Intraventricular Drug Delivery and Sampling for Pharmacokinetics and Pharmacodynamics Study

Abstract

Although the blood-brain barrier (BBB) protects the brain from foreign entities, it also prevents some therapeutics from crossing into the central nervous system (CNS) to ameliorate diseases or infections. Drugs are administered directly into the CNS in animals and humans to circumvent the BBB. The present protocol describes a unique way of treating brain infections through intraventricular delivery of antibiotics, i.e., polymyxins, the last-line antibiotics to treat multi-drug resistant Gram-negative bacteria. A straightforward stereotaxic surgery protocol was developed to implant a guide cannula reaching into the lateral ventricle in rats. After a recovery period of 24 h, rats can be injected consciously and repeatedly through a cannula that is fitted to the guide. Injections can be delivered manually as a bolus or infusion using a microinjection pump to obtain a slow and controlled flow rate. The intraventricular injection was successfully confirmed with Evans Blue dye. Cerebrospinal fluid (CSF) can be drained, and the brain and other organs can be collected. This approach is highly amenable for studies involving drug delivery to the CNS and subsequent assessment of pharmacokinetic and pharmacodynamic activity.

Introduction

The blood-brain barrier (BBB) is a crucial protective mechanism for the central nervous system (CNS). The selectively-permeable, anatomic barrier separates the circulating blood and its solutes from the brain’s extracellular fluid, thus preventing most molecules from entering the brain^{1 , 2 , 3 , 4}, depending on their size, lipophilicity⁵, and the availability of an active transport mechanism².

This protective barrier is beneficial for the effective regulation of intricate brain homeostasis and CNS health^{4 , 6}. However, it also makes it difficult to deliver drugs to treat infections in the brain or other CNS diseases^{4 , 7}. Apart from disrupting the BBB using a variety of methods^{8 , 9}, the primary approach to circumvent the BBB is to deliver a drug directly into the brain by releasing it into the cerebrospinal fluid (CSF)⁴. Even though it is a relatively invasive practice, it has been used successfully to deliver targeted therapeutics to patients and laboratory animals. In humans, drugs can be delivered into the intraventricular system or CSF and subsequently sampled using the Ommaya reservoir, a reservoir residing under the scalp, attached to a catheter inserted into the lateral ventricle^{10 , 11}. Similar techniques have been established in laboratory animals such as rodents to achieve equivalent goals. Micro-osmotic pumps were implanted in mice^{12 , 13 , 14 , 15} and rats^{16 , 17} for continuous drug delivery into the ventricular system or brain parenchyma. Additionally, direct intracerebroventricular injections were conducted in anesthetized mice using a disposable needle^{18 , 19} and conscious rats via a surgically implanted cannula^{20 , 21 , 22 , 23}. Drug delivery to the CNS has been an invaluable method to enhance understanding in various fields^{20 , 24 , 25 , 26 , 27 , 28}.

CNS infections are one such field that urgently needs new therapeutics and an enhanced understanding of existing anti-infective therapies. CNS Infections caused by multi-drug resistant Gram-negative bacteria are particularly concerning⁷. Polymyxins are the last-line antibiotics increasingly used to treat infections due to these ‘superbugs’²⁹. When polymyxins are administered intravenously as per the current dosing guidelines³⁰, their penetration into the CNS is very low, while higher doses increase the risk of nephrotoxicity. Therefore, intravenous polymyxin therapy is of little use to treat CNS infections⁷. Establishing a safe and effective dosage regimen for polymyxins delivery to the CNS is an urgent unmet medical need^{31 , 32 , 33}. Therefore, the present protocol was established and is described with a focus on injecting antibiotics directly into the CSF of rats. It can, however, be used to administer any drug that is not neurotoxic and where therapeutic concentrations can be administered in small volumes (e.g., up to 10 μL in rats). The techniques described can also be modified to target different brain regions and deliver multiple injections.

The present protocol presents a straightforward surgery and injection technique that allows for efficient pharmacokinetics and distribution post-ICV administration of drugs. The surgery involves implanting a guide cannula. As it is a less invasive procedure than the implantation of a micro-osmotic pump^{12 , 13 , 14 , 15 , 16 , 17}, this is an advanced option suitable for the short-term administration of drugs into CSF. This protocol is simplified and can produce very high survival rates and stable body weights 24 h post-surgery, which is an improvement compared to existing methods³⁴. After surgery, conscious rats received either a manual bolus ICV injection or slower delivery using a micropump to lower the peak plasma concentrations. At the same time, they could freely move in their cage. To establish safe and effective drug dosage regimens, samples of CSF, brain, spinal cord, kidney, plasma, etc., were then used to study pharmacokinetics and drug distribution following intracerebroventricular (ICV) administration. Drug distribution can also be investigated visually, e.g., using immunohistochemistry or matrix-assisted laser desorption/ionization mass spectrometry imaging (MALDI-MSI). If necessary, a bilateral cannula can be implanted, e.g., to inject drugs that would otherwise distribute unilaterally into both hemispheres.

Protocol

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 × 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 dentalcement 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.

Table of Materials

Name of Material/Equipment	Company	Catalog Number	Comments/Description
1 ml syringes	Terumo, Japan	SS+01T
5 ml syringes	Terumo, Japan	SS+05S
Acetone	Merck, Germany	67641
Bench protector sheets	Halyard, USA	2765-C
Betadine®	Mundipharma, Netherlands	1015695
Buprenorphine; Temgesic®	Clifford Hallam Healthcare, Australia	1238366
Carprofen	Zoetis, Australia	10001132
Chlorhexidine	Tasman Chemicals, Australia	890401
Chux® superwipes (or equivalent)	Chux, Australia	n/a	autoclaved
Clippers	n/a	n/a
Cotton swabs	LP Italiana, Italy	112191	autoclaved
Dental cement powder (Vertex Self cure powder)	Henry Schein, USA	VX-SC500GVD5
Dental cement solvent (Vertex Self cure liquid)	Henry Schein, USA	VX-SC250MLLQ
Disposable needles: 18G, 26G, 30G	Terumo, Japan	NN+2525RL
Disposable surgical blades	Westlab, Australia	663-255
Dummy cannulas	Bio Scientific, Australia	C313DC/SPC	cut to fit 4mm cannula
Ethanol 80%	Merck, Australia	10107
Eye lube	Clifford Hallam Healthcare, Australia	2070491
Felt tip pen	Sharpie, USA	D-4236
Flattened needle (18G) or similar to apply superglue	n/a	n/a
Glass pipettes, pulled	Hirschmann Laborgeraete, Germany	9100175
Glass syringe 10ul	Hamilton, USA	701 LT and 1701 LT
Guide cannulas	Bio Scientific, Australia	C313G/SPC	cut 4mm below the pedestal for lateral ventricle cannulation in adult Sprague Dawley rats
Heat bead steriliser	Inotech, Switzerland	IS-250
Heat pad	n/a	n/a
Hydrogen peroxide 3%	Perrigo, Australia	11383
Injector cannula	Bio Scientific, Australia	C313I/SPC	cut to fit 4mm cannula + 0.5mm projection
Isoflurane	Clifford Hallam Healthcare, Australia	2093803
Isoflurane vaporiser and appropriate scavenging system	n/a	n/a
Medium size weighing boats	n/a	n/a
Metal spatula	Met-App, Australia	n/a
Micro syringe pump	New Era, USA	NE-300
Microdrill	RWD Life Science Co, China	87001
Protein LoBind® tubes, 0.5ml	Eppendorf, Germany	Z666491
Ropivacaine; Naropin®	AstraZeneca, UK	PS09634
Scissors	F.S.T.	14079–10
Screwdriver	n/a	n/a
Screws	Mr. Specs, Australia	n/a
Stereotaxic frame	RWD Life Science Co, China	n/a	Necessary components: rat ear bars, tooth bar, anaesthesia nose cone, arm with digital readout (X, Y, Z) and cannula holder
Sterile saline 0.9%	Baxter, USA	AHB1323
Super etch (37% phosphoric acid) gel	SDI Limited, Australia	8100045
Superglue	UHU, Germany	n/a
Tubing, PE-50	Bio Scientific, Australia	C313CT

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 × 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.

Figure 1: Collection of CSF after injection of Evans Blue dye (1.1%) in the anesthetized rat to confirm cannula location.

CSF is extracted using a pulled glass pipette (A) and then collected in a tube for snap freezing and storage (B).

Figure 2: Tracing of injection materials in the brain ventricles with Evan’s blue dye.

Whole brains are sliced with a blade at the injection site (A,B) or at more posterior locations (C) to confirm the successful injection in the ventricular system. Scale bar (B) = 1 mm.

Representative Results

The surgical protocol presented is highly successful, with trained surgeons reaching >99.8% survival rate and animals showing stable body weight post-surgery on Day 1, compared to their pre-surgery weight on Day 0 (mean ± SD of 315.8 g ± 42.1 g for Day 0 and 314.1 g ± 43.0 g for Day 1, Figure 3).

Figure 3: Bodyweight of representative animals.

Average body weight (+SD) of n = 174 representative animals (19 cohorts) before surgery (Day 0) and on the day of ICV injection (Day 1).

Before collecting CSF, an injection of 1.1% Evans Blue dye into the implanted cannula can aid as confirmation of the injection having been delivered into the intended location. CSF collected will be blue (Figure 1), as will the ventricular system in the brain tissue (Figure 2).

The method was beneficial for a complete study of pharmacokinetics, with samples taken at different time points post-surgery.

Discussion

Researchers and clinicians employ ICV injections to circumvent the protective mechanism of the BBB and deliver drugs directly into the CNS^{12 , 18 , 19 , 21 , 24}. The present work is a complete ICV protocol for delivering drugs efficiently into the CNS and extracting CSF for pharmacokinetic analysis. At the start of the experimentation, when this protocol is being established in the laboratory, the injection location can be confirmed by administering Evans Blue dye through the implanted cannula. This is especially useful and critical if a different strain or age of rats is employed, as different coordinates may be used. An alternative option that can be used in experimental animals is injecting Angiotensin II and observing the animals’ drinking behavior after that; however, this method is less reliable³⁸.

Drug-induced neurotoxicity can be a limiting factor for ICV delivery³⁹; however, decreasing the drug delivery rate can significantly reduce adverse events⁴⁰. This method is not suitable for drugs where a therapeutic dose cannot be administered in small volumes over short periods, such as <10 μL in <1 h in rats. The dose regiment can be adjusted to administer therapeutics in a single dose or over multiple days.

Imaging studies of whole brains or analyzing dissected brain sections can reveal distribution characteristics of the drug. Studying the distribution of the intraventricularly administered drug is crucial to pharmacokinetic analysis such as LC-MS. The exact flow dynamics of CSF are still under investigation and debated⁴¹. However, anatomical characteristics within the CNS and the physiochemical properties of the injected drug can impede drugs from distributing equally within the CNS and even between different CSF compartments⁴². Also, the condition of interest itself can influence the distribution, e.g., the blood-CSF/blood-brain barrier can become leaky in patients and rats with meningitis^{42 , 43}. Thus, it is recommended that each drug and its distribution be investigated for each relevant condition. If the therapeutic under investigation is shown to distribute unilaterally, i.e., does not efficiently cross to the other hemisphere, researchers may consider implanting bilateral cannulas by modifying the described surgery method.

With this protocol, experienced surgeons can achieve very high survival rates and stable body weights in animals. This is an enhancement compared to previous protocols, one of which suggested inhalation anesthesia as a potential improvement for the wellbeing of animals over the reported median non-survival rate of 2%³⁴. This progress can be further combined with minimally invasive analgesia self-administration through food^{35 , 36 , 37}. This results in an optimized prerequisite for good animal welfare and understanding the behavior of a therapeutic agent. With stress being one of the main confounding variables in animal research^{44 , 45 , 46 , 47}, the optimization of invasive protocols is crucial for the ‘refinement’ factor of the three R’s⁴⁸ and the ability to obtain cleaner data that feeds into the ‘reduce’ component.

Studying pharmacokinetics is crucial for the safe and effective delivery of any new or repurposed therapeutic. Injecting one-time or repeatedly into the ventricular brain system is an irreplaceable method in translational neuropharmacology. This protocol can aid in studying a wide variety of drugs. It can be complemented by injecting dyes for imaging or can be used to manipulate the CNS environment, e.g., by inducing specific diseases such as brain infections for investigating the therapeutic effect of antibiotics.