Isolation and Purification of Murine Cardiac Pericytes

We have optimized a protocol to isolate and purify murine cardiac pericytes for basic research and investigation of their biology and therapeutic potential.

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Our protocol is optimized for the mouse model and utilizes materials that are readily available to all investigators. This technique can be applied to healthy, disease, or genetically altered mouse models. Our protocol will enable investigators to answer any questions about cardiac pericyte biology from any mouse model of disease or genetic variation.

This procedure will provide insight into cardiac pericyte biology and help us understand their contribution to cardiac homeostasis and hemodynamics in both health and disease. Place the anesthetized mouse in the supine position, and tape down its forelimbs. Carefully open the chest cavity, and cannulate the descending aorta using a 25-gauge butterfly needle.

Make a nick in the right atrium. Then, perfuse the heart with at least 20 milliliters of 250-units-per-milliliter heparinized, calcium-magnesium-free Dulbecco's phosphate buffered saline at two milliliters per minute with a variable-flow peristaltic pump. Perfusion is complete when clean PBS comes out of the right atria.

Cut the heart out at the aorta, and place it into the ice-cold calcium-magnesium-free DPBS. Then, transfer the heart into a 15-by-15-centimeter Petri dish. Cut the heart into tiny pieces using spring scissors and fine-point forceps with enough enzyme solution to cover the pieces.

Now, transfer the pieces and solution into a 50-milliliter conical tube, and seal with paraffin plastic film. Incubate at 37 degrees Celsius on an orbital shaker at 120 rpm for 75 minutes. After collagenase digestion with the enzyme solution, decant the liquid through a 100-micron cell strainer into a new 50-milliliter tube, leaving enough solution so that the pieces do not dry out.

Using fine-point forceps, take the tissue from the tube and place a few pieces on a microscope slide. Then, grind the tissue between two microscope slides to break up the tissue. Rinse the slides with enzyme-free culture media into a new 50-milliliter conical tube.

Repeat this step until all tissue pieces are dissociated. Combine the strained solution and the ground-up tissue into one tube. Strain the resulting suspension through a 100-micron cell strainer into a new 50-milliliter conical tube.

Centrifuge the sample at 220 times g and four degrees Celsius for five minutes. Aspirate off the previous solution, and gently resuspend the cell pellet in cold FACS staining buffer containing DPBS and bovine serum albumin. Count the cells, and check viability using a cell counter.

Dilute the cells to one million cells per milliliter with cold FACS staining buffer containing DPBS and bovine serum albumin. The cells are now ready to be stained and sorted. Prepare and label five-milliliter FACS tubes for all controls and cell samples.

Aliquot one milliliter of cells per tube for an unstained sample, fluorescence minus one controls, and isotype-matched controls. Use the remaining cells for the sort. All controls and samples can be prepared and stained at the same time.

Also, prepare and label five-milliliter FACS tubes for a total of nine compensation controls, two kinds of unstained beads plus seven different fluorochromes from the marker panel. To optimize fluorescence compensation controls, add one drop of compensation beads from the squeeze vial to each tube. Then, add one microliter of antibody to the beads.

Repeat for each antibody from the marker panel. Use FMO controls to optimize background staining due to spectral overlap. To the cells in the FMO control tubes, add all antibodies from the marker panel at a one-to-100 dilution, but exclude one antibody.

Repeat for each antibody for a total of seven controls. Use isotype-matched control antibodies for nonspecific staining. To the cells in the isotype-matched labeled tubes, add the isotype-matched control antibody at a one-to-100 dilution.

Next, prepare the cells to be sorted. To the freshly isolated cells, add an antibody cocktail at a one-to-100 dilution. Also, add cell viability dye at a one-to-1, 000 dilution.

Vigorously mix the compensation controls by pulse vortexing. Incubate for 30 minutes at four degrees Celsius, protected from light, except for the cell viability beads, which can be left at room temperature, protected from light. Gently mix the FMO controls, the isotype-matched controls, and the cells to be sorted by pulse vortexing.

Incubate for 30 minutes at four degrees Celsius, protected from light. Add three milliliters of FACS staining buffer to each compensation control, FMO control, and isotype control. Centrifuge the tubes at 300 times g for five minutes at four degrees Celsius.

Aspirate off the solution, and resuspend each pellet in 400 microliters of FACS staining buffer. The compensation controls, FMO controls, and isotype controls are now ready to be used. Following staining, wash the cells with FACS staining buffer by centrifugation at 300 times g for five minutes at four degrees Celsius.

Aspirate off the solution, and resuspend the cell pellet in FACS staining buffer to 0.5 million cells per milliliter. Using new FACS tubes that have 35-micron filter tops, pipette the stained cell samples onto the lids and gravity filtrate to obtain single-cell suspensions. Keep on ice.

Use a cell sorter to purify the cells. Run the unstained cells on the cell sorter to set voltages and correct for the background signal. Run each single-color compensation bead sample one at a time to adjust voltages for each channel and adjust gates for the positive signal.

Collect the data. Use the software to calculate for spectral overlap by calculating the compensation matrix. All voltages are ready and set.

Run each isotype control one at a time. This data can be used to adjust gates for nonspecific binding if there are any. Run each FMO sample one at time.

Adjust voltages for each channel to correct for spectral bleed-through due to a multicolor panel. Run the stained cell samples in the cell sorter. Collect cells in 10-milliliter enzyme-free culture media in a 15-milliliter conical collection tube.

Use the following gating strategy. First, gate for single cells. Then, gate for live cells.

Next, gate for CD45-negative cells. Gate for CD34 and CD31-negative cells. Then, gate for NG2-positive cells.

And finally, gate for CD146 and CD140b-positive cells. To culture pericytes, seed the freshly obtained cells in enzyme-free culture media on a coated 24-well plate. Culture the cells in a cell incubator set at 37 degrees Celsius, 5%carbon dioxide, and 95%oxygen.

After enzymatic digestion and dissociation of the whole heart and before FACS purification of the cells, cells are a crude mixture that contain many different cell types from the heart. After FACS purification and culturing, cells are homogenous. They are single nucleated, quite flat, and have the typical pericyte rhomboid morphology.

Using FACS, cells are purified to homogeny. First, debris and doublets were gated out based on forward and side scatter distributions. Then, dead cells were gated out due to their amine reaction with the dye, which produces a signal greater and more intense than live cells.

Of the live cells, hematopoietic cells were gated out by being CD45-positive. To further remove hematopoietic and endothelial cells, CD34-positive and CD31-positive cells were gated out. Finally, NG2-positive and CD140b/CD146-positive cells were selected for being perivascular cells with expression of typical pericyte markers.

When compared with human brain pericytes, the cells had a similar morphology. Compared with mouse and human smooth muscle cells, the cells had a different morphology Phenotypic characterization of cells at passage seven by immunocytochemistry for pericyte markers showed no observed changes in morphology or marker expression. Similarly, analysis by flow cytometry of the pericytes at passage seven confirmed that the population remained homogenous.

Cell viability is critical to obtain a good yield. Keep tissue cold during procurement, and keep cells cold during cell staining. Also, make sure that the enzymatic solution is prepared fresh each time.

To ensure isolation of the correct cells post-culturing, characterize them with flow cytometry and immunofluorescent staining. These cells can be used in coculture experiments with endothelial cells for functional barrier studies, as well as assays to study their biological and physiological function and characteristics. Cardiac pericytes play a fundamental role in vascular integrity and stability, and their dysfunction is consequential to the global cardiac function.

With our technique, researchers can explore the therapeutic potential of cardiac pericytes.

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Journal (Medicine)

Journal (Medicine)

Isolation and Purification of Murine Cardiac Pericytes

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