Chemical Amputation and Regeneration of the Pharynx in the Planarian Schmidtea mediterranea

1. Preparation

1. Preparation of planaria water¹⁷
   1. Maintain planarians in a 1X Montjuïc salt solution. To prepare planarian water, make individual stock solutions of 1 M CaCl₂, 1 M MgSO₄, 1 M MgCl₂, 1 M KCl and 5 M NaCl in ultrapure water. Filter-sterilize with a 0.2 µm bottle-top filterfor long-term storage.
      Note: Use only ultrapure deionized water (with a resistivity of 18.2 MΩ at 25 °C) to prepare Montjuïc salts.
   2. To prepare a 1 L stock of 5X salt solution, combine 5 mL of 1 M CaCl₂, 5 mL of 1 M MgSO₄, 0.5 mL of MgCl₂, 0.5 mL of KCl and 1.6 mL of 5 M NaCl solutions in 900 mL ultrapure water. To this solution, add 0.504 g of NaHCO₃ and stir to mix. Adjust the pH to 7.0 with hydrochloric acid.
   3. Dilute this 5X stock solution to a 1X working concentration in ultrapure water in a sterile container such as a large capacity carboy (see the Table of Materials). Use this 1X solution for maintaining asexual planarians.
      Note: As an alternative to Montjuïc salt solution, use locally-purchased spring water or ultrapure water containing a commercially available aquarium salt mix (see Table of Materials) at a concentration of0.5 g/L¹⁸.
   4. To prevent bacterial infection in static culture¹⁹, maintain planarians in water containing an antibiotic. Prepare a 50 mg/mL stock solution of gentamicin sulfate in ultrapure water and filter-sterilize. To containers where animals are maintained, add gentamicin sulfate to a final concentration of 50 µg/mL (1:1000 dilution).
2. Preparation of liver paste
   1. As planarians thrive on a diet of organic, grass-fed beef liver, purchase fresh liver and process within 24 h. Remove the membranous capsule encapsulating the liver by peeling it off gently. Cut the liver into ~1-cm cubes, and use a blade to scrape off and discard all hepatic veins and arteries.
   2. Macerate liver pieces using a food mill or a food processor and then pass it through a wire mesh food strainer. Combine into pastry bags, and dispense into syringes or 35-mm Petri dishes. Prior to feeding, centrifuge gently to remove air bubbles.
   3. Store aliquots of liver at -80 °C for up to a year and thaw prior to using. After thawing, re-freeze any leftover liver once, or store at 4 °C for up to 24 h.
3. Maintenance of animals
   1. Planarians will grow and shrink to various sizes depending on the frequency of feeding. Feed planarians every other week (once every 14 days). For long-term bulk cultures, use plastic containers of various sizes (see Table of Materials).
   2. To feed animals, use a metal spatula or plastic transfer pipet to place a pea-sized drop of liver in a box. Allow animals to eat for 1-2 h. Remove remaining food before cleaning the box.
   3. Clean worms twice a week if fed (once directly after feeding and once two days later), or once a week if not fed. To clean, drain water into a plastic beaker by carefully pouring out the water while retaining planarians in the box. Wipe box surface with a paper towel, then repeat on all sides until the box is clean.
   4. Replace water (to approximately ¾ of the box volume) and add gentamicin to a final concentration of 50 µg/mL. Store animals in a dark cabinet or in a 20 °C incubator.
4. Selection of worms
   1. Select worms that were fed 5-7 days prior.
      Note: It is significantly more difficult to amputate pharynges from worms that have been recently fed (1-2 days prior to amputation). Fed animals are also more sensitive to sodium azide.
   2. Select worms that are roughly 6 mm in length. To estimate the size of the worm, transfer it to a Petri dish using a plastic transfer pipette. Place either a flat 6-in ruler or 5 mm x 5 mm graph paper underneath the Petri dish and measure the worm length while it crawls.
      Note: Worms smaller than 6 mm in length are more difficult to amputate.
5. Preparation of sodium azide
   1. Prepare a 100 mM solution by dissolving sodium azide powder in planaria water.
      CAUTION: Sodium azide is toxic and should be handled carefully. Do not discard sodium azide down the drain. Following use, collect it separately for disposal as hazardous waste.

2. Pharynx Amputation

1. Place worms in a 35-mm diameter Petri dish. Carefully remove all planarian water from the dish by using a plastic transfer pipet.
   Note: A maximum of 20 6-mm worms can be comfortably accommodated in a dish of this size.
2. Place the dish under the microscope and adjust magnification so that multiple worms can be viewed simultaneously (e.g. 10X magnification). Replace planarian water with 100 mM sodium azide solution.
   Note: For a 35-mm Petri dish, approximately 5 mL of sodium azide is sufficient. The solution should completely submerge planarians. Adjust solution volume for larger dishes as necessary.
3. Monitor the animals under the microscope and refrain from moving the dish for the first 3-4 min.
4. Observe the extrusion of the pharynx through the microscope; the pharynx will protrude from the pharyngeal pouch and extend fully (approximately 1 mm in length) as shown in Figure 1B.
   Note: It is important to wait until the pharynx has finished extending completely before proceeding further. Once the pharynx emerges from the animal, full extension takes roughly 1 min.
5. Use a plastic transfer pipet to squirt animals around the dish. Suck the animals into the pipet and forcibly release them into the dish a couple of times. If the pharynx is fully extended, this vigorous pipetting will cause it to detach.
6. Alternatively, grasp the pharynx either vertically along its length, or horizontally along its circumference using a pair of fine forceps (see Table of Materials) as shown in Figure 1C. With the pharynx pinched in the forceps, lift the animal upwards towards the meniscus. As the animal is raised, surface tension will cause the pharynx to detach from the animal.
   Note: This method can be used to remove the pharynges of smaller animals (1-3 mm in length) where forceful swirling is not adequate. Avoid poking or otherwise injuring the body of the animal.
7. Using a transfer pipet, move amputated animals to a new dish containing fresh planaria water.
8. Once transferred, rinse amputated animals thoroughly in planaria water, completely exchanging the water three times. Transfer to a new dish containing planaria water with 50µg/mL of gentamicin.
   Note: Complete buffer exchange during these three washes is essential to ensure removal of sodium azide.
9. The next day, replace water in the dish with fresh planaria water containing 50 µg/mL of gentamicin. Repeat every other day thereafter.

3. Assessment of Pharynx Removal After Amputation

1. Examine animals under a microscope (with 10-20X magnification) to determine whether a dark spot has appeared after successful removal of the pharynx, as shown in Figure 2A.
2. Alternatively, fix and bleach animals according to the protocol described by Pearson et al.²⁰. Soak animals overnight in 4',6-diamidino-2-phenylindole (DAPI) or fluorescently-conjugated streptavidin (1 mg/mL diluted 1:500 in 1X Phosphate Buffered Saline containing 0.3% Triton-X). Mount animals on slides in 80% glycerol/20% 1X Phosphate Buffered Saline and image under a fluorescent stereomicroscope (Figure 2B).
   Note: All images in this manuscript were captured on a fluorescent stereomicroscope with a 1.0X objective.

4. Assessment of Pharynx Regeneration by Measuring Feeding Behavior

1. Preparation of liver
   1. Transfer required quantity of liver paste into a microcentrifuge or conical tube. Spin briefly to remove air bubbles. Estimate the volume of liver, and then add planaria water to 1/5 of its volume, and 2% of the total volume in red food coloring. For example, to 1000 µL of liver paste, add 200 µL of planaria water and 24 µL of food coloring.
   2. Using a plastic pestle, pipette tip or metal spatula, mix thoroughly until food coloring is evenly distributed in the liver paste. Spin again briefly. Liver paste can be stored at 4 °C for 24 h or frozen in aliquots at -80 °C for long-term storage.
   3. If testing up to 25 animals, prepare 25 µL of liver paste per dish (approximately 1 µL of liver paste per animal).
2. Feeding animals
   1. Seven days after chemical amputation, transfer animals to a new Petri dish. If testing 10-15 animals, use a 35 mm dish; more than 15 animals should be tested in a 60-mm Petri dish. Keep animals in the dark, undisturbed, for approximately 1 h prior to feeding.
   2. Using scissors, increase the width of a P200 pipette tip by trimming roughly ½ cm off of the narrow end. Pipet 25 µL of red liver paste into the dish.
      Note: Trimming the end makes pipetting the viscous liver paste easier. To prevent the liver from floating on the surface, touch the tip to the bottom of the dish while dispensing.
   3. Allow animals to feed for 30 min.
   4. Score the number of animals that have eaten by placing the animals on a white background or examining them under a microscope with 10-20X magnification.
   5. After feeding, remove liver from the dish and clean.