This work was supported by the NIDA Intramural Research Program (Bruce T. Hope, Yavin Shaham). F.J.R. was supported by an appointment to the NIDA Research Participation Program sponsored by the National Institutes of Health and administered by the Oak Ridge Institute for Science and Education, and received additional financial support from a Becas-Chile scholarship managed by CONICYT and the Universidad de los Andes, Santiago, Chile. The Johns Hopkins FACS Core facility was supported by Award P30AR053503 from the National Institute of Arthritis and Musculoskeletal and Skin Diseases of the National Institute of Health.

All experiments were performed in accordance with the Institutional Animal Care and Use Committee (IACUC) of the National Institutes of Health Guide for the Care and Use of Laboratory animals 16.
Note: All the steps below use low-binding centrifuge tubes that were kept on ice unless otherwise specified.
1. Preparation Before Tissue Collection
<ol>
	<li>Set the centrifuge to 4 oC.</li>
	<li>Fire polish a set of three glass Pasteur pipettes with decreasing diameters of approximately 1.3, 0.8, and 0.4 mm for each sample.</li>
	<li>Prepare labeled 1.7 ml-tubes containing 1 ml cold Buffer A and keep the tubes on ice.</li>
	<li>Prepare an ice tray containing the brain slicing matrix, spatulas and glass plates (or inverted glass Petri dish) on which to perform the dissection. Pre-chill two or more razor blades on the glass plates and use tissues paper to dry all instruments that touch the tissue.</li>
	<li>Thaw the Enzyme solution at room temperature (RT) for 30 min before tissue collection.</li>
</ol>
2. Tissue Collection and Dissection
<ol>
	<li>Initiate the behavioral or pharmacological treatment 90 min before tissue collection. 
	NOTE: For the results shown here, acute intraperitoneal injections of 5 mg/kg methamphetamine on rats in a novel environment (test condition) and na&#239;ve rats kept in their home cages (control condition) were performed.</li>
	<li>Anesthetize the rat by placing it in a glass desiccator jar with saturated isoflurane and decapitate the rat 30 - 60 sec later using a guillotine. 
	Use scissors to remove the skin and muscle from the head and expose the skull. Use rongeurs to cut the skull and open up the foramen magnum and remove the back part of the skull. Use rongeurs to cut along the top edges of the skull to expose the brain. Be careful not to damage the brain. Use a small spatula to gently scoop under and elevate the brain. Raise the brain and cut the nerves until the brain is free 17.</li>
	<li>Dissect tissue using razor blades. 
	NOTE: Obtain tissue samples using either of the following methods: (2.3.1) freshly dissected tissue; (2.3.2) frozen tissue after dissection; or (2.3.3) frozen tissue dissected from frozen whole brain. Keep the brain slicing matrix, razor blades and glass plate dry throughout the dissection and mincing processes as condensed water can lead to hypotonic lysing of cells. Use magnifying glasses to ensure accurate dissection.
	<ol>
		<li>For freshly dissected tissue, place the freshly extracted brain into a brain slicing matrix (a rat brain-shaped metal mold with slots for the razor blades at 1 mm intervals) that has been chilled on ice. Insert two or more pre-chilled razor blades into the slots to cut coronal slices that contain the brain region(s) of interest. Place the cut slice on the chilled glass plate. 
		Note: Dissect the brain regions of interest on a chilled glass plate.</li>
		<li>For frozen tissue after dissection, dissect the brain region(s) of interest from slices of freshly extracted brain, similar to that described above in 2.3.1. Place the dissected tissue into a microtube and rapidly freeze the tissue by submerging the microtube in -40 &#186;C isopentane for 20 sec. Keep dissected tissue frozen at all times in a -80 &#186;C freezer until further processing.</li>
		<li>For frozen tissue, immediately freeze the freshly extracted brain in -40 &#186;C isopentane and store in a sealed bag at -80 &#186;C for up to 6 months.
		<ol>
			<li>On the day of dissection, place the frozen brain in a cryostat set at approximately -20 &#186;C (no warmer than -18 &#186;C) for approximately 2 hr to equilibrate the temperature. 
			NOTE: Brains can be cut easily at this temperature, while much cooler temperatures make the brain too brittle to be cut safely.</li>
			<li>Use razor blades to cut 1 - 2 mm coronal slices of the frozen brains in a cryostat. Use a blunted 12- to 16-gauge needle to obtain tissue punches from these slices under freezing conditions. Keep dissected tissue frozen at all times until further processing.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Add 1 - 2 drops of Buffer A to cover the dissected tissue before mincing. Remove most of the white matter from the tissue using two razor blades to prevent loss of neurons during the rest of the trituration process. If starting with frozen tissue, then allow the tissue to thaw on the cold glass plate for no more than 1 min before applying Buffer A solution.</li>
	<li>Mince the tissue 100 times in each orthogonal direction with a razor blade on a chilled glass plate. Hold the razor blade vertical to the glass plate when mincing. 
	NOTE: Thorough mincing is critical for all protocol steps.</li>
	<li>Use razor blades to transfer the minced tissue into microcentrifuge tubes containing 1 ml of cold Buffer A. Invert the tube 3 - 5 times to keep all minced tissue in the solution.</li>
</ol>
3. Cell Dissociation
NOTE: Use gentle end over end mixing for all of the following steps. Do not use a vortex mixer and avoid air bubbles that cause cell damage.
<ol>
	<li>Centrifuge the sample tubes at 110 x g for 2 min at 4 &#186;C. Discard supernatant. Slowly add 1 ml of cold freshly thawed Enzyme solution (containing a mix of proteolytic enzymes) down the inner wall of the microtube. Immediately suck up the entire pellet and gently pipette up and down only 4 times with a moderately large tip diameter pipette to disperse the minced tissue pellet.</li>
	<li>Invert the microtubes immediately to prevent the pellet from sticking to the bottom and incubate the samples with end-over-end mixing for 30 min at 4 oC.</li>
	<li>After enzymatic tissue digestion, centrifuge the tubes at 960 x g for 2 min at 4 &#186;C. Remove the supernatant and add 0.6 ml of cold Buffer A. Immediately after adding the Buffer A, use the same pipette tip to disperse the pellet by sucking up the entire pellet and gently pipette up and down 5 times.</li>
	<li>Mechanically triturate the digested tissue and collect in 15 ml tubes using the following steps. For each sample, use a separate set of 3 fire-polished glass Pasteur pipettes with descending diameters of approximately 1.3, 0.8 and 0.4 mm attached to latex bulbs.
	<ol>
		<li>Gently triturate each sample 10 times with the 1.3 mm glass pipette. Let the sample settle for 2 min on ice (the debris and undissociated cells will settle to the bottom). Collect the supernatant (~ 0.6 ml) and transfer to a 15 ml conical tube (tube #1).</li>
		<li>Add 0.6 ml Buffer A solution to the remaining pellet. Triturate the sample 10 times with the 0.8 mm glass pipette. Let the sample settle for 2 min on ice. Collect the supernatant (~ 0.6 ml) and transfer to the same 15 ml conical tube #1.</li>
		<li>Add 0.6 ml Buffer A solution to the remaining pellet. Triturate the sample 10 times with the 0.4 mm glass pipette. Let the sample settle for 2 min on ice. Collect the supernatant (~ 0.6 ml; without touching the pellet with non-dissociated cells) and transfer to the same 15 ml conical tube #1. 
		NOTE: Keep the 0.4 mm diameter pipette in the tube #1 for the following trituration steps.</li>
		<li>Repeat step 3.4.3 three more times using the smallest glass pipette (~ 0.4 mm diameter), and collect the supernatant into a separate 15 ml conical tube (tube #2).</li>
		<li>Triturate the collected cell suspensions in tubes #1 and #2 for 10 more times using the 0.4 mm glass pipette and keep on ice.</li>
	</ol>
	</li>
</ol>
4. Cell Fixation and Permeabilization
<ol>
	<li>Prepare 4 microtubes per sample (two microtubes for cells from tube #1 and two for cells from tube #2) by adding 800 &#181;l of 100% cold ethanol (stored at -20 oC before use) into each tube and keep them on ice. Note: The final ethanol concentration will be 50%.</li>
	<li>Pipette ~ 800 &#181;l of the cell suspensions (from tube#1 and tube#2) into each of the 4 tubes containing cold ethanol and invert the tubes to mix the samples.</li>
	<li>Incubate tubes on ice for 15 min while inverting the tubes every 5 min.</li>
	<li>After fixation/permeabilization, centrifuge the tubes at 1,700 x g for 4 min at 4 &#186;C and discard the supernatant. Leave 50 &#181;l of solution to prevent cell loss. 
	NOTE: The pellets at this stage are white and stick less to the wall of the tubes than before permeabilization. Therefore, remove the supernatant carefully by touching the wall of the microtube where there is no pellet and draw up the supernatant slowly using a micropipette.</li>
</ol>
5. Cell Filtration
<ol>
	<li>Re-suspend the pellets from step 4.4 with cold PBS as follows:
	<ol>
		<li>Add 550 &#956;l of cold PBS to one of the two microtubes coming from tube #1, and use a moderately large diameter pipette tip to gently pipette the cell suspension up and down 5 times. Repeat the same for one microtube coming from tube #2.</li>
		<li>For cells from tube #1, using the same pipette, transfer the first cell suspension to the second pellet. Resuspend the second pellet by gently pipetting the combined cell suspension up and down 5 times.</li>
		<li>For cells from tube #2, resuspend and combine the pellets (i.e., third and fourth microtubes) as described in 5.1.2.</li>
	</ol>
	</li>
	<li>Filter the two cell suspensions from steps 5.1.2 and 5.1.3 separately using two different pairs of cell strainers (100 &#181;m and 40 &#181;m pore sizes) as follows:
	<ol>
		<li>Pre-wet each cell strainer by adding PBS to the inside of the strainer. Use a pipette to remove the excess PBS from the outside of the strainer.</li>
		<li>Pipette the cell suspension from step 5.1.2 evenly onto the first cell strainer with 100 &#181;m pore size. Collect the flow-through in a 50 ml tube on ice. Use the pipette to collect the flow- through attached on the outer side bottom of the cell strainer.</li>
		<li>Use the same pipette to transfer the flow-through from the 100 &#956;m cell strainer to the 40 &#956;m cell strainer. Collect this flow-through in another 50 ml tube on ice. When transferring the flow through from 40 &#956;m cell strainer, change to new pipette tips to avoid contamination from 100 &#956;m cell strainer.</li>
		<li>Repeat these steps for the cell suspension from step 5.1.3 using a new set of filters.</li>
	</ol>
	</li>
	<li>Combine the two flow-through from each sample for a final volume of approximately 1 ml per sample.</li>
</ol>
6. Incubation with Antibodies
NOTE: Use small aliquots of the brain cell suspensions to prepare multiple control samples for setting appropriate flow cytometer settings prior to running the main sample. For each antibody labeling, prepare control samples that include no antibodies, only the secondary antibody (or other fluorescent label), and then both primary and secondary antibodies (or other fluorescent label). Use these controls to set the gates that separate specific versus non-specific labeling in the flow cytometer. When possible, use fluorochromes that do not require compensation. Increase the final volume of the gating control samples to a final volume of 700 &#181;l by adding PBS prior to adding the primary antibodies.
<ol>
	<li>Prepare light-scattering and DAPI control sample by transferring a 100 &#181;l sample of the filtered cells to a 1.7 ml microtube. Add 600 &#181;l of cold PBS with 1 &#181;g/ml DAPI for nuclei staining. Incubate for 10 min with end-over-end mixing, and wash (as indicated in steps 6.4.2 to 6.4.5) before passing through the flow cytometer. 
	NOTE: This sample is used only to set the cell/nuclei gate prior to sorting the remaining cells in the FACS machine.</li>
	<li>Prepare single immunolabeled control samples by transferring 100 &#181;l of the filtered cells to a 1.7 ml microtube. Add 600 &#181;l of cold PBS with one antibody (e.g., NeuN antibody). Add 600 &#181;l of cold PBS to a microtube with another antibody (e.g., Fos antibody). Add the corresponding secondary antibody if it is not a direct-conjugated primary antibody. 
	NOTE: These samples are used to determine the appropriate photomultiplier tube (PMT) voltage, and to assess the overlap of the two fluorescent signals if necessary. Antibody concentrations should be titrated to achieve the best signal-to-noise ratios in all channels. Overlapping fluorescent signals can also be compensated by internal calibration of different fluorescent channels within the flow cytometer.</li>
	<li>Prepare double fluorescent-labeled negative control sample by transferring 100 &#181;l of the filtered cells to a 1.7 ml microtube. Add 600 &#181;l of cold PBS with the secondary antibodies alone or IgG direct-conjugated primary antibodies. 
	NOTE: This control sample can be used every time before sorting to determine the background fluorescence for each fluorophore.</li>
	<li>Prepare main sample to be sorted.
	<ol>
		<li>Transfer 700 &#956;l of the filtered cells into a 1.7 ml microtube. Add 7 &#181;l (1:100) of primary Fos antibody conjugated directly to Alexa Fluor 647 (anti-Fos-AF647) and 1.4 &#181;l (1:500) of primary NeuN antibody conjugated directly to Phycoerythrin (anti-NeuN-PE). Incubate tubes for 30 min at 4 &#186;C with end-over-end mixing.</li>
		<li>At the end of the incubation, add 800 &#181;l of cold PBS to each microtube, including the main sample and the control samples, and mix by inversion.</li>
		<li>Centrifuge microtubes at 1,300 x g for 3 min at 4 &#186;C. Discard the supernatant, leaving 50 &#181;l supernatant to prevent cell loss.</li>
		<li>Add 1 ml of cold PBS to the pellet and resuspend cells using a pipette with a moderately large tip diameter.</li>
		<li>Centrifuge at 1,300 x g for 3 min at 4 &#186;C. Discard the supernatant.</li>
		<li>Re-suspend the pellet in 500 &#181;l cold PBS (adjust final volume depending the size of the pellet).</li>
		<li>Transfer and filter the suspension into a round-bottom FACS sample tube with a cell strainer cap (40 &#181;m). Keep the immunolabeled cells on ice and immediately perform FACS. Note: For filtering, touch the pipette tip vertically on the strainer cap and push the cell suspension through it.</li>
	</ol>
	</li>
	<li>Set flow cytometer gates using control samples.
	<ol>
		<li>Use the light-scattering and DAPI control sample to identify the neuronal cell population from the total events and cellular debris. 
		NOTE: Cell bodies in this preparation are only 1 - 2% of the total events (the rest are debris and broken cellular processes). Based on DAPI fluorescent signal (indicating nuclei) and Forward scatter (size of the cell) it is possible to locate cell bodies and gate backwards based on the Forward and Side light-scattering properties of the cells (as shown in Figure 1 A,B).</li>
		<li>Use the single immunolabeled control samples to determine the PMT voltage settings, and to analyze spillover of emission from one specific fluorochrome into another filter/channel (e.g., Alexa Fluor 488/FITC or Phycoerythrine). If fluorescent signals overlap, correct the overlap by adjusting compensation levels manually.</li>
		<li>Use the double fluorescent-labeled negative control to set up the threshold for the positive population. 
		NOTE: The signal coming from this sample is thought to be due to auto-fluorescence and the cells of the main sample located above this threshold can be considered as positive. Typically, the threshold parameters for sorting are more stringent and approximately the top 2/3 of the labeled cells above the negative control are actually sorted.</li>
	</ol>
	</li>
	<li>Sort the neurons from the main sample by FACS into 1.7 ml microtubes. Collect the cells into 50 &#181;l of RNA extraction buffer. After sorting, vortex and centrifuge the tube and mix the suspension by pipetting up and down 10 times to recover all sorted cells from the walls of the tube.</li>
	<li>Incubate the sorted cell suspension at 42 &#186;C for 30 min to ensure that all cells are lysed prior to RNA extraction, and then centrifuge at 2,800 x g for 2 min at 4 &#186;C.</li>
	<li>Transfer the supernatant to an RNase-free tube for long-term storage at -80 &#186;C or immediately process the sample for RNA isolation, cDNA synthesis, target gene pre-amplification and qPCR, as described previously 13-15.</li>
</ol>

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The authors declare that they have no competing financial interests.

FACS can be used to sort neurons and other cell types from either fresh or frozen adult brain tissue. As mentioned in the introduction, the ability to use frozen tissue allows optimal utilization of samples from animals that have undergone complex and protracted behavioral procedures, such as self-administration and relapse studies in addiction research. These behavioral procedures usually takes 1 - 2 hr or longer, and require all animals (10 - 20 total) be tested on the same day 13,18. It takes ~ 4 hr to proc...

During learning, animals form associations between complex sets of highly specific stimuli. This high-resolution information is thought to be encoded by alterations within specific patterns of sparsely distributed neurons called neuronal ensembles. Neuronal ensembles have recently been identified by the induction of immediately early genes (IEGs) such as Fos, Arc, and Zif268 and their protein products in neurons that were strongly activated during behavior or cue exposure. Fos-expressing neurons in particular have been shown to play causal roles in context and cue-specific learned behaviors 1-4. Thus, unique molecular neuroadaptations within these activated Fos-expressing neurons are top candidates for the neural mechanisms that encode learned associations formed during normal learning and abnormal learning disorders, such as addiction and post-traumatic stress disorder (PTSD) 5.
Fluorescence Activated Cell Sorting (FACS) has recently allowed analysis of unique molecular neuroadaptations within Fos-expressing neurons. Flow cytometry and cell sorting were developed in the 1960s 6,7 to characterize and isolate cells according to their light-scattering and immunofluorescent characteristics, and have long been used in immunology and cancer research. However flow cytometry and FACS requires dissociated single cells that are difficult to obtain from adult brain tissue. FACS was first used to isolate and analyze Green Fluorescent Protein (GFP)-expressing striatal neurons from transgenic mice that did not require antibody labeling 8,9. We developed an antibody-based FACS method 10 to isolate and assess molecular alterations in Fos-expressing neurons activated by drug and/or cues in wild type animals 11-15. In this method, neurons are labeled with an antibody against the general neuronal marker NeuN, while strongly activated neurons are labeled with an antibody against Fos. Although our initial method required pooling of up to 10 rats per sample for fresh tissue, subsequent modifications of the protocol allowed FACS isolation of Fos-expressing neurons and quantitative Polymerase Chain Reaction (qPCR) analysis of discrete brain areas from a single rat 13-15. Overall, unique molecular alterations were found in Fos-expressing neurons activated during a variety of context- and cue-activated behaviors in addiction research 12,14,15.
A major logistical problem with performing FACS on fresh tissue is that it takes one whole day to dissociate the tissue and process by FACS. In addition, only about four samples can be processed per day. This usually means that only one brain area can be assessed from each brain and the remaining brain areas have to be discarded. This is a major problem for low throughput behavioral procedures such as self-administration and extinction training that requires surgery and many weeks of intensive training. Furthermore, long and complicated behavioral procedures on test day makes it difficult to perform FACS on the same day. It would be a significant advantage to be able to freeze the brains from the animals immediately after behavioral testing, and then isolate Fos-expressing neurons from one or more brain areas at different times of the investigators&#39; choosing.
Here we demonstrate that our FACS protocol can be used to isolate Fos-expressing neurons (and other cell types) from both fresh and frozen brain tissue. As an example, we isolated Fos-expressing neurons from rat striatum after acute methamphetamine injections and from na&#239;ve rats without injections (control condition). However, this FACS protocol can be used following any behavioral or pharmacological treatment. Subsequent qPCR analysis of our samples indicated that gene expression from these cell types could be assessed with similar efficiency from both fresh and frozen tissue.

<table border="0" cellpadding="0" cellspacing="0" width="1274">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:455px;">Brain matrix</td>
			<td style="width:191px;">CellPoint Scientific</td>
			<td style="width:241px;">69-2160-1</td>
			<td style="width:388px;">to obtain coronal brain slices</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hibernate A low fluorescence&#160;</td>
			<td>Brain Bits</td>
			<td>HA-lf&#160;</td>
			<td>Buffer A&#39; in the protocol is used for processing tissue and cells from the dissociation to fixation steps</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Accutase&#160;</td>
			<td>Millipore</td>
			<td>SCR005</td>
			<td>Enzyme solution&#39; in the protocol is used for enzymatic digestion of tissue prior to trituration</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Protein LoBind Tube 1.5 mL, PCR clean</td>
			<td>Eppendorf</td>
			<td>22431081</td>
			<td>to prevent cell lost during the protocol</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cell Strainer, 40 &#181;m</td>
			<td>BD Falcon</td>
			<td>352340</td>
			<td>to filter cell suspension</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cell Strainer, 100 &#181;m</td>
			<td>BD Falcon</td>
			<td>352360</td>
			<td>to filter cell suspension</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Falcon 5 ml round-bottom polystyrene test tube with cell strainer snap cap</td>
			<td>BD Bioscience</td>
			<td>352235</td>
			<td>to filter cell suspension before passing though the flow cytometer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pasteur Pipet, Glass</td>
			<td>NIH supply</td>
			<td>6640-00-782-6008</td>
			<td>to do tissue trituration</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Milli-Mark Anti-NeuN-PE, Clone A60 (mouse monoclonal) antibody</td>
			<td>EMD Millipore</td>
			<td>FCMAB317PE</td>
			<td>antibody to detect neurons</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Phospho-c-Fos (Ser32) (D82C12) XP Rabbit (Alexa Fluor 647 Conjugate)&#160;</td>
			<td>Cell Signaling Technology&#160;</td>
			<td>8677</td>
			<td>antibody to detect Fos-expressing cells</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DAPI</td>
			<td>Sigma</td>
			<td>D8417</td>
			<td>to label nuclei</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PicoPure RNA Isolation Kit</td>
			<td>Applied Biosystems.</td>
			<td>KIT0204</td>
			<td>The kit includes the RNA extraction buffer for step 6.14. It is used to collect sorted cells</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Superscript III first strand cDNA synthesis system</td>
			<td>Invitrogen</td>
			<td>18080-051</td>
			<td>to synthesize cDNA from RNA</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">TaqMan PreAmp Master Mix</td>
			<td>Applied Biosystems.</td>
			<td>4391128</td>
			<td>to do targeted-preamplification from cDNA</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">TaqMan Advance Fast Master&#160;</td>
			<td>Applied Biosystems.</td>
			<td>4444963</td>
			<td>to do PCR using TaqMan probes</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fos TaqMan probe</td>
			<td>Applied Biosystems.</td>
			<td>Rn00487426_g1</td>
			<td>TaqMan probe/primers</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NeuN TaqMan probe</td>
			<td>Applied Biosystems.</td>
			<td>CACTCCAACAGCGTGAC</td>
			<td>nucleotide sequence for neuronal gene marker</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NeuN Forward primer</td>
			<td>Applied Biosystems.</td>
			<td>GGCCCCTGGCAGAAAGTAG</td>
			<td>nucleotide sequence for neuronal gene marker</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NeuN Reverse primer</td>
			<td>Applied Biosystems.</td>
			<td>TTCCCCCTGGTCCTTCTGA</td>
			<td>nucleotide sequence for neuronal gene marker</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gfap TaqMan probe</td>
			<td>Applied Biosystems.</td>
			<td>Rn00566603_m1</td>
			<td>TaqMan probe/primers for astrocityc gene marker</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">iba-1 TaqMan probe</td>
			<td>Applied Biosystems.</td>
			<td>Rn00574125_g1</td>
			<td>TaqMan probe/primers for microglya gene marker</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gapdh TaqMan probe</td>
			<td>Applied Biosystems.</td>
			<td>CTCATGACCACAGTCCA</td>
			<td>nucleotide sequence for reference/housekeeping gene</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gapdh Forward primer</td>
			<td>Applied Biosystems.</td>
			<td>GACAACTTTGGCATCGTGGAA</td>
			<td>nucleotide sequence for reference/housekeeping gene</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gapdh Reverse primer</td>
			<td>Applied Biosystems.</td>
			<td>CACAGTCTTCTGAGTGGCAGTGA</td>
			<td>nucleotide sequence for reference/housekeeping gene</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Oligo2 TaqMan probe</td>
			<td>Applied Biosystems.</td>
			<td>Rn01767116_m1&#160;</td>
			<td>TaqMan probe/primers for oligodendrocytic gene marker</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">P1000 pipettor</td>
			<td>Rainin</td>
			<td>17014382</td>
			<td>It is refered to as the pipette with a large tip diameter in steps 3.1 and 3.3 for mild tissue trituration and step 6.7 to resuspend cells</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">7500 Fast Real-time PCR system</td>
			<td>Applied Biosystems.</td>
			<td>446985</td>
			<td>for quantitative PCR</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">FACSAria I Cell Sorter</td>
			<td>BD Biosciences</td>
			<td></td>
			<td>for FACS sorting</td>
		</tr>
	</tbody>
</table>

fluorescence activated cell sorting (facs) and gene expression analysis of fos-expressing neurons from fresh and frozen rat brain tissue

The study of neuroplasticity and molecular alterations in learned behaviors is switching from the study of whole brain regions to the study of specific sets of sparsely distributed activated neurons called neuronal ensembles that mediate learned associations. Fluorescence Activated Cell Sorting (FACS) has recently been optimized for adult rat brain tissue and allowed isolation of activated neurons using antibodies against the neuronal marker NeuN and Fos protein, a marker of strongly activated neurons. Until now, Fos-expressing neurons and other cell types were isolated from fresh tissue, which entailed long processing days and allowed very limited numbers of brain samples to be assessed after lengthy and complex behavioral procedures. Here we found that yields of Fos-expressing neurons and Fos mRNA from dorsal striatum were similar between freshly dissected tissue and tissue frozen at -80 &#186;C for 3 - 21 days. In addition, we confirmed the phenotype of the NeuN-positive and NeuN-negative sorted cells by assessing gene expression of neuronal (NeuN), astrocytic (GFAP), oligodendrocytic (Oligo2) and microgial (Iba1) markers, which indicates that frozen tissue can also be used for FACS isolation of glial cell types. Overall, it is possible to collect, dissect and freeze brain tissue for multiple FACS sessions. This maximizes the amount of data obtained from valuable animal subjects that have often undergone long and complex behavioral procedures.

Sorting Fos-positive and Fos-negative neurons from fresh and frozen dorsal striatum tissue from single rats after acute methamphetamine injections.
The protocol described above was used to sort Fos-positive and Fos-negative neurons from a single rat dorsal striatum 90 min after an intraperitoneal injection of methamphetamine (5 mg/kg). Na&#239;ve rats in their home cages were used as controls. Dorsal striatum tissue was processe...

Watch this Scientific Journal Video about Fluorescence Activated Cell Sorting FACS and Gene Expression Analysis of Fos-expressing Neurons from Fresh and Frozen Rat Brain Tissue at JoVE.com

Fluorescence Activated Cell Sorting FACS and Gene Expression Analysis of Fos-expressing Neurons from Fresh and Frozen Rat Brain Tissue

Here we present a Fluorescence Activated Cell Sorting (FACS) protocol to study molecular alterations in Fos-expressing neuronal ensembles from both fresh and frozen brain tissue. The use of frozen tissue allows FACS isolation of many brain areas over multiple sessions to maximize the use of valuable animal subjects.

Fluorescence Activated Cell Sorting (FACS) and Gene Expression Analysis of Fos-expressing Neurons from Fresh and Frozen Rat Brain Tissue

The study of neuroplasticity and molecular alterations in learned behaviors is switching from the study of whole brain regions to the study of specific ...

fluorescence-activated-cell-sorting-facs-gene-expression-analysis-fos

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

29.1K Views.  NIDA, NIH, DHHS. Here we present a Fluorescence Activated Cell Sorting (FACS) protocol to study molecular alterations in Fos-expressing neuronal ensembles from both fresh and frozen brain tissue. The use of frozen tissue allows FACS isolation of many brain areas over multiple sessions to maximize the use of valuable animal subjects.

Video: Fluorescence Activated Cell Sorting FACS and Gene Expression Analysis of Fos-expressing Neurons from Fresh and Frozen Rat Brain Tissue

Double Fluorescence in situ Hybridization in Fresh Brain Sections

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh frozen brain sections. Our group has successfully used this approach to study gene co-regulation. More specifically, we have used this dFISH method to explore the anatomical organization, neurochemical properties, and the impact of sensory experience in central sensory circuits, at single cell resolution. This protocol has been validated in brain tissue from mice, rats and songbirds but is expected to be easily adaptable to other vertebrate species, as well as to an array of non-neural tissues. In this film we provide a detailed demonstration of the main steps of this procedure.

Double Fluorescence in situ Hybridization in Fresh Brain Sections

This protocol involves a non-radioactive in-situ hybridization procedure that enables the simultaneous identification of two transcript species, at a single cell resolution, in thin sections of the vertebrate brain.

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh ...

Optimized Analysis of DNA Methylation and Gene Expression from Small, Anatomically-defined Areas of the Brain

Exposure to diet, drugs and early life adversity during sensitive windows of life 1,2 can lead to lasting changes in gene expression that contribute to the display of physiological and behavioural phenotypes. Such environmental programming is likely to increase the susceptibility to metabolic, cardiovascular and mental diseases 3,4.
DNA methylation and histone modifications are considered key processes in the mediation of the gene-environment dialogue and appear also to underlay environmental programming 5. In mammals, DNA methylation typically comprises the covalent addition of a methyl group at the 5-position of cytosine within the context of CpG dinucleotides.
CpG methylation occurs in a highly tissue- and cell-specific manner making it a challenge to study discrete, small regions of the brain where cellular heterogeneity is high and tissue quantity limited. Moreover, because gene expression and methylation are closely linked events, increased value can be gained by comparing both parameters in the same sample.
Here, a step-by-step protocol (Figure 1) for the investigation of epigenetic programming in the brain is presented using the 'maternal separation' paradigm of early life adversity for illustrative purposes. The protocol describes the preparation of micropunches from differentially-aged mouse brains from which DNA and RNA can be simultaneously isolated, thus allowing DNA methylation and gene expression analyses in the same sample.

A streamlined workflow to study DNA methylation and gene expression changes upon early-life stress is shown. Starting from maternal separation of newborn mice and isolation of discrete brain tissues, we represent a protocol to simultaneously isolate DNA and RNA from brain tissue punches for subsequent bisulfite sequencing and RT-PCR analysis.

Exposure to diet, drugs and early life adversity during sensitive windows of life 1,2 can lead to lasting changes in gene expression that contribute to ...

An Optimized Procedure for Fluorescence-activated Cell Sorting (FACS) Isolation of Autonomic Neural Progenitors from Visceral Organs of Fetal Mice

During development neural crest (NC)-derived neuronal progenitors migrate away from the neural tube to form autonomic ganglia in visceral organs like the intestine and lower urinary tract. Both during development and in mature tissues these cells are often widely dispersed throughout tissues so that isolation of discrete populations using methods like laser capture micro-dissection is difficult. They can however be directly visualized by expression of fluorescent reporters driven from regulatory regions of neuron-specific genes like Tyrosine hydroxylase (TH). We describe a method optimized for high yields of viable TH+ neuronal progenitors from fetal mouse visceral tissues, including intestine and lower urogenital tract (LUT), based on dissociation and fluorescence-activated cell sorting (FACS). 
The Th gene encodes the rate-limiting enzyme for production of catecholamines. Enteric neuronal progenitors begin to express TH during their migration in the fetal intestine1 and TH is also present in a subset of adult pelvic ganglia neurons2-4 . The first appearance of this lineage and the distribution of these neurons in other aspects of the LUT, and their isolation has not been described. Neuronal progenitors expressing TH can be readily visualized by expression of EGFP in mice carrying the transgene construct Tg(Th-EGFP)DJ76Gsat/Mmnc1. We imaged expression of this transgene in fetal mice to document the distribution of TH+ cells in the developing LUT at 15.5 days post coitus (dpc), designating the morning of plug detection as 0.5 dpc, and observed that a subset of neuronal progenitors in the coalescing pelvic ganglia express EGFP. 
To isolate LUT TH+ neuronal progenitors, we optimized methods that were initially used to purify neural crest stem cells from fetal mouse intestine2-6. Prior efforts to isolate NC-derived populations relied upon digestion with a cocktail of collagenase and trypsin to obtain cell suspensions for flow cytometry. In our hands these methods produced cell suspensions from the LUT with relatively low viability. Given the already low incidence of neuronal progenitors in fetal LUT tissues, we set out to optimize dissociation methods such that cell survival in the final dissociates would be increased. We determined that gentle dissociation in Accumax (Innovative Cell Technologies, Inc), manual filtering, and flow sorting at low pressures allowed us to achieve consistently greater survival (&gt;70% of total cells) with subsequent yields of neuronal progenitors sufficient for downstream analysis. The method we describe can be broadly applied to isolate a variety of neuronal populations from either fetal or adult murine tissues.

An Optimized Procedure for Fluorescence-activated Cell Sorting FACS Isolation of Autonomic Neural Progenitors from Visceral Organs of Fetal Mice

An optimized procedure to purify neural crest-derived neuronal progenitors from fetal mouse tissues is described. This method takes advantage of expression from fluorescent reporter alleles to isolate discrete populations by fluorescence-activated cell sorting (FACS). The technique can be applied to isolate neuronal subpopulations throughout development or from adult tissues.

During development neural crest (NC)-derived neuronal progenitors migrate away from the neural tube to form autonomic ganglia in visceral organs like the ...

Laser Capture Microdissection of Enriched Populations of Neurons or Single Neurons for Gene Expression Analysis After Traumatic Brain Injury

Long-term cognitive disability after TBI is associated with injury-induced neurodegeneration in the hippocampus-a region in the medial temporal lobe that is critical for learning, memory and executive function.1,2 Hence our studies focus on gene expression analysis of specific neuronal populations in distinct subregions of the hippocampus. The technique of laser capture microdissection (LCM), introduced in 1996 by Emmert-Buck, et al.,3 has allowed for significant advances in gene expression analysis of single cells and enriched populations of cells from heterogeneous tissues such as the mammalian brain that contains thousands of functional cell types.4 We use LCM and a well established rat model of traumatic brain injury (TBI) to investigate the molecular mechanisms that underlie the pathogenesis of TBI. Following fluid-percussion TBI, brains are removed at pre-determined times post-injury, immediately frozen on dry ice, and prepared for sectioning in a cryostat. The rat brains can be embedded in OCT and sectioned immediately, or stored several months at -80 &deg;C before sectioning for laser capture microdissection. Additionally, we use LCM to study the effects of TBI on circadian rhythms. For this, we capture neurons from the suprachiasmatic nuclei that contain the master clock of the mammalian brain. Here, we demonstrate the use of LCM to obtain single identified neurons (injured and degenerating, Fluoro-Jade-positive, or uninjured, Fluoro-Jade-negative) and enriched populations of hippocampal neurons for subsequent gene expression analysis by real time PCR and/or whole-genome microarrays. These LCM-enabled studies have revealed that the selective vulnerability of anatomically distinct regions of the rat hippocampus are reflected in the different gene expression profiles of different populations of neurons obtained by LCM from these distinct regions. The results from our single-cell studies, where we compare the transcriptional profiles of dying and adjacent surviving hippocampal neurons, suggest the existence of a cell survival rheostat that regulates cell death and survival after TBI.

We describe how to use laser capture microdissection (LCM) to obtain enriched populations of hippocampal neurons or single neurons from frozen sections of the injured rat brain for subsequent gene expression analysis using quantitative real time PCR and/or whole-genome microarrays.

Long-term cognitive disability after TBI is associated with injury-induced neurodegeneration in the hippocampus-a region in the medial temporal lobe that ...

RNA-Seq Analysis of Differential Gene Expression in Electroporated Chick Embryonic Spinal Cord

In ovo electroporation of the chick neural tube is a fast and inexpensive method for identification of gene function during neural development. Genome wide analysis of differentially expressed transcripts after such an experimental manipulation has the potential to uncover an almost complete picture of the downstream effects caused by the transfected construct. This work describes a simple method for comparing transcriptomes from samples of transfected embryonic spinal cords comprising all steps between electroporation and identification of differentially expressed transcripts. The first stage consists of guidelines for electroporation and instructions for dissection of transfected spinal cord halves from HH23 embryos in ribonuclease-free environment and extraction of high-quality RNA samples suitable for transcriptome sequencing. The next stage is that of bioinformatic analysis with general guidelines for filtering and comparison of RNA-Seq datasets in the Galaxy public server, which eliminates the need of a local computational structure for small to medium scale experiments. The representative results show that the dissection methods generate high quality RNA samples and that the transcriptomes obtained from two control samples are essentially the same, an important requirement for detection of differential expression genes in experimental samples. Furthermore, one example is provided where experimental overexpression of a DNA construct can be visually verified after comparison with control samples. The application of this method may be a powerful tool to facilitate new discoveries on the function of neural factors involved in spinal cord early development.

This video shows the basic steps for performing whole transcriptome analysis on dissected chick embryonic spinal cord samples after transfection with in ovo electroporation.

In ovo electroporation of the chick neural tube is a fast and inexpensive method for identification of gene function during neural development. Genome ...

Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective treatment for several movement disorders that have failed to respond to medication. Recent progress in the field of DBS surgery has begun to extend the application of this surgical technique to other conditions as diverse as morbid obesity, depression and obsessive compulsive disorder. Despite these expanding indications, little is known about the underlying physiological mechanisms that facilitate the beneficial effects of DBS surgery. One approach to this question is to perform gene expression analysis in neurons that receive the electrical stimulation. Previous studies have shown that neurogenesis in the rat dentate gyrus is elicited in DBS targeting of the anterior nucleus of the thalamus1. DBS surgery targeting the ATN is used widely for treatment refractory epilepsy. It is thus of much interest for us to explore the transcriptional changes induced by electrically stimulating the ATN. In this manuscript, we describe our methodologies for stereotactically-guided DBS surgery targeting the ATN in adult male Wistar rats. We also discuss the subsequent steps for tissue dissection, RNA isolation, cDNA preparation and quantitative RT-PCR for measuring gene expression changes. This method could be applied and modified for stimulating the basal ganglia and other regions of the brain commonly clinically targeted. The gene expression study described here assumes a candidate target gene approach for discovering molecular players that could be directing the mechanism for DBS.

The mechanism underlying the therapeutic effects of Deep Brain Stimulation (DBS) surgery needs investigation. The methods presented in this manuscript describe an experimental approach to examine the cellular events triggered by DBS by analyzing the gene expression profile of candidate genes that can facilitate neurogenesis post DBS surgery.

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective ...

Using Fluorescence Activated Cell Sorting to Examine Cell-Type-Specific Gene Expression in Rat Brain Tissue

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant cell type in the brain, neurons are the most widely studied of these cell types given their direct role in impacting behaviors. Other cell types in the brain also impact neuronal function and behavior via the signaling molecules they produce. Neuroscientists must understand the interactions between the cell types in the brain to better understand how these interactions impact neural function and disease. To date, the most common method of analyzing protein or gene expression utilizes the homogenization of whole tissue samples, usually with blood, and without regard for cell type. This approach is an informative approach for examining general changes in gene or protein expression that may influence neural function and behavior; however, this method of analysis does not lend itself to a greater understanding of cell-type-specific gene expression and the effect of cell-to-cell communication on neural function. Analysis of behavioral epigenetics has been an area of growing focus which examines how modifications of the deoxyribonucleic acid (DNA) structure impact long-term gene expression and behavior; however, this information may only be relevant if analyzed in a cell-type-specific manner given the differential lineage and thus epigenetic markers that may be present on certain genes of individual neural cell types. The Fluorescence Activated Cell Sorting (FACS) technique described below provides a simple and effective way to isolate individual neural cells for the subsequent analysis of gene expression, protein expression, or epigenetic modifications of DNA. This technique can also be modified to isolate more specific neural cell types in the brain for subsequent cell-type-specific analysis.

The goal of this protocol is to use the fluorescence activated cell sorting (FACS) technique to sort specific types of neural cells for subsequent analysis of cell-type-specific gene expression, epigenetic markers, and or protein expression.

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant ...

Isolation of Cerebral Capillaries from Fresh Human Brain Tissue

Understanding blood-brain barrier function under physiological and pathophysiological conditions is critical for the development of new therapeutic strategies that hold the promise to enhance brain drug delivery, improve brain protection, and treat brain disorders. However, studying the human blood-brain barrier function is challenging. Thus, there is a critical need for appropriate models. In this regard, brain capillaries isolated from human brain tissue represent a unique tool to study barrier function as close to the human in vivo situation as possible. Here, we describe an optimized protocol to isolate capillaries from human brain tissue at a high yield and with consistent quality and purity. Capillaries are isolated from fresh human brain tissue using mechanical homogenization, density-gradient centrifugation, and filtration. After the isolation, the human brain capillaries can be used for various applications including leakage assays, live cell imaging, and immune-based assays to study protein expression and function, enzyme activity, or intracellular signaling. Isolated human brain capillaries are a unique model to elucidate the regulation of the human blood-brain barrier function. This model can provide insights into central nervous system (CNS) pathogenesis, which will help the development of therapeutic strategies for treating CNS disorders.

Isolated brain capillaries from human brain tissue can be used as a preclinical model to study barrier function under physiological and pathophysiological conditions. Here, we present an optimized protocol to isolate brain capillaries from fresh human brain tissue.

Understanding blood-brain barrier function under physiological and pathophysiological conditions is critical for the development of new therapeutic ...

Optimization of Laser-Capture Microdissection for the Isolation of Enteric Ganglia from Fresh-Frozen Human Tissue

The purpose of this method is to obtain high-integrity RNA samples from enteric ganglia collected from unfixed, freshly-resected human intestinal tissue using laser capture microdissection (LCM). We have identified five steps in the workflow that are crucial for obtaining RNA isolates from enteric ganglia with sufficiently high quality and quantity for RNA-seq. First, when preparing intestinal tissue, each sample must have all excess liquid removed by blotting prior to flattening the serosa as much as possible across the bottom of large base molds. Samples are then quickly frozen atop a slurry of dry ice and 2-methylbutane. Second, when sectioning the tissue, it is important to position cryomolds so that intestinal sections parallel the full plane of the myenteric plexus, thereby yielding the greatest surface area of enteric ganglia per slide. Third, during LCM, polyethylene napthalate (PEN)-membrane slides offer the greatest speed and flexibility in outlining the non-uniform shapes of enteric ganglia when collecting enteric ganglia. Fourth, for distinct visualization of enteric ganglia within sections, ethanol-compatible dyes, like Cresyl Violet, offer excellent preservation of RNA integrity relative to aqueous dyes. Finally, for the extraction of RNA from captured ganglia, we observed differences between commercial RNA extraction kits that yielded superior RNA quantity and quality, while eliminating DNA contamination. Optimization of these factors in the current protocol greatly accelerates the workflow and yields enteric ganglia samples with exceptional RNA quality and quantity.

The goal of this protocol is to obtain high-integrity RNA samples from enteric ganglia isolated from unfixed, freshly-resected human intestinal tissue using laser capture microdissection (LCM). This protocol involves preparing flash-frozen samples of human intestinal tissue, cryosectioning, ethanolic staining and dehydration, LCM, and RNA extraction.

The purpose of this method is to obtain high-integrity RNA samples from enteric ganglia collected from unfixed, freshly-resected human intestinal tissue ...

Primary Cell Culture of Purified GABAergic or Glutamatergic Neurons Established through Fluorescence-activated Cell Sorting

The overall goal of this protocol is to generate purified neuronal cultures derived from either GABAergic or glutamatergic neurons. Purified neurons can be cultured in defined media for 16 days in vitro and are amenable to any analyses typically performed on dissociated cultures, including electrophysiological, morphological and survival analyses. The major advantage of these cultures is that specific cell types can be selectively studied in the absence of complex external influences, such as those arising from glial cells or other neuron types. When planning experiments with purified cells, however, it is important to note that neurons strongly depend on glia-conditioned media for their growth and survival. In addition, glutamatergic neurons further depend on glia-secreted factors for the establishment of synaptic transmission. We therefore also describe a method for co-culturing neurons and glial cells in a non-contact arrangement. Using these methods, we have identified major differences between the development of GABAergic and glutamatergic neuronal networks. Thus, studying cultures of purified neurons has great potential for furthering our understanding of how the nervous system develops and functions. Moreover, purified cultures may be useful for investigating the direct action of pharmacological agents, growth factors or for exploring the consequences of genetic manipulations on specific cell types. As more and more transgenic animals become available, labeling additional specific cell types of interest, we expect that the protocols described here will grow in their applicability and potential.

This protocol describes a cell sorting based method for the purification and culture of fluorescent GABAergic or glutamatergic neurons from the neocortex and hippocampus of postnatal mice or rats.

The overall goal of this protocol is to generate purified neuronal cultures derived from either GABAergic or glutamatergic neurons. Purified neurons can ...

Fluorescence Activated Cell Sorting (FACS) and Gene Expression Analysis of Fos-expressing Neurons from Fresh and Frozen Rat Brain Tissue

Summary

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Chapters in this video

Double Fluorescence in situ Hybridization in Fresh Brain Sections

Optimized Analysis of DNA Methylation and Gene Expression from Small, Anatomically-defined Areas of the Brain

An Optimized Procedure for Fluorescence-activated Cell Sorting (FACS) Isolation of Autonomic Neural Progenitors from Visceral Organs of Fetal Mice

Laser Capture Microdissection of Enriched Populations of Neurons or Single Neurons for Gene Expression Analysis After Traumatic Brain Injury

RNA-Seq Analysis of Differential Gene Expression in Electroporated Chick Embryonic Spinal Cord

Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

Using Fluorescence Activated Cell Sorting to Examine Cell-Type-Specific Gene Expression in Rat Brain Tissue

Isolation of Cerebral Capillaries from Fresh Human Brain Tissue

Optimization of Laser-Capture Microdissection for the Isolation of Enteric Ganglia from Fresh-Frozen Human Tissue

Primary Cell Culture of Purified GABAergic or Glutamatergic Neurons Established through Fluorescence-activated Cell Sorting