1. Materials
   • Timed pregnant mice at E13-E13.5 for isolating embryonic salivary glands
   • 70% ethanol spray bottle
   • Standard forceps (e.g. Fine Science Tools, 11000-12)
   • Fine sharp scissors (Fine Science Tools, 14060-10)
   • DMEM/F-12 (e.g., Thermo Fisher, 11039047)
   • 100x PenStrep (10,000 units/mL penicillin, 10,000 µg/mL streptomycin; e.g., Thermo Fisher, 15140163)
   • DMEM/F-12 + 1x PenStrep (referred to as “medium” throughout this protocol): add 5 mL 100x PenStrep to 500 mL DMEM/F-12
   • Dissecting microscope (e.g., Leica M80; any common dissecting microscopes should work)
   • Dumont #5 forceps (Fine Science Tools, 11251-20)
   • Dumont #5 fine forceps (Fine Science Tools, 11254-20)
   • Scalpel (Fine Science Tools, 10011-00 and 10003-12)
   • 10-cm dish (e.g., Corning 430167)
   • 35-mm dish (e.g., Corning, 430165)
   • Polycarbonate filter, 13 mm diameter 0.1 µm pore size (MilliporeSigma, WHA110405)
   • 75 mg/mL vitamin C (MilliporeSigma, A7506)
   • 25 mg/mL transferrin (MilliporeSigma, T8158)
   • MatTek dish (MatTek, P50G-1.5-14-F)
2. Isolate the string of embryos from timed pregnant mice.
   • Euthanize the timed pregnant mouse (or mice) at E13-E13.5 by CO₂ following standard operating procedures of your animal facility.
   • With the ventral side up, the euthanized mouse was sprayed with 70% ethanol around the abdomen.
   • Use the standard forceps to grab and pull the skin towards the bottom, and then make a horizontal cut with fine sharp scissors.
   • Next, cut upwards along the midline of the abdomen to expose the abdominal cavity.
   • The string of embryos should be easily visible. While using forceps to grab between embryos and pull up the string, use scissors to cut off the 3 connections (cervix, left and right ovaries).
   • Put the string of embryos in ~10 mL medium in a 10-cm dish.
3. Dissect out embryos from the uterus tube (the string of embryos) in the 10-cm dish.
   • From one end of the string, use one pair of Dumont #5 forceps to pierce through the uterus membrane between the last and the second to the last embryos, and use another pair to tear the uterus membrane and squeeze out the embryo. The embryo is still inside the amniotic sac.
   • Use one pair of Dumont #5 forceps to pierce through the amniotic membrane surrounding the embryo, and use another pair to tear the amniotic membrane to release the embryo.
   • Repeat above steps to release all embryos.
4. Isolate mandibles (with tongues attached) from embryos. This step does not need to be under a dissecting scope.
   • Clean a glass plate with 70% ethanol to have a clean working surface. You can also use the glass working surface of the dissecting scope.
   • Pre-fill a 35-mm dish with 3 mL medium to store isolated mandibles.
   • On the cleaned glass plate, use a scalpel to decapitate the mouse embryo.
   • While the detached head was held on its side with one prong of forceps pierced through the top, a scalpel was used to slice across the mouth opening to isolate the mandible and the attached tongue, between which a pair of submandibular glands were sandwiched.
     • The anatomy of the tissue will naturally guide the scalpel to find its way through. It is rare to damage the tongue without any extra caution.
   • Collect the mandible into the 35-mm dish with medium.
   • Repeat above steps for all embryos.
5. Isolate submandibular salivary glands from the mandibles (with tongues attached). This step needs to be under a dissecting scope.
   • Clean a glass plate with 70% ethanol to have a clean working surface. You can also use the glass working surface of the dissecting scope.
   • Pre-fill a 35-mm dish with 3 mL medium to store isolated glands.
   • Under a dissecting microscope, use forceps to move a detached mandible tissue from the dish onto a clean glass plate with the tongue facing down.
   • Use a pair of forceps to slice through the midline of the mandible tissue to expose the tongue. The two submandibular salivary glands attached to the base of the tongue should be visible at this point.
   • Use forceps to remove surrounding tissues and detach the glands. Collect them into the 35-mm dish with medium.
   • Repeat above steps to process all mandibles.
6. Culture isolated salivary glands on floating filters.
   • Usually ~6 glands are cultured on one 13-mm diameter filter, and each filter takes 180-200 µL organ culture medium.
   • Prepare enough organ culture medium: DMEM/F-12 supplemented with 1x PenStrep, 150 µg/mL vitamin C (1:500 dilution from stock) and 50 µg/mL transferrin (1:500 dilution from stock). Inhibitors and solvent control can be added when preparing the organ culture medium.
   • Add 180 µL organ culture medium into the glass bottom area of a 50-mm MatTek dish.
   • Place a polycarbonate filter on top of the medium, with the shinier side up. If glands will be followed up at different time points by imaging, use fine sharp scissors to cut a small notch at the edge of the filter as a marker to track gland identities.
   • Use forceps to transfer ~6 glands onto each floating filter. I found it is the easiest to use custom-modified forceps to transfer glands. Briefly, one tip of the forceps is bent toward the other tip, while the other tip was bent sideways. See bent-tip forceps in Fig. 2B of this paper.
   • Culture at 37°C with 5% CO₂.