Many thanks to CSIC and PEDECIBA for the financial support given to this work; to Natalia Rosano for her manuscript corrections; to Marcelo Casacuberta that makes the video and to Nicol&#225;s Bolatto for lending his voice for it.

All animal procedures were performed according to the guidelines of the National Law N&#176; 18611 for Care of Animals Used for Experimental Purposes. The protocol was approved by the Institutional Ethical Committee (CEUA IIBCE, Protocol Number 004/09/2015).
1. Muscle dissection (Day 1)
NOTE: Before starting, make 40 mL of 0.5% paraformaldehyde (PFA), pH 7.4 in Dulbecco&#180;s phosphate saline (DPBS). Optionally, make 20 mL of 4% PFA. Prepare 5 mL aliquots and freeze at -20 &#176;C. On the day of dissection, defrost a 4% aliquot and add 35 mL of DPBS to obtain 40 mL of 0.5% PFA.
<ol>
	<li>Isolation of EDL (fast-twitch muscle)
	<ol>
		<li>Euthanize a rat. Place the animal with the abdomen facing upwards. Make an initial incision using a surgical blade between the toes towards the hind limb. 
		NOTE: In this case, an intraperitoneal injection of 90:10 mg/Kg ketamine:xylazine was given to euthanize the rat, but other options of anesthesia are also allowed.</li>
		<li>Peel off the skin, pulling it upwards until the rat knee is exposed.</li>
		<li>To find the EDL, follow the foot tendons up to the annular ligament. This ligament circles two tendons. Cut the ligament between the two tendons with uniband scissors (or similar). Identify the EDL tendon by lifting both and choose the one that makes the toes move upwards.</li>
		<li>Cut the tendon with uniband scissors. Then, while holding the tendon with fine biological tweezers, begin to slowly separate the EDL muscle from the tibialis anterior, the peroneus brevis, and peroneus longus40 (see Figure 1A). 
		NOTE: Ensure that the muscle is separated without any damage. Do this by cutting the rest of the lateral muscles to open a path between them (the EDL does not have a superficial location) while holding and lifting the EDL muscle.</li>
		<li>To completely isolate the muscle, cut the tendon attached to the rat knee with uniband scissors.</li>
		<li>Immerse the dissected muscle in 5 mL of 0.5% PFA and leave for 24 h at 4 &#176;C. Optionally, staple the muscle in a piece of cardboard and turn it with the muscle facing down. Then immerse it completely in 8 mL of the fixative solution. This step helps in maintaining the muscle elongated during the fixation process.</li>
	</ol>
	</li>
	<li>Isolation of soleus (slow-twitch muscle)
	<ol>
		<li>Flip the animal (the abdomen is now facing downwards). Through the skin, cut the calcaneal tendon using a surgical blade.</li>
		<li>With the help of the biological tweezers and uniband scissors, separate the gastrocnemius muscle from the bones, creating a &#34;muscular lid&#34;. The soleus will be on the internal side of the muscular lid. It can be identified because it is red in color and has a flat morphology.</li>
		<li>With a pair of biological tweezers, reach and lift the soleus tendon that lies above the gastrocnemius (see Figure 1B).</li>
		<li>Cut the tendon with uniband scissors. Lift the whole muscle while cutting some of the weak attachment points (i.e., neurovascular elements). Finally, to completely free the soleus muscle, cut the soleus fascicle that forms the calcaneal tendon with uniband scissors.</li>
		<li>Repeat step 1.1.6 to fix dissected soleus muscle.</li>
	</ol>
	</li>
</ol>
2. Teased fiber preparation (Day 2)
<ol>
	<li>After 24 h of fixation, rinse the muscles with 6 mL of DPBS solution 3x for 10 min each before isolating the muscle fibers.</li>
	<li>To isolate the fibers, put a muscle on a microscope slide. Using a stereomicroscope, gently hold one of the tendons with one pair of biological tweezers. Then, with the other biological tweezers, begin to pinch the tendon slowly to separate the muscle fibers. After that slowly pull the pinched tissue&#160;upwards towards the opposite muscle tendon. It will be necessary to repeat this action several times until obtaining multiple small, isolated bundles.</li>
	<li>Place them carefully on a pre-treated slide (silanized). It is necessary to keep all the fibers in order so that they do not overlap. At this point, air-dry the fibers for 24 h.</li>
</ol>
3. Immunofluorescence
<ol>
	<li>Primary antibody incubation (Day 3) 
	NOTE: All steps were performed at room temperature, except if otherwise stated. The most efficient way to remove the different solutions without detaching fibers from the slide is to use an insulin syringe.
	<ol>
		<li>To start the immunostaining process, encircle the dried isolated muscle fibers with a pap pen to create a waterproof barrier.</li>
		<li>To hydrate the muscle fibers, add 200 &#181;L of distilled water for 5 min, and then change the water to 200 &#181;L of DPBS for the next 5 min incubation. At this point, start the immunofluorescence labeling.</li>
		<li>Incubate fibers with 300 &#181;L of a blocking buffer (BB) containing 50 mM glycine, 1% BSA, and 1% Triton X-100 for 30 min.</li>
		<li>Replace BB with the primary antibody at a concentration suggested by manufacturers. Use the BB to dilute the antibody. This experiment used 400 &#181;L of SMI 32 or SMI 31 that recognized non-phosphorylated (Nf) or phosphorylated (Phos-Nf) neurofilament H&#160;at 1/800 dilution to detect the NMJ presynaptic component. 
		NOTE: When deciding to recognize the whole presynaptic element instead of evaluating neurofilament phosphorylation, choose antibodies against synapsin, synaptophysin or SV2a that were previously successfully employed.</li>
		<li>Incubate the fibers with the primary antibody dilution at 4 &#176;C overnight (optionally, incubation can be performed at 37 &#176;C for 1 h). In both cases, it will be important to perform the incubation using a humid chamber.</li>
	</ol>
	</li>
	<li>Immunofluorescence: Secondary antibody incubation (Day 4)
	<ol>
		<li>Remove the primary antibody and rinse the fibers 3x for 10 min each with 500 &#181;L of DPBS and the last time with BB.</li>
		<li>Remove this last wash and add the fluorescently labeled secondary antibody diluted in BB. For this experiment, 500 &#181;L of goat anti-mouse Alexa Fluor 488 at 1/1000 dilution in BB was used. To view the postsynaptic element of the NMJs, 1:300 &#945;-Bungarotoxin (Btx)&#160;biotin-XX conjugate was added to the secondary antibody dilution. 
		&#8203;NOTE: The addition of Btx-biotin XX&#160;allows better signal-to-noise images once detected with streptavidin that amplifies the signal. In addition, streptavidin conjugation to different fluorophores increased color combinations in co-labeling assays. Alternatively, fluorescently labeled Btx allows obtaining images of similar quality.</li>
		<li>Incubate the secondary antibody and the Btx for 2 h at room temperature or 1 h at 37 &#176;C protected from light.</li>
		<li>Remove the secondary antibody and the Btx. Wash 3x for 10 min each with 500 &#181;L of DPBS and the last rinse with BB.</li>
		<li>Incubate with 500 &#181;L of Streptavidin Alexa Fluor 555 at 1/1000 dilution with BB for 1 h at room temperature. At this point, if desired, add a probe to counterstain the nuclei, such as diaminophenylindole at a final concentration of 1 &#181;g/mL.</li>
		<li>Remove the incubation solution and wash fibers 3x for 10 min each with 500 &#181;L of DPBS. Then, mount the fibers.</li>
	</ol>
	</li>
	<li>Mounting
	<ol>
		<li>Place 4 dots of transparent nail polish on the microscope slide as if they were the vertices of a square. Let them dry 1-2 min. These will be the points where the coverslips will rest, helping to avoid tissue crushing.</li>
		<li>Remove the last DPBS wash and add enough mounting media (~ 200 &#181;L) to cover the fibers; avoid drying. 
		NOTE: To prepare 10 mL of the mounting media mix, use Tris-HCl 1,5 M pH 8,8: glycerol in a 4:1 (v:v).</li>
		<li>Use a 200 &#181;L pipette tip to carefully spread the mounting media over the fibers to ensure a complete immersion of all of them. Before placing the cover glass, remove air bubbles.</li>
		<li>Place the coverslip onto the samples trying to prevent the generation of air bubbles. This movement can be done with the aid of forceps.</li>
	</ol>
	</li>
</ol>
4. Microscopy and morphometric analysis
<ol>
	<li>Image the teased fiber preparations using laser confocal microscopy.</li>
	<li>Take a series of optical sections (30 &#181;m Z stacks) across the entire synapse using a one-micrometer interval. To set the stack, use the Btx signal. Image at least 15-20 NMJs for each muscle condition with a 2048 px x 2048 px resolution. This method was chosen to obtain images with sufficient optical quality and resolution for the morphometric analysis.</li>
	<li>To perform the measurements, use the projection images of stacks with any analysis software. 
	NOTE: Although the explained morphometric analysis was done manually, there are two recently developed Image J-based tools for studying many different aspects of pre and postsynaptic NMJ morphology33,34.</li>
	<li>To obtain the Total NMJ area values, select the external border (external outline of the stained area, see Figure 2) of the endplate using the Photoshop&#160;Magnetic lasso tool and determine the number of selected pixels in the Histogram window. Then, the pixel areas can be converted to square micrometers using the scale specified by the confocal software.</li>
	<li>With the same lasso tool, select the internal border (outlines of the stained area within the endplate, see Figure 2) and determine the non-stained area in the whole structure (or empty area) as detailed above (step 4.4.). The Postsynaptic Area values will be obtained after subtracting the total area from the empty area of each NMJ.</li>
	<li>Obtain the Presynaptic Area, that is defined as the area with anti-neurofilament immunofluorescence, in a similar way that the Total NMJ area. 
	NOTE: All these parameters were used to determine the Post-Synaptic Density index (Postsynaptic Area/Total Area) and the NMJ Coverage (Presynaptic Area/Postsynaptic Area). A higher Postsynaptic Density index implies a denser -or more compact- endplate, whereas a smaller NMJ Coverage reflects a much poorer interaction between nerve terminal and endplate (both compatible with a denervation process). In the case shown here, since phosphorylated and non-phosphorylated forms of neurofilaments were detected at the nerve terminal level, the phosphorylated/non-phosphorylated presynaptic signal ratio and the phosphorylated/non-phosphorylated coverage ratio were calculated.</li>
</ol>

The authors have nothing to disclose.

In this article, we present a detailed protocol for the dissection of two rat skeletal muscles (one slow-twitch and the other fast-twitch), fiber muscle isolation and immunofluorescence detection of pre and postsynaptic markers to quantitatively assess&#160;NMJ changes as well as assembly/disassembly processes. This kind of protocol can be useful in rodent models41,42 to evaluate NMJ during physiological or pathological processes as exemplified here in a model of...

The mammal neuromuscular junction (NMJ) is a large cholinergic tripartite synapse made up of the motor neuron nerve ending, the postsynaptic membrane on the skeletal muscle fiber, and the terminal Schwann cells1,2,3. This synapse exhibits high morphological and functional plasticity4,5,6,7,8, even during adulthood when NMJs can undergo dynamic structural modifications. For example, some researchers have shown that motor nerve endings continually change their shape at the micrometer scale9. It has also been reported that the morphology of the NMJ responds to functional requirements, altered use, aging, exercise, or variations in locomotor activity4,10,11,12,13,14,15. Thus, training and lack of use represent&#160;essential stimulus to modify some characteristics of the NMJ, such as its size, length, dispersion of synaptic vesicles and receptors, as well as nerve terminal branching14,16,17,18,19,20.
Furthermore, it has been shown that any structural change or degeneration of this vital junction could result in motor neuron cell death and muscle atrophy21. It is also thought that altered communication among nerves and muscles could be responsible for the physiological age-related NMJ changes and possibly for its destruction in pathological states. Neuromuscular junction dismantling plays a crucial role in the onset of Amyotrophic Lateral Sclerosis (ALS), a neurodegenerative disease that constitutes one of the best examples of impaired muscle-nerve interplay3. Despite the numerous studies conducted on motor neuron dysfunction, it is still debated whether the deterioration observed in ALS occurs due to the direct damage into the motor neuron and then extends to the cortico-spinal projections22; or if it should be considered as a distal axonopathy where degeneration begins in the nerve endings and progresses toward the motor neuron somas23,24. Given the complexity of ALS pathology, it is logical to consider that a mix of independent processes occurs. As NMJ is the central player of the physiopathological interplay between muscle and nerve, its destabilization represents a pivotal point in the origin of the disease that is relevant to be analyzed.
The mammal neuromuscular system is functionally organized into discrete motor units, consisting of a motor neuron and the muscle fibers that are exclusively innervated by its nerve terminal. Each motor unit has fibers with similar or identical structural and functional properties25. Motor neuron selective recruitment allows optimizing muscle response to functional demands. Now it is clear that mammalian skeletal muscles are composed of four different fiber types.&#160;Some muscles are named according to the characteristics of their most abundant fiber type. For example, the soleus (a posterior muscle of the hind limb involved in the maintenance of the body posture) bears a majority of slow-twitch units (type 1) and is recognized as a slow muscle. Instead, extensor digitorum longus&#160;(EDL) is essentially composed of units with similar fast-twitch properties (type 2 fibers) and is known as a fast muscle specialized for phasic movements needed for locomotion. In other words, although adult muscles are plastic in nature due to the hormonal and neural influences, its fiber composition determines the capacity to perform different activities, as seen in soleus that experiences continuous low-intensity activity and EDL that exhibits a more rapid single twitch. Other features that are variable among different types of muscle fibers are related to their structure (mitochondrial content, extension of sarcoplasmic reticulum, thickness of the Z line), myosin ATPase content, and myosin heavy chain composition26,27,28,29.
For rodent NMJs, there are significant differences among muscles28,29. Morphometric analyses performed in soleus and EDL from rats revealed a positive correlation between the synaptic area and fiber diameter (i.e., the synaptic area in soleus slow fibers is greater than in EDL fast fibers) but the ratio between NMJ area and fiber size is similar in both muscles30,31. Also, in relation to the nerve terminals, the endplate absolute areas in type 1 fibers were lower than in type 2 fibers, whereas the normalization by fiber diameter made areas of nerve terminals in type 1 fibers the largest32.
However, very few studies focus on morphometric analysis to show the evidence of changes in some of the NMJ components33,34. Thus, due to the relevance of the NMJ in the function of the organism, whose morphology and physiology are altered in various pathologies, it is important to optimize dissection protocols of different types of muscles with quality enough to allow the visualization of the whole NMJ structure. It is also necessary to evaluate the occurrence of pre or postsynaptic changes in different experimental situations or conditions such as aging or exercise35,36,37,38. In addition, it can be helpful to evidence more subtle alterations in NMJ components such as altered neurofilament phosphorylation in the terminal nerve endings as reported in ALS39.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Stereomicroscope with cool light illumination</td>
			<td>Nikon</td>
			<td>SMZ-10A</td>
			<td></td>
		</tr>
		<tr>
			<td>Rocking platform</td>
			<td>Biometra (WT 16)</td>
			<td>042-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover glasses (24 x 32 mm)</td>
			<td>Deltalab</td>
			<td>D102432</td>
			<td></td>
		</tr>
		<tr>
			<td>Premium (Plus) microscope slides</td>
			<td>PORLAB</td>
			<td>PC-201-16</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers</td>
			<td>F.S.T</td>
			<td>11253-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Uniband LA-4C Scissors 125mm</td>
			<td>E.M.S</td>
			<td>77910-26</td>
			<td></td>
		</tr>
		<tr>
			<td>Disponsable surgical blades #10</td>
			<td>Sakira Medical</td>
			<td>1567</td>
			<td></td>
		</tr>
		<tr>
			<td>Disponsable sterile syringe (1 ml)</td>
			<td>Sakira Medical</td>
			<td>1569</td>
			<td></td>
		</tr>
		<tr>
			<td>Super PAP pen</td>
			<td>E.M.S</td>
			<td>71310</td>
			<td></td>
		</tr>
		<tr>
			<td>100 &#956;l or 200 &#956;l pipette</td>
			<td>Finnpipette</td>
			<td>9400130</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM 800 - AiryScan</td>
			<td></td>
		</tr>
		<tr>
			<td>NTac:SD-TgN(SOD1G93A)L26H rats</td>
			<td>Taconic</td>
			<td>2148-M</td>
			<td></td>
		</tr>
		<tr>
			<td>1X PBS (Dulbecco)</td>
			<td>Gibco</td>
			<td>21600-010</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma</td>
			<td>158127</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>T8787</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycine</td>
			<td>Amresco</td>
			<td>167</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Bio Basic INC.</td>
			<td>9048-46-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Mallinckrodt</td>
			<td>5092</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris</td>
			<td>Amresco</td>
			<td>497</td>
			<td></td>
		</tr>
		<tr>
			<td>Purified anti-Neurofilament H (NF-H), Phosphorylated Antibody</td>
			<td>BioLegend</td>
			<td>801601</td>
			<td>Previously Covance # SMI 31P</td>
		</tr>
		<tr>
			<td>Purified anti-Neurofilament H (NF-H), Nonphosphorylated Antibody</td>
			<td>BioLegend</td>
			<td>801701</td>
			<td>Previously Covance # SMI-32P</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 goat anti-Mouse IgG (H+L)</td>
			<td>Thermo Scientific</td>
			<td>A11029</td>
			<td></td>
		</tr>
		<tr>
			<td>&#945;-Bungarotoxin, biotin-XX conjugate</td>
			<td>Invitrogen</td>
			<td>B1196</td>
			<td></td>
		</tr>
		<tr>
			<td>Streptavidin, Alexa Fluor 555 conjugate</td>
			<td>Invitrogen</td>
			<td>S32355</td>
			<td></td>
		</tr>
		<tr>
			<td>Diaminophenylindole (DAPI)</td>
			<td>Sigma</td>
			<td>D8417</td>
			<td></td>
		</tr>
	</tbody>
</table>

dissection of single skeletal muscle fibers for immunofluorescent and morphometric analyses of whole-mount neuromuscular junctions

The neuromuscular junction (NMJ) is a specialized point of contact between the motor nerve and the skeletal muscle. This peripheral synapse exhibits high morphological and functional plasticity. In numerous nervous system disorders, NMJ is an early pathological target resulting in neurotransmission failure, weakness, atrophy, and even in muscle fiber death. Due to its relevance, the possibility to quantitatively assess certain aspects of the relationship between NMJ components can help to understand the processes associated with its assembly/disassembly. The first obstacle when working with muscles is to gain the technical expertise to quickly identify and dissect without damaging their fibers. The second challenge is to utilize high-quality detection methods to obtain NMJ images that can be used to perform quantitative analysis. This article presents a step-by-step protocol for dissecting extensor digitorum longus and soleus muscles from rats. It also explains the use of immunofluorescence to visualize pre and postsynaptic elements of whole-mount NMJs. Results obtained demonstrate that this technique can be used to establish the microscopic anatomy of the synapsis and identify subtle changes in the status of some of its components under physiological or pathological conditions.

This protocol offers a straightforward method to isolate and immunostain muscle fibers from two different types of muscles (fast- and slow-twitch muscles, see Figure 1). Using the correct markers and / or probes, NMJ components can be detected and evaluated since a quantitative point of view to assess some of the morphological changes that can occur as consequence of illness progression or a specific drug treatment. In the present study, presynaptic and postsynaptic components of the NMJ wer...

Watch this Scientific Journal Video about Dissection of Single Skeletal Muscle Fibers for Immunofluorescent and Morphometric Analyses of Whole-Mount Neuromuscular Junctions at JoVE.com

Dissection of Single Skeletal Muscle Fibers for Immunofluorescent and Morphometric Analyses of Whole-Mount Neuromuscular Junctions

The ability to accurately detect neuromuscular junction components is crucial in evaluating modifications in its architecture because of pathological or developmental processes. Here we present a complete description of a straightforward method to obtain high-quality images of whole-mount neuromuscular junctions that can be used to perform quantitative measurements.

The neuromuscular junction (NMJ) is a specialized point of contact between the motor nerve and the skeletal muscle. This peripheral synapse exhibits high ...

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8.1K Views.  Universidad de la República. The ability to accurately detect neuromuscular junction components is crucial in evaluating modifications in its architecture because of pathological or developmental processes. Here we present a complete description of a straightforward method to obtain high-quality images of whole-mount neuromuscular junctions that can be used to perform quantitative measurements.

Video: Dissection of Single Skeletal Muscle Fibers for Immunofluorescent and Morphometric Analyses of Whole-Mount Neuromuscular Junctions

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 ...

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Morphometric Analyses of Retinal Sections

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the retinal ganglion cell layer (RGCL) in rat models with N-methyl-D-aspartate (NMDA)&#x2013;induced excitotoxicity1, retinal ischemia-reperfusion injury2 and glaucoma3. Reduction of INL and inner plexiform layer (IPL) thicknesses were reversed with citicoline treatment in rats' eyes subjected to kainic acid-mediated glutamate excitotoxicity4. Alteration of RGC density and soma sizes were observed with different drug treatments in eyes with elevated intraocular pressure3,5,6. Therefore, having objective methods of analyzing the retinal morphometries may be of great significance in evaluating retinal pathologies and the effectiveness of therapeutic strategies.
The retinal structure is multi-layers and several different kinds of neurons exist in the retina. The morphometric parameters of retina such as cell number, cell size and thickness of different layers are more complex than the cell culture system. Early on, these parameters can be detected using other commercial imaging software. The values are normally of relative value, and changing to the precise value may need further accurate calculation. Also, the tracing of the cell size and morphology may not be accurate and sensitive enough for statistic analysis, especially in the chronic glaucoma model. The measurements used in this protocol provided a more precise and easy way. And the absolute length of the line and size of the cell can be reported directly and easy to be copied to other files. For example, we traced the margin of the inner and outer most nuclei in the INL and formed a line then using the software to draw a 90 degree angle to measure the thickness. While without the help of the software, the line maybe oblique and the changing of retinal thickness may not be repeatable among individual observers. In addition, the number and density of RGCs can also be quantified. This protocol successfully decreases the variability in quantitating features of the retina, increases the sensitivity in detecting minimal changes.
 	This video will demonstrate three types of morphometric analyses of the retinal sections. They include measuring the INL thickness, quantifying the number of RGCs and measuring the sizes of RGCs in absolute value. These three analyses are carried out with Stereo Investigator (MBF Bioscience &mdash; MicroBrightField, Inc.). The technique can offer a simple but scientific platform for morphometric analyses.

This video demonstrates three types of morphometric analyses of the retina, which include measuring the inner nuclear layer thickness, quantifying the number of retinal ganglion cells (RGCs) and measuring the sizes of RGCs. The technique can offer a simple but scientific platform for morphometric analyses.

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the ...

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

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

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

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

Whole Mount Immunofluorescent Staining of the Neonatal Mouse Retina to Investigate Angiogenesis In vivo

Angiogenesis is the complex process of new blood vessel formation defined by the sprouting of new blood vessels from a pre-existing vessel network. Angiogenesis plays a key role not only in normal development of organs and tissues, but also in many diseases in which blood vessel formation is dysregulated, such as cancer, blindness and ischemic diseases. In adult life, blood vessels are generally quiescent so angiogenesis is an important target for novel drug development to try and regulate new vessel formation specifically in disease. In order to better understand angiogenesis and to develop appropriate strategies to regulate it, models are required that accurately reflect the different biological steps that are involved. The mouse neonatal retina provides an excellent model of angiogenesis because arteries, veins and capillaries develop to form a vascular plexus during the first week after birth. This model also has the advantage of having a two-dimensional (2D) structure making analysis straightforward compared with the complex 3D anatomy of other vascular networks. By analyzing the retinal vascular plexus at different times after birth, it is possible to observe the various stages of angiogenesis under the microscope. This article demonstrates a straightforward procedure for analyzing the vasculature of a mouse retina using fluorescent staining with isolectin and vascular specific antibodies.

Whole Mount Immunofluorescent Staining of the Neonatal Mouse Retina to Investigate Angiogenesis In vivo

The neonatal murine retina provides a well characterized physiological model of angiogenesis, which permits investigations of the roles of different genes or drugs that modulate angiogenesis in an in vivo context. Immunofluorescent staining to accurately visualize the vascular plexus is pivotal to the success of these types of studies.

Angiogenesis  is the complex process of new blood vessel formation defined by the sprouting  of new blood vessels from a pre-existing vessel network. ...

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

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

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

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

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

Whole Mount 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 ...

Isolation of Primary Murine Skeletal Muscle Microvascular Endothelial Cells

The endothelial cells of skeletal muscle capillaries (muscle microvascular endothelial cells, MMEC) build up the barrier between blood stream and skeletal muscles regulating the exchange of fluids and nutrients as well as the immune response against infectious agents by controlling immune cell migration. For these functions, MMEC form a functional &#34;myovascular unit&#34; (MVU), with further cell types, such as fibroblasts, pericytes and skeletal muscle cells. Consequently, a dysfunction of MMEC and therefore the MVU contributes to a vast variety of myopathies. However, regulatory mechanisms of MMEC in health and disease remain insufficiently understood and their elucidation precedes more specific treatments for myopathies. The isolation and in-depth investigation of primary MMEC functions in the context of the MVU might facilitate a better understanding of these processes.
This article provides a protocol to isolate primary murine MMEC of the skeletal muscle by mechanical and enzymatic dissociation including purification and culture maintenance steps.

Microvascular endothelial cells of skeletal muscles (MMEC) shape the inner wall of muscle capillaries and regulate both, exchange of fluids/molecules and migration of (immune) cells between muscle tissue and blood. Isolation of primary murine MMEC, as described here, enables comprehensive in vitro investigations of the &#34;myovascular unit&#34;.

The endothelial cells of skeletal muscle capillaries (muscle microvascular endothelial cells, MMEC) build up the barrier between blood stream and skeletal ...

Muscle Velocity Recovery Cycles to Examine Muscle Membrane Properties

Although conventional nerve conduction studies (NCS) and electromyography (EMG) are suitable for the diagnosis of neuromuscular disorders, they provide limited information about muscle fiber membrane properties and underlying disease mechanisms. Muscle velocity recovery cycles (MVRCs) illustrate how the velocity of a muscle action potential depends on the time after a preceding action potential. MVRCs are closely related to changes in membrane potential that follow an action potential, thereby providing information about muscle fiber membrane properties. MVRCs may be recorded quickly and easily by direct stimulation and recording from multi-fiber bundles in vivo. MVRCs have been helpful in understanding disease mechanisms in several neuromuscular disorders. Studies in patients with channelopathies have demonstrated the different effects of specific ion channel mutations on muscle excitability. MVRCs have been previously tested in patients with neurogenic muscles. In this prior study, muscle relative refraction period (MRRP) was prolonged, and early supernormality (ESN) and late supernormality (LSN) were reduced in patients compared to healthy controls. Thereby, MVRCs can provide in vivo evidence of membrane depolarization in intact human muscle fibers that underlie their reduced excitability. The protocol presented here describes how to record MVRCs and analyze the recordings. MVRCs can serve as a fast, simple, and useful method for revealing disease mechanisms across a broad range of neuromuscular disorders.

Presented here is a protocol for the recording of muscle velocity recovery cycles (MVRCs), a new method of examining muscle membrane properties. MVRCs enable in vivo assessment of muscle membrane potential and alterations in muscle ion channel function in relation to pathology, and it enables the demonstration of muscle depolarization in neurogenic muscles.

Although conventional nerve conduction studies (NCS) and electromyography (EMG) are suitable for the diagnosis of neuromuscular disorders, they provide ...