Von Kossa Staining of Aortic Vascular Smooth Muscle Cells: An In Vitro Technique for Assessment of Arterial Medial Layer Calcification

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Preparation of Reagents

1. Isolation and Calcification of Murine Aortic VSMCs
   1. Aortic Digestion Solution:
      1. Prepare a fresh solution (~3-5 ml per aorta harvested) with Hank's Balanced Salt Solution (HBSS) containing 175 U/ml type 2 collagenase and 1.25 U/ml elastase. Sterilize the solution with a 0.22 µm vacuum-driven filtration system and keep the solution on ice until use.
   2. Cell Media:
      1. Supplement 500 ml of Dulbecco's Modified Eagle Medium (DMEM) with 10% fetal bovine serum, 100 units/ml penicillin, and 100 µg/ml streptomycin. Warm the media to 37 °C prior to use.
   3. Calcification Media:
      1. Calcification Media A (NaPhos; used in mouse cell lines):
         1. Supplement 100-500 ml of DMEM (volume as needed) with 10% fetal bovine serum, 2 mM sodium phosphate, 100 units/ml penicillin, and 100 µg/ml streptomycin. Warm the media to 37 °C prior to use.
            OR
      2. Calcification Media B (βGP/Asc/DEX; used in either mouse or human cell lines):
         1. Supplement 100-500 ml of DMEM (volume as needed) with 10% fetal bovine serum, 10 mM β-glycerophosphate disodium, 50 µg/ml L-ascorbic acid, 10 nM dexamethasone, 100 units/ml penicillin, and 100 µg/ml streptomycin. Warm the media to 37 °C prior to use.

2. Mouse Dissection

1. Euthanize mouse with a 200 mg/kg intraperitoneal pentobarbital injection.
2. Lay the animal supine on the dissection board and stabilize by taping each paw to the board. Using a dissecting microscope and small scissors, make a midline incision extending from the lower abdomen to the upper thorax.
3. Peel back the skin with forceps and remove the peritoneum, revealing the abdominal organs. Remove the gastrointestinal organs, taking care not to transect the aorta.
4. Make a lateral incision in the anterior diaphragm and continue the incision across the abdomen. Using dissection scissors, release the ribcage by cutting through the sides of the ribs and removing the soft tissue adherent to the superior portion of the sternum. Remove the ribcage, revealing the lungs.
5. Leave the heart in place initially (to aid in identifying and dissecting the proximal aorta) and carefully remove the lungs. Remove the thymus, trachea, and esophagus with care, ensuring that the aorta remains intact.
6. Using straight fine forceps and micro-dissecting scissors, remove the soft tissue surrounding the aorta from the iliac bifurcation to the aortic arch, paying careful attention when removing the peri-aortic fat (Figure 1A). Remove the remaining fat and soft tissue surrounding the large branches of the aortic arch (i.e., brachiocephalic, common carotid and subclavian arteries, Figure 1B).
   NOTE: It is important to remove the fat from the aorta because fat can increase the background signal when performing fluorescence imaging.
7. Remove the heart from the thoracic cavity, carefully detaching it from the proximal aorta, and discard. Transect the distal aorta at the iliac bifurcation. Using an insulin needle, inject normal saline into the aorta from the aortic arch to wash out remaining blood cells. Detach the aorta along with the aortic arch vessels, completely removing it from the body.
8. Place the aorta in normal saline solution on ice until ready for imaging.

3. Isolation of Primary Murine Aortic Vascular Smooth Muscle Cells

1. Place aortas in cold HBSS until the dissections are complete. Carefully cut away any remaining periaortic fat and soft tissue, leaving only the aorta.
2. Under a sterile tissue culture hood, transfer the aortas to the Aortic Digestion Solution in 35 mm x 10 mm tissue culture dishes. Place in an incubator at 37 °C for 30 min with gentle intermittent rocking. After digestion, the aortas exhibit a stretched or frayed appearance.
3. With the dissection microscope and sterile forceps, remove the outer adventitial layer of the aorta while keeping the medial layer intact. One technique for removing the adventitia is to peel away the outer layer of the aorta at one end and remove it from the underlying medial layer like a sock could be peeled back and removed.
4. Once the adventitial layer has been removed, place the remaining aorta into a new tissue culture dish with cell culture media and store at 37 °C with 5% CO₂ for 2-4 hr.
5. Under a sterile hood and using sterile 3 mm micro-dissection scissors, cut the aorta into 1-2 mm wide rings.
6. Place these rings in a new tissue culture dish with Aortic Digestion Solution and incubate at 37 °C with gentle intermittent rocking for 120 min. Pipette the solution up and down several times during this incubation to resuspend cells.
7. Add 5 ml of warm cell culture media to the digestion solution and transfer to a 15 ml conical tube.
8. Centrifuge the tube for 5 min at 200 x g.
9. Aspirate the media and resuspend cells in the desired volume of cell culture media (e.g., 5 ml).
10. Plate the entire amount of isolated cells from each aorta in a 25 cm² cell culture flask and incubate at 37 °C with 5% CO₂. Propagate cells using standard techniques, as previously described. During the initial 7-10 days of incubation, change the media every 72-96 hr. As the cells approach confluence, replenish media more frequently (every 48 hr).
    NOTE: It may take many weeks to grow a sufficient quantity of cells.
11. Once confluent, passage cells with trypsin that is warmed to 37 ^°C^.
    1. Add 0.5-1.0 ml of trypsin to each culture flask and incubate for 3-5 min; gently tap the side of the flask every 30-60 sec as needed to detach cells from the surface.
    2. Once the cells detach from the bottom of the flask, add 10 ml of cell media to the cells in trypsin. Centrifuge the cells at 200 x g for 5 min. Aspirate the media and trypsin from the cell pellet. Resuspend the cells in the desired amount of fresh cell media (e.g., 5-10 ml) and transfer to a new flask (with some cells transferred to a chamber slide).
12. At the first passage of cells, confirm the smooth muscle cell lineage with standard immunocytochemistry techniques, as previously described, using an antibody directed against α-smooth muscle actin.

4. Inducing Calcification of Cultured Smooth Muscle Cells

1. Plate cells obtained from 3.12 in a 6-well format.
   NOTE: Starting with 1 x 10⁵ cells/well in a total volume of 2.0 ml of cell media per well is recommended.
2. Allow cells to grow in Calcification Media A or B for at least 7 days in a 6-well plate format. Incubate cells at 37 °C with 5% CO_2.
3. Change cell media every 48 hr.

5. Assessing VSMC Calcification Using the von Kossa Staining Method

NOTE: The von Kossa method for measuring extracellular matrix calcification of tissues or cultured cells is based on the substitution of phosphate-bound calcium ions with silver ions. In the presence of light and organic compounds, the silver ions are reduced and visualized as metallic silver. Any unreacted silver is removed by treatment with sodium thiosulfate.The protocol for von Kossa staining is as follows:

1. Aspirate media from cell culture plates.
2. Fix cells by placing them in 1 ml of 10% formalin at room temperature for 20 min.
3. Remove the formalin and wash the fixed cells with distilled water for 5 min.
4. Incubate cells in 1 ml of 5% silver nitrate solution under a 60-100 W bulb for 1-2 hr.
5. Aspirate the silver nitrate solution and wash with distilled water for 5 min.
6. Remove unreacted silver by placing the cells in 1 ml of 5% sodium thiosulfate (w/v) solution in distilled water for 5 min.
7. Rinse cells with distilled water for 5 min. Repeat washes 3x. The von Kossa stain is ready for imaging with standard inverted light microscopy.
8. Optional Step: Counterstain with 1ml of nuclear fast red for 5 min. Follow this with three washes with distilled water (5 min each).