在大鼠脑组织用荧光活化细胞分选，以检查细胞类型特异性基因表达

摘要

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.

摘要

大脑是由四个主要类型的细胞，包括神经元，星形胶质细胞，小胶质细胞和少突胶质细胞。虽然它们不是在大脑中最丰富的细胞类型，神经元是最广泛研究因为它们在影响行为直接作用这些细胞类型。在大脑中的其它细胞类型也影响通过它们产生的信号分子的神经元功能和行为。神经科学家必须了解的细胞类型之间的相互作用在大脑更好地了解这些相互作​​用如何影响神经功能和疾病。迄今为止，分析蛋白质或基因表达的最常见方法是利用整个组织样品均质化，通常是用血，而不考虑的细胞类型。这种方法是用于检查基因或可能影响神经功能和行为表达一般的变化信息的方式;然而，这种分析方法不适合于电池的更深入的了解-type特异性基因表达和细胞 - 细胞通信的对神经功能的影响。行为表观遗传学分析已经日益焦点其中审查的区域如何的脱氧核糖核酸（DNA）的结构影响长期基因表达和行为的修改;然而，该信息可以仅是相关的，如果在给定的鉴别谱系，从而后生标记可存在于个体的神经细胞类型的某些基因的细胞类型特异性的方式进行分析。荧光激活细胞分选如下所述（FACS）技术提供了一种简单而有效的方法，以分离单个神经细胞的基因表达，蛋白质表达，或DNA的表观遗传修饰的随后的分析。这种技术也可以被修改，以在脑中为随后的细胞类型特异性分析隔离更具体的神经细胞类型。

引言

The purpose of the protocol described below is to isolate individual cell types from a heterogeneous population of neural tissue for the subsequent analysis of cell-type-specific gene expression, protein expression, or even epigenetic markers. The brain is comprised of many cell types that are derived from distinct progenitor cells and that have cell-type-specific properties and functions. Despite these differences, these distinct neural cell types can express similar receptors and intracellular signaling molecules which make the analysis of these more ubiquitous proteins difficult to measure or interpret in a cell-type-specific manner using conventional methods. Neuroscientists must identify the function and activation of these distinct cell types in the brain and how they can individually impact behavior as this will ultimately be the first step to identifying more specific drugs and therapeutic targets for neurological and neuropsychiatric diseases. Despite this overarching goal of neuroscience research, it can been difficult to isolate individual neural cell types from the brain for the analysis of gene or protein expression, the activation of signaling molecules, or the modification of epigenetic markers on DNA. Of the techniques that are currently used, immunohistochemistry can identify the expression of proteins in a cell-type-specific manner when combined with additional staining of a cell-type-specific marker, though this relies on specific antibodies that can properly stain for the proteins of interest and it can be difficult to quantify. In situ hybridization can identify the specific localization of messenger ribonucleic acid (mRNA) in individual cells in the brain, but this is a laborious process that also limits the co-analysis of specific cell types and only allows for the analysis of one or maybe a few genes of interest. Laser capture micro-dissection uses a laser to isolate subpopulations of cells that are visualized via microscopy; however the time-consuming nature of this process and the relatively low yield can significantly limit the subsequent analysis of proteins or mRNA levels, particularly if the expression of these molecules is low to begin with. Fluorescence activated cell sorting (FACS) is a relatively novel technique in the field of neuroscience to isolate individual cell types from the brain for subsequent analysis of gene expression¹ and/or epigenetic targets². This process can also be used to sort specific types of neural cells for subsequent analysis of cell-type-specific gene expression, protein expression, or epigenetic markers. FACS has been used in a number of medical research fields such as cancer and immunology for decades to count and sort different cells based on either physical or biochemical characteristics³. In addition, flow cytometry has classically been used to analyze protein expression on a per cell basis, using specific antibodies. The procedure described below, takes advantage of classical flow cytometry techniques to isolate individual cell types for subsequent analysis of molecular biology endpoints. The flow cytometer can analyze several thousand cells in a second, which makes it a quick and efficient alternative to the techniques described above. In addition, cells can be isolated based on the cellular expression of a specific protein (for example a neurotransmitter receptor) or a combination of two or more proteins (colocalization of multiple proteins in a specific cell type). This allows the user to isolate very selective neural cell types based on their molecular properties to identify their function in the brain.

To perform FACS, neural cells are prepared into a single-cell suspension which is passed through a flow cell that carries and aligns the cells so that they pass single-file through a light beam and lasers for analysis. A computer acquires the data from each cell and plots it on a histogram for analysis of specified parameters (size, granularity, and fluorescence). Based on these parameters, the cells can immediately be sorted into separate tubes for their recollection and subsequent analysis of any endpoint desired. The protocol described below utilizes three antibodies to sort neurons (using a Thymocyte antigen 1, Thy1 antibody), astrocytes (using a glial glutamate transporter, GLT1 antibody), and microglia (using a cluster of differentiation molecule 11B, CD11b antibody). This protocol can be used as described below or modified with different antibodies depending on the cell type that one would like to isolate for his own experiments.

There are a few caveats to consider when determining whether this protocol is appropriate for specific experiments. One major caveat may concern the specific cell type that one would like to isolate. In this protocol, the three antibodies that are used are extracellular antibodies, which allow the experimenter to keep the cell types intact during the staining procedure, thus preserving the integrity of the RNA, DNA and proteins inside. It is possible that one may wish to isolate a specific neural cell type using an antibody that identifies a protein that is only expressed inside that particular cell. For example, one might want to isolate dopaminergic neurons using an antibody to tyrosine hydroxylase or isolate acetylcholine neurons using an antibody for choline acetyltransferase. These proteins are intracellular and thus would require fixation and permeabilization of the cell membrane for subsequent staining with the appropriate antibodies. While this has been done before ^4,5, this process may significantly decrease the yield of RNA or DNA from these permeabilized cells. Another caveat may be that not all antibodies are appropriate for FACS. For example, one may currently use an antibody that works very well for western blot, a technique that requires the denaturation of proteins. This antibody may not necessarily be suitable for identification of these cell types using FACS given that the proteins are not denatured at any point in this protocol and thus the antibody may have no way to bind to its inherent antigen. Companies provide specification sheets which identify the applications for which an antibody has been approved. If an antibody has not been approved for flow cytometry, it should not discourage one from trying this protocol with a particular antibody; however, one should be aware that it isn’t guaranteed to work for FACS. A third caveat of this protocol has to do with the number of cells that one is trying to isolate. FACS is an excellent technique to yield the most cells possible from even a small piece of tissue, but it is also possible that if one would like to isolate a relatively sparse population of cells from a relatively small brain region, the yield from one animal will be inherently low. In this case, it may be necessary to pool the brain tissue from a few animals in the same treatment group in order to yield the number of cells necessary for subsequent analysis; however, a recent publication has used gene-targeted pre-amplification of cDNA for subsequent analysis of gene expression from a small number of activated neurons (5-6%) from a larger set sorted neurons, indicating that it is possible to analyze gene expression from even small subsets of neural cells without pooling large numbers of animals⁶A final caveat of this technique is that one should have access to a cell sorter that is not too far away. The cell sorter is a complicated machine that requires significant training in order to use it properly. Thus these machines are often run by a qualified technician in a core facility. In addition, the goal of this procedure is to dissociate the neural tissue, stain it for specific antibodies, and immediately take those samples to a sorter within a short period of time (perhaps a half day). This timeframe will help to increase the survival and yield of isolated cells and maintain the integrity of the cells for subsequent processing and analysis. If all of these parameters described above are met, FACS is an excellent method to analyze cell-type specific expression of genes and proteins from neural tissue.

All experiments were performed in accordance with the Institutional Animal Care and Use Committee (IACUC) of the University of Delaware and the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

研究方案

1. Preparation for Tissue Collection (15 – 30 min)

1. Fire polish three sets of glass Pasteur pipettes in decreasing diameters and number them “1” = approximately 1 mm diameter, “2” = approximately 0.75 mm diameter, and “3” = approximately 0.50 mm diameter.
2. Resuspend lyophilized Enzyme A with all of Buffer A according to the protocol provided with the Neural Dissociation Kit (see Table of Materials). Do not vortex. Aliquot the solution into 50 µl aliquots and store at -20 ºC for later use.
3. Make 1 L of myelin removal buffer and vacuum filter for sterilization [myelin removal buffer = 1,000 ml of 0.1 M phosphate-buffered saline (PBS), 2 ml of 500 mM Ethylenediaminetetraacetic acid (EDTA), and 10 g of Bovine Serum Albumin (BSA)].
4. Cut nylon mesh sheet into small 1  x 1 inch (2.5 x 2.5 cm) squares that will be used for filtering cellular debris and autoclave in a small beaker covered with aluminum foil.
5. Prepare 1 L of wash buffer [wash buffer = 1,000 ml of 0.1 M PBS, 2 ml of 500 mM EDTA, and 10 g BSA].

2. Tissue Collection

1. Turn on water bath to 37 ºC.
2. Prepare Buffer X from the Neural Dissociation Kit by adding beta-mercaptoethanol to Buffer X to a final concentration of 0.067 mM (for 4 samples, add 13.5 µl of 50 mM beta-mercaptoethanol to 10 ml of Buffer X).
3. Prepare Enzyme Mix 1 by mixing 50 µl of Enzyme P with 1,900 µl of Buffer X for each sample as described in the protocol for the Neural Dissociation Kit For Example, for 4 samples, mix 200 µl of Enzyme P with 7,600 µl of Buffer X.
4. Place Buffer X in the water bath at 37 ºC.
5. Place 2 ml of cold Hank’s Buffered Salt Solution (HBSS) (without Ca⁺⁺ and Mg⁺⁺) into one well of a sterile 6-well culture plate for each sample that will be collected and keep on ice for tissue dissection.

3. Tissue Dissection (45 min – 1 hr)

1. Inject the animal with appropriate volume of euthanasia solution (commercial barbiturate solution, e.g., Euthasol), according to Institutional Animal Care and Use Committee (IACUC) guidelines, and perfuse out all blood with cold 0.9% saline solution.
   1. Administer Rats with an overdose of euthanasia solution.
   2. Monitor anesthesia via breathing and via toe-pinch. When the animal is sufficiently anesthetized (no response to the toe-pinch and deep, slow breathing), the rat can be perfused.
   3. To perfuse the rat, cut the chest cavity open with a pair of heavy-duty scissors. Cut the diaphragm to allow access up to the heart. Cut the ribs along both sides up to towards the top of the chest, to allow full access to the heart. At that point, the rib cage can be held off to the side using a pair of hemostats.
   4. Perform cardiac perfusion by inserting an 18 G needle (for adult rats) attached to an electric pump into the left ventricle of the heart. The pump is drawing up cold 0.9% saline solution.
   5. Then cut open the right atrium with a small pair of scissors to allow for the blood and liquid perfusate to drain from the body.
   6. Perfuse rats with approximately 10 ml (for adult rats) of 0.9% saline to remove all peripheral immune cells and factors.
2. Dissect the desired part of the brain region using sterile razor blades and appropriate dissection, then place the tissue pieces into the cold HBSS without Ca²⁺ and Mg²⁺.
   NOTE: The protocol requires HBSS without Ca²⁺ and Mg²⁺ in the early steps of the protocol as these ions may interfere with the enzymes required for neural dissociation.
3. After all tissue samples have been collected, return to the clean bench or sterile culture hood for tissue processing.

4. Neural Dissociation (1 hr)

1. Dice each tissue sample into small pieces using sterile razor blades on the lid of the 6-well culture plate. This protocol has been used for tissue samples of approximately 30 - 100 mg, but is approved for tissue samples of up to 400 mg. For tissue samples larger than 400 mg, the reagent volumes must be scaled up appropriately.
2. Use 1 ml of fresh HBSS without Ca²⁺ and Mg²⁺ to transfer the diced pieces of tissue with a pipette into a sterile 2 ml centrifuge tube. Repeat steps 4.1 and 4.2 for all sample until all samples have been transferred into a 2ml centrifuge tube.
3. Centrifuge the samples at 300 x g for 2 min at RT and aspirate supernatant.
4. Add 1,900 µl of warm Enzyme Mix 1, as prepared in Steps 2.2 and 2.3, to each sample and close the tubes.
5. Incubate the samples for 15 min in the 37 ºC water bath, inverting the tubes several times every 5 min to re-suspend the settled pieces of tissue.
6. Meanwhile, prepare Enzyme Mix 2 by mixing 20 µl of Buffer Y with 10 µl of thawed Enzyme A (for 4 samples, mix 80 µl of Buffer Y with 40 µl of thawed Enzyme A).
7. Add 30 µl of Enzyme Mix 2 to each sample and invert gently. Do not vortex.
8. Dissociate each tissue sample with Pasteur pipette “1”, triturating up and down 30 times.
9. Incubate the samples for 15 min in the 37 ºC water bath; invert the tubes several times every 5 min to re-suspend the settled pieces of tissue.
10. Dissociate each tissue sample with Pasteur pipette “2”, triturating up and down 30 times. Avoid generating bubbles.
11. Dissociate each tissue sample with Pasteur pipette “3”, triturating up and down 30 times. Avoid generating bubbles.
12. Incubate the samples for 10 min in the 37 ºC water bath; invert the tubes several times every 5 min to re-suspend the settled cells.
13. Apply single-cell suspension to an 80 micron cell strain placed on top of a 15 ml falcon tube to remove any large pieces of debris and wash the filter and cells with 10 ml of HBSS (with Ca²⁺ and Mg²⁺) to stop the enzyme reactions. Repeat until each sample is transferred into a clean 15 ml falcon tube.
14. Centrifuge the samples at 300 x g for 10 min at RT and aspirate off the supernatant.

5. Myelin Depletion (45 min)

1. Thoroughly re-suspend the pellet of cells with 400 µl of myelin removal buffer.
2. Add specified amount of the myelin removal beads and pipette up and down to thoroughly mix.
   NOTE: The volume of myelin removal beads that is used will depend upon the size of the tissue, the age of the animal, and the amount of myelin expected in that particular brain region. e.g., for an adult rat hippocampus sample (approximately 300 mg), incubate the sample with 100 µl of myelin removal beads.
3. Incubate for 15 min in the refrigerator at 4 ºC.
4. Wash each sample with 5 ml of myelin removal buffer.
5. Centrifuge the samples at 300 x g for 10 min at RT.
6. While centrifuging the samples, place 1 column for each sample in the magnetic field of the magnetic sorter and place a clean 80 micron filter on top of each column.
7. Prepare the columns and filter by rinsing each column with 1 ml of myelin removal buffer, 3 times (3 ml total). Collect all flow-through in a waste container such as the bottom of an empty tip box.
8. Position 5 ml polystyrene round bottom tube directly underneath each column in preparation of sample collection.
9. When cells have finished centrifugation, aspirate supernatant and thoroughly re-suspend in 500 µl of myelin removal buffer. Immediately apply cell suspension to the column and collect the cells in the tube below.
10. Wash the filter and column with 1 ml myelin removal buffer, 4 times (4 ml total).
11. Centrifuge the cell collection at 300 x g for 10 min at RT.
12. Aspirate the supernatant and the cells are now ready for staining.

6. Staining Live Cells for FACS (1 hr for staining)

1. Briefly vortex (2 sec) the cells to separate the pellet and immediately add 5 µl of Fc block (anti- clusters of differentiation 32, CD32), vortex again, and incubate in the refrigerator at 4 ºC for 5 min.
2. Meanwhile, prepare the primary antibody mixture by adding the three primary antibodies in wash buffer as follows: 0.125 µl of allophycocyanin (APC) -conjugated CD11b antibody (1:800), 1 µl of anti-GLT1 antibody (1:100), and 0.4 µl of Fluorescein isothiocyanate (FITC) – conjugated Thy1 antibody (1:250) for every 100 µl of wash buffer. Each sample will receive 100 µl of primary antibody mixture for incubation; however, if your sample is much larger than 400 mg of tissue, you may want to consider increasing the volume of your primary antibody mixture accordingly
3. Prepare tubes for single staining controls by removing just 2 µl of cells from each sample tube and adding it to each single stain control tube. The “single stain” controls include: APC-CD11b only, GLT1 only, FITC-Thy1 only, PE secondary antibody only, and the “blank” or unstained control.
   NOTE: All samples should be represented in all staining controls, thus 2 µl of cells from each sample should be added to each staining control tube.
4. Briefly vortex the tubes (2 sec) and add 100 µl of primary antibody mixture to each sample tube and 100 µl of appropriate primary antibody to each staining control tube, vortex the tubes again, cover the tubes with aluminum foil to protect the fluorescent antibodies from any light, and incubate the tubes in the refrigerator at 4 ºC for 20 min. The samples should be kept cool (at 4 ºC) for the remainder of the procedure to maintain the integrity of the samples and the antibodies.
5. Briefly vortex the tubes and add 2 ml of wash buffer to each tube.
6. Centrifuge tubes at 350 x g for 5 min at 4 ºC.
7. Meanwhile, prepare the secondary antibody mixture by adding 0.2 µl of PE-conjugated anti-rabbit secondary antibody for every 100 µl of wash buffer. Each sample will get 100 µl of secondary antibody mixture; however, if your sample is much larger than 400 mg of tissue, you may want to consider increasing the volume of your primary antibody mixture accordingly.
8. When the centrifugation is complete, dump the supernatant off of each sample into the sink and blot the tubes upside-down on a paper towel.
9. Briefly vortex the tubes and add 100 µl of secondary antibody mixture to each sample tube and the appropriate secondary antibody to each staining control, vortex the tubes again, cover the tubes with aluminum foil, and incubate the tubes in the refrigerator at 4 ºC for 15 min.
   NOTE: The GLT1-only staining control tube should get the secondary antibody mixture.
10. After the incubation, briefly vortex the tubes and add 2 ml of wash buffer to each tube.
11. Centrifuge the tubes at 350 x g for 5 min at 4 ºC.
12. When the centrifugation is complete, dump the supernatant off of each sample into the sink and blot the tubes upside-down on a paper towel.
13. Vortex the tubes and add 0.25 ml of sterile PBS to each tube. The samples are ready for sorting.
    NOTE: More cells may require 0.5 ml of sterile PBS; however it is best to begin with a lower volume as the sample can always be diluted if it is too concentrated with cells.
    NOTE: Optionally add DNase to the tube to prevent clumping of the cells and thus clogging of the sorter. Samples can also be filtered using a sterile 80 micron filter just prior to sorting in order to avoid clogging the sorter.
14. Take cells to the sorter on ice. Collect the cell populations in 2 ml nuclease-free centrifuge tubes with 0.5 ml of sterile PBS in each tube.
15. After the sort, spin the cells down at 300 x g for 2 min at 4 ºC. Aspirate off most of the PBS and flash freeze the cells in the -80 ºC freezer until further processing for either RNA extraction (see 1 for further protocol details), DNA extraction (see 2 for further protocol details) or cell lysis for protein analysis.

结果

The Importance of Myelin Depletion and Tissue Perfusion

Figure 1 depicts the importance of myelin depletion. Myelin depletion (Steps 5.1 through 5.12 of the protocol above) occurs when the single-cell suspension is incubated in the Myelin Removal Beads, washed, and subsequently passed through the column on the magnetic sorter. The purpose of these steps is to reduce the amount of cellular debris present in each sample. As neurons are dissociated and triturated, it can shear of...

讨论

Similar to many other tissues and systems, the brain is comprised of a heterogeneous cell population that functions together to impact behavior. The analysis of gene expression, protein expression, or epigenetic modifications from individual cells within that heterogeneous population has the potential to reveal information about the function of the system as a whole, to identify cellular processes regulating both normal behavior and disease processes, and to provide cell-type-specific targets for potential therapies. It ...

材料

Name	Company	Catalog Number	Comments
Neural Dissociation Kit (P)	Miltenyi Biotec	130-092-628
Myelin Removal Beads II	Miltenyi Biotec	130-096-733
LS Columns	Miltenyi Biotec	130-042-401
QuadroMACS Separator	Miltenyi Biotec	130-090-976
MACS MultiStand	Miltenyi Biotec	130-042-303
Nylon Mesh Sheet	Amazon	CMN-0074-10YD	40 inch width, 80 micron size mesh
Fc Block / anti-CD32	BD Biosciences	BDB550270	reactivity for rat
APC-conjugated CD11b antibody	Biolegend	201809	reactivity for rat
Rabbit anti-GLT1	Novus Biologicals	NBP1-20136	reactivity for rat or human
PE-conjugated anti-rabbit secondary antibody	eBioscience	1037259	secondary antibody for anti-GLT1
FITC-conjugated anti-rat CD90 (Thy1) mouse antibody	Biolegend	202504	reactivity for rat