The author would like to acknowledge use of the Children's Research Institute Imaging Core for the confocal microscopy and Dr. Jian Zhang for the suggestion to mount sections using agarose beads. This work was funded by N.I.H. grant DC010387.

1. Isolation and fixation of inner ears
<ol>
 <li>For analysis of cellular patterning in the organ of Corti, euthanize appropriately staged embryos or adults.</li>
 <li>Dissect out the otic capsule, containing the membranous inner ear. In the mouse, this can be easily done on embryos at embryonic (E) day 14.5 and older, in which E0.5 is the day at which a vaginal plug was detected. Dissection of the otic capsule is accomplished first by decapitation and opening of the skull at the midline. Remove brain to expose each ear (Fig. 1A). Inner ears can be shelled out intact using forceps (such as Dumont #5, Fig. 1B). If imaging the organ of Corti, careful removal of all neuronal and connective tissue associated with the otic capsule is not necessary.</li>
 <li>Fix whole inner ears by immersion in 4% paraformaldehyde (PFA) with gentle rocking at 4&deg;C overnight in 1.8 ml screw-capped NUNC Cryo-tube vials, or other appropriately sized container. PFA solution should be made fresh in ddH20, then stored at -20&deg;C until needed.</li>
 <li>The next day, wash out the fixative with three changes of 1X phosphate buffered saline (PBS). Tissue can be stored in sterile 1X PBS for days up to months until ready for processing. For adult tissue, decalcify in 10% EDTA (0.27 M) at 4&deg;C for 5 days, followed by rinsing and storage in 1X PBS.</li>
 
</ol>

2. Cutting vibratome sections

<ol>
 <li> Prepare 4% Low Melting Point agarose in 1X PBS. Add low melting point agarose to 1X PBS and microwave until agarose is in solution. Take care that the solution does not bubble over - take out of the microwave periodically and swirl to mix. Keep solution in a 55&deg;C water bath until ready to use.</li>
 <li>Using forceps, remove inner ear from 1X PBS solution and position at the bottom of a Peel-A-Way embedding mold. Pipet away any excess 1X PBS from the mold. Fill mold to cover the tissue with plenty of liquid 4% Low Melting Point agarose (Fig. 2A).</li>
 <li>Position the inner ear for sectioning in the appropriate plane. I have obtained good cross-sections of the organ of Corti by approximating the position of the otic capsule as if it had been left in situ in the head and sectioning in the transverse plane. For E14.5 embryos to adults, position the inner ear so that it is viewed laterally, with the lateral semi-circular canal visible (Fig. 2B). Angle the inner ear so that the line formed by the posterior side of the otic capsule is approximately at a 45-degree angle from the intended plane of sectioning (Fig. 2A, B). When the otic capsule is viewed so that the dorsal portion of the otic capsule is at the top, the basal turn of the cochlea should be located at the bottom (arrowhead, Fig. 2B) and the anterior semi-circular canal (asc) should be at the top (Fig. 2B). </li>
 <li>Agarose will set at RT within 30 mins. Transfer Peel-Away molds containing embedded tissue to 4&deg;C until ready to section. If sectioning will be put off until the next day, add 1X PBS to cover the agarose in the mold and store in a box containing moist paper towels soaked in 1X PBS.</li>
 <li>Peel away the plastic mold. Using a razor blade, cut out an agarose box containing the inner ear tissue. Take care to ensure that the plane of section is straight and parallel to the opposite side of the block, which will be the surface on which the agarose block is glued down. Since the agarose does not infiltrate the tissue, do not trim too much agarose from around the tissue, so that the tissue will be well-supported.</li>
 <li>Use superglue to attach the block to the cutting surface.</li>
 <li>Section at 40 to 100 &mu;m thickness by vibratome. Save sections in a tissue-culture dish filled with 1X PBS, 0.1% TritonX-100. A representative section through the cochlear spiral is shown in Fig. 2C.</li>
 <li>Use forceps to dissect away the agarose from the tissue. Agarose will not penetrate the tissue, but will just be stuck to the outer surface of the inner ear.</li>
 <li>If desired, pool agarose-free, sectioned tissue using a pipet with a large-hole opening to prevent excessive shearing of the tissue. If performing multiple antibody stains on multiple different tissues, sections can be organized in a 16- or 24-well tissue culture dish.</li> 
</ol>
3. Antibody staining vibratome sections
Proceed with antibody stain. Process sections in whole-mount, free-floating in solution. Below is an example antibody staining protocol for the S100 marker of the pillar cells of the organ of Corti, but the protocol can be used for any antibody. All washes are conducted at room temperature unless otherwise noted.
<ol>
 <li>In a multi-well tissue culture dish, incubate free-floating sections with gentle rocking in PBT (1X PBS, 1% bovine serum albumin, 0.1% Triton X-100) for 30 mins.</li>
 <li>Pipet off old solution and block free-floating sections with gentle rocking in PBT + normal goat serum (50 &mu;l NGS/ 1 ml PBT) for 30 mins.</li>
 <li>Pipet off old solution and add primary antibody diluted in PBT + NGS (1:200 dilution of S100 in PBT+NGS). Incubate overnight at 4&deg;C with gentle rocking.</li>
 <li>Pipet off primary antibody solution. Wash 3X, 5 mins. with PBT, then 4X, 30 mins. with PBT.</li>
 <li>Block with PBT + NGS, 30 mins.</li>
 <li>Incubate with fluorescently-conjugated secondary antibody (raised in goat) diluted in PBT + NGS. Incubate overnight at 4&deg;C, or a few hours, RT, gentle rocking.</li>
 <li>Pipet off secondary antibody solution. Wash 3X, 5 mins., then 4X, 30 mins. with PBT.</li>
 <li>Pipet off last PBT wash and replace with appropriate aqueous mounting medium containing a nuclear counterstain.</li>
</ol>
4. Mounting sections on slides and confocal microscopy
To prevent crushing the thick sections with the coverslip, when mounting individual sections, a spacer must be inserted between the microscope slide and the coverslip. Below are two alternative methods of mounting thick vibratome sections:
<ol>
 <li>
 <ol type="a">
 <li>A spacer can be created by placing a rim of vacuum grease in a square around the tissue section. Thus, when laying down the coverslip, the edges of the coverslip will be sealed with vacuum grease. Press down on the edges of the coverslip with your thumb to firmly adhere it onto the slide. If the vacuum grease is neatly and uniformly present around all edges of the coverslip, the slide will be sealed sufficiently for use on an inverted confocal microscope.</li>
 <li>Alternatively, a spacer can be created using agarose chromatography resin of known mesh size (such as Affi-Gel Blue Gel, Bio-Rad). Pipet 1 &mu;l of resin onto the microscope slide in each of the four corners where the coverslip will be placed. Pipet one tissue section and mounting media into the middle of the four dots of resin. Coverslip. If using an inverted confocal microscope, seal the coverslip onto the microscope slide using nailpolish.</li> 
 </ol> 
 </li>
 <li>Obtain optical sections of the stained vibratome slices using confocal microscopy. To quantify various cell types within the organ of Corti, obtain a z-stack at a defined step size (between 1.5 and 3 &mu;m). Sections should be counter-stained with a nuclear dye (step 3.8). Images can be examined on any computer screen after confocal image collection. Cells can be counted manually be observing the appearance and disappearance of nuclei as images are examined sequentially through the z-stack.</li>
</ol>
5. Representative Results:
A representative confocal z-stack through a vibratome section, stained for S100 protein, which is present in pillar and Deiters' cells, and counterstained with a nuclear dye (Fig. 3A-F). Each panel is the confocal image taken at 3 &mu;m-steps, with panel A the confocal image obtained closest to the surface of the vibratome section and panel F the confocal image obtained most internally within the vibratome section. The morphology of the organ of Corti appears minimally disrupted. In particular, the cytoplasmic extensions of the pillar cells are not broken. By noting the appearance and disappearance of pillar cell nuclei through the z-stack (Fig. 3A to 3F), both inner and outer pillar cell number can be counted (see numbering, Fig. 3A - F). Due to the small number of each cell type per confocal image, this approach can be used to count most cell types within the organ of Corti.
<img src="/files/ftp_upload/2793/2793fig1.jpg" alt="Figure 1"/> 
Figure 1. Intact, 15-week-old, adult inner ears encased in the otic capsule. (A) The right otic capsule (bracketed) encased in the skull, viewed from a vantage point of inside the head, after removal of the brain. Anterior is to the right. (B) Intact, 15-week-old inner ears after dissection from the skull. Lateral surface of the left inner ear is visible; medial surface of the right inner ear is visible. Labels indicate the locations of the anterior semi-circular canal (asc), co (cochlea), and ow (oval window).
<img src="/files/ftp_upload/2793/2793fig2.jpg" alt="Figure 2"/> 
Figure 2. (A) Whole inner ear from a 15-week-old adult, embedded in agarose in the center of a Peel-Away mold. Intended plane of section is indicated. (B) Close-up image of a 15-week-old, whole inner ear prior to embedding. A portion of the surrounding bone has been dissected away so the membranous labyrinth could a viewed more clearly. The lateral semi-circular canal (lsc), anterior semi-circular canal (asc), and cochlea (co) are traced (white curves). The basal turn of the cochlea (arrowhead) and intended plane of section (dashed line) are indicated. (C) Representative cross-section through the cochlea. In one cross-section through the cochlear duct, the organ of Corti (oC) is boxed, the stria vascularis (sv) is bracketed, and the Reissner's membrane (Rm) is indicated.
<img src="/files/ftp_upload/2793/2793fig3.jpg" alt="Figure 3"/> 
Figure 3. The organ of Corti in a wild-type mouse at P21. (A-F) Sequential confocal series, at a 3 &mu;m step-size, through a S100 antibody-stained organ of Corti to visualize the pillar cells and Deiters' cells. Counts of inner and outer pillar cells from the start to the end of the series are numbered. Abbreviations: IHC (inner hair cell), aIHC (adjacent inner hair cell), OHC (outer hair cell), PC (pillar cell), DC (Deiters' cell).

<ol>
	<li>Lim, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+structure+of+the+organ+of+Corti:+a+review.">Functional structure of the organ of Corti: a review.</a> Hear Res. 22, 117-146 (1986).</li><li>Raphael, Y., Altschuler, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+and+innervation+of+the+cochlea.">Structure and innervation of the cochlea.</a> Brain Res Bull. 60, 397-422 (2003).</li><li>Chung, W. S., Shin, C. H., Stainier, D. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bmp2+signaling+regulates+the+hepatic+versus+pancreatic+fate+decision.">Bmp2 signaling regulates the hepatic versus pancreatic fate decision.</a> Dev Cell. 15, 738-748 (2008).</li><li>Sun, X., Mariani, F. V., Martin, G. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functions+of+FGF+signalling+from+the+apical+ectodermal+ridge+in+limb+development.">Functions of FGF signalling from the apical ectodermal ridge in limb development.</a> Nature. 418, 501-508 (2002).</li><li>Trinh, L. A., Stainier, D. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fibronectin+regulates+epithelial+organization+during+myocardial+migration+in+zebrafish.">Fibronectin regulates epithelial organization during myocardial migration in zebrafish.</a> Dev Cell. 6, 371-382 (2004).</li><li>Yin, C., Kikuchi, K., Hochgreb, T., Poss, K. D., Stainier, D. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hand2+regulates+extracellular+matrix+remodeling+essential+for+gut-looping+morphogenesis+in+zebrafish.">Hand2 regulates extracellular matrix remodeling essential for gut-looping morphogenesis in zebrafish.</a> Dev Cell. 18, 973-984 (2010).</li><li>Shim, K., Minowada, G., Coling, D. E., Martin, G. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sprouty2,+a+mouse+deafness+gene,+regulates+cell+fate+decisions+in+the+auditory+sensory+epithelium+by+antagonizing+FGF+signaling.">Sprouty2, a mouse deafness gene, regulates cell fate decisions in the auditory sensory epithelium by antagonizing FGF signaling.</a> Dev Cell. 8, 553-564 (2005).</li></ol>

Experiments on animals were performed in accordance with the guidelines and regulations set forth by the Medical College of Wisconsin IACUC.

The procedure of vibratome sectioning, followed by immunohistochemistry and confocal imaging allows for the visualization of the organ of Corti with minimal tissue damage. Clearly, confocal images taken from internal regions in the vibratome section show excellent preservation of cellular morphology that is limited only by fixation artifacts. Confocal images taken close to the two cut-surfaces of a vibratome section are often indistinguishable from images from internal regions, although occasional cellular disruptions, s...

<table border="1">
	<tbody>
 	<tr>
 	<th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments (optional)</th>
 </tr>
 <tr>
 	<td>Leica VT1000S Vibrating Blade Microtome</td>
 <td>Leica </td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Zeiss LSM510 laser scanning confocal microscope</td>
 <td>Zeiss</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Student Dumont #5 Forceps</td>
 <td>Fine Science Tools</td>
 <td>91150-20</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>UltraPure Low Melting Point Agarose</td>
 <td>Invitrogen</td>
 <td>16520-050</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Peel-A-Way embedding mold</td>
 <td>Polysciences</td>
 <td>18986 or 18646A</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>bovine serum albumin</td>
 <td>Jackson ImmunoResearch</td>
 <td>001-000-162</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Goat Serum</td>
 <td>Invitrogen</td>
 <td>16210-064</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>S100</td>
 <td>Dako</td>
 <td>Z0311</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Alexa Fluor 568 goat anti-rabbit IgG</td>
 <td>Invitrogen</td>
 <td>A-11036</td>
 <td>1:1000 dilution</td>
 </tr>
 <tr>
 	<td>Vectashield</td>
 <td>Vector Labs</td>
 <td>H-1000</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>YO-PRO-1</td>
 <td>Invitrogen</td>
 <td>Y3603</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>vacuum grease</td>
 <td>Fisher</td>
 <td>S41718</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Affi-Gel Blue Gel</td>
 <td>Bio-Rad</td>
 <td>153-7301</td>
 <td>&nbsp;</td>
 </tr>
 
 </tbody>
</table>

vibratome sectioning for enhanced preservation of the cytoarchitecture of the mammalian organ of corti

The mammalian organ of Corti is a highly ordered cellular mosaic of mechanosensory hair and nonsensory supporting cells (reviewed in 1,2).Visualization of this cellular mosaic often requires that the organ of Corti is cross-sectioned. In particular, the nonsensory pillar and Deiters' cells, whose nuclei are located basally with respect to the hair cells, cannot be visualized without cross-sectioning the organ of Corti. However, the delicate cytoarchitecture of the mammalian organ of Corti, including the fine cytoplasmic processes of the pillar and Deiters' cells, is difficult to preserve by routine histological procedures such as paraffin and cryo-sectioning, which are compatible with standard immunohistochemical staining techniques.
Here I describe a simple and robust procedure consisting of vibratome sectioning of the cochlea, immunohistochemical staining of these vibratome sections in whole mount, followed by confocal microscopy. This procedure has been used widely for immunhistochemical analysis of multiple organs, including the mouse limb bud, zebrafish gut, liver, pancreas, and heart (see 3-6 for selected examples). In addition, this procedure was sucessful for both imaging and quantitificaton of pillar cell number in mutant and control organs of Corti in both embryos and adult mice 7. This method, however, is currently not widely used to examine the mammalian organ of Corti. The potential for this procedure to both provide enhanced preservation of the fine cytoarchitecture of the adult organ of Corti and allow for quantification of various cell types is described.

Watch this Scientific Journal Video about Vibratome Sectioning for Enhanced Preservation of the Cytoarchitecture of the Mammalian Organ of Corti at JoVE.com

Vibratome Sectioning for Enhanced Preservation of the Cytoarchitecture of the Mammalian Organ of Corti

A simple procedure of vibratome sectioning the organ of Corti, followed by immunohistochemistry and confocal microscopy is described. This procedure allows for improved preservation of the fine cytoarchitecture of the mammalian organ of Corti, and consequently allows for accurate quantification of cell types.

The mammalian organ of Corti is a highly ordered cellular mosaic of mechanosensory hair and nonsensory supporting cells (reviewed in 1,2).Visualization of ...

vibratome-sectioning-for-enhanced-preservation-cytoarchitecture

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24.2K 조회수.  Medical College of Wisconsin. immunohistochemistry과 공촛점 현미경 다음 Corti의 기관을 sectioning vibratome의 간단한 절차 설명합니다. 이 절차는 Corti의 포유 동물 장기의 벌금 cytoarchitecture 개선 보존을 허용하고, 결과적으로 세포 유형의 정확한 양을 정함을 허용합니다.

비디오: Vibratome Sectioning for Enhanced Preservation of the Cytoarchitecture of the Mammalian Organ of Corti

Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals through the vitreous fluid ensures that they act upon the entire RGC population. Viral vectors, plasmids or short interfering RNAs (siRNAs) can also be delivered to the vitreous chamber in order to infect or transfect retinal cells 5-12. The high tropism of Adeno-Associated Virus (AAV) vectors is beneficial to target RGCs, with an infection rate approaching 90% of cells near the injection site 6, 7, 13-15. Moreover, RGCs can be selectively transfected by applying siRNAs, plasmids, or viral vectors to the cut end of the optic nerve 16-19 or injecting vectors into their target the superior colliculus 10. This allows researchers to study apoptotic mechanisms in the injured neuronal population without confounding effects on other bystander neurons or surrounding glia. RGC apoptosis has a characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a window for experimental manipulations directed against pathways involved in apoptosis. Manipulations that directly target RGCs from the transected optic nerve stump are performed at the time of axotomy, immediately after cutting the nerve. In contrast, when substances are delivered via an intraocular route, they can be injected prior to surgery or within the first 3 days after surgery, preceding the initiation of apoptosis in axotomized RGCs. In the present article, we demonstrate several methods for experimental manipulations after optic nerve transection.

Optic Nerve transection is a widely used model of adult CNS injury. This model is ideal for performing a number of experimental manipulations that target the retina globally or directly target the injured neuronal population of retinal ganglion cells.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Vibratome Sectioning Mouse Retina to Prepare Photoreceptor Cultures

The retina is a part of the central nervous system that has organized architecture, with neurons in layers from the photoreceptors, both rods and cones in contact with the retinal pigmented epithelium in the most distant part on the retina considering the direction of light, and the ganglion cells in the most proximal distance. This architecture allows the isolation of the photoreceptor layer by vibratome sectioning. The dissected neural retina of a mouse aged 8 days is flat-embedded in 4% gelatin on top of a slice of 20% gelatin photoreceptor layer facing down. Using a vibratome and a double edged razor blade, the 100 &#181;m thick inner retina is sectioned. This section contains the ganglion cells and the inner layer with notably the bipolar cells. An intermediary section of 15 &#181;m is discarded before 200 &#181;m of the outer retina containing the photoreceptors is recovered. The gelatin is removed by heating at 37 &#176;C. Pieces of outer layer are incubated in 500 &#181;l of Ringer&#39;s solution with 2 units of activated papain for 20 min at 37 &#176;C. The reaction is stopped by adding 500 &#181;l 10% fetal calf serum (FCS) in Dulbecco&#39;s Modified Eagle Medium (DMEM), then 25 units of DNAse I is added before centrifugation at RT, washed several times to remove serum and the cells are resuspended in 500 &#181;l of DMEM and seeded at 1 x 105 cells/cm2. The cells are grown to 5 days in vitro and their viability scored using live/dead assay. The purity of the culture is first determined by microscopic observation during the experiment. The purity is then validated by seeding and fixing cells on a histological slide and analyzing using a rabbit polyclonal anti-SAG, a photoreceptor marker and mouse monoclonal anti-RHO, a rod photoreceptor specific marker. Alternatively, the photoreceptor layer (97% rods) can be used for gene or protein expression analysis and for transplantation.

Neural retina of a mouse aged 8 days is on top of a 4% gelatin block. After isolation of the photoreceptor layer (200 &#181;m) by vibratome, the photoreceptors are seeded after mechanical and enzymatic dissociation for culture. The photoreceptor layer can be used for molecular, biochemical analyses or transplantation.

The retina is a part of the central nervous system that has organized architecture, with neurons in layers from the photoreceptors, both rods and cones in ...

In-depth Physiological Analysis of Defined Cell Populations in Acute Tissue Slices of the Mouse Vomeronasal Organ

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory neurons (VSNs) express a specific type of G protein-coupled receptor (GPCR) from at least three different chemoreceptor gene families allowing sensitive and specific detection of chemosensory cues. These families comprise the V1r and V2r gene families as well as the formyl peptide receptor (FPR)-related sequence (Fpr-rs) family of putative chemoreceptor genes. In order to understand the physiology of vomeronasal receptor-ligand interactions and downstream signaling, it is essential to identify the biophysical properties inherent to each specific class of VSNs.
The physiological approach described here allows identification and in-depth analysis of a defined population of sensory neurons using a transgenic mouse line (Fpr-rs3-i-Venus). The use of this protocol, however, is not restricted to this specific line and thus can easily be extended to other genetically modified lines or wild type animals.

Here, we describe a physiological approach that allows identification and in-depth analysis of a defined population of sensory neurons in acute coronal tissue slices of the mouse vomeronasal organ using whole-cell patch-clamp recordings.

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory ...

Sublimation of DAN Matrix for the Detection and Visualization of Gangliosides in Rat Brain Tissue for MALDI Imaging Mass Spectrometry

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass Spectrometry (IMS) experiments. Determining the appropriate protocol to follow throughout the sample preparation process can be difficult as each step must be optimized to comply with the unique characteristics of the analytes of interest. This process involves not only finding a compatible matrix that can desorb and ionize the molecules of interest efficiently, but also selecting the appropriate matrix deposition technique. For example, a wet matrix deposition technique, which entails dissolving a matrix in solvent, is superior for desorption of most proteins and peptides, whereas dry matrix deposition techniques are particularly effective for ionization of lipids. Sublimation has been reported as a highly efficient method of dry matrix deposition for the detection of lipids in tissue by MALDI IMS due to the homogeneity of matrix crystal deposition and minimal analyte delocalization as compared to many wet deposition methods 1,2. Broadly, it involves placing a sample and powdered matrix in a vacuum-sealed chamber with the samples pressed against a cold surface. The apparatus is then lowered into a heated bath (sand or oil), resulting in sublimation of the powdered matrix onto the cooled tissue sample surface. Here we describe a sublimation protocol using 1,5-diaminonaphthalene (DAN) matrix for the detection and visualization of gangliosides in the rat brain using MALDI IMS.

A protocol for the sublimation of DAN matrix onto rat brain tissue for the detection of gangliosides using MALDI Imaging Mass Spectrometry is presented.

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass ...

Visualization of Cortical Modules in Flattened Mammalian Cortices

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and can be delineated by certain histochemical and immunohistochemical methods. In this study, we highlight a method to isolate the cortex from mammalian brains and flatten them to obtain sections parallel to the cortical sheet. We further highlight selected histochemical and immunohistochemical methods to process these flattened tangential sections to visualize cortical modules. In the somatosensory cortex of various mammals, we perform cytochrome oxidase histochemistry to reveal body maps or cortical modules representing different parts of the body of the animal. In the medial entorhinal cortex, an area where grid cells are generated, we utilize immunohistochemical methods to highlight modules of genetically determined neurons which are arranged in a grid-pattern in the cortical sheet across several species. Overall, we provide a framework to isolate and prepare layer-wise flattened cortical sections, and visualize cortical modules using histochemical and immunohistochemical methods in a wide variety of mammalian brains.

This article describes a detailed methodology to obtain flattened tangential sections from mammalian cortices and visualize cortical modules using histochemical and immunohistochemical methods.

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and ...

A Volumetric Method for Quantification of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor determining patient outcome and is therefore frequently taken as a study endpoint. However, in small animal studies on SAH, quantification of cerebral vasospasm is a major challenge. Here, an ex vivo method is presented that allows quantification of volumes of entire vessel segments, which can be used as an objective measure to quantify cerebral vasospasm. In a first step, endovascular casting of the cerebral vasculature is performed using a radiopaque casting agent. Then, cross-sectional imaging data are acquired by micro computed tomography. The final step involves 3-dimensional reconstruction of the virtual vascular tree, followed by an algorithm to calculate center lines and volumes of the selected vessel segments. The method resulted in a highly accurate virtual reconstruction of the cerebrovascular tree shown by a diameter-based comparison of anatomical samples with their virtual reconstructions. Compared with vessel diameters alone, the vessel volumes highlight the differences between vasospastic and non-vasospastic vessels shown in a series of SAH and sham-operated mice.

The goal of this article is to present a method that allows a 3-dimensional reconstruction of the cerebrovascular tree in mice after micro computed tomography and determination of volumes of entire vessel segments that can be used to quantify cerebral vasospasm in murine models of subarachnoid hemorrhage.

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...