Name: diolistic labeling of neurons from rodent and non-human primate brain slices
Uploaded: 2010-07-06T17:00:00
Duration: 9 min 21 s
Description: Watch this Scientific Journal Video about DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices at JoVE.com

We would like to acknowledge Michael Feyder and Terrell Holloway for their assistance during the initial set up of the technique and Dr. Fumi Ono s laboratory for the access to their confocal microscope. This research was funded by the National Institute of Health through the intramural program of NIAAA and NINDs.

1. Preparing DiI/Tungsten Bead Bullets: DiI (1-1'-Dioctadecyl-3,3,3',3'- tetramethylindocarbocyanine perchlorate) bullets should be prepared in advance of cutting brain slices. The procedure takes approximately 2-4 hours depending on how many bullets are prepared at once.
<ol>
	<li>If starting from a new batch of tungsten beads, prepare aliquots of 300 mg each. Aliquots are prepared by suspending 6 grams of tungsten in 2 ml of methylene chloride. Then, pipette 100 &mu;l of bead solution into microfuge tubes to produce aliquots of 300 mg tungsten beads. Store aliquots at room temperature. (Note: methylene chloride evaporates extremely quickly. Alcohol can also be used minimize evaporation).</li>
 <li>When starting from aliquots of beads, resuspend each aliquot (300 mg tungsten beads) in 300 &mu;l of methylene chloride and cover quickly to prevent evaporation. This quantity is enough for 3 batches of bullets.</li>
 <li>Prepare dye solution by weighing 13.5 mg of lipophilic dye DiI and adding 450 &mu;l of methylene chloride (final concentration = 3mg/100&mu;l). </li>
 <li>Coat the beads with dye. Place a glass slide on a piece of wax-coated weigh paper and pipette 100 &mu;l of beads onto the slide. Add 100 &mu;l of DiI solution, mix thoroughly using pipette tip, and air dry until beads turn light grey.</li>
 <li>Use a razor blade to scrape beads off the glass slide and dice the beads into a fine powder. (Note: use a new blade if doing more than one color of bullets so as to not contaminate dyes).</li>
 <li>Scrape beads onto the weigh paper and funnel beads into a 15 ml conical tube. Add 3 ml of water and sonicate in water bath for 10-30 min at room temperature. (Note: dicing of the powder and sonication are applied to minimize the formation of clumps of coated beads that will disrupt the sparse labeling of individual neurons). </li>
</ol>
2. Preparing polyvinylpyrrolidone (PVP) solution and tubing: To improve bead attachment to the bullet tubing (TEZFEL tubing), coat it with polyvinylpyrrolidone (PVP) solution.
<ol>
 <li>Prepare 10 ml of PVP solution in water at a concentration of 10 mg PVP/ml. </li>
 <li>Use a 12 ml syringe with tubing adaptor (flexible tubing with slightly larger diameter) to pass the PVP solution through the bullet tubing and then expel it out the other end. Reuse solution to coat more bullet tubing if several batches are prepared simultaneously. Discard PVP solution after use.</li>
 <li>To add the beads to the bullet tubing, vortex the beads to produce a homogenous solution and use the syringe to pull the bead solution through the tubing so that beads are evenly distributed throughout the tubing. ((Note:) try to minimize the number of air bubbles in the tubing).</li>
 <li>Feed the tubing through the prep station and allow the beads to settle. Use the syringe to gently remove the water so that only the beads remain.</li>
 <li> Spin the tubing in the prep station and dry with nitrogen gas until water droplets are no longer visible. This step will take approximately 10-20 min.</li>
 <li>Cut bullets with tubing cutter into 13 mm lengths and place in scintillation vials containing Drierite or some kind of anhydrous calcium sulfate. Wrap vials in foil to protect from light.</li>
</ol>
3. DiI Staining: This labeling technique can be used to stain neurons in brain tissue from diverse species; in this particular case, from mouse (3-6 month old) and non-human primates (9-15 years old).
It is preferable that slices are prepared from fixed brain tissue obtained by transcardiac perfusion of fixative solution because tissue preservation is expected to be best under this condition. However, fixation by perfusion is not always feasible (as was the case with the monkey tissue) and DiOLISTIC labeling can still be successfully applied under different procedures for tissue preparation. Here we describe three different procedures for tissue preparation:
<ol class="lalpha">
 <li>Fix brain by transcardiac perfusion &rarr; remove brain &rarr; postfix for 10 min &rarr; cut slice &rarr; stain </li>
 <li>Remove brain &rarr; cut block of tissue &rarr; fix block &rarr; wash in PBS &rarr; cut slice &rarr; stain</li>
 <li>Remove brain &rarr; cut slices &rarr; fix &rarr; wash &rarr; stain</li>
</ol>
In the present study, the monkey brains were obtained from subjects that were transcardially perfused with ice cold artificial cerebral spinal fluid (ACSF) prior to brain harvest and a block (4 mm thick) of tissue containing the caudate and putamen was fixed for 60 min at room temperature. For all procedures, fixative solution contains 4% paraformaldehyde and 4% sucrose in PBS (prepared fresh or used within 15 days). It is important to note that fixation times are critical for successful staining. Overfixation can affect staining by disrupting the integrity of the plasma membrane and causing dye to leak out of the cell (see more information under results).
<ol>
	<li>For procedure a and b, slices (200&mu;m) are cut from fixed brain or tissue block and slices placed in 24-well plate with PBS. For procedure c, fresh acute brain slices (200 &mu;m) are cut with a vibratome in cold cutting solution containing (in mM) 110 Choline-Cl, 25 NaHCO3, 1.25 NaH2PO4, 2.5 KCl, 0.5 CaCl2 , 7 MgCl2, 25 glucose, 11.6 ascorbic acid, 3.1 pyruvic acid (osmolarity = 310 mOsmols) and immediately after slices are placed in 24-well plates and fixed for 30 min with fixative solution at room temperature.</li>
 <li>Wash the slices with PBS, 3 times.</li>
 <li>Before shooting slices, remove the PBS from the wells and, using a paintbrush, place the slice in the center of the well. </li>
 <li>Shoot DiI bullets through 3.0 &mu;m filter paper using a Helios Gene Gun system at 120-180 psi helium gas pressure by placing the gun at a distance of 1.5 cm between the sample and the end of the barrel. (Important note: Only neurons within 50 mm of the slice surface will be labeled by this method so it is important to keep the same side of the slice facing up during the washes and for mounting. In this way, you will make sure that the labeled neurons will be within focal distance when imaging with a confocal microscope).</li>
 <li>Wash slices with PBS 3 times and store them in PBS for several hours to allow the DiI to spread along the dendrites. </li>
 <li>Mount slices onto a glass slide with ProLong Gold Antifade and cover with an 18X18 mm No.1 coverslip. (Note: for procedure c, it is best to mount slices on the same day as the integrity of the slices is not well maintained overnight). </li>
 <li>Seal with clear nail polish after mounting media has dried.</li>
 <li>Alternatively, the slices can be stained with antibodies before mounting as described below. (Note: Slides can be used at least 6 months to a year if stored in the dark at 4&deg;C. DiI is light sensitive and long-term exposure to light will cause the stain to fade).</li>
</ol>
4. Antibody Staining: Immunostaining should be performed after step 3.4 and before mounting.
<ol>
 <li>Permeabilization: Incubate slices in 0.01% Triton X-100 in PBS solution for 15 min at room temperature (300-500 &mu;l per well). NOTE: Do not use higher concentrations of detergent as this will dissolve DiI and significantly reduce the fluorescent staining. Try to minimize the time in detergent as extensive incubations will also reduce the DiI labeling. If extended incubations are necessary, keep slices in PBS rather than in blocking solution.</li>
 <li>Blocking: Incubate slices in blocking solution containing 10% goat serum, 0.01% Triton X-100 in PBS for 30 min at room temperature.</li>
 <li> Primary antibody: Add antibody at the desired dilution in blocking solution (e.g. rabbit anti-GFP antibody at 1:1000 dilution). Add 300-500 &mu;l per well and incubate for 1-2 hrs. (Note: if overnight incubations are necessary, remove detergent from blocking solution).</li>
 <li>Wash slices with PBS for 5 - 15 min, 3 times.</li>
 <li>Secondary antibody: Incubate with secondary antibody for 30 min at room temperature. Prepare secondary antibody solution at desired dilution in blocking solution (e.g. Alexa-Fluor 488 conjugated antibody at 1:1000 dilution).</li>
 <li>Wash slices with PBS for 5-15 min, 4 times.</li>
 <li>Mount slices on slides with ProLong Gold Antifade and cover with an 18x18 mm No.1 coverslip. Seal with nail polish after mounting media has dried.</li>
</ol>
5. Representative Results: 
 <ol>
 <li>Checking for correct labeling: Section can be visualized under a fluorescent dissecting scope equipped with a mercury lamp and red fluorescent filter cube to confirm successful DiOLISTIC labeling. Figure 1 shows exemplary images of nicely labeled brain sections obtained at low magnification. Two important characteristics to look for are a sparse labeling pattern (Figure 1A) and the ability to identify individual cellular components (Figure 1B-C). </li>
 <li>Troubleshooting DiOLISTIC labeling: Figure 2 shows examples of sections in which the DiOLISTIC labeling worked incorrectly. In our experience, there are three main causes for inefficient labeling:
 <ul> 
		<li>Labeling is too sparse or dense: Very few cells are labeled per section in one case or too many cells are labeled preventing the isolation and study of single cellular elements. Solution: adjust the concentration of coated beads in the final solution used for coating the TEFZEL bullet tubing (section 1.6). Decrease or increase the volume of water (3 ml initial value) used to dissolve the beads in order to correct for too sparse or too dense labeling, respectively. In addition, low efficiency of labeling might be caused by less than optimal gas pressure when shooting the coated beads onto the sections. This possibility can be ruled out by making sure that the TEFZEL bullet tubing has released most of the coated beads and it appears clear after shooting. Otherwise, gas pressure for shooting should be increased.</li>
		<li>Bad bullets: large clumps or clusters of dye-coated tungsten beads are formed during the bullet preparation. Sections will resemble examples in Figure 2A-B. Solution: make new bullets paying particular attention to section 1.5 and/or extend sonication time in section 1.6.</li>
		<li>Over-fixation: tissue is kept in paraformaldehyde solution for longer than the optimally required time for fixing (30 minutes for 200-300 &mu;m sections, 1 hour for 4 mm sections). This compromises the integrity of the plasma membrane and produces labeled sections that look like those shown in Figure 2C-D in which labeling seems out of focus due to dispersion of dye out of the cells. Solution: reduce fixation time.</li>
 </ul>
 </li>
 <li>Confocal imaging: Image stacks are acquired using a confocal microscope (Zeiss LSM 510 META) with a 20x objective for the whole cell and 63x water immersion objective for dendrite segments. DiI was excited using a DPS 561 nm laser line and green fluorescence from the secondary antibody was excited using a 488 nm laser line. The optical sectioning of the labeled sample achieved when using confocal microscopy further allows for isolation of single labeled cells in the sample. Figure 3A shows a striatal medium spiny neuron from mouse brain and its complex dendrite branch pattern. High magnification images of dendrite branches of these neurons from rodent (Figure 3B) and non-human primate (Figure 3C) demonstrate the degree of staining in dendrites and dendritic spines using this technique in both species. Dendrites and their protrusions (spines) appear well defined, even when considering the long, thin spine heads so abundant in medium spiny neurons. When combining DiOLISTICS with immunostaining, confocal imaging is also helpful in unequivocally localizing the stain to the same focal plane and the same cell. Figure 4 shows an example of colocalization of DiI labeling and immunostaining for GFP in a single cell. The image is one stack (0.7 &mu;m z-section) obtained from a striatal slice from a transgenic mouse expressing the fluorescent protein GFP under the D1 dopamine receptor promoter.</li>
</ol>
<img src="/files/ftp_upload/2081/2081fig1.jpg" alt="Figure 1" /> Figure 1. DiI labeling of neurons in brain slices from non-human primates. (A) Low magnification image of a stained brain slice containing the caudate nucleus from a cynomolgus monkey that shows sparse labeling of neurons with DiI. (B-C) Isolated medium spiny neurons can be easily identified using a fluorescent dissecting scope.
<img src="/files/ftp_upload/2081/2081fig2.jpg" alt="Figure 2" /> Figure 2. Troubleshooting DiOLISTIC labeling. (A-B) Stained slices from dorsal striatum of monkey (A) and mouse (B) showing large clumps of dye-coated beads and as a consequence, no individual cellular elements can be distinguished. (C-D) Images that exemplify the result of applying DiOLISTIC labeling to sections that were kept in fixative solution for prolonged periods of time or where tissue preservation failed in monkey (C) and mouse (D) tissue.
<img src="/files/ftp_upload/2081/2081fig3.jpg" alt="Figure 3" /> Figure 3. Confocal image of striatal medium spiny neuron stained with DiI. (A), Image stack of a whole cell allows for morphological analysis of the dendritic branches. (B-C) Dendrite segments from mouse (B) and monkey (C) medium spiny neurons show clear labeling of dendrite and spines.
<img src="/files/ftp_upload/2081/2081fig4.jpg" alt="Figure 4" /> Figure 4. Combining DiOLISTIC labeling with immunostaining. (A) DiI labeled medium spiny neuron from striatum of BAC transgenic mouse expressing GFP under the D1 dopamine receptor promoter. (B) Immunostaining using anti-GFP antibody as described by methods adapted from Lee et al. (2006). Same field as A. (C) Composite image of red and green fluorescence identifies DiI-labeled neuron as a GFP-positive neuron.

<ol>
	<li>Gan, W. B., Grutzendler, J., Wong, W. T., Wong, R. O., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multicolor+"DiOLISTIC"+labeling+of+the+nervous+system+using+lipophilic+dye+combinations.">Multicolor "DiOLISTIC" labeling of the nervous system using lipophilic dye combinations.</a> Neuron. 27, 219-225 (2000).</li><li>Gan, W. B., Grutzendler, J., Wong, R. O., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ballistic+delivery+of+dyes+for+structural+and+functional+studies+of+the+nervous+system.">Ballistic delivery of dyes for structural and functional studies of the nervous system.</a> Cold Spring Harb Protoc. , pdb.prot5202-pdb.prot5202 (2009).</li><li>Grutzendler, J., Tsai, J., Gan, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+labeling+of+neuronal+populations+by+ballistic+delivery+of+fluorescent+dyes.">Rapid labeling of neuronal populations by ballistic delivery of fluorescent dyes.</a> Methods. 30, 79-85 (2003).</li><li>Lee, K. W., Kim, Y., Kim, A. M., Helmin, K., Nairn, A. C., Greengard, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cocaine-induced+dendritic+spine+formation+in+D1+and+D2+dopamine+receptor-containing+medium+spiny+neurons+in+nucleus+accumbens.">Cocaine-induced dendritic spine formation in D1 and D2 dopamine receptor-containing medium spiny neurons in nucleus accumbens.</a> Proc. Natl. Acad. Sci. U S A. 103, 3399-3404 (2006).</li><li>Matsubayashi, Y., Iwai, L., Kawasaki, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+double-labeling+with+carbocyanine+neuronal+tracing+and+immunohistochemistry+using+a+cholesterol-specific+detergent+digitonin.">Fluorescent double-labeling with carbocyanine neuronal tracing and immunohistochemistry using a cholesterol-specific detergent digitonin.</a> J Neurosci Methods. 174, 71-81 (2008).</li><li>Neely, S. t. a. n. w. o. o. d., Deutch, G. D., Y, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Combination+of+diOlistic+labeling+with+retrograde+tract+tracing+and+immunohistochemistry.">Combination of diOlistic labeling with retrograde tract tracing and immunohistochemistry.</a> J Neurosci Methods. 184, 332-336 (2009).</li><li>O'Brien, J. A., Lummis, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diolistics:+incorporating+fluorescent+dyes+into+biological+samples+using+a+gene+gun.">Diolistics: incorporating fluorescent dyes into biological samples using a gene gun.</a> Trends Biotechnol. 25, 530-534 (2007).</li><li>Shen, H. W., Toda, S., Moussawi, K., Bouknight, A., Zahm, D. S., Kalivas, P. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Altered+dendritic+spine+plasticity+in+cocaine-+withdrawn+rats.">Altered dendritic spine plasticity in cocaine- withdrawn rats.</a> J. Neurosci. 29, 2876-2884 (2009).</li></ol>

All animal procedures were performed following guidance from the Animal Care and Use Committee at NIAAA and the Oregon National Primate Research Center. The authors have nothing to disclose.

DiOLISTIC labeling is one of the most versatile techniques available for fluorescently labeling cells because it can be applied to tissue sections from diverse species, to a wide range of ages, and to tissue obtained fresh or from fixative-perfused animals (see also Gan et al., 2009). The process is relatively fast as it takes 1-2 days and it can be combined with other more classical labeling approaches such as immunostaining (Lee et al., 2006). Special attention should be paid to avoid over fixing and the use of high detergent concentrations in incubation solutions because these will comprise the integrity of the lipophilic membrane and cause the dye to leak out of the cells. Antibody penetration can be facilitated by low concentrations of Triton X-100 (Lee et al., 2006), as well as digitonin or saponin in the incubation solution (Matsubayashi et al., 2008). Some modifications that can be applied to this technique include the use of bullets with multiple dyes as described by Gan et al. (2000) or changing the length and type of antibody exposure (Neely et al., 2009).
Traditionally, DiI has been used to trace neuronal projections in the brain. DiOLISTIC labeling expands its application by describing a method useful to examine cell morphology. Neuronal morphology is of great interest because of the large diversity found in the brain and the speculation that cell shape might reflect on the function multiplicity of neuronal populations in the nervous system. One example of this is the fact that many neurons in the mammalian nervous system display small protrusion called dendritic spines that are the site of glutamatergic synapses. In this way, the density of glutamatergic synapses onto a cell can be correlated to the density of dendritic spines, which can be measured using the labeling technique described herein. Furthermore, other morphological parameters such as dendrite total length, branching pattern, dendritic spine shape and density can be quantified and studied.
The use of fluorescent labeling for studying neuronal morphology has many advantages when compared to more traditional techniques that rely on bright field microscopy (e.g. Golgi staining) because it allows for higher resolution confocal imaging. Another advantage of using DiI as a fluorophore in experiments aimed at measuring dendritic spine density and morphology is its lipophilic properties. DiI partitions into the plasma membrane and provides a well defined outline of the neuronal processes and dendritic protrusions. Given the small volume of most dendritic spines (less than 1 femtoliter), membrane staining is more efficient and allows for better visualization of small, thin protrusions than cytoplasmic staining.

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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>1-1&#x2019;-dioctadecyl-3,3,3&#x2019;,3&#x2019;- tetramethyl-indocarbocyanine perchlorate (DiI)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>D-282</td>
<td>&nbsp;</td>
<tr>
<td>Methylene chloride</td>
<td>&nbsp;</td>
<td>Mallinckrodt Chemicals</td>
<td>H485-06</td>
<td>&nbsp;</td>
<tr>
<td>Tungsten beads</td>
<td>&nbsp;</td>
<td>Bio-Rad</td>
<td>165-2269</td>
<td>1.7 &mu;m diameter</td>
</tr>
<tr>
<td>Polyvinylpyrrolidone </td>
<td>&nbsp;</td>
<td>Calbiochem</td>
<td>529504</td>
<td>&nbsp;</td>
<tr>
<td>Tefzel bullet tubing</td>
<td>&nbsp;</td>
<td>Bio-Rad</td>
<td>165-2441</td>
<td>&nbsp;</td>
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<td>Tubing cutter</td>
<td>&nbsp;</td>
<td>Bio-Rad</td>
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<td>&nbsp;</td>
<tr>
<td>Helios Gene Gun</td>
<td>&nbsp;</td>
<td>Bio-Rad</td>
<td>165-2431</td>
<td>&nbsp;</td>
<tr>
<td>Isopore membrane filter paper</td>
<td>&nbsp;</td>
<td>Millipore</td>
<td>TSTP02500</td>
<td>3.0 &mu;m pore size</td>
</tr>
<tr>
<td>ProLong Gold Antifade</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>P36930</td>
<td>&nbsp;</td>
<tr>
<td>Paraformaldehyde</td>
<td>&nbsp;</td>
<td>Alfa Aesar</td>
<td>30525-89-4</td>
<td>(16% w/v)</td>
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diolistic labeling of neurons from rodent and non-human primate brain slices

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; Gan et al., 2000). Here we provide a detailed description of each step required together with exemplary images of good and bad outcomes that will help when setting up the technique. In our experience, a few steps proved critical for the successful application of DiOLISTICS. These considerations include the quality of the DiI-coated bullets, the extent of fixative exposure, and the concentration of detergent used in the incubation solutions. Tips and solutions for common problems are provided. This is a versatile labeling technique that can be applied to multiple animal species at a wide range of ages. Unlike other fluorescent labeling techniques that are limited to preparations from young animals or restricted to mice because they rely on the expression of a fluorescent transgene, DiOLISTIC labeling can be applied to animals of all ages, species and genotypes and it can be used in combination with immunostaining to identify a specific subpopulation of cells. Here we demonstrate the use of DiOLISTICS to label neurons in brain slices from adult mice and adult non-human primates with the purpose of quantifying dendrite branching and dendritic spine morphology.

Watch this Scientific Journal Video about DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices at JoVE.com

DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices

We demonstrate the use of the gene gun to introduce fluorescent dyes, such as DiI, into neurons in brain slices from rodents and non-human primates of different ages. In this particular case, we use adult mice (3-6 months old) and adult cynomologus monkeys (9-15 years old). This technique, originally described by the laboratory of Dr. Lichtman (Gan et al., 2000), is well suited for the study of dendritic branching and dendritic spine morphology and can be combined with traditional immunostaining, if detergents are kept at a low concentration.

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; ...

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Research

JoVE Journal

Neuroscience

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Imaging Calcium Responses in GFP-tagged Neurons of Hypothalamic Mouse Brain Slices

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the precise molecular steps as well as the activity of specific neurons in multi-functional nuclei of brain areas such as the hypothalamus remain. This problem includes identification of the molecular components involved in the regulation of various neurohormone signal transduction cascades. Elevations of intracellular Ca2+ play an important role in regulating the sensitivity of neurons, both at the level of signal transduction and at synaptic sites.
New tools have emerged to help identify neurons in the myriad of brain neurons by expressing green fluorescent protein (GFP) under the control of a particular promoter. To monitor both spatially and temporally stimulus-induced Ca2+ responses in GFP-tagged neurons, a non-green fluorescent Ca2+ indicator dye needs to be used. In addition, confocal microscopy is a favorite method of imaging individual neurons in tissue slices due to its ability to visualize neurons in distinct planes of depth within the tissue and to limit out-of-focus fluorescence. The ratiometric Ca2+ indicator fura-2 has been used in combination with GFP-tagged neurons1. However, the dye is excited by ultraviolet (UV) light. The cost of the laser and the limited optical penetration depth of UV light hindered its use in many laboratories. Moreover, GFP fluorescence may interfere with the fura-2 signals2. Therefore, we decided to use a red fluorescent Ca2+ indicator dye. The huge Stokes shift of fura-red permits multicolor analysis of the red fluorescence in combination with GFP using a single excitation wavelength. We had previously good results using fura-red in combination with GFP-tagged olfactory neurons3. The protocols for olfactory tissue slices seemed to work equally well in hypothalamic neurons4. Fura-red based Ca2+ imaging was also successfully combined with GFP-tagged pancreatic &beta;-cells and GFP-tagged receptors expressed in HEK cells5,6. A little quirk of fura-red is that its fluorescence intensity at 650 nm decreases once the indicator binds calcium7. Therefore, the fluorescence of resting neurons with low Ca2+ concentration has relatively high intensity. It should be noted, that other red Ca2+-indicator dyes exist or are currently being developed, that might give better or improved results in different neurons and brain areas.

In this protocol, we update recent progress in imaging Ca2+ signals of GFP-tagged neurons in brain tissue slices using a red fluorescent Ca2+ indicator dye.

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Growing Neural Stem Cells from Conventional and Nonconventional Regions of the Adult Rodent Brain

Recent work demonstrates that central nervous system (CNS) regeneration and tumorigenesis involves populations of stem cells (SCs) resident within the adult brain. However, the mechanisms these normally quiescent cells employ to ensure proper functioning of neural networks, as well as their role in recovery from injury and mitigation of neurodegenerative processes are little understood. These cells reside in regions referred to as &#34;niches&#34; that provide a sustaining environment involving modulatory signals from both the vascular and immune systems. The isolation, maintenance, and differentiation of CNS SCs under defined culture conditions which exclude unknown factors, makes them accessible to treatment by pharmacological or genetic means, thus providing insight into their in vivo behavior. Here we offer detailed information on the methods for generating cultures of CNS SCs from distinct regions of the adult brain and approaches to assess their differentiation potential into neurons, astrocytes, and oligodendrocytes in vitro. This technique yields a homogeneous cell population as a monolayer culture that can be visualized to study individual SCs and their progeny. Furthermore, it can be applied across different animal model systems and clinical samples, being used previously to predict regenerative responses in the damaged adult nervous system.

Neural stem cells harvested from the adult brain are increasing utilized in applications ranging from the basic research of nervous system development to exploring potential clinical applications in regenerative medicine. This makes rigorous control in the isolation and culturing conditions used to grow these cells critical to sound experimental outcomes.

Recent work demonstrates that central nervous system (CNS) regeneration and tumorigenesis involves populations of stem cells (SCs) resident within the ...

Isolation, Culture and Long-Term Maintenance of Primary Mesencephalic Dopaminergic Neurons From Embryonic Rodent Brains

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes involved in physiological functions and vulnerability and death of these neurons is imparative to understanding the underlying causes and unraveling the cure for this common neurodegenerative disorder. Primary cultures of mesDA neurons provide a tool for investigation of the molecular, biochemical and electrophysiological properties, in order to understand the development, long-term survival and degeneration of these neurons during the course of disease. Here we present a detailed method for the isolation, culturing and maintenance of midbrain dopaminergic neurons from E12.5 mouse (or E14.5 rat) embryos. Optimized cell culture conditions in this protocol result in presence of axonal and dendritic projections, synaptic connections and other neuronal morphological properties, which make the cultures suitable for study of the physiological, cell biological and molecular characteristics of this neuronal population.

The causes of degeneration of midbrain dopaminergic neurons during Parkinson&#8217;s disease are not fully understood. Cellular culture systems provide an essential tool for study of the neurophysiological properties of these neurons. Here we present an optimized protocol, which can be utilized for in vitro modeling of neurodegeneration.

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes ...

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and characterize ultrastructural and morphological details of neurons, such as synaptic contacts. Good preservation of brain tissue for electron microscopy can be obtained by rigorous cryo-fixation methods, but these techniques are rather costly and limit the use of immunolabeling, which is crucial to understand the connectivity of identified neuronal systems. Freeze-substitution methods have been developed to allow the combination of cryo-fixation with immunolabeling. However, the reproducibility of these methodological approaches usually relies on costly freezing devices. Moreover, achieving reliable results with this technique is very time-consuming and skill-challenging. Hence, the traditional chemically fixed brain, particularly with acrolein fixative, remains a time-efficient and low-cost method to combine electron microscopy with immunohistochemistry. Here, we provide a reliable experimental protocol using chemical acrolein fixation that leads to the preservation of primate brain tissue and is compatible with pre-embedding immunohistochemistry and transmission electron microscopic examination.

Here, we provide an easy, low-cost, and time-efficient protocol to chemically fix primate brain tissue with acrolein fixative, allowing for long-term preservation that is compatible with pre-embedding immunohistochemistry for transmission electron microscopy.

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and ...

Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a challenging and low-throughput technique due to its complexity and high reliance on user operation and control. This manuscript demonstrates an image-guided automatic patch clamp system for in vitro whole-cell patch clamp experiments in acute brain slices. Our system implements a computer vision-based algorithm to detect fluorescently labeled cells and to target them for fully automatic patching using a micromanipulator and internal pipette pressure control. The entire process is highly automated, with minimal requirements for human intervention. Real-time experimental information, including electrical resistance and internal pipette pressure, are documented electronically for future analysis and for optimization to different cell types. Although our system is described in the context of acute brain slice recordings, it can also be applied to the automated image-guided patch clamp of dissociated neurons, organotypic slice cultures, and other non-neuronal cell types.

This protocol describes how to conduct automatic image-guided patch-clamp experiments using a system recently developed for standard in vitro electrophysiology equipment.

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is tightly regulated during development and neuronal maturation. Importantly, deficits in synapse number can lead to cognitive dysfunction. Therefore, the evaluation of synapse number is an integral part of neurobiology. However, as synapses are small and highly compact in the intact brain, the assessment of absolute number is challenging. This protocol describes a method to easily identify and evaluate synapses in hippocampal rodent slices using immunofluorescence microscopy. It includes a three-step procedure to evaluate synapses in high-quality confocal microscopy images by analyzing the co-localization of pre- and postsynaptic proteins in hippocampal slices. It also explains how the analysis is performed and gives representative examples from both excitatory and inhibitory synapses. This protocol provides a solid foundation for the analysis of synapses and can be applied to any research investigating the structure and function of the brain.

A protocol for accurately identifying and analyzing synapses in hippocampal slices using immunofluorescence is outlined in this article.

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is ...

Isolation and RNA Extraction of Neurons, Macrophages and Microglia from Larval Zebrafish Brains

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is important to isolate these cells without changing their gene expression profile. The zebrafish model provides a large number of transgenic fish lines in which specific cell types are labelled; for example neurons in the NBT:DsRed line or macrophages/microglia in the mpeg1:eGFP line. Furthermore, antibodies have been developed to stain specific cells, such as microglia with the 4C4 antibody.
Here, we describe the isolation of neurons, macrophages and microglia from larval zebrafish brains. Central to this protocol is the avoidance of an enzymatic tissue digestion at 37 &#176;C, which could modify cellular profiles. Instead a mechanical system of tissue homogenization at 4 &#176;C is used. This protocol entails homogenization of brains into cell suspension, their immuno-staining and the isolation of neurons, macrophages and microglia by FACS. Afterwards, we extracted RNA from those cells and evaluated their quality/quantity. We managed to obtain RNA of high quality (RNA Integrity Number (RIN) &#62; 7) to perform qPCR on macrophages/microglia and neurons, and transcriptomic analysis on microglia. This approach enables a better characterization of these cells, as well as a clearer understanding about their role in development and pathologies.

We present a protocol to isolate neurons, macrophages and microglia from larval zebrafish brains under physiological and pathological conditions. Upon isolation, RNA is extracted from these cells to analyze their gene expression profile. This protocol allows for the collection of high-quality RNA for performing downstream analysis like qPCR and transcriptomics.

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is ...

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) is a promising approach when pharmacological treatment fails or neurosurgery is not recommended. Acute brain slices coupled to microelectrode arrays (MEAs) represent a valuable tool to study neuronal network interactions and their modulation by electrical stimulation. As compared to conventional extracellular recording techniques, they provide the added advantages of a greater number of observation points and a known inter-electrode distance, which allow studying the propagation path and speed of electrophysiological signals. However, tissue oxygenation may be greatly impaired during MEA recording, requiring a high perfusion rate, which comes at the cost of decreased signal-to-noise ratio and higher oscillations in the experimental temperature. Electrical stimulation further stresses the brain tissue, making it difficult to pursue prolonged recording/stimulation epochs. Moreover, electrical modulation of brain slice activity needs to target specific structures/pathways within the brain slice, requiring that electrode mapping be easily and quickly performed live during the experiment. Here, we illustrate how to perform the recording and electrical modulation of 4-aminopyridine (4AP)-induced epileptiform activity in rodent brain slices using planar MEAs. We show that the brain tissue obtained from mice outperforms rat brain tissue and is thus better suited for MEA experiments. This protocol guarantees the generation and maintenance of a stable epileptiform pattern that faithfully reproduces the electrophysiological features observed with conventional field potential recording, persists for several hours, and outlasts sustained electrical stimulation for prolonged epochs. Tissue viability throughout the experiment is achieved thanks to the use of a small-volume custom recording chamber allowing for laminar flow and quick solution exchange even at low (1 mL/min) perfusion rates. Quick MEA mapping for real-time monitoring and selection of stimulating electrodes is performed by a custom graphic user interface (GUI).

We illustrate how to perform recording and electrical modulation of 4-aminopyridine-induced epileptiform activity in rodent brain slices using microelectrode arrays. A custom recording chamber maintains tissue viability throughout prolonged experimental sessions. Live electrode mapping and selection of stimulating pairs are performed by a custom graphical user interface.

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) ...