Direct Cannula Implantation in the Cisterna Magna of Pigs

Summary

This article presents a step-by-step protocol for the direct cannula implantation in the cisterna magna of pigs.

Abstract

The glymphatic system is a waste clearance system in the brain that relies on the flow of cerebrospinal fluid (CSF) in astrocyte-bound perivascular spaces and has been implicated in the clearance of neurotoxic peptides such as amyloid-beta. Impaired glymphatic function exacerbates disease pathology in animal models of neurodegenerative diseases, such as Alzheimer's, which highlights the importance of understanding this clearance system. The glymphatic system is often studied by cisterna magna cannulations (CMc), where tracers are delivered directly into the cerebrospinal fluid (CSF). Most studies, however, have been carried out in rodents. Here, we demonstrate an adaptation of the CMc technique in pigs. Using CMc in pigs, the glymphatic system can be studied at a high optical resolution in gyrencephalic brains and in doing so bridges the knowledge gap between rodent and human glymphatics.

Introduction

Cerebrospinal fluid (CSF) is an ultrafiltrate of blood which is found within and around the central nervous system (CNS)^1,2. Apart from giving buoyancy to the brain or absorbing damaging mechanical forces, CSF also plays a pivotal role in clearing metabolic waste from the CNS³. Waste clearance is facilitated by the recently characterized glymphatic system which permits the convective flow of CSF through the brain parenchyma via perivascular spaces (PVS), which encircle penetrating arteries^3,4,5. This process has been shown to be dependent on aquaporin-4 (AQP4), a water channel expressed primarily on the astrocytic endfeet, bound to the PVS^4,6. The study of the glymphatic system is achieved by both in vivo and ex vivo imaging, using either advanced light microscopy or magnetic resonance imaging (MRI), following the introduction of a fluorescent/radioactive tracer or contrast agent into the CSF^7,8,9,10,11.

An effective way to introduce a tracer into the CSF without incurring damage to the brain parenchyma is through cisterna magna cannulation (CMc)^12,13. A large majority of all glymphatic studies, thus far have been carried out in rodents and avoided in higher mammals because of the invasiveness of CMc coupled to the practical simplicity of working with a small mammal. Additionally, the thin skulls of mice permit in vivo imaging without the need for a cranial window and subsequently allow for an uncomplicated brain extraction^11,14. Experiments carried out in humans have yielded a valuable macroscopic in vivo data on the glymphatic function, but relied on intrathecal tracer injections in the distal lumbar spine and, furthermore, utilize MRI which does not yield sufficient resolution to capture the microanatomy the glymphatic system^7,15,16. Understanding the architecture and extent of the glymphatic system in higher mammals is essential for its translation to humans. In order to facilitate glymphatic translation to humans, it is important to apply techniques that are carried out in rodents to higher mammals so as to allow for direct comparisons of the glymphatic system across species of increasing cognition and brain complexity¹⁷. Pig and human brains are gyrencephalic, possessing a folded neuroarchitecture, while rodent brains are lissencephalic, thereby having substantial difference among each other. In terms of the overall size, pig brains are, also, more comparable to humans, being 10-15 times smaller than the human brain, while mouse brains are 3,000 times smaller¹⁸. By better understanding the glymphatic system in large mammals, it may be possible to utilize the human glymphatic system for future therapeutic intervention in conditions such as stroke, traumatic brain injury and neurodegeneration. Direct CMc in pigs in vivo is a method that allows for the high-resolution light microscopy of the glymphatic system in a higher mammal. Furthermore, due to the size of the pigs used, it is possible to apply monitoring systems similar to those used in human surgeries making it feasible to tightly document and regulate vital functions in order to assess how these contribute to the glymphatic function.

Protocol

All procedures were carried out in accordance with the European directive 2010/63/EU and were approved by Malmö-Lund Ethical committee on Animal Research (Dnr 5.8.18-05527/2019) and conducted according to the CODEX guidelines of the Swedish Research Council.

1. Preparation

1. Tracer
   1. Prepare artificial CSF (126 mM NaCl, 2.5 mM KCl, 1.25 mM NaH₂PO₄, 2 mM MgCl₂, 2 mM CaCl₂, 10 mM glucose, 26 mM NaHCO₃; pH 7.4)
   2. To 500 μL of artificial CSF, add 10 mg of albumin from bovine serum (BSA) conjugated with Alexa Fluor 647 (BSA-647).
   3. Centrifuge at 5,000 x g for 5 min and use the supernatant.
2. Cannula
   1. Attach a 1 mL syringe to the female Luer connection of the intravenous (IV) line, 3-ways tap with 10 cm extension.
   2. Attach an 18 G needle to the male end.
   3. Open the 3-way stop lock to allow for continuity from the needle to the syringe.
   4. Carefully unsheathe the needle and aspirate approximately 300 μL of the saline into the IV line.
   5. Remove the needle from the saline and proceed to introduce in some air to create a small air bubble (5-10 mm) in the IV line.
   6. Place the needle into the tracer and aspirate all 500 μL of the tracer. The saline in the IV line should be visibly separated by an air bubble.
   7. Discard the needle and close the 3-way stop lock.
3. Animal
   1. Sedate a pig by intramuscular (i.m.) injection of tiletamine (3.75 mg/kg) and zolazepam (3.75 mg/kg) and dexmedetomidine (37.5 μg/kg). Wait for it to become unconscious.
   2. Prepare an intravenous line by inserting a 20 G cannula into the ear vein.
      ​NOTE: Make sure the cannula is in the vein by injecting 5-10 mL of saline through the cannula. If the vein has been missed, this will be noticeable by small edema in the ear tissue.
   3. Intubate the pig to ensure that the breathing rate can be regulated throughout the surgery.
      NOTE: Ensure successful intubation by applying pressure on the pig's thorax and confirm that forcibly expired air is coming out of the intubation tube.
   4. Attach the intubation tube to a ventilator set to a breath rate of 14 breaths/min.
   5. Connect a pulse oximeter and cuff to the tail to monitor the heart rate (HR), blood pressure (BP), and oxygen saturation (sats). Insert a rectal thermometer to monitor the core temperature.
   6. Prepare an IV bag of ketamine (5 mg/kg/min), midazolam (0.25 mg/kg/min), and fentanyl (2.5 μg/kg/min), in saline and begin to infuse through the ear vein at approximately 2 drops/s.
      NOTE: Throughout the surgery, the infusion rate may need to be increased or decreased based on the animal's vitals.
   7. With the pig in the prone position, palpate the back of the head and neck of the animal to locate and mark the occipital crest and spine of the first thoracic vertebrae and the base of each ear.
   8. Draw a straight line between the crest and the vertebrae along the longitudinal axis. Draw two lines from the crest to the base of each ear by following the base of the skull (Figure 1A).
   9. Check that the animal is in a deep sleep by carefully clamping the tail and watching for the absence of a tail reflex.
      ​NOTE: If the animal is still reflexive, the anesthetic infusion rate should be incrementally increased until the animal no longer exhibits a reflex.

2. Surgery

NOTE: All through the surgery, it is necessary to have at least one assistant to suction the light bleeding and cauterize any severed vessels.

1. Using a scalpel with a # 21 blade, make a dermal incision along the longitudinal line down to the muscle.
2. Extend two perpendicular dermal incisions further along the shoulders, 10-15 cm in length.
3. From the occipital crests, make dermal incisions along the line down to the base of each ear.
4. Gripping the skin corners formed at the occipital crest with anatomical forceps, carefully separate the skin from the underlying muscle by lightly running the scalpel blade over the fascia, moving from the rostral to caudal. Once the skin has been resected following each of the five incisions, parts of the trapezius muscles should then be visible.
5. Make a longitudinal incision with the scalpel, approximately 1 cm deep, where the trapezius comes together at the midline.
   NOTE: When cutting through the muscles, there is an increased propensity for bleeding, so the cauterizer should be ready. If a larger vessel is severed, one person should quickly compress it with the gauze, while the other person uses the cauterizer.
6. Using a combination of straight and curved surgical forceps, perform blunt dissection working along the longitudinal cut in the muscles. This will separate the bellies of the trapezius, as well as the underlying semispinalis capitus biventer muscle.
7. Sever any persisting muscle fibers with a scalpel and continue blunt dissection until semispinalis capitus complexus becomes visible.
8. Sever the origins of the trapezius and semispinalis capitus biventer muscles along the posterior aspect of the skull. Carefully separate them longitudinally with the scalpel performing blunt dissection until the semispinalis capitus complexus is fully visible.
9. Retract the trapezius and semispinalis capitus biventer muscles using self-retaining retractors.
10. Where the bellies of the semispinalis capitus complexus come together in the midline, make a longitudinal incision with the scalpel approximately 1 cm deep.
    NOTE: Be aware for any additional bleeding here. Bleeding can be managed using a combination of cotton swabs and cauterization.
11. Using surgical forceps, perform a blunt dissection working along the longitudinal cut between the muscle bellies until the dorsal aspect of the atlas (CI) is palpable.
12. Sever the origins of the semispinalis capitus complexus muscles along the posterior aspect of the skull and separate it longitudinally from the underlying vertebrae by scalpel and blunt dissection.
13. Retract the semispinalis capitus complexus muscles using another set of self-retaining retractors.
14. Using a scalpel, carefully remove any remaining tissue overlying the region where the atlas meets the skull base.
15. Placing one arm under the animal's neck and one finger at the juncture of the atlas and skull, simultaneously elevate the head and flex the neck while palpating with the finger to reveal the cisterna magna using the other hand.
    ​NOTE: The cisterna magna is recognizable when palpating as a strong elastic structure with a small amount of rebound as pressure is released with the finger.

3. Cannulation and injection

NOTE: This step also requires at least two people and is carried out with the animal's head elevated and neck flexed.

1. Ensure that one person elevates and flexes the head and neck of the animal whilst the other palpates for the cisterna magna making a note of its anatomical location.
2. Slowly and carefully introduce a 22 G cannula through the dura and into the cisterna magna at an oblique angle to the longitudinal axis.
   NOTE: Do not insert the cannula too deep, as this can cause damage to the brain. Knowing how far to insert the cannula comes with experience in understanding how it feels for the cannula to pierce the dura. Essentially, just as the dura has been pierced, the cannula is then deep enough for a successful tracer injection. This depth is approximately 3-5 mm but will differ based on the size or age of the animal. Successful cannulation should be immediately evident through the visualization of clear, pulsatile CSF ascending the cannula. For the best outcome, it is recommended to practice several cannulations beforehand in euthanized animals to get one's understanding of dural piercing.
3. Retract the needle from the cannula and place a cap on the lock.
4. First, apply superglue and an accelerator where the cannula enters the tissue, followed by the application of the dental cement. Wait for 5 min for the cement to harden.
5. Carefully remove the cap from the cannula and attach the male end of the previously prepared IV line tap with 10 cm extension, with the tracer, to the cannula.
6. Slowly inject the tracer by hand or using a micro-infusion pump at a rate of 100 μL/min. Remove the 3-ways IV line tap with 10 cm extension and replace it with the cap. Tracer should now be visible pulsating at the base of the cannula (Supplementary Video 1).
   NOTE: If injecting by hand, do this until the tracer is just still visible in the cannula shaft, approximately 1-2 mm above where the dental cement is covering the shaft.
7. Following injection, place sandbags under the neck to maintain some flexion. The head may then be released, and the animal is left in a resting prone position.
8. Release the self-retaining retractors and place muscles as they lay before. Bring the skin together over the muscles using surgical towel clamps.
9. Cover towel clamps and incision with gauze and then a blanket to limit heat loss.
10. Allow the tracer to circulate for the desired time before euthanizing the animal by i.v. Pentobarbital injection (140 mg/kg). Confirm euthanasia by the absence of heart sounds upon auscultation with a stethoscope.

4. Brain extraction and processing

1. Using a scalpel with 20-blade, extend the longitudinal dermal incision from the occipital crest to approximately 7 cm above the nose.
2. Reflect the skin overlying the dorsal aspect of the skull using the scalpel.
   NOTE: There are several ways to cut and remove the dorsal aspect of the pig skull on an animal-by-animal basis. What follows is the procedure that has worked most often for this experiment.
3. Using a handheld compact saw, make a coronal cut in the skull, approximately 3 cm above the two large veins seen exiting the skull. Extend to two further vertical cuts from the coronal cuts and two more further cuts to bring the vertical cuts together in the midline.
   NOTE: Maintain a firm grip of the saw when making the skull bone cuts as it will tend to pull away upon the first contact with the bone or tissue, which can lead to a severe injury.
4. Ensure the skull cuts are through the entire thickness of the bone by following up with a hammer and narrow chisel (10 mm) to each of the cuts.
5. Using the hammer, finally knock a wide chisel (25-30 mm) into the coronal cut. With one person supporting the head, ensure that the other person applies leverage on the chisel to wince open the dorsal skull.
6. Once the dorsal skull fragment has been removed, dissect out the overlying dura mater using curved surgical scissors.
7. Use a spatula to severe the spinal cord from the cerebellum at the rostral aspect. Then proceed to guide the spatula under the brain from the front, severing the olfactory bulbs, pituitary gland, and cranial nerves.
8. Place the spatula behind the cerebellum and apply a fair amount of pressure to dislodge the brain from the cranial cavity, carefully lifting it out once loose.
9. Immediately fix the whole brain by tissue immersion in 4% paraformaldehyde overnight.
   NOTE: After this step, it is possible to carry out the whole brain imaging using a stereoscope (Figure 1E).
10. The next day, make coronal slices of the brain using a salmon knife and fix the slices overnight by tissue immersion in 4% paraformaldehyde.
11. Finally, place the slices in 0.01% azide in PBS for long-term storage.

Results

Once the pig is unconscious, it is palpated, and its surface anatomy is marked, starting at the occipital crest (OC) and working towards the thoracic vertebrae (TV) and each ear base (EB). It is along these lines that the dermal incisions are made (Figure 1A). The three muscle layers including trapezius, semispinalis capitus biventer and semispinalis capitus complexus are resected and held open by two sets of self-retaining retractors to expose the cisterna magna (CM) (F...

Discussion

Herein, is described, a detailed protocol to perform the direct cannulation of the cisterna magna in pigs, including the necessary preparation, surgical procedure, tracer infusion and extraction of the brain. This requires someone with experience and certification for working with large animals. If carried out correctly, this allows for the delivery of desired molecules with surety directly into the CSF, after which a series of different advanced light imaging modalities can be used to explore CSF distribution and glymph...

Materials

Name	Company	Catalog Number	Comments
0.01% azide in PBS	Sigmaaldrich	S2002
18G needle	Mediq
1ml Syringe	FischerSci	15849152
20G cannula	Mediq	NA
22G cannula	Mediq	NA
4% paraformaldehyde	Sigmaaldrich	P6148
Anatomical forceps	NA	NA
Bovine serum albumin Alexa-Fluor 647 Conjugate	ThermoFischer	A34785	2 vials (10mg)
CaCl2	Sigmaaldrich	C1016
Chisel	ClasOhlson	40-8870
Dental cement	Agnthos	7508
compact saw	ClasOhlson	40-9517
Glucose	Sigmaaldrich	G8270
Hammer	ClasOhlson	40-7694
Insta-Set CA Accelerator	BSI-Inc	BSI-151
IV line TAP, 3-WAYS with 10cm extension	Bbraun	NA
KCl	Sigmaaldrich	P9333
Marker pen	NA	NA
MgCl2	Sigmaaldrich	M8266
MilliQ water	NA	NA
NaCL	Sigmaaldrich	S7653
NaH2PO4	Sigmaaldrich	S8282
NaHCO3	Sigmaaldrich	S5761
No. 20 scalpel blade	Agnthos	BB520
No. 21 Scalpel blade	Agnthos	BB521
No. 4 Scalpel handle	Agnthos	10004-13
Saline	Mediq	NA
Salmon knife	Fiskers	NA
Self-retaining retractors	NA	NA
Superglue	NA	NA
Surgical curved scissors	NA	NA
Surgical forceps	NA	NA
Surgical towel clamps	NA	NA