and 5% CO2. If cultures are to be maintained for more than 48 hours, half of the culture media should be changed every other day. Cultures can be maintained for a week or longer.</li>
</ol>
2. Adenovirus Infection of Supporting Cells

Important: Working with adenovirus requires Biosafety Level 2 (BSL2) procedures and certification. Check with your Institutional Biosafety Officer for guidance and training on BSL2 procedures.
<ol>
 <li>Dissect utricles as above in dissection media (no serum). Remove otoconia as above.</li>
 <li>When ready to infect, transfer 1 utricle (hair cells up) into each well of a Nunc mini-tray (Fisher #12-565-68) containing 15 &mu;L serum-free DMEM F12 (this is important because serum can inactivate the virus). This transfer is easier with a 1.5 mm microcurette (#10082-15 from Fine Science Tools).</li>
 <li>Our Ad-GFP is serotype 5 with the viral E1 and E3 genes deleted and the human CMV promoter driving the transgene (GFP). This virus was obtained from Vector BioLabs (Philadelphia, PA <a href="http://www.vectorbiolabs.com/">vectorbiolabs.com</a>). Stock viruses are provided at titers of 1.0-4.0&times;1010 PFU/ml. Add 0.5-2 &mu;l of stock virus to each well containing a utricle (Figure 1L). The actual amount used varies depending on the virus and stock titer. We usually infect each utricle with 1.0-4.0&times;107 PFU. </li>
 <li>Culture the utricles in virus-containing media at 37 &deg;C/5% CO2 for 2 hours. After 2 hours, transfer the utricles back to the 24-well tissue culture plate containing culture media with serum. Culture the utricles overnight at 37 &deg;C/5% CO2.</li>
 <li>Utricles can be used the next day for live imaging experiments, or they can be fixed for hair cell immunochemistry (see below). Viral transgene expression increases over time, so if transgene expression is low at 24 hours post-infection it may beneficial to culture cells an additional 24 hours in serum-containing media. If you see hair cell toxicity, reduce the amount of virus used. </li>
</ol>
3. Utricle Fixation, Otoconia Removal, and Immunochemistry
<ol>
 <li> At the end of the culture period, fix utricles in 4% paraformaldehyde for either 3 hours at room temp (on a rocking platform) or overnight at 4 &deg;C. Wash utricles (3 times, 15 min each) in 0.1 M (1X) Sorensen's phosphate buffer pH 7.4. These steps are performed in a 24-well tissue culture plate. </li>
 <li>Otoconia removal: Note: Otoconia must be removed prior to adenovirus infection (see step 1.11 above). However, utricles that are not to be infected can be cultured and fixed with the otoconia still intact. In this case, use the steps described here to remove the otoconia after fixation. Inspect utricles for any remaining pigmented roof epithelium. Carefully remove any remaining roof epithelium using two #55 forceps. To remove the otoconia, replace Sorensen's phosphate buffer with 750-1000 &mu;l Cal-Ex decalcifier (Fisher #CS510-1D). Leave Cal-Ex on utricles for 1 minute and 40 seconds (do not exceed 2 minutes). Replace Cal-Ex with 0.1 M Sorensen's phosphate buffer and wash (5 times, 5 min each).</li>
 <li>Incubate utricles in sodium borohydride (Sigma #452882, 1% in deionized water) for 10 mins. Wash utricles in 0.1 M Sorensen's phosphate buffer (5 times, 5 min each).</li>
 <li>Place utricles in Blocking Solution (PBS + 2% bovine serum albumin + 0.8% normal goat serum + 0.4% Triton X-100) for 3 hours at room temperature on a rocking platform.</li>
 <li>Add primary antibodies and incubate overnight at 4 &deg;C. Anti-Myosin 7a (either Proteus Biosciences #25-6790 or #MYO7A 138-1 from the Developmental Studies Hybridoma Bank) at 1:100 in blocking solution labels all hair cells. To separately label striolar and non-striolar hair cells, we have used a double-label protocol with monoclonal anti-calmodulin (Sigma #C 3545; 1:150) and polyclonal anti-calbindin (Chemicon #AB1778, Temecula, CA, USA; 1:200). Secondary antibodies (usually Alexa fluor conjugated secondary antibodies from Invitrogen, Carlsbad, CA, USA) are diluted 1:500 in blocking solution and incubated for 4 hours in the dark at room temperature on a rocking platform.</li>
 <li>Mount utricles on glass slides using Fluoromount-G (Southern Biotech, Birmingham, AL, USA). </li>
</ol>
4. Representative Results
Utricles cultured using this method retain full complements of both hair cells and supporting cells (Figure 2). Hair cells in healthy cultures show round nuclear profiles surrounded by thin cytoplasmic regions that are myosin 7a-positive (Figure 2, top panel). Supporting cells (labeled with anti-Sox2) are smaller and more densely packed than hair cells (Figure 2, lower panel). Adenovirus infects 25-50% of the supporting cells in the utricle, and no hair cells are infected (Figure 2, Ad-GFP panels). It should be noted that the optimal working titer of each adenovirus must be determined empirically, since hair cell death is possible if the viral titer is too high. In addition, regions of mechanical damage (caused during the dissection) will take up large amounts of virus. These regions of mechanical damage are easily distinguishable by a continuous line of cells with very high levels of GFP expression and missing hair cells. This is in contrast to the scattered GFP expression in supporting cells of an undamaged culture (Figure 2). 
<img src="/files/ftp_upload/3734/3734fig1.jpg" alt="Figure 1"/> 
Figure 1. Utricle Dissection. A: Auditory bulla (AB). B: The bulla is broken using two sturdy forceps. C: Bony cochlea (RW = round window, S = stapedial artery, C = cochlea, ST = stapes). D: A scalpel blade (SB) is used to cut off the apex of the cochlea apical to the VIIIth cranial nerve (8th N). ASC = anterior semicircular canal. E: The preparation is held firmly using a #3 forceps with one fork inserted into the anterior semicircular canal (ASC) and the other on the outermost bone (arrows). The membranous labyrinth is removed using forceps. F: With the membranous labyrinth removed, the basal turn of the osseous spiral lamina is visible near the hook region. A fine probe is inserted beneath this bony shelf to lift the shelf up and remove it (arrow). G: After removal of the saccule, utricle (U) is visible immediately adjacent to the stapes footplate (SF). H: Utricle (U) is removed using a #55 forceps (arrow). SF = stapes footplate. I: Utricle with otoconia (O) partially removed and sensory epithelium (SE) visible underneath. J: Otoconia (O) are removed from the utricle using a stream of media delivered by a syringe fitted with a 26-gauge needle. The shadow of the needle (S) is visible at the bottom of the image. K: Utricle with otoconia removed and sensory epithelium (SE) visible. L: Adenoviral infection: each utricle (U) is placed in an individual well of a mini-tray. Supporting cells are infected with adenovirus by pipetting virus into the well (P = Pipet tip).
<img src="/files/ftp_upload/3734/3734fig2.2.jpg" alt="Figure 2"/> 
Figure 2. Adenovirus-mediated infection of supporting cells. Utricles were infected with adenovirus driving expression of green fluorescent protein (Ad-GFP). Shown are confocal images of the hair cell layer and the supporting cell layer in the same area of one utricle. Hair cells are labeled with an antibody against Myosin 7a (magenta). Supporting cells are labeled with an antibody against Sox2 (red). Schematic shows the structure of the utricle sensory epithelium and the locations of the hair cells (HC) and supporting cells (SC). Locations of confocal (optical) sections shown in the upper (U) and lower (L) panels are indicated by dashed lines in the schematic. Upper panels: In confocal images taken at the level of the hair cell nuclei, Ad-GFP signal appears in the spaces between hair cells and does not overlap with the hair cell marker Myosin 7a. Lower panels: In confocal sections at the level of the supporting cell nuclei, Ad-GFP signal colocalizes with the supporting cell marker Sox-2. Ad-GFP infection results in GFP expression in supporting cells only, and no hair cells are infected.
<img src="/files/ftp_upload/3734/3734fig3.jpg" alt="Figure 3"/> 
Figure 3. Adenoviral infection does not result in death of hair cells or supporting cells. Utricles were infected with Ad-GFP at 4&times;108 PFU/ml. Utricles were labeled with antibodies against Myosin 7a (hair cell marker) and Sox2 (supporting cell marker). Hair cell and supporting cell counts were equal for control utricles and Ad-GFP infected utricles.

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No conflicts of interest declared.

Sensory hair cells are susceptible to death caused by a variety of stresses, including aging, noise trauma, and exposure to ototoxic drugs, including the aminoglycoside antibiotics and the antineoplastic agent cisplatin. In mammals hair cell death results in permanent hearing loss and/or balance disturbance. In vitro model systems are critical tools for studies aimed at determining the cellular and molecular mechanisms underlying hair cell death, as well as those aimed at preventing or reversing hair cell...

A correction was made to <a href="http://www.jove.com/video/3734">Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection</a>. There was an error with a figure, the authors middle names were omitted, and the acknowledgements were updated.

Erratum: Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection

<table border="1">
 <tbody>
 <tr>
 <td>Reagent</td>
 <td>Company</td>
 <td>Catalog number</td>
 </tr>
 <tr>
 <td>Medium 199</td>
 <td>Gibco/Invitrogen</td>
 <td>12350</td>
 </tr>
 <tr>
 <td>DMEM/F12</td>
 <td>Gibco/Invitrogen</td>
 <td>11320</td>
 </tr>
 <tr>
 <td>Fine forceps (#55)</td>
 <td>Fine Science Tools</td>
 <td>11255-20</td>
 </tr>
 <tr>
 <td>Fine forceps (#5)</td>
 <td>Fine Science Tools</td>
 <td>11252-30</td>
 </tr>
 <tr>
 <td>Forceps (#3)</td>
 <td>Fine Science Tools</td>
 <td>11231-30</td>
 </tr>
 <tr>
 <td>Penicillin G</td>
 <td>Sigma</td>
 <td>P3032</td>
 </tr>
 <tr>
 <td>Fetal bovine serum </td>
 <td>Invitrogen</td>
 <td>10082-139</td>
 </tr>
 <tr>
 <td>Nunc mini-tray</td>
 <td>Fisher</td>
 <td>12-565-68</td>
 </tr>
 <tr>
 <td>Adenovirus-GFP</td>
 <td>Vector Biolabs</td>
 <td>1060*</td>
 </tr>
 <tr>
 <td>Cal-Ex decalcifier</td>
 <td>Fisher</td>
 <td>CS510-1D</td>
 </tr>
 <tr>
 <td>sodium borohydride</td>
 <td>Sigma</td>
 <td>452882</td>
 </tr>
 <tr>
 <td>Anti-Myosin 7a (polyclonal)</td>
 <td>Proteus Biosciences</td>
 <td>25-6790</td>
 </tr>
 <tr>
 <td>Anti-Myosin 7a (monoclonal)</td>
 <td>Developmental Studies Hybridoma Bank</td>
 <td>MYO7A 138-1</td>
 </tr>
 <tr>
 <td>Anti-Calmodulin</td>
 <td>Sigma</td>
 <td>C 3545</td>
 </tr>
 <tr>
 <td>Anti-Calbindin</td>
 <td>Chemicon</td>
 <td>AB1778</td>
 </tr>
 <tr>
 <td>Fluoromount G</td>
 <td>Southern Biotechnology Associates</td>
 <td>0100-01 </td>
 </tr>
 </tbody>
 </table>
Table 1. Reagents and tools used in utricle culture and adenovirus infection.	 
* Note: Ad-GFP is normally provided by Vector Biolabs at a stock titer of 1x1010 PFU/ml. We requested a custom amplification in order to provide the stock virus at a titer of 1.2&times;1011 PFU/ml.

dissection of adult mouse utricle and adenovirus-mediated supporting-cell infection

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear. Since there is no cell line that satisfactorily represents mammalian hair cells, research on hair cells relies on primary organ cultures. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice (Figure 1) 1-6. The utricle is a vestibular organ, and the hair cells of the utricle are similar in both structure and function to the hair cells in the auditory organ, the organ of Corti. The adult mouse utricle preparation represents a mature sensory epithelium for studies of the molecular signals that regulate the survival, homeostasis, and death of these cells.
Mammalian cochlear hair cells are terminally differentiated and are not regenerated when they are lost. In non-mammalian vertebrates, auditory or vestibular hair cell death is followed by robust regeneration which restores hearing and balance functions 7, 8. Hair cell regeneration is mediated by glia-like supporting cells, which contact the basolateral surfaces of hair cells in the sensory epithelium 9, 10. Supporting cells are also important mediators of hair cell survival and death 11. We have recently developed a technique for infection of supporting cells in cultured utricles using adenovirus. Using adenovirus type 5 (dE1/E3) to deliver a transgene containing GFP under the control of the CMV promoter, we find that adenovirus specifically and efficiently infects supporting cells. Supporting cell infection efficiency is approximately 25-50%, and hair cells are not infected (Figure 2). Importantly, we find that adenoviral infection of supporting cells does not result in toxicity to hair cells or supporting cells, as cell counts in Ad-GFP infected utricles are equivalent to those in non-infected utricles (Figure 3). Thus adenovirus-mediated gene expression in supporting cells of cultured utricles provides a powerful tool to study the roles of supporting cells as mediators of hair cell survival, death, and regeneration.

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Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection

Mechanosensory hair cells are the receptor cells of the inner ear. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice. We present the dissection of the adult mouse utricle, and we demonstrate adenovirus-mediated infection of supporting cells in cultured utricles.

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear.  Since there ...

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JoVE Journal

Neuroscience

23.6K Görüntüleme.  Medical University of South Carolina. Mechanosensory saç hücreleri iç kulak reseptör hücrelerdir. En-karakterize In vitro model sistem yetişkin farelerin utricles organ kültürleri kullanır. Biz yetişkin fare kırbacık diseksiyonu mevcut ve biz kültürlü utricles hücreleri destekleyici adenovirüs aracılı enfeksiyonu gösterir.

Video: Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

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.

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 ...

Cerebrovascular Casting of the Adult Mouse for 3D Imaging and Morphological Analysis

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). Animal models are necessary for studying the etiopathology and potential therapies of cerebrovascular diseases. Imaging the vasculature in large animals is relatively easy. However, developing vessel imaging methods of murine brain disease models is desirable due to the cost and availability of genetically-modified mouse lines. Imaging the murine cerebral vascular tree is a challenge. In humans and larger animals, the gold standard for assessing the angioarchitecture at the macrovascular (conductance) level is x-ray catheter contrast-based angiography, a method not suited for small rodents.
In this article, we present a method of cerebrovascular casting that produces a durable skeleton of the entire vascular bed, including arteries, veins, and capillaries that may be analyzed using many different modalities. Complete casting of the microvessels of the mouse cerebrovasculature can be difficult; however, these challenges are addressed in this step-by-step protocol. Through intracardial perfusion of the vascular casting material, all vessels of the body are casted. The brain can then be removed and clarified using the organic solvent methyl salicylate. Three dimensional imaging of the brain blood vessels can be visualized simply and inexpensively with any conventional brightfield microscope or dissecting microscope. The casted cerebrovasculature can also be imaged and quantified using micro-computed tomography (micro-CT)1. In addition, after being imaged, the casted brain can be embedded in paraffin for histological analysis.
The benefit of this vascular casting method as compared to other techniques is its broad adaptation to various analytic tools, including brightfield microscopic analysis, CT scanning due to the radiopaque characteristic of the material, as well as histological and immunohistochemical analysis. This efficient use of tissue can save animal usage and reduce costs. We have recently demonstrated application of this method to visualize the irregular blood vessels in a mouse model of adult BAVM at a microscopic level2, and provide additional images of the malformed vessels imaged by micro-CT scan. Although this method has drawbacks and may not be ideal for all types of analyses, it is a simple, practical technique that can be easily learned and widely applied to vascular casting of blood vessels throughout the body.

In this article, we present a simple, practical technique for cerebrovascular casting that is easy to perform and can be utilized to image the vascular tree of the adult mouse brain.

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). ...

Dissection and Culture of Mouse Dopaminergic and Striatal Explants in Three-Dimensional Collagen Matrix Assays

Midbrain dopamine (mdDA) neurons project via the medial forebrain bundle towards several areas in the telencephalon, including the striatum1. Reciprocally, medium spiny neurons in the striatum that give rise to the striatonigral (direct) pathway innervate the substantia nigra2. The development of these axon tracts is dependent upon the combinatorial actions of a plethora of axon growth and guidance cues including molecules that are released by neurites or by (intermediate) target regions3,4. These soluble factors can be studied in vitro by culturing mdDA and/or striatal explants in a collagen matrix which provides a three-dimensional substrate for the axons mimicking the extracellular environment. In addition, the collagen matrix allows for the formation of relatively stable gradients of proteins released by other explants or cells placed in the vicinity (e.g. see references 5 and 6). Here we describe methods for the purification of rat tail collagen, microdissection of dopaminergic and striatal explants, their culture in collagen gels and subsequent immunohistochemical and quantitative analysis. First, the brains of E14.5 mouse embryos are isolated and dopaminergic and striatal explants are microdissected. These explants are then (co)cultured in collagen gels on coverslips for 48 to 72 hours in vitro. Subsequently, axonal projections are visualized using neuronal markers (e.g. tyrosine hydroxylase, DARPP32, or &beta;III tubulin) and axon growth and attractive or repulsive axon responses are quantified. This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of mesostriatal and striatonigral axon growth and guidance during development. Using this assay, it is also possible to assess other (intermediate) targets for dopaminergic and striatal axons or to test specific molecular cues.

Explants from the midbrain dopamine system and striatum are used in a collagen matrix assay for the in vitro analysis of mesostriatal and striatonigral pathway development. In this assay axonal outgrowth and guidance can be manipulated and quantified. It can also be modified for assessing other regions or molecular cues.

Midbrain dopamine (mdDA) neurons project via the medial forebrain bundle towards several areas in the telencephalon, including the striatum1. ...

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

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 ...

Adult Mouse DRG Explant and Dissociated Cell Models to Investigate Neuroplasticity and Responses to Environmental Insults Including Viral Infection

This protocol describes an ex vivo model of mouse-derived dorsal root ganglia (DRG) explant and in vitro DRG-derived co-culture of dissociated sensory neurons and glial satellite cells. These are useful and versatile models to investigate a variety of biological responses associated with physiological and pathological conditions of the peripheral nervous system (PNS) ranging from neuron-glial interaction, neuroplasticity, neuroinflammation, and viral infection. The usage of DRG explant is scientifically advantageous compared to simplistic single cells models for multiple reasons. For instance, as an organotypic culture, the DRG explant allows ex vivo transfer of an entire neuronal network including the extracellular microenvironment that play a significant role in all the neuronal and glial functions. Further, DRG explants can also be maintained ex vivo for several days and the culture conditions can be perturbed as desired. In addition, the harvested DRG can be further dissociated into an in vitro co-culture of primary sensory neurons and satellite glial cells to investigate neuronal-glial interaction, neuritogenesis, axonal cone interaction with the extracellular microenvironment, and more general, any aspect associated with the neuronal metabolism. Therefore, the DRG-explant system offers a great deal of flexibility to study a wide array of events related to biological, physiological, and pathological conditions in a cost-effective manner.

In this report, the advantages of organotypic cultures and dissociated primary cultures of mouse-derived dorsal root ganglia are highlighted to investigate a wide range of mechanisms associated with neuron-glial interaction, neuroplasticity, neuroinflammation, and response to viral infection.

This protocol describes an ex vivo model of mouse-derived dorsal root ganglia (DRG) explant and in vitro DRG-derived co-culture of dissociated sensory ...

Cochlear Surface Preparation in the Adult Mouse

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process. Loss of sensory hair cells is a common pathological feature of the varieties of acquired hearing loss. Additionally, the inner hair cell synapse can be damaged by mild insults. Therefore, surface preparations of cochlear epithelia, in combination with immunolabeling techniques and confocal imagery, are a very useful tool for the investigation of cochlear pathologies, including losses of ribbon synapses and sensory hair cells, changes in protein levels in hair cells and supporting cells, hair cell regeneration, and determination of report gene expression (i.e., GFP) for verification of successful transduction and identification of transduced cell types. The cochlea, a bony spiral-shaped structure in the inner ear, holds the auditory sensory end organ, the organ of Corti (OC). Sensory hair cells and surrounding supporting cells in the OC are contained in the cochlear duct and rest on the basilar membrane, organized in a tonotopic fashion with high-frequency detection occurring in the base and low-frequency in the apex. With the availability of molecular and genetic information and the ability to manipulate genes by knockout and knock-in techniques, mice have been widely used in biological research, including in hearing science. However, the adult mouse cochlea is miniscule, and the cochlear epithelium is encapsulated in a bony labyrinth, making microdissection difficult. Although dissection techniques have been developed and used in many laboratories, this modified microdissection method using cell and tissue adhesive is easier and more convenient. It can be used in all types of adult mouse cochleae following decalcification.

This article presents a modified cochlear surface preparation method that requires decalcification and use of a cell and tissue adhesive to adhere the pieces of cochlear epithelia to 10 mm round cover slips for immunohistochemistry in adult mouse cochleae.

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous ...

Cryo-section Dissection of the Adult Subependymal Zone for Accurate and Deep Quantitative Proteome Analysis

The subependymal neurogenic niche consists of a paraventricular ribbon of the lateral ventricular wall of the lateral ventricle. The subependymal zone (SEZ) is a thin and distinct region exposed to the ventricles and cerebrospinal fluid. The isolation of this niche allows the analysis of a neurogenic stem cell microenvironment. However, extraction of small tissues for proteome analysis&#160;is challenging, especially for the maintenance of considerable measurement depth and the achievement of reliable robustness. A new method termed cryo-section-dissection (CSD), combining high precision with minimal tissue perturbation, was developed to address these challenges. The method is compatible with state-of-the-art mass spectrometry (MS) methods that allow the detection of low-abundant niche regulators. This study compared the CSD and its proteome data to the method and data obtained by laser-capture-microdissection (LCM) and a standard wholemount dissection. The CSD method resulted in twice the quantification depth in less than half the preparation time compared to the LCM and simultaneously clearly outperformed the dissection precision of the wholemount dissection. Hence, CSD is a superior method for collecting the SEZ for proteome analysis.

Cryo-section-dissection allows fresh, frozen preparation of the largest neurogenic niche in the murine brain for deep quantitative proteome analysis. The method is precise, efficient, and causes minimal tissue perturbation. Therefore, it is ideally suited for studying the molecular microenvironment of this niche, as well as other organs, regions, and species.

The subependymal neurogenic niche consists of a paraventricular ribbon of the lateral ventricular wall of the lateral ventricle. The subependymal zone ...