We are grateful to Austin Ferro and Juao-Guilherme Rosa in the Cvetanovic lab for their help in troubleshooting dissections and in RNA extraction and RTqPCR. This research is funded by M. Cvetanovic, R01 NS197387; HHS | National Institutes of Health (NIH).

1. Setup
<ol>
	<li>Gather necessary equipment including decapitation scissors, blunt forceps, dissection scissors, vascular scissors, microspatula, sagittal mouse brain matrix, razor blades, 200 &#181;L&#160;pipet tips, glass petri dish, glass slide, and ice bucket. Lay out all equipment on an absorbent pad.</li>
	<li>Place petri dish, glass plate, and brain matrix on ice.</li>
	<li>Using a razor blade at a perpendicular angle, trim about 5 mm off the tip of one 200 &#181;L&#160;pipet tip. This makes the size of the opening at the end of the tip about 1 mm wide. This should be sufficient to punch out the DCN. But this can also be adjusted as needed depending on the dissection. Have many tips ready for easy replacement and adjustment.</li>
	<li>Label 1.5 mL microfuge tubes with animal identification and cerebellar region (four tubes per animal).</li>
	<li>Fill cryosafe container with liquid nitrogen to flash freeze tissue after extraction.</li>
</ol>
2. Brain extraction and dissection
All experiments were conducted in accordance with the guidelines of the Animal Care Committees of the University of Minnesota.
<ol>
	<li>Euthanize the mouse using 5% CO2 exposure. Once breathing has ceased, perform cervical dislocation. Decapitate the mouse with the decapitation scissors and discard the carcass in the appropriate receptacle.</li>
	<li>Make an incision with a razor blade along the medial sagittal line of the head starting at the nose and continuing all the way back. Separate the skin, parting to either side of the midline. Use the razor blade to cut away the muscle on each side, cutting down past the ear canals.</li>
	<li>Using dissecting scissors, trim any spinal cord regions, up to where the brain stem meets the cerebellum, careful to not damage the cerebellum.</li>
	<li>Insert one of the vascular scissor blades into the space between the brainstem and vertebral column and cut toward the ear canal, lifting up on the scissors to cleanly cut the bone but limit damage to the tissue.</li>
	<li>Continue to cut along the edge of the skull up toward the olfactory bulbs, continuing to lift up while cutting to limit damage to the brain tissue.</li>
	<li>Using the blunt forceps, gently peel off the back of the skull uncovering the posterior region of the brain and cerebellum.</li>
	<li>Using the blunt forceps along the edge of the skull that was just cut, peel the rest of the skull up and over the brain. This step should remove the majority of the skull cap, revealing the brain.</li>
	<li>Trim the rest of the skull with the vascular scissors and blunt forceps, clearing most of the skull from the top of the brain.</li>
	<li>Using the microspatula, lift the brain slightly and scoop under and slide up to remove the olfactory bulbs from the remaining skull and disconnect optic tract fibers. Brain should come free easily at this point.</li>
	<li>Place the brain into the petri dish sitting on ice and remove any remaining skull or other debris.</li>
	<li>Using the microspatula, gently place the brain into the brain matrix with dorsal side up. Take time to make sure it is set level in the matrix, especially that the midline falls center in the matrix. This matrix is designed for adult mouse brain tissue, tissue from younger or diseased animals may rest lower in the matrix but it should still be possible to achieve reproducible results across samples.</li>
	<li>Place one razor blade along the sagittal midline, making sure that the blade pushes all the way to the bottom of the matrix (Figure 1B).</li>
	<li>Place another razor blade 1 mm to the side of the first blade (Figure 1B). Place two more blades 1 mm apart from each other. The end result should be three blades placed on one side of the brain, all 1mm apart. Do the same to the other side. In total, 7 blades will be placed 1mm apart (Figure 1C).</li>
	<li>Carefully, grab the front and back ends of the razor blades and lift straight up out of the matrix. The tissue on the outside of the razor blades can be discarded.</li>
	<li>Slowly, separate one razor blade at a time from the others, being careful not to damage the tissue sections.</li>
	<li>Carefully slide the tissue section off of the razor blade and onto the glass slide with the microspatula. In total there will be six sagittal brain sections (Figure 1D).</li>
	<li>The 4 most lateral sections will have DCN visible (Purple box, Figure 1D). To isolate the DCN, hold the trimmed 200 &#181;L&#160;pipet tip perpendicularly over the DCN and push down through the tissue firmly, rocking in all directions to fully dissect the DCN from surrounding tissue. Lift straight back up to cleanly remove the DCN, and visually confirm the presence of the tissue in the tip.</li>
	<li>Place one finger at the top of the tip and push down, causing the tissue to bulge out. Place the tip into correctly labeled microfuge tube and ensure that tissue punch is placed in the bottom of the tube. Repeat 2.17 for the remaining three sections, placing the DCN punches in the same tube. Place the tube into liquid nitrogen to flash freeze. Representation of size of each punch in Figure 1E.</li>
	<li>Sections that had DCN extracted qualify as cerebellar hemisphere. Push away the rest of the brain tissue around the cerebellum in these sections. Use blunt forceps and gently pick up these hemisphere cerebellar cortex sections and place into respective microfuge tube. Flash freeze.</li>
	<li>For the last two vermal sections (light blue box, Figure1D), push away surrounding brain tissue leaving only the cerebellum. Using a razor blade, make a cut separating the anterior lobules from the posterior lobules. Cut should be just after the formation of lobule 6 and should not include lobule 10 (Figure 1F).</li>
	<li>Using blunt forceps, carefully place the anterior cerebellar cortex sections and posterior cerebellar cortex sections into their respective microfuge tubes and flash freeze by leaving the tubes in liquid nitrogen for 5 minutes. From here move forward to RNA extraction, or may &#160;store the tubes at -80 &#730;C.</li>
</ol>
3. RNA extraction 
NOTE: This protocol is modified from the Cold Spring Harbor Protocol for RNA Extraction with TRIzol20. TRIzol solubilizes biological material, making it possible to extract RNA.
<ol>
	<li>Place the microfuge tubes in ice to keep the tissue from thawing too quickly, apply 150 &#181;L&#160;of cold TRIzol in the microfuge tube. Homogenize with a sterilized pestle. Once tissue is homogenized, pipet the solution up and down to ensure there is no remaining tissue intact. Further break up any small tissue pieces by pulling it up into an insulin syringe a few times.</li>
	<li>Add another 350 &#181;L&#160;of TRIzol and pipet up and down to mix thoroughly. Let sit at room temp for 5 minutes.</li>
	<li>Add 150 &#181;L&#160;of chloroform to the tube and shake vigorously and then let rest for 2-3 minutes. The chloroform separates the homogenized tissue solution into phases (RNA, DNA, and protein).</li>
	<li>Centrifuge at 12,000 x g, at 15&#730;C, for 10 minutes. Ensure that all tubes are at the same orientation.</li>
	<li>Carefully remove the tubes and set the temperature of the centrifuge to 4&#730;C. Remove only the clear aqueous phase into a new tube (this is the RNA), careful not to disrupt the opaque interphase (the DNA). &#160;The lowest phase will be red and contains protein. Remaining solution in the tubes can be saved or discarded.</li>
	<li>Add 100% isopropyl alcohol at a 1:2 ratio (if removed 200 ul of aqueous phase, add 100 &#181;L&#160;of isopropyl alcohol). Mix thoroughly by pipetting up and down. Let rest at room temperature for 10 minutes. The isopropyl alcohol precipitates the RNA out of solution.</li>
	<li>Centrifuge at 12,000 x g, at 4&#730;C, for 10 minutes. Be sure to place all the tubes at the same orientation to make it easier to visualize the pellet. The resulting pellet will be the extracted RNA.</li>
	<li>Carefully remove the tubes, remove the supernatant with a pipet being careful not to disrupt the pellet. The pellet is somewhat gel-like and will be difficult to see, but should be able to estimate where it is based on the orientation of the tubes in the centrifuge.</li>
	<li>After removing all of the supernatant, add 500 &#181;L&#160;of 75% ethanol, vortex briefly, and centrifuge at 7500 x g, at 4&#730;C, for 5 minutes. The ethanol further washes the pellet.</li>
	<li>Remove the supernatant carefully, without disrupting the pellet. Leave caps open to dry the sample. This usually takes 5-10 minutes but can vary depending on how much ethanol was left on the pellet. Do not over dry.</li>
	<li>Once dry, resuspend the pellet in DNase free water. Add 20 &#181;L&#160;to samples for DCN, and 30 &#181;L&#160;for all others.</li>
	<li>Once resuspended, samples can be stored at -80 &#730;C or proceed to further testing.</li>
</ol>
4. Real Time quantitative Polymerase Chain Reaction (RTqPCR)
<ol>
	<li>Before this step it will be necessary to DNase treat the RNA to remove any genomic DNA, and generate cDNA utilizing iScript from BioRad. Be sure to normalize the concentration of RNA before making the cDNA. In addition, it is important to optimize primers for qPCR. Follow the methods described here to complete this step21. Brief description of the qPCR below.</li>
	<li>Primers are all purchased from IDT. Forward and reverse sequences are both in the same sample tube, and stored at 20x concentration. 
	Rps18 (control gene): 
	Forward &#8211; 5&#8217;-CCTGAGAAGTTCCAGCACAT-3&#8217; 
	Reverse &#8211; 5&#8217;-ACACCACATGAGCATATCTCC-3&#39; 
	Parvalbumin (Calcium binding protein, inhibitory cells): 
	Forward &#8211; &#8216;5-ATGAGGTGAAGAAGGTGTTCC-3&#8217; 
	Reverse &#8211; &#8216;5-AGCGTCTTTGTTTCTTTAGCAG-3&#8217; 
	Kcng4 (Potassium channel subunit): 
	Forward &#8211; 5&#8217;-CTGTCTTTTCCTGGTCAGTGA-3&#8217; 
	Reverse &#8211; 5&#8217;-GCATTGCCTCAGACTGTCAG-3&#8217; 
	Aldolase C (Zebrin II, differentially expressed across the cerebellar cortex): 
	Forward &#8211; 5&#8217;-AGAGGACAAAGGGATAATGCTG-3&#8217; 
	Reverse &#8211; 5&#8217;-TCAGTAGGCATGGTTGGC-3&#8217;</li>
	<li>qPCR conditions were as follows. Pre-incubation period, 5 minutes, at 95&#730;C. Amplification period, 50 cycles: 10 minutes at 95&#730;C, 10 minutes at 62&#730;C, and 10 minutes at 72&#730;C. Melting curve period, 5 seconds at 95&#730;C, one minute at 65&#730;C, and set ramp rate to .07&#730;C/second until target temp of 97&#730;C. Then cooling period for 10 minutes to 40&#730;C. &#160;All qPCR reactions were performed in triplicate and gene expression was analyzed using 2- &#916;&#916;Ct with Rps18 as a loading control to normalize gene expression. Bulk cerebellar extract was used as a reference.</li>
</ol>

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D., Caplan, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cognition,+enotion,+and+the+cerebellum.">Cognition, enotion, and the cerebellum.</a> Brain. 129 (2), 290-292 (2006).</li><li>Badura, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Normal+cognitive+and+social+development+require+posterior+cerebellar+activity.">Normal cognitive and social development require posterior cerebellar activity.</a> eLife. 7, 36401 (2018).</li><li>Diedrichsen, J., King, M., Hernandez-Castillo, C., Sereno, M., Ivry, R. 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The authors have nothing to disclose.

The method described here makes it possible to assess the underlying gene expression and molecular mechanisms within four distinct cerebellar regions &#8211; the deep cerebellar nuclei (DCN), the anterior cerebellar cortex of the vermis (CCaV), the posterior cerebellar cortex of the vermis (CCpV), and the cerebellar cortex of the hemispheres (CCH). The ability to assess these regions separately will expand our knowledge of the heterogeneity of specific cerebellar regions and possibly shed light on their contribution to v...

The cerebellum contains over half of the neurons in the brain and has historically been referred to as a motor control and balance center in the brain1. More recently, studies have demonstrated that the cerebellum plays a key role in various other functions including cognition, reward processing, and affect2,3,4,5.
The cerebellum has well-described anatomy: the cortex region is composed of granule, Purkinje, and molecular layers. Granule cells form the granule cell layer and send input via parallel fibers to the Purkinje cell dendrites of the molecular layer which also receive input from climbing fibers that originated in the inferior olive. Purkinje cells send inhibitory projections to cells in the deep cerebellar nuclei (DCN), which serves as the main output from the cerebellum. The output of this cerebellar circuit is further modulated by the activity of the inhibitory interneurons in the cerebellar cortex, including Golgi, stellate, and basket cells4. This cerebellar functional unit is distributed throughout all the lobules of the cerebellar cortex. Despite this relatively uniform circuitry across &#160;the cerebellum, evidence from human neuroimaging literature and patient studies indicates functional heterogeneity of the cerebellum6,7.
The cerebellar cortex can be divided into two main regions: the midline-defined vermis, and the lateral hemispheres. The vermis can be further divided into anterior and posterior lobules. These distinct regions of the cerebellum have been implicated in contributing to different behaviors. Task-evoked or task-free activity patterns implicated that anterior regions of the vermis contribute more to motor function while posterior vermis contributes more to cognition6,7. The vermis is also linked with affect and emotions, while cerebellar hemispheres contribute to executive, visual-spatial, language, and other mnemonic functions8. In addition, anatomical studies provided evidence that functionally distinct cerebellar regions are connected with different cortical regions9. Lesion-symptom mapping revealed that patients with strokes affecting the anterior lobules (extending into lobule VI) had poorer performance on fine motor tasks, while patients with damage to posterior lobe regions and hemispheres exhibited cognitive deficits in the absence of cerebellar motor syndrome10. Finally, regional cerebellar pathology in disease indicates that functionally distinct cerebellar regions are also differently susceptible to disease11,12.
While much less explored, preliminary evidence demonstrates distinct gene expression signatures across cerebellar cortical regions. Purkinje cell expression of Zebrin II shows region specific patterning in the vermis such that there are more Zebrin II positive cells in the posterior lobules and fewer in the anterior lobules13. This also correlates with regionally distinct physiological function as Zebrin II negative Purkinje cells display higher frequency of tonic firing than Purkinje cells that are Zebrin II positive14.
In addition to the cerebellar cortex, the cerebellum includes the deep cerebellar nuclei (DCN) which serve as the primary output for the cerebellum. The nuclei are made up of the medial (MN), interposed (IN), and lateral nuclei (LN). Functional imaging and patient studies have demonstrated that the DCN also participate in various behaviors15, but very few studies examine gene expression change in DCN.
Advances in molecular techniques have made it possible to assess regional gene expression in the brain and have uncovered heterogeneity across and within different brain regions in both physiological and disease states16. Such studies implicate that the cerebellum is different from other brain regions. For example, the ratio of neurons to glial cells is inverted in the cerebellum&#160; compared to other brain regions1. Even in normal physiological conditions, the expression of proinflammatory genes is upregulated in the cerebellum compared to the other brain regions17. Molecular techniques have also been very useful in identifying the pathways that contribute to the pathogenesis of cerebellar diseases. For instance, RNA sequencing of the whole cerebellar extracts identified genes altered in a Purkinje cell specific transgenic mouse model of spinocerebellar ataxia type 1 (SCA1) as compared to their wild type controls. Such evidence has revealed key molecular pathways underlying pathogenesis in cerebellar Purkinje cells and has helped identify potential therapeutic targets18. However, recent studies suggest that there are differences in the vulnerability to diseases across the cerebellar regions11,12,19. This could indicate that there are key changes occurring in distinct cerebellar regions, which may be masked or undetected with whole cerebellar extracts. Thus, there is a need to develop techniques which allow researchers to examine molecular profiles in different cerebellar regions.
The technique proposed here describes a reproducible method to dissect four distinct regions of the mouse cerebellum in order to isolate RNA from those regions and explore regional differences in gene expression. The schematic of the mouse cerebellum in Figure 1A highlights the vermis in blue, and hemispheres in yellow. Specifically, this technique makes it possible to isolate four regions:&#160; deep cerebellar nuclei (DCN) (red-dotted boxes in Figure 1A), the cerebellar cortex of anterior vermis (CCaV) (dark blue in Figure 1A), the cerebellar cortex of the posterior vermis (CCpV) (light blue in Figure 1A), and the cerebellar cortex of the hemispheres (CCH) (yellow in Figure 1A). By assessing gene expression of these regions separately, it will be possible to investigate molecular mechanisms underlying discrete functions of these different regions as well as potential differences in their vulnerability in disease.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >1.5 Microcentrifuge tubes</td>
			<td >ThermoScietific</td>
			<td >3456</td>
			<td ></td>
		</tr>
		<tr >
			<td >100% Isopropyl Alcohol</td>
			<td >VWR Life sciences</td>
			<td >1106C361</td>
			<td></td>
		</tr>
		<tr >
			<td >200 ul Pipet tips</td>
			<td >GeneMate</td>
			<td >P-1237-200</td>
			<td></td>
		</tr>
		<tr >
			<td >Adult Mouse Brain Matrix Sagittal</td>
			<td >Kent Scientific Corporation</td>
			<td >RBMA-200S</td>
			<td></td>
		</tr>
		<tr >
			<td >Blunt forceps</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Chloroform</td>
			<td >Macron</td>
			<td >220905</td>
			<td></td>
		</tr>
		<tr >
			<td >Decapitation Scissors</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Dissecting Scissors</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Ethyl Alcohol</td>
			<td >Pharmco</td>
			<td >111000200</td>
			<td></td>
		</tr>
		<tr >
			<td >Glass Slide (for electrophoresis)</td>
			<td >BIORAD</td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Homogenizer</td>
			<td >Kimble</td>
			<td >6HAZ6</td>
			<td></td>
		</tr>
		<tr >
			<td >Ice Bucket</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Insulin Syringe (.5ml)</td>
			<td >BD</td>
			<td >329461</td>
			<td></td>
		</tr>
		<tr >
			<td >iScript Adv cDNA kit for RT-qPCR</td>
			<td >BIORAD</td>
			<td >1725037</td>
			<td></td>
		</tr>
		<tr >
			<td >Micro Spatula</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Needle Nose forceps</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Petri Dish</td>
			<td >Pyrex</td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Primetime Primer for Aldolase C</td>
			<td >IDT</td>
			<td >Mm.PT.58&#62;43415246</td>
			<td></td>
		</tr>
		<tr >
			<td >Primetime Primer for Kcng4</td>
			<td >IDT</td>
			<td >Mm.PT.56a.9448518</td>
			<td></td>
		</tr>
		<tr >
			<td >Primetime Primer for Parvalbumin</td>
			<td >IDT</td>
			<td >Mm.PT.58.7596729</td>
			<td></td>
		</tr>
		<tr >
			<td >Primetime Primer Rps18&#160;</td>
			<td >IDT</td>
			<td >Mm.PT.58.12109666</td>
			<td></td>
		</tr>
		<tr >
			<td >Single Edge Rzor Blades</td>
			<td >Personna GEM</td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Sterile, sigle-use pestles</td>
			<td >FisherScientific</td>
			<td >12141364</td>
			<td></td>
		</tr>
		<tr >
			<td >TRIzol Reagent</td>
			<td >Ambion by Life technologies</td>
			<td >15596018</td>
			<td></td>
		</tr>
		<tr >
			<td >Vascular Scissors</td>
			<td ></td>
			<td ></td>
			<td></td>
		</tr>
	</tbody>
</table>

cerebellar regional dissection for molecular analysis

Cerebellum plays an important role in several key functions including control of movement, balance, cognition, reward, and affect. Imaging studies indicate that distinct cerebellar regions contribute to these different functions. Molecular studies examining regional cerebellar differences are lagging as they are mostly done on whole cerebellar extracts thereby masking any distinctions across specific cerebellar regions. Here we describe a technique to reproducibly and quickly dissect four different cerebellar regions: the deep cerebellar nuclei (DCN), anterior and posterior vermal cerebellar cortex, and the cerebellar cortex of the hemispheres. Dissecting out these distinct regions allows for the exploration of molecular mechanisms that may underlie their unique contributions to balance, movement, affect and cognition. This technique may also be used to explore differences in pathological susceptibility of these specific regions across various mouse disease models.

For these experiments, four eleven-week-old female wild type C57/Black6 mice were used. One mouse was used to conduct a full cerebellar dissection which is referred to as &#39;bulk cerebellum&#39; and allowed for the comparison of RNA levels in dissected regions to a full dissection. The other three mice were used to conduct the cerebellar dissection described in this protocol. Using three mice makes it possible to ensure that the trends detected in the levels of RNA are reproducible acro...

Watch this Scientific Journal Video about Cerebellar Regional Dissection for Molecular Analysis at JoVE.com

Cerebellar Regional Dissection for Molecular Analysis

Different cerebellar regions have been implicated to play a role in distinct behavioral outputs, yet the underlying molecular mechanisms remain unknown. This work describes a method to reproducibly and quickly dissect cerebellar cortex of the hemispheres, anterior and posterior regions of the vermis, and the deep cerebellar nuclei in order to probe for molecular differences by isolating RNA and testing for differences in gene expression.&#160;

Cerebellum plays an important role in several key functions including control of movement, balance, cognition, reward, and affect. Imaging studies ...

cerebellar-regional-dissection-for-molecular-analysis

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JoVE Journal

Neuroscience

3.8K Views.  University of Minnesota. Different cerebellar regions have been implicated to play a role in distinct behavioral outputs, yet the underlying molecular mechanisms remain unknown. This work describes a method to reproducibly and quickly dissect cerebellar cortex of the hemispheres, anterior and posterior regions of the vermis, and the deep cerebellar nuclei in order to probe for molecular differences by isolating RNA and testing for differences in gene expression. 

Video: Cerebellar Regional Dissection for Molecular Analysis

A Simple Composite Phenotype Scoring System for Evaluating Mouse Models of Cerebellar Ataxia

We describe a protocol for the rapid and sensitive quantification of disease severity in mouse models of cerebella ataxia. It is derived from previously published phenotype assessments in several disease models, including spinocerebellar ataxias, Huntington s disease and spinobulbar muscular atrophy. Measures include hind limb clasping, ledge test, gait and kyphosis. Each measure is recorded on a scale of 0-3, with a combined total of 0-12 for all four measures. The results effectively discriminate between affected and non-affected individuals, while also quantifying the temporal progression of neurodegenerative disease phenotypes. Measures may be analyzed individually or combined into a composite phenotype score for greater statistical power. The ideal combination of the four described measures will depend upon the disorder in question. We present an example of the protocol used to assess disease severity in a transgenic mouse model of spinocerebellar ataxia type 7 (SCA7).
Albert R. La Spada and Gwenn A. Garden contributed to this manuscript equally.

We describe a protocol for the rapid and sensitive quantification of disease severity in mouse models of cerebellar ataxia. Measures include hind limb clasping, ledge test, gait and kyphosis. This protocol effectively discriminates between affected and non-affected individuals, and detects the progression of affected individuals over time.

We describe a protocol for the rapid and sensitive quantification of disease severity in mouse models of cerebella ataxia. It is derived from previously ...

Organotypic Cerebellar Cultures: Apoptotic Challenges and Detection

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. Since then, the technique was developed further and currently there are many different ways to prepare organotypic cultures. The method presented here was adapted from the one described by Stoppini et al. for the preparation of the slices and from Gogolla et al. for the staining procedure 3,4.
A unique feature of this technique is that it allows you to study different parts of the brain such as hippocampus or cerebellum in their original structure, providing a big advantage over dissociated cultures in which all the cellular organization and neuronal networks are disrupted. In the case of the cerebellum it is even more advantageous because it allows the study of Purkinje cells, extremely difficult to obtain as dissociated primary culture. This method can be used to study certain developmental features of the cerebellum in vitro, as well as for electrophysiological and pharmacological experiments in both wild type and mutant mice.
The method described here was designed to study the effect of apoptotic stimuli such as Fas ligand in the developing cerebellum, using TUNEL staining to measure apoptotic cell death. If TUNEL staining is combined with cell type specific markers, such as Calbindin for Purkinje cells, it is possible to evaluate cell death in a cell population specific manner. The Calbindin staining also serves the purpose of evaluating the quality of the cerebellar cultures.

This method describes the generation of organotypic cerebellar cultures and the effect of certain apoptotic stimuli on the viability of different cerebellar cell types.

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. ...

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 ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

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

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

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

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

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

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

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 ...

Laser Nanosurgery of Cerebellar Axons In Vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to understand the mechanism that promotes neuronal regeneration, an in-depth analysis of the neuronal types that can remodel after injury is needed. Several studies showed that damaged climbing fibers are capable of regrowing also in adult animals1,2. The investigation of the time-lapse dynamics of degeneration and regeneration of these axons within their complex environment can be performed by time-lapse two-photon fluorescence (TPF) imaging in vivo3,4. This technique is here combined with laser surgery, which proved to be a highly selective tool to disrupt fluorescent structures in the intact mouse cortex5-9.
This protocol describes how to perform TPF time-lapse imaging and laser nanosurgery of single axonal branches in the cerebellum in vivo. Olivocerebellar neurons are labeled by anterograde tracing with a dextran-conjugated dye and then monitored by TPF imaging through a cranial window. The terminal portion of their axons are then dissected by irradiation with a Ti:Sapphire laser at high power. The degeneration and potential regrowth of the damaged neuron are monitored by TPF in vivo imaging during the days following the injury.

Laser Nanosurgery of Cerebellar Axons In Vivo

Two-photon imaging, coupled to laser nanodissection, are useful tools to study degenerative and regenerative processes in the central nervous system with subcellular resolution. This protocol shows how to label, image, and dissect single climbing fibers in the cerebellar cortex in vivo.

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to ...

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 ...