Intraventricular Drug Delivery and Sampling for Pharmacokinetics and Pharmacodynamics Study

All experiments were conducted following the Australian code for the care and use of animals for scientific purposes. Experiments were approved by the University of Melbourne ethics committee (application #1914890). 8-14 weeks old male and female Sprague-Dawley rats were used for the experiments.

1. Stereotaxic surgery for lateral ventricle cannulation

1. Use autoclaved cotton tips and surgical drapes during the surgery.
2. Set up the following for the surgery.
   1. Set up stereotaxic frame, anesthesia delivery system, drugs, chemicals, and assisting materials (see Table of Materials). Clean the stereotaxic frame, fiber optic light source, handheld drill, etc., with 80% ethanol.
   2. Soak the 22 G guide cannula and the associated dummy cannula (see Table of Materials) in 80% ethanol for sterilization.
      NOTE: The manufacturer pre-cut the 22 G guide and the dummy cannula to ~4 mm length below the pedestal.
   3. Sterilize the surgery instruments (see Table of Materials) using the heat bead sterilizer for 20 s and then spray with 80% ethanol. Place the sterilized instruments on the sterile drape.
   4. Prepare a recovery box (a rat housing box lined with an underpad) with half of the box placed over a heating pad.
3. Perform the surgery.
   1. Screw the ethanol-cleaned 22 G guide cannula into the cannula holder on the stereotaxic frame.
   2. Place the rat into an induction chamber (300 mm x 200 mm, see Table of Materials) and induce anesthesia with 5% isoflurane at 1 L/min of oxygen.
   3. Once the rat is deeply anesthetized (no pedal reflex), move the rat over to the stereotaxic frame and reduce isoflurane to 2%-3% in 1 L/min of oxygen for maintenance through the nose cone. Hook teeth onto the bite bar and carefully pull the nose cone over the nose – pull back gently on the rat to check that it is secure.
   4. Apply protective eye lube on both eyes to avoid drying out.
   5. Inject Carprofen (5 mg/kg), Buprenorphine (0.05 mg/kg) in saline, and 3 mL of saline subcutaneously (s.c.) for pain management and to aid post-surgery recovery.
   6. Pinch toes to check for pedal reflex. Once absent, fix the rat's skull in the frame. Place one ear bar into the ear canal and tighten. Repeat on the other side.
   7. Move ear bars laterally to ensure that the numbers given on the ear bars are equal on both sides. The head should not move when pressing down on the skull.
   8. Shave the top of the head with a clipper. Wipe the hair away with a tissue and saline. If necessary, re-apply eye lube on both eyes to avoid drying out.
   9. Swab the scalp with 4% chlorhexidine and alcoholic skin preparation solution, using sterile cotton swabs for each step (see Table of Materials). Start in the center of the skull and move in circles outwards until all surface is cleaned.
   10. Inject ~150 µL of Ropivacaine (1%) s.c. (see Table of Materials) along the intended incision site.
   11. Use the scalpel blade to make a 10-15 mm incision along the midline of the head, from in between the eyes to the base of the skull.
   12. Use the cotton tips and the scalpel to scrape away connective tissue and expose the skull.
   13. Dip a sterile cotton tip into a 3% hydrogen peroxide solution and apply it to the surface of the skull. Wait for 5 s for the chemical to react with the skin, and then clean the area with a dry cotton tip.
   14. Apply 3% hydrogen peroxide for a second time. This will make the suture lines of the skull appear more clearly. Wait for 10 s for the chemical to react with the skin, and then clean the area with a dry cotton tip. Wash with saline and dry with a clean cotton tip.
   15. Carefully apply a small amount of Super etch gel on a cotton tip and then apply it to the surface of the skull. This creates a more porous surface for the dental cement to adhere to.
   16. Apply a generous amount of saline to the skull to wash off the super etchant and dry it with a clean cotton tip.
   17. Identify bregma and mark it using the felt tip pen.
   18. Drill an indentation for the anchor screw in a spot that will not obstruct the placement of the guide cannula.
       NOTE: If the right lateral ventricle is cannulated, drill on the left hemisphere and posteriorly relative to the ventricle.
   19. Hold the drill at a ~75°-80° angle to the skull and slowly drill an indent with a depth of approximately half of the thickness of the skull. Use saline on a cotton tip to wipe away bone dust.
   20. Carefully insert the screw by holding it firmly in the indent with forceps and screwing it into the skull with the screwdriver, avoiding breaking in the skull. After each complete turn, test if the screw sits tightly.
   21. Confirm that the skull is flat. Using the mobile arms of the frame, position the guide cannula to touch the skull at bregma. Zero the DV (dorsal-ventral) representing the Z plane coordinates by pressing the button on the frame controlling monitor.
   22. Raise the guide cannula (22 G and 4 mm in length) using the mobile arms of the frame and move it to lambda. Lower it so that it touches the surface of the skull and note the DV display again.
       NOTE: The difference between DV at bregma and DV at lambda should be less than 0.2 mm. If necessary, adjust the nose cone accordingly, i.e., move the rat's head up or down as needed, and repeat the measurements.
   23. Move the guide cannula back to touch bregma and re-zero all three coordinates: DV, AP (anterior-posterior), and ML (medial-lateral) by pressing the buttons on the monitor.
   24. Move the guide cannula to the pre-established coordinates of choice.
       NOTE: For cannulation of the right lateral ventricle in a 300-350 g adult rat, the following coordinates were used: -0.7 mm AP, -1.4 mm ML, and -4.00 mm DV (the same length as the guide cannula).
   25. Mark the final cannula position by coloring the end of the cannula using the marker pen and then lowering it down to the skull surface to touch and mark the position.
   26. Drill a hole of 2.3 mm in diameter for the 22 G guide cannula by holding the drill shaft in a straight vertical position, i.e., at a 90° angle to the scull. Be careful to slowly and progressively move deeper into the skull, i.e., avoid drilling into brain tissue.
       NOTE: If necessary, lower the cannula into the hole using the mobile stereotaxic arm to test whether the cannula can be placed successfully or if further drilling is required.
   27. Once the cannula has been lowered into the brain, lift it back up, carefully apply super glue to the underside of the cannula's plastic pedestal and lower the cannula back into the hole using the mobile arm of the stereotaxic frame.
   28. Leave in place to allow the glue to set.
   29. While waiting, mix dental solvent and powder (see Table of Materials) in a weigh boat. Pour dental cement powder into the weigh boat, use a 1 mL syringe to add ~300 µL of solvent, and mix using the same syringe until it thickens and is of a usable consistency. If necessary, add more powder to increase viscosity or more solvent to reduce viscosity.
   30. Release the cannula from the cannula holder using the dedicated screw and carefully lift the arm.
       NOTE: The cannula should be glued to the skull at a 90° angle.
   31. Apply the fresh dental cement onto the exposed skull by drawing it up into the disposable 1 mL syringe and applying it around the cannula. Avoid any dental cement on the skin or cannula opening. Let it set for a few seconds, depending on its consistency.
   32. When the dental cement becomes thicker, use a spatula to apply more dental cement and form the final head mount covering the screw entirely.
       NOTE: Be careful not to cover too much of the cannula so that the dummy cannula can still be screwed on. Remove any excess dental cement from the skin.
   33. Insert a sterile dummy cannula once the dental cement is dry and firm.
   34. Turn off the isoflurane, release the ear bars, remove the rat from the frame and place it into the recovery box.
   35. Once the animal has recovered ~15-30 min, house it in a clean housing box.
       NOTE: After surgery, rats need to be single-housed. Consider replacing standard grid cage tops where the food hopper lowers into the cage with dedicated high-raised grid cage tops to minimize the risk of interference with the head mount. Alternatively, block off any parts of the cage that are much lower than the rearing height of the animal.
   36. Fill out the monitoring checklist and monitor as per monitoring guidelines outlined in the protocol.
       NOTE: Consider placing some food pellets on the floor for easier reach. If it does not interfere with the experiment, additional pain relief can be provided in sweet food to replace additional post-surgery s.c. injections, e.g., Buprenorphine mixed with sweet nut chocolate paste^35,36,37. In this case, some of the same sweet was provided as a reward before and after surgery to avoid food neophobia when offering pain relief. The nut paste can be provided on tape attached to the cage wall.
   37. Stock more dummy cannulas than guide cannulas or injection cannulas, as they could get lost amongst the animal cage bedding if they come off accidentally. In case of such an event, replace the cannula with a new, sterile dummy cannula.

2. ICV injections

1. Prepare the drug and vehicle, e.g., 30 mg/mL of Polymyxin B (see Table of Materials) in 0.9% sterile saline. Keep the injection volume between 5-10 µL in adult rats with a bodyweight ranging between 300-400 g.
2. At a minimum of 24 h after surgery, weigh the rat and transport it to the procedure room.
3. Calculate the injection volume using the rat body weight.
4. Remove the cage lid and unscrew the dummy cannula.
   NOTE: Rats usually sit still when gently holding them in their cage and do not need restraint. A tea towel can be used for a curious or nervous rat or a tickle of the cheekbones.
5. For a bolus injection, attach PE-50 tubing (see Table of Materials) of at least 40 cm length to a 10 µL gas-tight microsyringe by pulling one end of the tubing over the fixed or removable needle on the syringe and ensuring a tight seal. Attach the fitted injector cannula on the other end of the tubing.
   1. Draw up the required amount through the cannula. Insert the cannula into the guide until fitted and inject. Once all liquid is injected, hold the syringe plunger in place for at least an additional minute to avoid backflow.
6. For a slower delivery rate, inject the drugs using a microinjection pump (see Table of Materials).
   1. First, use filtered water and a disposable 1 mL syringe with a 23 G drawing up needle to fill the PE-50 tubing fully. Then remove the disposable syringe from the drawing up needle and replace it with the microsyringe.
   2. Draw back 1-3 µL to create a small but visible air bubble. Then, draw up the required amount of drugs. Optionally, mark the air bubble with a marker to aid in the visibility of drug flow. Lock the syringe in place on the pump.
      NOTE: Ensure correct settings for the drug and equipment used, i.e., set the delivery speed and the internal diameter of the syringe used according to the user manual.
7. Once all the liquid is injected, stop the pump if it does not stop automatically. Leave the needle inserted for at least an additional minute to avoid backflow.
8. Slowly remove the injector cannula and screw on the guide cannula.
9. Clean the equipment by drawing up and ejecting ethanol and DI water three times each.

3. CSF and tissue sampling

1. Transport the rat to a procedure room.
2. Place the rat in the induction chamber and induce anesthesia with 5% isoflurane in 2 L/min oxygen until breathing has slowed (~ 5 min).
3. Move the rat over to the stereotaxic frame and maintain the same level of deep anesthesia through the nose cone.
4. Check for pedal reflex. Once absent, fix the skull in the frame using the ear bars.
   NOTE: This can be a simpler stereotaxic frame without display, rounded ear bars, or movable arms. This is a non-recovery procedure; the single necessary feature is any rat ear bars.
5. Using the nose cone, angle the rat's nose downwards at about 45° to expose the neck to a good working position.
6. Using small, blunt, curved scissors, feel the skull and move posteriorly to the end of the skull. At this position, cut the outermost muscle layer right through the midline of the neck, about 2-3 cm in length. Carefully cut through all other muscle layers.
   1. Use a small spring retractor to hold open the wound and allow good visibility. Cut until the cisterna magna membrane is visible. Use a cotton tip to aid in gently removing any more skin.
7. If bleeding occurs, use gauze to clean and stop the bleeding.
8. If necessary, use small strips of rolled tissue and place it around the cisterna magna to avoid any blood from contaminating the CSF.
   NOTE: To confirm the success of injection using dye, inject 2-3 µL of 1.1% Evans Blue dye (see Table of Materials) into the anesthetized rat before collecting CSF using the fitted cannula and a disposable 1 mL syringe.
9. To prepare the tool for extracting CSF, fill the syringe with ~500 µL of filtered water, attach a 23 G drawing up needle and pull tubing over the needle. Attach a pulled glass pipette to the other end of the tubing.
10. To extract CSF, slowly insert the pulled pipette into the exposed cisterna magna.
    NOTE: CSF will slowly and passively flow into the pipette and be actively drawn up using the syringe. If the dye is injected, the CSF would be colored blue (Figure 1).
11. Add the CSF into a labeled tube and immediately place it on dry ice. CSF can then be stored at -80 °C.
12. Collect any other tissue of interest for analysis. For plasma, collect at least 3 mL of cardiac blood.
    1. Place the deeply anesthetized rat on its back and open the chest cavity to expose the heart. Use a hemostat to clamp the rib cage and place it on the animal's throat to aid visibility. Use tissue forceps with hooks to hold the right ventricle and insert a 5 mL disposable syringe with an 18 G needle into the left ventricle parallelly to the septum. Slowly withdraw blood into the syringe. Once 3-5 mL of blood were collected, remove the needle from the heart.
13. Remove the needle from the syringe and place it in a sharp container. Transfer the blood to heparinized tubes and spin at 2,739 x g and 4 °C for 15 min before collecting the supernatant with a 200 µL pipette and snap freeze on dry ice.
14. Euthanize the rat by removing the heart with scissors. Turn off the isoflurane supply.
15. Collect any other abdominal organs of interest, such as the kidney or spleen. Identify the organ, hold the organ of interest with tissue forceps with hooks, lift it and cut any attachments. Place the organ into a labeled tube and snap freeze on dry ice.
16. To remove the brain, decapitate the animal with a pair of sharp, large scissors by cutting at the base of the skull. Pull off the head mount by hand. Cut through the remaining skin that is covering the skull.
17. Using dissector scissors, cut the skull at the midline from the base to lambda, i.e., cut the supraoccipital bone in half. Use a hemostat to grasp a half and ply it away. Repeat on the other side to expose the cerebellum fully.
18. Proceed to cut along the sagittal suture up to the frontal bone. Use a hemostat to ply away each of the parietal bones. If required, cut the frontal bone further and remove more bone to expose the olfactory bulbs.
19. Hold the head upside down and carefully pry away the brain with a spatula, detaching it from the optical and trigeminal nerves by severing them with the spatula. Collect the brain into a tube and snap freeze on dry ice.
20. The brain ventricles will be stained in blue if the dye is injected. Use a disposable blade to cut the brain at the injection site (Figure 2A,B) and ~1 cm posterior (Figure 2C) to investigate dye distribution in the ventricles.
21. To remove the spinal cord, expose the vertebral column by cutting the skin along the midline of the rat's dorsal surface. Excise the vertebrate column by making a bilateral incision around it and cutting it transversely at the caudal end of the lumbar spinal cord.
22. Cut the lamina bilaterally along the column using scissors until the entire spinal cord is exposed. Lift the spinal cord from one end, e.g., the rostral end, and extract it by cleaving the spinal nerves in a caudal direction until all nerves are cut. Collect the spinal cord into a tube and snap freeze on dry ice.
    NOTE: Weigh all tubes before and after organ collection for data analysis. If necessary and appropriate, the ventricular system can be washed after CSF collection and before brain collection to remove residual drugs in the ventricular compartments. Use a 1 mL syringe filled with saline attached to the tubing and an injector cannula and insert it into the guide cannula. Inject 2-3 mL of saline. The saline should exit from the opening in the cisterna magna. Guide cannulas, dummy cannulas, and screws can be reused by soaking them in acetone for 24 h, followed by soaking in and cleaning with medical detergent and water. Once dried, residual dental cement can be removed using fine forceps.