Isolation of Mouse Interstitial Valve Cells to Study the Calcification of the Aortic Valve In Vitro

Calcified aortic valve disease (CAVD) is the most prevalent valvular heart disease in western populations, affecting nearly 2.5% of elderly individuals over 65 years of age¹. CAVD affects over six million Americans and is associated with changes in the mechanical properties of the leaflets that impair normal blood flow-through^1,2. Currently, there is no pharmacological treatment to stop the progression of the disease or to activate mineral regression. The only effective therapy to treat CAVD is aortic valve replacement by surgery or transcatheter aortic valve replacement³. It is therefore imperative to investigate the molecular mechanisms leading to valve mineralization to identify new pharmacological targets. Indeed, non-treated aortic stenosis has several adverse consequences such as left ventricle dysfunction and heart failure⁴.

The aortic valve consists of three layers known as fibrosa, spongiosa, and ventricularis, which contain VICs as the predominant cell type⁵. The fibrosa and the ventricularis are covered by a layer of vascular endothelial cells (VECs)⁵. The VECs regulate the permeability of inflammatory cells as well as paracrine signals. Increased mechanical stress may affect the integrity of the VECs and disturb the homeostasis of the aortic valve, leading to inflammatory cell invasion⁶. Scanning electron microscopy analyses showed disrupted endothelium in a human calcified aortic valve⁷.

Histological analyses of calcified tissue reveal the presence of osteoblasts and osteoclasts. Furthermore, osteogenic differentiation of VICs was observed both in vitro and in human valve tissue⁸. This process is mainly orchestrated by the Runt-related transcription factor 2 (Runx2) and the bone morphogenetic proteins (BMPs)^8,9.

NOTE: All animal procedures described here have been approved by Icahn School of Medicine at Mount Sinai institutional core and use committee.

1. Preparation before valve cell isolation from adult mice

1. Clean and sterilize all the surgical instruments shown in Figure 1A by using 70% v/v ethanol and subsequently autoclaving them for 30 min. clean the surgical workspace with 70% ethanol.
2. Add 500 µL of penicillin-streptomycin to 50 mL of 10 mm HEPES. Prepare an aliquot of 50 mL of 1x phosphate-buffered saline (PBS). Keep the solutions on ice.
3. Prepare 1 mg/mL and 4.5 mg/mL collagenase solutions, and use 5 mL of each solution in 15 mL tubes to perform the entire procedure. To prepare 5 mL of 1 mg/mL collagenase, mix 5 mg of collagenase with 2.5 mL of Dulbecco's Modified Eagle Medium (DMEM, fetal bovine serum (FBS)-free) and 2.5 mL of 10 mM HEPES supplemented with antibiotics (1% penicillin-streptomycin from step 1.2). Filter the solutions through a 0.22 µm filter to remove any contamination.
   NOTE: Keep the solutions on ice to protect the enzymes.
4. Warm the DMEM solution to 37 °C before use in all the steps described below. Prepare complete medium by supplementing DMEM with 1% penicillin-streptomycin, 1% sodium pyruvate, 5 mL of 200 mM L-glutamine, 1 mL of mycoplasma elimination reagent (see the Table of Materials), and 10% FBS.

2. Isolation of valve cells

1. To obtain 10⁶ cells for the experiment, use five 8-week-old mice (minimum of three). Place the mouse in an induction chamber along with a small piece of tissue paper soaked with 1 mL of isoflurane, but do not allow contact with the tissue. To confirm that the animal is fully anesthetized; check for toe pinch reflex, and then euthanize the mouse by cervical dislocation. Use isoflurane to alleviate any pain prior to the cervical dislocation as the procedure described below is terminal.
2. Place the mouse on a dissecting platform, and fix the paws with cannulas to hold it in place. Clean the chest and the abdomen with ethanol; open the abdomen and the chest with scissors. With small surgical scissors, cut between the left atrium and the left ventricle to exsanguinate the mouse. Perfuse the heart with 10 mL of cold 1x PBS to remove blood from the heart.
3. Cut the heart, and keep 3 mm from the ascending aorta as shown in Figure 1B. Dissect the aortic valve under a stereomicroscope. Cut the heart horizontally in the middle of the ventricles (Figure 1C). Cut the left ventricle toward the aorta, and carefully dissect the aortic valve (Figure 1D-F). Pool the valves together in a small 35 mm tissue culture dish.
4. Wash the isolated valves in a 75 mm cell culture dish with 5 mL of cold HEPES (10 mM) supplemented with antibiotics (1% penicillin-streptomycin) to remove blood (Figure 2). Prepare two 15 mL tubes of collagenase 1 mg/mL and 4.5 mg/mL as described above in step 1.3.
   NOTE: After the dissection, manipulate the isolated valves in a sterile biosafety hood to minimize contamination.
5. Incubate the valves in collagenase type I (1 mg/mL) for 30 min at 37 °C with continuous shaking (Figure 2). Centrifuge the tube for 5 min at 150 × g, wash the pellet once with 2 mL of HEPES (10 mM), and vortex for 30 s at high speed. Pour the contents of this tube into a 35 mm culture dish, and carefully transfer the fragments of tissue using thin tweezers into a new tube.
   NOTE: At this stage, the VICs are still not dissociated from the tissue, and the pellet contains pieces of tissue. To avoid contamination with endothelial cells, do not centrifuge after vortexing in step 2.5.
6. Incubate the pellet in a 15 mL tube with 5 mL of collagenase type I (4.5 mg/mL) at 37 °C under continuous agitation for 35 min. Re-suspend the cells with a 1 mL pipette to separate the cells, and centrifuge at 150 × g for 5 min at 4 °C.
7. Discard the supernatant, and re-suspend the pellet in 2 mL of complete DMEM. Centrifuge at 150 × g for 5 min at 4 °C. Repeat this step twice to clean the cells.
   NOTE: The pellet will still have some tissue fragments.
8. Re-suspend the pellet in 1 mL of complete medium, and plate the cells in one well of a 6-well cell culture dish in a minimum amount of medium to facilitate their attachment to the culture dish. Leave the cells, undisturbed, in a 37 °C incubator with 5% carbon dioxide.
9. After 3 days, check the cells under the microscope to verify good growth close to the tissue debris. Once 1,000 cells are visible under the microscope, carefully remove the tissue debris with autoclaved tweezers, and change the medium.
   NOTE: The plate should not be disturbed; if the required number of cells are not observed, place the cell culture dish back in the incubator for another 2 days.
10. When the cells are 70% confluent (2.5 × 10⁵), trypsinize and then transfer them to a 75 mm tissue culture dish.

3. Analysis of cell identity and morphology

NOTE: Immunofluorescence staining was used to study cell morphology and endothelial cell contamination.

1. Clean the hood with 70% v/v/ ethanol. Place sterile coverslips (22 mm x 22 mm) in 6-well plates.
   NOTE: To sterilize the coverslips, wash them with 70% ethanol, and keep them in the hood overnight under ultraviolet light.
2. Seed 100,000 cells per well in a 6-well plate. After 24 h, wash the cells twice in 1x PBS, and fix them in 4% paraformaldehyde (PFA) for 20 min. Wash the cells again twice with 1x PBS.
   NOTE: At this point, the cells could be kept in PBS at 4 °C until the start of the staining procedure.
3. To verify the purity of the VICs, use alpha-smooth muscle actin (αSMA), vimentin, and cluster of differentiation 31 (CD31) to detect contamination with VECs.
4. Prepare an aliquot of blocking buffer by mixing 500 µL of normal serum (the same species as the secondary antibody), 9.5 mL of 1x PBS, and 30 µL of Triton X-100. Incubate the cells in 2 mL of the blocking buffer for 1 h.
5. Prepare the antibody dilution buffer containing 30 µL of Triton X-100, 10 mL of 1x PBS, and 0.1 g of bovine serum albumin (BSA).
6. Take an empty tips box, fill half of the box with water to create a humid chamber. Cover the tip holder with a wet tissue and then with a sheet of parafilm.
7. Take 1 µL of the primary antibody, and mix it with 100 µL of the dilution buffer prepared in step 3.5. Place 50 µL of the diluted antibody on the parafilm. Take the coverslips from the wells, flip them over, and place them on the top of the drops of antibody; incubate the cells overnight with the antibody.
8. Add 1 mL of PBS in the 6-well plate. Carefully take out the coverslip from the parafilm, flip it over, and place it in the well. Wash the cells with a continuous gentle agitation for 5 min. Replace the PBS with fresh PBS; wash the cells 3 times.
9. Incubate the cells with the diluted secondary antibody (1/500) (Alexa-488, Alexa-555) for 1 h. Add 1 µL of the secondary antibody to 500 µL of the antibody dilution buffer (prepared in step 3.5). Cover the plate with aluminum foil. Wash the cells 3 times with 1 mL of 1x PBS with continuous agitation.
10. Mount the coverslips with 50 µL of 4′,6-diamidino-2-phenylindole (DAPI)-mounting medium, and observe the cells under the microscope to analyze the morphology of cells and VEC contamination.

4. In vitro calcification assay

1. Clean the hood with 70% ethanol, warm the DMEM medium to 37 °C.
2. Seed 100,000 cells/condition into 6-well plates in complete DMEM, and culture for 24 h at 37 °C.
3. Prepare the calcifying medium by mixing 2 mM of NaH₂PO₄, 10^-7 M insulin, and 50 µg/mL ascorbic acid in DMEM with 5% FBS. For 93 mL of DMEM, add 5 mL of FBS, 1 mL of antibiotics (final concentration 1%), 1 mL of sodium pyruvate (100 mM), 27.5 mg of NaH₂PO_4, 5.8 µL of insulin, and 5 mg of ascorbic acid.
   NOTE: Filter the solution using a 0.22 µm filter before use.
4. After 24 h, replace the supernatant medium with the calcifying medium. Incubate the cells for 7 days at 37 °C. On the 3^rd day, replace with fresh calcifying medium, and place the plate back in the incubator to complete the 7 days of treatment.
5. After 7 days, remove the medium, and wash the cells twice with 2 mL of 1x PBS. Incubate the cells in 1 mL of 0.6 N hydrochloric acid (HCl) for 24 h at 37 °C. Collect the HCl in a 1.5 mL tube, and evaporate it in a rotary evaporator. Re-suspend the contents of all the tubes in 60 µL of HCl.
   NOTE: The drying procedure is important to concentrate the solution and to have the same volume for each condition.
6. Use a 96-well plate to measure calcium concentration by using Arsenazo III reagent, available in a ready-to-use kit (see the Table of Materials for more details).
7. Prepare a calcium standard solution of 10 mg/dL concentration. Weigh 10 mg of calcium hydroxide (Ca(OH)₂) and dissolve in 100 mL of distilled water.
8. In a clear 96 well plate, pipet 2 µL of blank solution (HCl, 0.6 N), the standard solution, the sample per well (10 mg/dL), and the samples. Perform the experiment in triplicate to verify the pipetting variability. Add 200 µL of the reagent for each condition.
   NOTE: Samples above 15 mg/dL should be diluted 1:1 with saline, re-assayed, and the result multiplied by two.
9. Incubate the reaction for 15 min at room temperature.
   NOTE: The reaction is stable for 60 min.
10. Read and record the absorbance of the plate at 650 nm. Use the following formula to calculate the amount of calcium in the samples:
    Calcium (mg/mL) = (Absorbance of sample/absorbance of standard) × Concentration of standard

As murine aortic valves are typically 1 mm in diameter, at least three valves must be pooled to collect a million viable cells for different experimental procedures. The different steps of the VIC isolation process are shown in Figure 1 and Figure 2. As it is difficult to manually scrape the valve tissue, it is preferable to use shear stress created by vortexing to remove the VECs. Indeed, the CD31 immunofluorescence staining results showed the absence of endothelial cells contamination (Figure 3D). In addition, mouse VICs express vimentin and α-SMA, which are the major markers of valve cells (Figure 3B,C).

Cell mineralization in vitro
A calcium reagent kit was used to measure the calcium concentration; cells treated with calcifying medium have higher calcium concentration compared to non-treated cells (Figure 4A). The concentration of calcium was normalized with the total protein concentration. Alizarin red staining confirmed the calcium-reagent kit measurements by showing red positive calcium nodes (Figure 4B).

Figure 1: Description of valve dissection. (A) Representative image of all the surgical instruments needed for the dissection, scissors 2 is needed to open the skin of the mouse and scissors 3 to open the chest. Tweezers 5 and 6 are needed to hold the skin and open the chest. (B) Leave 3 mm of tissue from the aorta (black arrow). (C) Cut the heart in the middle of the ventricles with scissors number 4. (D) Open the heart toward the aortic valve with scissors 3. Use the thin tweezers 7 and 8 to carefully dissect the aortic valve. The valve is visible and has some black dots that are characteristic of mice valve tissue (blue arrow). (E) Increase the magnification to better visualize the aortic valve. Isolate the valve with the small scissors 4; (F) maintain the tissue with tweezers 7. Please click here to view a larger version of this figure.

Figure 2: Representative description of mouse valve cell isolation. Abbreviations: HEPES = 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid; RT = room temperature; DMEM = Dulbecco's modified Eagle medium; FBS = fetal bovine serum. Please click here to view a larger version of this figure.

Figure 3: Mouse valve cell phenotype. Microscopic view of (A) freshly isolated valve cells. Immunofluorescence staining showing (B) vimentin-positive cells and (C) α-SMA. Cells are negative for (D) CD31 staining. Scale bars = 200 µm. Abbreviations: DAPI = 4′,6-diamidino-2-phenylindole; CD31 = cluster of differentiation 31; α-SMA = alpha-smooth muscle actin. Please click here to view a larger version of this figure.

Figure 4: In vitro calcification assay. (A) Phosphate-rich calcifying medium induced VIC calcification in vitro, which was measured with a reagent kit. (B) Microscopic image showing red positive staining (right) for calcium nodes. (C) Alizarin red staining showed positive calcium nodes (black arrow) of VICs in response to calcifying medium. Scale bars = 100 µm. Abbreviations: CTL- = Control; mVICs = mouse valvular interstitial cells. Please click here to view a larger version of this figure.

This article presents a detailed protocol of mouse valve cell isolation for primary culture. Three aortic valves from 8-week-old mice were pooled to obtain an adequate number of cells. In addition, this protocol describes the characterization of VIC phenotype and the in vitro mineralization assay. The method was adapted from the previously described protocol from Mathieu et al.⁷.

During the isolation of aortic valves, care must be taken to avoid all sources of possible contagion to protect the cells from bacterial or mycoplasma contamination. Indeed, it is crucial to autoclave all the surgical tools prior to starting the experiments. The HEPES solution should be supplemented with 1% antibiotics to minimize bacterial infection. Furthermore, mycoplasma may cause cytopathology and consequently interfere with every parameter measured in cell culture¹⁰.

Plating cells in small culture dishes with lower volume of culture medium is critical for VIC growth and proliferation. Letting the tissue settle and adhere to the cell culture dish permits cell migration from the tissue to the dish wall. Given that isolated cells from young mice proliferate faster, it is recommended to transfer cells to a larger culture dish of 75 cm² after 5 days of culture. Maintaining cells to 80% confluence is crucial to minimize the differentiation of VICs to a myofibroblast phenotype⁸.

As shown by immunofluorescence imaging, the isolated valve cells show a fibroblast-like phenotype. VICs have an elongated cytoplasm and express both vimentin and αSMA as described by previous studies. The present work confirmed that the mouse VIC phenotype is similar to that previously described for porcine VICs¹¹ and human VICs¹². Most in vitro studies on aortic stenosis are performed on cells from large animals^8,11. The key disadvantage of porcine VICs is their spontaneous differentiation to an osteoblast phenotype in vitro even in normal media¹³. However, mouse VICs do not calcify spontaneously even at higher passages.

Mouse VICs differentiate to the osteoblast phenotype in response to calcifying medium using ascorbic acid, insulin, and phosphate stimulation. This article describes a quantitative method of calcium measurement using a kit and a qualitative method using Alizarin red staining. Both methods showed significant increase of calcification in response to calcifying medium treatment. The calcium measurement kit is the gold standard method, which offers an exact quantitative calcium measurement¹⁴.

In the Arsenazo III reagent, magnesium interference is prevented by the inclusion of 8-hydroxyquinoline sulfonate. Calcium reacts with the reagent to form a purple-colored complex, which absorbs at 650 nm. The intensity of the color is proportional to the calcium concentration. The accuracy of the Arsenazo-III reagent was previously validated with atomic absorption spectrophotometry. The same method is used in clinical laboratories to measure total calcium concentration in biological fluids¹⁴. The calcification in aortic stenosis is mainly hydroxyapatite, as shown with dispersive x-ray energy scanning electron microscopy analysis^7,12,15. Indeed, it is important to analyze the calcification of the cell membrane rather than free calcium to more accurately mimic the calcification of the aortic valve tissue.

Mice represent a good source of VICs for the study of molecular mechanisms leading to aortic valve calcification. However, keep in mind that VICs in vitro are not similar to VICs in living valves. Another limitation is the fact that a pool of valves from 3-5 mice is needed to make a single cell culture. The pool should be from littermate mice to minimize variations. In addition, experiments should be performed in triplicate to confirm all findings. However, the use of the entire aortic valve in the culture can alleviate this limitation. Nevertheless, these in vitro studies must be validated in human tissue to strengthen the findings.