The research was funded by The Lundbeck Foundation, Denmark.

The protocol was optimized and demonstrated on adult male albino rats. In all experimental procedures, animals were treated according to the regulations in the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. Animals were euthanized by carbon dioxide and subsequent cervical dislocation.
1. Rat Retinal Tissue Preparations
<ol>
	<li>Cryo-section
	<ol>
		<li>Make posterior and anterior ~0.5 cm slits of in the rat eyelid with a scalpel.
		<ol>
			<li>Optional: Using a diathermy burner, mark the eye at the inner angle to orientate the eye in a uniform fashion during embedding and allow for vertical cryostat sectioning through the optic nerve.</li>
		</ol>
		</li>
		<li>Grab the eye with forceps and tilt carefully to the side to expose the surrounding tissue. Enucleate the eye by making cuts with dissection scissors in the connective and muscular tissue. 
		CAUTION: Do not pull the eye too hard as excessive pressure on the optic nerve may cause retinal detachment.</li>
		<li>Put the eye briefly in 4% formaldehyde in phosphate-buffered saline (PBS) to stabilize before making an initial hole at the corneal limbus by applying light pressure with the tip of a scalpel.</li>
		<li>Under a microscope, cut along the corneal limbus with dissection scissors to remove the cornea and remove the lens with forceps before submerging in 4% formaldehyde in PBS for 2&#8211;4 h.</li>
		<li>Rinse sequentially in S&#246;rensen&#39;s phosphate buffer with 10% sucrose and 25% sucrose.</li>
		<li>Embed in Yazulla medium prepared with 3% gelatin from porcine skin and 30% albumin from chicken egg white. 
		NOTE: The protocol can be paused here with tissue storage at -20&#160;&#176;C.</li>
	</ol>
	</li>
	<li>Whole-mount
	<ol>
		<li>Make posterior and anterior ~0.5 cm slits in the rat eyelid with a scalpel.</li>
		<li>Grab the eye with forceps and tilt carefully to the side to expose the surrounding tissue. Enucleate the eye by making cuts with dissection scissors in the connective and muscular tissue. 
		CAUTION: Do not pull the eye too hard as excessive pressure on the optic nerve may cause retinal detachment.</li>
		<li>Put the eye briefly in 4% formaldehyde in PBS to stabilize before making an initial hole at the corneal limbus by applying light pressure with the tip of a scalpel.</li>
		<li>Under a microscope, cut along the corneal limbus with dissection scissors to remove the cornea and remove the lens with forceps.</li>
		<li>Separate the retina from the retinal pigment epithelium towards the optic nerve with forceps using small opening movements to avoid major tearing.</li>
		<li>Free the retina at the optic nerve with dissection scissors and make four slits of a few millimeters length from the retinal periphery towards the optic nerve head.</li>
		<li>Spread the retina onto a glass slide and allow drying for 5&#8211;10 min.</li>
		<li>Fix in 4% formaldehyde for 20&#8211;30 min by dripping formaldehyde onto the retina. 
		CAUTION: Do not apply directly on the retina as it may detach from the glass.</li>
		<li>Rinse with PBS. For optimal results, immuno-stain directly after rinsing.</li>
	</ol>
	</li>
	<li>Hypotonic isolation
	<ol>
		<li>Make posterior and anterior ~0.5 cm slits in the rat eyelid with a scalpel.</li>
		<li>Grab the eye with forceps and tilt carefully to the side to expose the surrounding tissue. Enucleate the eye by making cuts with dissection scissors in the connective and muscular tissue. 
		CAUTION: Do not pull the eye too hard as excessive pressure on the optic nerve may cause retinal detachment.</li>
		<li>Make an initial hole at the corneal limbus by applying light pressure with the tip of a scalpel.</li>
		<li>Under a microscope, cut along the corneal limbus with dissection scissors to remove the cornea and remove the lens with forceps.</li>
		<li>Separate the retina from the retinal pigment epithelium towards the optic nerve head with forceps using small opening movements to avoid major tearing.</li>
		<li>Free the retina at the optic nerve with dissection scissors, place the retina in 1 mL of deionized water in a 24-well plate and shake at 200 rpm with a 1.5 mm vibration orbit for 1 h at room temperature. 
		NOTE: Hereafter, the retina will appear less defined at the edges.</li>
		<li>Add 200 U DNAse 1 to dissociate the lysed cell debris from the retinal vasculature and shake for another 30 min at room temperature. 
		NOTE: Debris might start to form in the wells.</li>
		<li>Rinse minimum 3 times in deionized water for 5 min with shaking at 150&#8211;300 rpm to remove neuronal cell debris. The retina should become more transparent with each rinse indicative of the removal of neuronal cellular debris.
		<ol>
			<li>Use a dark background to look into the 24-well plate to clearly see the diaphanous isolated retinal vasculature.</li>
			<li>(Optional):&#160;If the vasculature does not appear free of neuronal layers (semi-transparent) at this point either add more rinse steps, increase the shaking speed or use a pipet to aspirate liquid onto the vasculature. 
			CAUTION: Either of the optional steps may damage the vasculature.</li>
		</ol>
		</li>
		<li>Fix 10 min in 1 mL of 4% paraformaldehyde in PBS at room temperature and rinse 3 times in PBS. 
		NOTE: The protocol can be paused here with tissue storage at 4&#160;&#176;C.</li>
	</ol>
	</li>
</ol>
2. Immunohistochemistry
<ol>
	<li>Staining of cryo-sections
	<ol>
		<li>Cut 10 &#181;m cryo-sections of the gelatin-embedded retina as vertical sectioning through the optic nerve and place the cryo-sections on a glass slide and let dry (minimum 1 h).</li>
		<li>Submerge the glass slide in PBS with 0.25 % Triton X-100 (PBS-T) for 15 min.</li>
		<li>Drip 1:100 PDGFR&#946; and 1:500 NG2 primary antibodies diluted in PBS-T + 1% BSA onto the cryo-section and incubate in incubation chambers at 4&#160;&#176;C overnight.</li>
		<li>Submerge the glass slide 2 times in PBS-T for 15 min and drip 1:100 anti-mouse Alexa Fluor 594-linked and 1:100 anti-rabbit FITC-linked secondary antibodies diluted in PBS-T with 3% BSA onto the cryo-sections.</li>
		<li>Incubate the glass slide 1 h at room temperature in the dark.</li>
		<li>Rinse the glass slide in PBS-T 2x 15 min. 
		NOTE: Optional: For double and triple immunofluorescent staining, sequential staining can be performed by repeating the procedure from 2.1.3 to 2.1.6 two and three times, respectively.</li>
		<li>Mount the stained cryo-sections with anti-fading mounting medium containing DAPI and a coverslip.</li>
	</ol>
	</li>
	<li>Staining of whole-mount
	<ol>
		<li>Drip PBS-T onto the whole-mount and incubate at room temperature for 15 min.</li>
		<li>Pour off and drip 1:100 PDGFR&#946; and 1:500 NG2 primary antibodies diluted in PBS-T + 1% BSA and incubate in moist chambers at 4&#160;&#176;C overnight.</li>
		<li>Pour off and drip on PBS-T to rinse the glass slide in 2x 15 min.</li>
		<li>Pour off and drip on 1:100 anti-rabbit Cy2- and 1:100 anti-mouse Cy3-linked secondary antibody diluted in PBS-T with 3% BSA to incubate 1 h in a moist chamber at room temperature in the dark.</li>
		<li>Pour off and drip on PBS-T to rinse in 2x 15 min in the dark. 
		NOTE: Optional: For double and triple immunofluorescent staining, sequential staining can be performed by repeating the procedure from 2.2.2 to 2.2.5 two and three times, respectively.</li>
		<li>Mount the stained whole-mount with anti-fading mounting medium containing DAPI and a coverslip.</li>
	</ol>
	</li>
	<li>Staining of hypotonic isolated vasculature
	<ol>
		<li>Block the hypotonic isolated vasculature 1 h with shaking at 100 rpm and room temperature with 500 &#181;L/well of 10 % donkey serum diluted in PBS.</li>
		<li>Incubate overnight at room temperature and shake at 100 rpm with 600 &#181;L/well of 1:100 PDGFR&#946; and 1:500 NG2 primary antibodies diluted in 10% donkey serum in PBS.</li>
		<li>Rinse the retinal network 3x in PBS for 5 min and incubate in 1:100 anti-mouse Alexa Fluor 594-linked and 1:100 anti-rabbit FITC-linked secondary antibodies diluted in 10% donkey serum in PBS shaking at 100 rpm and room temperature for 1 h in the dark.</li>
		<li>Rinse in PBS-T for 5 min and incubate in 0.2 ng/mL DAPI in PBS-T for 15 min followed by 3x 5 min rinses in PBS-T in the dark.</li>
		<li>Cut the tip of a plastic Pasteur pipet, moisten it with PBS-T and use it to transfer the retinal network to a 4-well glass chamber slide. 
		NOTE: The moistening step is important to avoid the retinal vasculature sticking to the inside of the Pasteur pipet.</li>
		<li>Unfold the retinal vasculature. Avoid touching the retinal vasculature with forceps as this can cause the vasculature to tangle. 
		NOTE: Unfolding can be done by tilting the chamber slide back and forth or aspirating drops of liquid onto the retinal vasculature.</li>
		<li>Remove the medium from the wells. The surface tension of the liquid will flatten the vasculature onto the bottom of the slide.</li>
		<li>Ensure correct unfolding under a microscope before removing the plastic wells from the chamber slide.</li>
		<li>Mount the stained vasculature with anti-fading mounting medium and a coverslip.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

We present three retinal preparation techniques that can be applied in the study of microvascular retinal pericytes. Below, we provide a comparison between each of the methods and highlight critical steps in the protocols.
With cryo-sectioning, the retina is cut in sagittal sections and hence, it is possible to obtain numerous specimens from the same retina. The numeral sections resulting from this method make it an ideal choice for antibody specificity and titration testing as it prevents unn...

Retinal pericytes are the focus of many research laboratories as these cells play a major role in the integrity of the vasculature. Pathological conditions such as diabetic retinopathy1, ischemia2, and glaucoma3 have vascular characteristics that involve the function of pericytes. Pericytes are found in the inner retinal capillary plexuses. The central retinal artery that supplies the inner retina branches into two layers of capillary plexuses. The inner vascular bed is situated between the ganglion cell and inner nuclear layers. The deeper layer is more dense and complex and is localized between the inner and outer nuclear layers4,5. In addition, some parts of the retina also contain a third network termed the radial parapapillary capillaries. These are long, straight capillaries that lie among the nerve fibers and rarely anastomose with one another or the other two plexuses6. Within the capillary wall, pericytes are embedded in the basement membrane and line the abluminal side of vascular endothelial cells.
To this date, there is no unique biological marker of these pericytes that can differentiate them from other vascular cells. Platelet-derived growth factor receptor &#946; (PDGFR&#946;) and nerve/glial antigen 2 (NG2) are commonly used markers which both present on pericytes but also other vascular cells. Identification of pericytes is further complicated by the existence of pericyte subsets that vary in morphology and protein expression7. Currently, the best identification relies on a combination of protein markers and the characteristic positioning of the pericyte in the vascular wall. We demonstrate here three different tissue preparation techniques for immunohistochemical PDGFR&#946;/NG2 staining of rat retinal microvascular pericytes, i.e., cryo-sections, whole-mounts, and hypotonic isolation of the vascular network.
With cryo-sections, the retina and sclera are cut through the optic nerve. This allows for the visualization of all layered structures of neurons. The distinct ten layers of the retina are apparent as interchanging nuclear and axonal/dendritic structures that can be visualized with stains such as hematoxylin/eosin or fluorescent nuclear 4',6-diamidino-2-phenylindole (DAPI)8. The metabolic requirements differ between the layers9 and it provides a method to determine the thickness or total absence of a specific layer (e.g., the loss of retinal ganglion cells is one of the hallmarks of retinal ischemia10,11). The vasculature is evident as transverse cuts through the retina, making it possible to separately study the capillary plexuses within the respective retinal layers12,13.
More traditionally, the investigations of the retinal vasculature network are performed in retinal whole-mounts. With this tissue preparation, the retina is cut and flattened as a flower-shaped structure. The method is a relatively fast tissue preparation technique that can highlight the overall architecture retinal vasculature and it is therefore often applied in the investigation of neovascularization in the murine retina. Successful visualization of the microvasculature in whole-mounted retinas is also reported in the developing neonatal mouse and rat retina14,15,16,17,18,19. These studies reveal a more defined pericytic activity with larger capillary-free areas in the adult compared to the neonatal retina14.
Another way of visualizing is the retinal microvasculature after hypotonic isolation. This tissue preparation technique results in retinal blood vessels and capillaries being freed of the neuronal cells. This type of two-dimensional imaging of the isolated retinal vascular network is usually performed after retinal trypsin digestion20 and used to assess the vascular abnormalities of diabetic retinopathy including pericyte loss and capillary degeneration20,21,22. The hypotonic isolation method offers the investigations of retinal vascular gene and protein regulatory responses as they has been done with RT-PCR and western blotting23,24,25. We provide here a protocol for the free-float immunohistochemical staining of the hypotonic isolated retinal vasculature as an alternative to trypsin digestion to examine the microvascular pericytes.

<table border="0" cellpadding="0" cellspacing="0" width="902">
	<tbody>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Geletin from porcine skin</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:165px;">G2625-500G</td>
			<td style="width:171px;"></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Albumin from chicken egg white</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:165px;">A5253-500G</td>
			<td style="width:171px;"></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Deoxyribonuclease (DNAse) I from bovine pancreas</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:165px;">D5025-15KU</td>
			<td style="width:171px;">Dissolved in 0.15 M NaCl</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Bovine serum albumin (BSA)</td>
			<td style="width:175px;">VWR</td>
			<td style="width:165px;">0332-100G</td>
			<td style="width:171px;"></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Normal donkey serum</td>
			<td style="width:175px;">Jackson ImmunoResearch</td>
			<td style="width:165px;">017-000-121, lot 129348</td>
			<td style="width:171px;"></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Rabbit anti-PDGFR&#946;</td>
			<td style="width:175px;">Santa Cruz</td>
			<td style="width:165px;">sc-432</td>
			<td style="width:171px;">1:100</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Mouse anti-NG2</td>
			<td style="width:175px;">Abcam</td>
			<td style="width:165px;">ab50009</td>
			<td style="width:171px;">1:500</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Alexa Fluor 594 AffiniPure Donkey Anti-Rabbit IgG</td>
			<td style="width:175px;">Jackson ImmunoResearch</td>
			<td style="width:165px;">711-585-152</td>
			<td style="width:171px;">1:100</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Fluorescein (FITC) AffiniPure Donkey Anti-Mouse IgG (H+L)</td>
			<td style="width:175px;">Jackson ImmunoResearch</td>
			<td style="width:165px;">715-095-151</td>
			<td style="width:171px;">1:100</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Cy2 AffiniPure Donkey Anti-Rabbit IgG (H+L)</td>
			<td style="width:175px;">Jackson ImmunoResearch</td>
			<td style="width:165px;">711-225-152</td>
			<td style="width:171px;">1:100</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Cy3 AffiniPure Donkey Anti-Mouse IgG (H+L)</td>
			<td style="width:175px;">Jackson ImmunoResearch</td>
			<td style="width:165px;">715-165-150</td>
			<td style="width:171px;">1:100</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">4&#39;,6-diamidino-2-phenylindole (DAPI)</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:165px;">D9542-1MG</td>
			<td style="width:171px;">Dissolved in DMSO</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Anti-fading mounting medium</td>
			<td style="width:175px;">Vector Laboratories</td>
			<td style="width:165px;">H-1000</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Anti-fading mounting medium with DAPI</td>
			<td style="width:175px;">Vector Laboratories</td>
			<td style="width:165px;">H-1200</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:392px;">Nunc Lab-Tek II 4-well chamber slide</td>
			<td style="width:175px;">Thermo Fisher Scientific</td>
			<td style="width:165px;">154526</td>
			<td></td>
		</tr>
	</tbody>
</table>

retinal cryo-sections, whole-mounts, and hypotonic isolated vasculature preparations for immunohistochemical visualization of microvascular pericytes

Retinal pericytes play an important role in many diseases of the eye. Immunohistochemical staining techniques of retinal vessels and microvascular pericytes are central to ophthalmological research. It is vital to choose an appropriate method of visualizing the microvascular pericytes. We describe retinal microvascular pericyte immunohistochemical staining in cryo-sections, whole-mounts, and hypotonic isolated vasculature using antibodies for platelet-derived growth factor receptor &#946; (PDGFR&#946;) and nerve/glial antigen 2 (NG2). This allows us to highlight advantages and shortcomings of each of the three tissue preparations for the visualization of the retinal microvascular pericytes. Cryo-sections provide transsectional visualization of all retinal layers but contain only a few occasional transverse cuts of the microvasculature. Whole-mount provides an overview of the entire retinal vasculature, but visualization of the microvasculature can be troublesome. Hypotonic isolation provides a method to visualize the entire retinal vasculature by the removal of neuronal cells, but this makes the tissue very fragile.

The successful protocols provide three different retinal preparations for visualizing microvascular pericytes. Each of these methods uses the PDGFR&#946; and NG2 immunoreactivity co-localization and the unique position of the pericytes that wrap around the capillary endothelium foridentification.
With cryo-sections, the neuronal layers can be identified by the fluorescent density of DAPI-labelled nuclei and the inner and deep ca...

Watch this Scientific Journal Video about Retinal Cryo-sections, Whole-Mounts, and Hypotonic Isolated Vasculature Preparations for Immunohistochemical Visualization of Microvascular Pericytes at JoVE.

Retinal Cryo-sections, Whole-Mounts, and Hypotonic Isolated Vasculature Preparations for Immunohistochemical Visualization of Microvascular Pericytes

We demonstrate three different tissue preparation techniques for immunohistochemical visualization of rat retinal microvascular pericytes, i.e., cryo-sections, whole-mounts, and hypotonic isolation of the vascular network.

Retinal pericytes play an important role in many diseases of the eye. Immunohistochemical staining techniques of retinal vessels and microvascular ...

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Research

JoVE Journal

Biology

9.9K Views.  Glostrup Research Institute. We demonstrate three different tissue preparation techniques for immunohistochemical visualization of rat retinal microvascular pericytes, i.e., cryo-sections, whole-mounts, and hypotonic isolation of the vascular network.

Video: Retinal Cryo-sections, Whole-Mounts, and Hypotonic Isolated Vasculature Preparations for Immunohistochemical Visualization of Microvascular Pericytes

Thin Sectioning of Slice Preparations for Immunohistochemistry

Many investigations in neuroscience, as well as other disciplines, involve studying small, yet macroscopic pieces or sections of tissue that have been preserved, freshly removed, or excised but kept viable, as in slice preparations of brain tissue.  Subsequent microscopic studies of this material can be challenging, as the tissue samples may be difficult to handle.  Demonstrated here is a method for obtaining thin cryostat sections of tissue with a thickness that may range from 0.2-5.0 mm.  We routinely cut 400 micron thick Vibratome brain slices serially into 5-10 micron coronal cryostat sections.  The slices are typically first used for electrophysiology experiments and then require microscopic analysis of the cytoarchitecture of the region from which the recordings were observed.  We have constructed a simple device that allows controlled and reproducible preparation and positioning of the tissue slice.  This device consists of a cylinder 5 cm in length with a diameter of 1.2 cm, which serves as a freezing stage for the slice.  A ring snugly slides over the cylinder providing walls around the slice allowing the tissue to be immersed in freezing compound (e.g., OCT).  This is then quickly frozen with crushed dry ice and the resulting  wafer  can be position easily for cryostat sectioning.  Thin sections can be thaw-mounted onto coated slides to allow further studies to be performed, such as various staining methods, in situ hybridization, or immunohistochemistry, as demonstrated here.

The present method allows reproducible cryostat sectioning of small, difficult-to-manage, tissue pieces, such as biopsies and brain slices. We utilize a simple aluminum freezing stage to facilitate handling of tissue and a standard cryostat to routinely produce 5-10 micron serial sections from 400 micron thick brain slices.

Many investigations in neuroscience, as well as other disciplines, involve studying small, yet macroscopic pieces or sections of tissue that have been ...

Using Laser Tweezers For Manipulating Isolated Neurons In Vitro

In this paper and video, we describe the protocols used in our laboratory to study the targeting preferences of regenerating cell processes of adult retinal neurons in vitro. Procedures for preparing retinal cell cultures start with the dissection, digestion and trituration of the retina, and end with the plating of isolated retinal cells on dishes made especially for use with laser tweezers. These dishes are divided into a cell adhesive half and a cell repellant half. The cell adhesive side is coated with a layer of Sal-1 antibodies, which provide a substrate upon which our cells grow. Other adhesive substrates could be used for other cell types. The cell repellant side is coated with a thin layer of poly-HEMA. The cells plated on the poly-HEMA side of the dish are trapped with the laser tweezers, transported and then placed adjacent to a cell on the Sal-1 side to create a pair. Formation of cell groups of any size should be possible with this technique. "Laser-tweezers-controlled micromanipulation" means that the investigator can choose which cells to move, and the desired distance between the cells can be standardized. Because the laser beam goes through transparent surfaces of the culture dish, cell selection and placement are done in an enclosed, sterile environment. Cells can be monitored by video time-lapse and used with any cell biological technique required. This technique may help investigations of cell-cell interactions.

This video describes the manipulation of cultured neurons using laser tweezers in vitro.

In this paper and video, we describe the protocols used in our laboratory to study the targeting preferences of regenerating cell processes of adult ...

Whole-mount Immunohistochemical Analysis for Embryonic Limb Skin Vasculature: a Model System to Study Vascular Branching Morphogenesis in Embryo

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching morphogenesis. We have developed the limb skin vasculature model to study vascular development in which a pre-existing primitive capillary plexus is reorganized into a hierarchically branched vascular network. Whole-mount confocal microscopy with multiple labelling allows for robust imaging of intact blood vessels as well as their cellular components including endothelial cells, pericytes and smooth muscle cells, using specific fluorescent markers. Advances in this limb skin vasculature model with genetic studies have improved understanding molecular mechanisms of vascular development and patterning. The limb skin vasculature model has been used to study how peripheral nerves provide a spatial template for the differentiation and patterning of arteries. This video article describes a simple and robust protocol to stain intact blood vessels with vascular specific antibodies and fluorescent secondary antibodies, which is applicable for vascularized embryonic organs where we are able to follow the process of vascular development.

We introduce a whole-mount immunohistochemistry and laser scanning confocal microscopy with multiple labelling for analyzing intricate vascular network formation in mouse embryonic limb skin.

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching ...

The ex vivo Isolated Skeletal Microvessel Preparation for Investigation of Vascular Reactivity

The isolated microvessel preparation is an ex vivo preparation that allows for examination of the different contributions of factors that control vessel diameter, and thus, perfusion resistance1-5. This is a classic experimental preparation that was, in large measure, initially described by Uchida et al.15 several decades ago. This initial description provided the basis for the techniques that was extensively modified and enhanced, primarily in the laboratory of Dr. Brian Duling at the University of Virginia6-8, and we present a current approach in the following pages. This preparation will specifically refer to the gracilis arteriole in a rat as the microvessel of choice, but the basic preparation can readily be applied to vessels isolated from nearly any other tissue or organ across species9-13. Mechanical (i.e., dimensional) changes in the isolated microvessels can easily be evaluated in response to a broad array of physiological (e.g., hypoxia, intravascular pressure, or shear) or pharmacological challenges, and can provide insight into mechanistic elements comprising integrated responses in an intact, although ex vivo, tissue. The significance of this method is that it allows for facile manipulation of the influences on the integrated regulation of microvessel diameter, while also allowing for the control of many of the contributions from other sources, including intravascular pressure (myogenic), autonomic innervation, hemodynamic (e.g., shear stress), endothelial dependent or independent stimuli, hormonal, and parenchymal influences, to provide a partial list. Under appropriate experimental conditions and with appropriate goals, this can serve as an advantage over in vivo or in situ tissue/organ preparations, which do not readily allow for the facile control of broader systemic variables. 
The major limitation of this preparation is essentially the consequence of its strengths. By definition, the behavior of these vessels is being studied under conditions where many of the most significant contributors to the regulation of vascular resistance have been removed, including neural, humoral, metabolic, etc. As such, the investigator is cautioned to avoid over-interpretation and extrapolation of the data that are collected utilizing this preparation. The other significant area of concern with regard to this preparation is that it can be very easy to damage cellular components such as the endothelial lining or the vascular smooth muscle, such that variable source of error can be introduced. It is strongly recommended that the individual investigator utilize appropriate measurements to ensure the quality of the preparation, both at the initiation of the experiment and periodically throughout the course of a protocol.

The ex vivo Isolated Skeletal Microvessel Preparation for Investigation of Vascular Reactivity

An ex vivo preparation is described for isolation of the largest gracilis muscle resistance arterioles for interrogation of both vascular responses to vasoactive stimuli and the assessment of basic structural properties via passive wall mechanics.

The isolated  microvessel preparation is an ex vivo preparation that allows for examination of the different contributions of  factors that control vessel ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Spatiotemporal Mapping of Motility in Ex Vivo Preparations of the Intestines

Multiple approaches have been used to record and evaluate gastrointestinal motility including: recording changes in muscle tension, intraluminal pressure, and membrane potential. All of these approaches depend on measurement of activity at one or multiple locations along the gut simultaneously which are then interpreted to provide a sense of overall motility patterns. Recently, the development of video recording and spatiotemporal mapping (STmap) techniques have made it possible to observe and analyze complex patterns in ex vivo whole segments of colon and intestine. Once recorded and digitized, video records can be converted to STmaps in which the luminal diameter is converted to grayscale or color [called diameter maps (Dmaps)]. STmaps can provide data on motility direction (i.e., stationary, peristaltic, antiperistaltic), velocity, duration, frequency and strength of contractile motility patterns. Advantages of this approach include: analysis of interaction or simultaneous development of different motility patterns in different regions of the same segment, visualization of motility pattern changes over time, and analysis of how activity in one region influences activity in another region. Video recordings can be replayed with different timescales and analysis parameters so that separate STmaps and motility patterns can be analyzed in more detail. This protocol specifically details the effects of intraluminal fluid distension and intraluminal stimuli that affect motility generation. The use of luminal receptor agonists and antagonists provides mechanistic information on how specific patterns are initiated and how one pattern can be converted into another pattern. The technique is limited by the ability to only measure motility that causes changes in luminal diameter, without providing data on intraluminal pressure changes or muscle tension, and by the generation of artifacts based upon experimental setup; although, analysis methods can account for these issues. When compared to previous techniques the video recording and STmap approach provides a more comprehensive understanding of gastrointestinal motility.

Spatiotemporal Mapping of Motility in Ex Vivo Preparations of the Intestines

Recently available video recording and spatiotemporal mapping (STmap) techniques make it possible to visualize and quantify both propagating and mixing patterns of intestinal motility. The goal of this protocol is to explain the generation and analysis of STmaps using the GastroIntestinal Motility Monitoring (GIMM) system.

Multiple approaches have been used to record and evaluate gastrointestinal motility including: recording changes in muscle tension, intraluminal pressure, ...

Improved Swiss-rolling Technique for Intestinal Tissue Preparation for Immunohistochemical and Immunofluorescent Analyses

Understanding the role of factors that regulate intestinal epithelial homeostasis and response to injury and regeneration is important. The current literature describes several different methodological approaches to obtain images of intestinal tissues for data validation. In this paper, we delineate a common protocol relating to the derivation and processing of mouse intestinal tissues. Proper fixation of intestinal tissues and Swiss-roll techniques that enhance intestinal epithelial morphology are discussed. Postresection processing and reorientation of embedded intestinal tissues are critical in obtaining paraffin-embedded blocks that display intact intestinal structural features after sectioning. The Swiss-rolling technique helps in histological assessment of the complete intestinal or colonic sections examined. An ability to differentiate intestinal structural features can be vital in quantitative measurements of intestinal inflammation and tumorigenesis along the entire length. Finally, paraffin-embedded sections are ideal for robust processing using both immunohistochemical and immunofluorescent detection methods. Nonfluorescent immunohistochemical sections provide a vibrant image of the tissue detailing different cellular structural features but do not provide flexibility for intracellular co-localization experiments. Multiple fluorescent channels can be appropriately utilized with immunofluorescent detection for co-localization experiments, lending support to mechanistic studies.

Accurate identification and location of epithelial cells along the intestinal mucosal lining are essential to define different cell lineages. Proper imaging of intestinal tissues is crucial for identification of protein expression patterns with maximum resolution. This study aims to delineate the optimal methods and conditions for processing mouse intestinal tissues.

Understanding the role of factors that regulate intestinal epithelial homeostasis and response to injury and regeneration is important. The current ...

Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated with specialization, development and increase in cellular size. In plants, endoreduplication seems to facilitate the growth and expansion of certain tissues and organs. Among them is the tuber of potato (Solanum tuberosum), which undergoes considerable cellular expansion in fulfilling its function of carbohydrate storage. Thus, endoreduplication may play an important role in how tubers are able to accommodate this abundance of carbon. However, the cellular debris resulting from crude nuclear isolation methods of tubers, methods that can be used effectively with leaves, precludes the estimation of the tuber endoreduplication index (EI). This article presents a technique for assessing tuber endoreduplication through the isolation of protoplasts while demonstrating representative results obtained from different genotypes and compartmentalized tuber tissues. The major limitations of the protocol are the time and reagent costs required for sample preparation as well as relatively short lifespan of samples after lysis of protoplasts. While the protocol is sensitive to technical variation, it represents an improvement over traditional methods of nuclear isolation from these large specialized cells. Possibilities for improvements to the protocol such as recycling enzyme, the use of fixatives, and other alterations are proposed.

The protocol described herein is a method for measuring endoreduplication within tubers of potato (Solanum tuberosum). It includes plasmolysis and protoplast extraction steps to decrease the noise and debris in downstream flow cytometric analysis.

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated ...