Using In Vivo and Tissue and Cell Explant Approaches to Study the Morphogenesis and Pathogenesis of the Embryonic and Perinatal Aorta

All mouse protocols are approved by the Institutional Animal Care and Use Committee at Yale University.

1. In Vivo Embryonic Fate Mapping and Clonal Analysis

Note: We have used these approaches widely to evaluate the origins of cells and their clonal architecture in development and disease models^7,8,9,10.

1. Set up mating between mice with a CreER and mice with a Cre reporter.
   NOTE: A CreER is used for SMC marking; Myh11-CreERT2 or Acta2-CreERT2 mice^11,12 is commonly used for this purpose. We have used ROSA26R-CreERT2 mice¹³ for broadly marking embryonic tissues.
   1. For clonal analysis, use a multi-color Cre reporter (e.g., Confetti or Rainbow [Rb] mice)^8,14,15 and for fate mapping, use a single-color Cre reporter (e.g., ROSA26R-YFP mice¹⁶) or the double-color Cre reporter ROSA26R-mTomato-mGFP (mTmG)¹⁷.
      NOTE: This breeding scheme uses adult mice that are 2 – 6 months old (generally ~30 g) and will generate embryos with a CreER and a Cre reporter.
2. Check for vaginal plugs in the morning using a metal probe and consider noon on the day of plug detection as embryonic day (E) 0.5.
3. Separate the female from the male when a plug is detected.
4. Prepare 4-OH-tamoxifen and tamoxifen working solutions.
   1. For 4-OH-tamoxifen, dissolve 5 mg into 50 µL of 100% ethanol, vortex for 2 min and then add 450 µL of corn oil. On ice, sonicate at output level 2 for 10 s cycles with resting between cycles until sample cools down. After sample is completely dissolved (usually ~ 4 – 5 cycles of sonication), take 200 µL of sonicated solution and dissolve in 1,800 µL of corn oil to make final working solution (4-OH-tamoxifen, 1 mg/mL).
   2. For tamoxifen, dilute to 50 mg/mL in 100% ethanol and vortex until it is dissolved. Dilute in corn oil to make the final working solution (tamoxifen, 10 mg/mL) and stir at 45 °C to ensure that it is completely dissolved.
5. For early embryonic induction, inject 4-OH-tamoxifen intraperitoneally into a pregnant dam.
   1. Use 4-OH-tamoxifen injections (up to ~150 µg per pregnant mouse, or roughly 5 mg/kg bodyweight) at E5.5 and thereafter as they yield viable dam, embryos, and pups.
   2. Use tamoxifen at doses of 0.5 – 1.5 mg (or 17 – 50 mg/kg) for dams pregnant with embryos at ~E9 and thereafter. Use a 30G needle for all injections.
       Note: Filtration through a 0.22 µm filter is typically used to sterilize all mixtures prior to injection.  
6. For clonal analysis, use intraperitoneal injections of high doses of 4-OH-tamoxifen (e.g., 5 mg/kg) or tamoxifen (e.g., 50 mg/kg) to mark multiple cells.
   1. To mark individual cells, titrate down the 4-OH-tamoxifen or tamoxifen dose such that in the tissue of interest (i.e., the aorta), almost all embryos analyzed (as described below at the end of this section) have either no cells labeled or only cells of a single color; this is the threshold dose.
7. To mark cells in the mid-late gestational period and to trace their fate or clonality in the postnatal mouse, inject progesterone into the intraperitoneal cavity of the pregnant dam concomitantly with tamoxifen in a 1:2 (progesterone:tamoxifen) dose ratio.
   NOTE: Tamoxifen doses are 17 – 50 mg/kg and progesterone doses are 8.5 – 25 mg/kg (i.e., half of the tamoxifen doses). The progesterone injection may delay labor by ~1 – 2 days.
8. Euthanize the dam pregnant with embryos of a desired age using an open drop method, where the dam is placed in a receptacle containing cotton or gauze soaked with undiluted isoflurane. Confirm death by opening the chest cavity to induce pneumothorax, by removing the vital organs, and/or by performing cervical dislocation.
9. Cleanse the abdomen with 70% ethanol. Use scissors to cut open the abdomen and dissect out the uterus. Remove the embryos from uterus and carefully separate the embryos from the placenta and yolk sac using forceps.
10. Euthanize the embryos at E15 or older by performing either cervical dislocation or decapitation with surgical scissors or a sharp blade.
    NOTE: Embryos younger than E15 rapidly die following euthanasia of the mother and/or the removal of the embryos from mother.
11. Place the embryos in ice-cold PBS and cut a small piece of tail using scissors to genotype for the CreER and Cre reporter.
12. Fix the embryos in 4% paraformaldehyde at 4 °C for 2 h. Wash the embryos three times in PBS and then incubate the embryos in 30% sucrose in PBS in 15-mL tubes, waiting until they sink, which may take up to two days.
13. Fill plastic freezing molds up to 2/3 of maximum level with optimum cutting temperature (OCT) compound. Place each embryo vertically in the mold, entirely submerged in OCT. Incubate for 10 min at room temperature (RT) to minimize bubbles.
14. Place freezing tissue blocks upright in a dry ice, 70% ethanol bath to freeze. Store the blocks at -80 °C.
15. Set the cryostat temperature to -22 °C and cut blocks to generate transverse sections through the entire aorta, 10 – 20 µm thick. Dry the slides at RT for 30 min prior to storing them at -80 °C.
16. Thaw the slides at RT, protected from light to avoid the bleaching of fluorophores. Wash the slides with PBS-Tween20 0.1%.
    1. For clonal analysis, stain the slides for nuclei (DAPI, 5 mg/mL) and directly image other fluorophores in the Rainbow Cre reporter (Cerulean: 433-nm excitation max, 475-nm emission max; mOrange: 548 nm, 562 nm; mCherry: 587 nm, 610 nm).
       1. Use the following filters for detecting fluorophores with an upright fluorescent microscope: DAPI (excitation max/bandwidth 350/50 nm, emission max/bandwith 460/50 nm), Cerulean (excitation 436/20 nm, emission 480/40 nm), texas red (excitation 560/40 nm, emission 630/75 nm) and custom filter to separate mCherry from mOrange (excitation 577/10 nm, emission 630/50 nm).
    2. For fate mapping with the mTmG Cre reporter, detect GFP (484 nm, 510 nm), either by directly imaging or by using immunostaining. For imaging with an upright fluorescent microscope use a GFP filter (excitation 470/40 nm, emission 525/50 nm).
    3. For immunostaining, incubate sections with primary antibodies overnight at 4 °C, wash with 0.1% Triton X-100 in PBS and then incubate with secondary antibodies for 1 h. Use anti-CD31 (final concentration 0.0016 mg/mL), anti-GFP (0.006 mg/mL) and Cy3-directly conjugated anti-SMA (1:500 final dilution) primary antibodies and use secondary antibodies directly conjugated to fluorophores (1:500 final dilution) (see the Table of Materials).
17. Image slides with an upright fluorescent microscope (magnification: 4X – 20X) or confocal microscope (10X -63X). For high magnification, use a confocal imaging 63x oil objective with a numerical aperture of 1.4-0.6 and a frame size of 1,024 x 1,024 pixels.

2. Explant Embryonic Aorta Culture

Note: This approach was previously used to evaluate the role of integrin beta3 in stenosis of the explanted Eln^(-/-) embryonic aorta^9,18.

1. Euthanize a timed-pregnant dam at E15.5 by excess isoflurane inhalation, as per step 1.8, above.
2. Harvest and euthanize the embryos, as per steps 1.9 – 1.10, above.
3. Position the embryo supinely and pin the extended limbs to the dissection board. Visualize with a dissection stereoscope and use scissors to cut a vertical incision from the abdomen through the sternum to the upper thorax. Dissect away the thymus, trachea, lungs, esophagus, liver, and intestine.
4. Gently pull the heart ventrally using forceps and use scissors to dissect the aorta away from the dorsal aspect of the thoracic and abdominal cavities. To release the aorta, cut it at the proximal root and the distal abdominal positions.
5. Place the aorta in ice-cold sterile PBS in a cell culture hood. Transfer the aorta to a 24-well plate and culture it in DMEM with 0.5% FBS for up to 24 h at 37 °C.
   NOTE: The culture medium can be supplemented with a blocking antibody (e.g., anti-integrin αvβ3 antibody at 0.02 mg/mL⁹; see the Table of Materials) or a pharmacological agent.
6. Wash the aorta twice with PBS and then fix it with 4% paraformaldehyde for 20 min.
7. Wash three times with PBS and transfer the aorta to a 1.5-mL tube with 30% sucrose in PBS. After the aorta sinks, make frozen tissue blocks, cut sections, and immunostain, as described in steps 1.13 – 1.15, above.

3. SMC Isolation from the Perinatal Aorta

Note: This approach is currently being used to compare the biology of aortic SMCs isolated from Eln^(-/-) and wildtype perinatal mice.

1. Euthanize a murine pup at postnatal day (P) 0.5, either by cervical dislocation or by decapitation with surgical scissors or a sharp blade, as described in step 1.10, above.
2. Perform step 2.3 and then, after opening the thorax and abdomen, puncture left ventricle with 23G needle. Use plastic tubing to connect the needle to sterile PBS-containing syringe held vertically at an elevated position so that PBS flows into the left ventricle by gravity. Allow infusion of sterile PBS into left ventricle until the liver blanches.
   1. Use forceps to remove adventitial tissue on the outside of the aorta and dissect the aorta as described in step 2.4.
3. Digest the aorta in a single solution of DMEM (500 µL) supplemented with 225 U/mL collagenase, 2.25 U/mL elastase, and 1x antibiotic-antimycotic for 45 min at 37 °C. Manually shake the tube every 5 – 10 min.
4. In the sterile cell culture hood, titurate the digested tissue by pipetting up and down with a 200-µL pipette to generate a single-cell suspension. Transfer the single-cell suspension to a 15-mL tube, add 2 mL of DMEM, and centrifuge at 920 x g for 5 min at 25°C.
5. Discard the supernatant. Resuspend the cell pellet in 3 mL of SMC culture medium (DMEM containing 10% FBS supplemented with 1 µg/mL rhFGF, 10 µg/mL rhEGF, 100 U/mL penicillin/streptomycin, and 2.5 µg/mL Amphotericin B). Transfer to a 35-mm culture plate.
   NOTE: Upon initial isolation, ~400,000 cells are obtained from a single P0.5 wildtype aorta; ~300,000 of these cells are SMCs.
6. Culture the cells at 37 °C and change to fresh SMC culture medium every 3 days. Using standard techniques with trypsin, passage the cells when confluent. For experiments, use cells from passages 3 – 7.

4. Subcutaneous Osmotic Mini-pump Placement in Pregnant (or Non-pregnant) Mice

Note: This approach was previously used to continuously deliver a pharmacological agent (e.g., integrin β3 and β5 inhibitor cilengitide) to embryos in utero⁹. The pump was inserted in the pregnant dam at E13.5 and maintained until parturition.

1. Anesthetize the mice with 2 – 5% isoflurane in oxygen (flow rate of 1 L/min) for induction and 1-3% for maintenance. Use the toe-pinch reflex to monitor the level of anesthesia, and adjust the anesthetic agent as needed. Administer subcutaneous buprenorphine (0.05 – 0.1 mg/kg) for analgesia.
2. Shave the lower back with clippers. Use sterile drapes, gloves, and instruments and maintain the mice on a heating surgical plate during surgery.
3. Place each mouse in prone position and scrub the back with betadine followed by isopropyl alcohol. Approximately 3 – 5 cm rostral to the base of the tail in the lower back, use a scalpel to make a horizontal skin incision 0.5 – 1.0 cm in length without injuring the underlying muscle.
4. Insert a hemostat into the incision and create a subcutaneous pocket rostrally for pump implantation by opening and closing hemostat jaws.
5. Fill an osmotic mini-pump with the agent of choice, as per the manufacturer's guidelines (see the Table of Materials). Insert the filled pump into the pocket and close the incision with clips or 6-0 non-absorbable sutures. Ensure that the total surgical time is less than 10 min.
6. Post-operatively, monitor the mice every 15 min until they have recovered from sternal recumbency. Administer subcutaneous buprenorphine (0.05 – 0.1 mg/kg) every 6 – 12 h for 48 h. When the mice have fully recovered from general anesthesia, monitor them daily. Remove the clips or sutures 7 – 10 days post-surgery.