Cannula Implantation into the Cisterna Magna of Rodents

Summary

Here we describe a protocol to perform cisterna magna cannulation (CMc), a minimally invasive way to deliver tracers, substrates and signaling molecules into the cerebrospinal fluid (CSF). Combined with different imaging modalities, CMc enables glymphatic system and CSF dynamics assessment, as well as brain-wide delivery of various compounds.

Abstract

Cisterna magna cannulation (CMc) is a straightforward procedure that enables direct access to the cerebrospinal fluid (CSF) without operative damage to the skull or the brain parenchyma. In anesthetized rodents, the exposure of the dura mater by blunt dissection of the neck muscles allows the insertion of a cannula into the cisterna magna (CM). The cannula, composed either by a fine beveled needle or borosilicate capillary, is attached via a polyethylene (PE) tube to a syringe. Using a syringe pump, molecules can then be injected at controlled rates directly into the CM, which is continuous with the subarachnoid space. From the subarachnoid space, we can trace CSF fluxes by convective flow into the perivascular space around penetrating arterioles, where solute exchange with the interstitial fluid (ISF) occurs. CMc can be performed for acute injections immediately following the surgery, or for chronic implantation, with later injection in anesthetized or awake, freely moving rodents. Quantitation of tracer distribution in the brain parenchyma can be performed by epifluorescence, 2-photon microscopy, and magnetic resonance imaging (MRI), depending on the physico-chemical properties of the injected molecules. Thus, CMc in conjunction with various imaging techniques offers a powerful tool for assessment of the glymphatic system and CSF dynamics and function. Furthermore, CMc can be utilized as a conduit for fast, brain-wide delivery of signaling molecules and metabolic substrates that could not otherwise cross the blood brain barrier (BBB).

Introduction

Cerebrospinal fluid (CSF) bathes the central nervous system (CNS) throughout the ventricular system and along the subarachnoid spaces, an anatomically defined space in continuum with the ventricles, which surrounds the brain and the spinal cord. One of the main functions of the CSF is to provide a route for clearance of metabolites and solutes from the brain parenchyma. Clearance is facilitated via the recently discovered glymphatic system¹, the brain analog to the peripheral lymphatic system. Herein, we describe and discuss the cisterna magna cannulation (CMc), a minimally invasive method for the direct delivery of molecules into the CSF. CMc is the key method for studying the glymphatic function. Furthermore, CMc can also be applied for the study of CSF dynamics and for a fast, brain-wide delivery of non-blood brain barrier (BBB) permeable molecules into the brain parenchyma, along the perivascular space.

The CMc exploits physiological principles of CSF movement dynamics through the CNS to deliver labeled tracer molecules or drugs into the CSF-filled space of the cisterna magna (CM). Molecules are injected through a cannula implanted into the atlanto-occipital dural membrane covering the CM. Molecules are then carried by CSF bulk flow into the brain parenchyma via the paravascular space¹. Tracer or contrast agent injected via the CMc follows the movement of CSF, which allows the assessment of CSF movement and glymphatic influx by quantifying intensity levels of labeled molecules that enter the brain parenchyma. CMc is compatible with different imaging techniques including epifluorescence, 2-photon microscopy, and magnetic resonance imaging (MRI). Also, this assessment can be performed both in vivo or ex vivo. Importantly, CMc allows for the visualization of the glymphatic system under anesthesia or during natural sleep, as well as in awake, freely moving animals.

The CMc technique can be utilized to study different aspects of fluid dynamics in the CSF, but has proven to be particularly useful for studying the glymphatic system. Glymphatic activity drives the convective flow of CSF from the periarterial space via aquaporin-4 (AQP-4) water channels, which are tethered in the membrane of astrocytic vascular-wrapping endfeet. The convective flow enables the interchange of CSF and interstitial fluid (ISF) within the brain parenchyma. CSF/ISF containing metabolic waste and solutes is then removed from the brain parenchyma via the perivenous space^2,3. Ultimately, CSF/ISF reaches the periphery via the recently described dural lymphatic vessels^4,5. The glymphatic system has been shown crucial for the clearance of harmful waste metabolites such as amyloid-β². Further, glymphatic clearance is impaired in aging⁶, after traumatic brain injury⁷, and in animal models of diabetes⁸ and Alzheimer's disease⁹. Notably, glymphatic activity is state dependent, showing significantly higher activity during sleep or anesthesia in comparison to wakefulness¹. Indeed, young anesthetized animals exhibit the highest glymphatic activity. Thus, experimental quantification of glymphatic activity is critical when studying its role in health and disease.

Several studies have addressed CSF dynamics and its interchange with interstitial fluid (ISF) in the brain parenchyma. However, the methods by which labeled molecules are delivered are rather invasive, triggering brain parenchyma damage and changes in the intracranial pressure (ICP) (see review¹⁰). Some examples are intraventricular or intraparenchymal injections which involve craniotomy or drilling of a burr hole in the skull. These procedures have been shown to alter ICP, thus disrupting glymphatic function². Also, such invasive methods induce astrogliosis and increase AQP-4 immunoreactivity in the brain parenchyma damaged area and its surroundings^11,12. As astrocytes and AQP-4 are key elements of the glymphatic system, the CMc is the method of choice for its studies. The major advantages of CMc in comparison to more invasive procedures are the maintenance of an intact skull and brain parenchyma, avoiding ICP alterations and astrogliosis, respectively. Thus, CMc in conjunction with different imaging tools opens for a wide range of possibilities to study not only the glymphatic system, but also the dynamics and mechanisms of fluid flow in homeostasis, as well as in animal models of neurological diseases.

The cisterna magna cannulation (CMc) procedure allows easy and direct access to the cerebrospinal fluid (CSF). By injecting different molecules (e.g. fluorescent tracers, MRI contrast agents) the experimenter can track their movement within the CSF compartment and assess the activity of the glymphatic system. The following protocol describes both the acute CMc, for injections immediately following the surgery, and chronic implantation of the cannula, in which the animal recovers from the surgical procedure for a later injection. The most important difference between the acute and chronic implantation is that the chronic implantation allows for the study of glymphatic activity in awake mice.

Protocol

All procedures were performed in accordance with the European Directive 2010/63/EU for animal research and were approved by the Animal Experiments Council under the Danish Ministry of Environment and Food (2015-15-0201-00535).

1. Procedure for Cannulation

1. Cannula preparation
   NOTE: Avoid touching the cannula with non-sterile gloves.
   1. Break off the beveled metal tip of a 30G dental needle using a needle holder.
   2. Using a needle holder, prepare the cannula by inserting the beveled metal tip (approximately 0.3 cm) into a 30-cm length of PE10 tubing (Polyethylene Tubing 0.024" OD x 0.011" ID) filled with aCSF (126 mM NaCl, 2.5 mM KCl, 1.25 mM NaH₂PO₄, 2 mM MgSO₄, 2 mM CaCl₂, 10 mM glucose, 26 mM NaHCO₃; pH 7.4 when gassed with 95% O₂ and 5% CO₂).
   3. Flush the cannula with aCSF using a 1 mL syringe fitted with a 30G needle (30 G x ½" 0.3 x 12).
2. Surgical procedure
   NOTE: The chronic CM cannulation allows the animals to recover from the surgical procedure. CM injections are made the day following cannula implantation and, importantly, can be performed in either anesthetized or awake animals. Since this is a recovery surgery, procedures should be carried out under sterile conditions.
   1. Weigh the mouse (C57BL/6JRj, both sexes, 8 weeks) and anesthetize it with a mixture of ketamine and xylazine (100 mg/kg; 10 mg/kg, respectively) via intraperitoneal (i.p.) injection. If necessary, redose the mouse with a half dose of ketamine (50 mg/kg) during the surgical procedure. Alternatively, for chronic cannulation, anesthetize mouse by placing it into an isoflurane induction chamber at 2.5 - 3% isoflurane, in circa 1 L/min O₂. In this case, use a nose cone to sustain isoflurane anesthesia at 1.5 - 2% throughout the surgery.
   2. When toe pinch reflexes cease, and the respiration becomes slow and steady, place the animal in a stereotaxic frame over a heating pad.
   3. Apply ophthalmic ointment. Repeat during surgery whenever necessary.
   4. Shave the head and neck of the mouse, remove fur, and sterilize exposed skin first with an alcohol swab and then two times with chlorhexidine (0.5%) or iodine solution (2%). Repeat the sterilization two more times.
      NOTE: Change the surgical drape to remove debris and hair after shaving. Then drape the animal to protect the sterile field.
   5. For the chronic cannulation, administer 0.5 - 1 ml lidocaine/bupivacaine (1 mg/ml and 0.25 mg/ml, respectively) subcutaneously (s.c.) at the incision site. Administer buprenorphine (0.05 mg/kg; s.c.) for post-surgical analgesia.
   6. Fix the mouse in the stereotaxic frame. After ensuring fixation, either intraural or by the zygomatic arch, tilt the head slightly so that it forms an angle of 120° to the body (Figure 1E).
   7. Find the part of the skull protruding immediately above the neck muscles - the occipital crest. Lift the overlying skin using a pair of tweezers, and cut an almond shaped piece of skin of approximately 1 cm along the midline. Use cotton swabs or eye spears to control any resultant bleeding.
   8. Using the occipital crest as a reference point, pull apart the superficial connective tissue to expose the neck muscles below.
   9. Separate the muscles at the midline by carefully running the forceps down the middle of the incision site in the anterior-to-posterior axis. With a pair of curved forceps in each hand, join the tips in the middle near the bottom of the skull and pull the muscles aside.
      NOTE: This should expose the CM, which appears as a tiny inverted triangle, outlined by the cerebellum above and medulla below, behind the translucent dural membrane (Figure 1B and 1C).
   10. Using a surgical eye spear or cotton swab, wipe the dural membrane covering the CM.
3. Insertion of the cannula into the CM
   1. Remove the cannula from the aCSF-filled syringe, keeping the 30G needle attached to the rear end of the tubing.
   2. Attach the 30G needle to a distilled water-filled 100 µL syringe connected to a syringe pump.
   3. Insert an air bubble of approximately 1 cm into the cannula by withdrawing air with the aid of a syringe pump.
   4. Using a syringe pump, withdraw 12 μL of desired CSF tracer into the cannula.
   5. Grasp the cannula, filled with the CSF tracer, near the tube-covered needle with a pair of curved tweezers held in the dominant hand. Rest the middle finger of the non-dominant hand on the ear bar of the opposite side and hold it steady for the later use as a rest for the cannula.
   6. Insert the cannula at an angle of 45° relative to the mouse head, passing into the center of the CM, identified by its triangular aspect seen through the dura. Avoid any penetration of the cerebellum or medulla. Ensure that the needle is only inserted to a depth of 1 - 2 mm, i.e. to the point where bevel is entirely under the dura. Release the tweezers holding the cannula, and let the cannula rest on the non-dominant hand.
      NOTE: The beveled end of the dental needles will require the application of some force to pierce the dural membrane covering the CM.
   7. If necessary, dry off any CSF leak upon penetration using a surgical eye spear or cotton swabs.
   8. Drop 2 - 3 drops of cyanoacrylate glue onto the dural membrane surrounding the cannula. Add a drop of glue accelerator to cure the glue immediately. Cover the skull and needle with a mixture of dental cement (approximately 0.5 mg) and cyanoacrylate glue (3 - 5 drops). Immediately after, apply a drop of glue accelerator to cure.
   9. For chronic cannulation, cut the tubing (leaving approximately 2 - 3 cm attached to the cannula) and seal it with a surgical weld to retain the intracranial pressure (ICP) levels, by preventing CSF leakage through the tubing.
   10. For chronic cannulation, administer carprofen (5 mg/kg; s.c.)
   11. For chronic cannulation, place the mouse in a cage, keeping it over a heating pad to maintain body temperature until the animal is fully recovered from anesthesia.
       NOTE: The animals should be singly housed in their cages to assure that the cannula remains intact. Ensure that the nose is clear of bedding by placing the mouse on a paper towel or other solid substrate.
       
       ​NOTE: The following day, when animals are recovered from the surgical procedure performed for cannula insertion, they can be injected with CSF tracers. For injection under anesthesia, administer a mixture of ketamine/xylazine (100 mg/kg; 10 mg/kg, respectively; i.p.) and proceed to steps described in section 2. For injection in awake animals, proceed to section 3.

2. Injection of CSF Tracers via Acute Implanted CM Cannula in Anesthetized Animals

NOTE: For injection of CSF tracers via acute implanted CM cannula in anesthetized animals, immediately after step 8 from previous section, proceed to CSF tracer injection as described below.

1. Using a syringe pump, start the injection into the CM of CSF tracers at a rate of 1 μL/min for 5 or 10 min, resulting in a total volume of 5 µL or 10 µL, respectively. At the end of the injection, allow the CSF tracer to circulate throughout the brain for 30 min with the cannula undisturbed.
2. After 30 min, cut the tubing connected to the cannula (approximately 4 cm distance from the needle tip) and seal its end using a surgical weld.
3. Under deep anesthesia, euthanize the animal by decapitation. Quickly dissect the brain and fix the tissue by immersion in 4% paraformaldehyde (PFA) diluted in phosphate buffered saline (PBS; 0.01M; pH 7.4) overnight (o/n) at 4 °C.

3. Injection of CSF Tracers via Chronically Implanted CM cannula in Awake Animals

1. Using a syringe pump, withdraw 7 µL or 12 µl of desired CSF tracer into a cannula composed of approximately 30 cm of PE10 tubing with a beveled dental needle tip of 0.5 cm.
2. Gently restrain the animal and cut of approximately 1 cm of the tubing attached to the cannula.
3. Still under gentle restraining, quickly connect the cannula filled with CSF tracer to the CM implanted cannula.
4. Using a syringe pump, start the injection into the CM of CSF tracers at a rate of 1 µL/min. Inject 7 µL or 12 µL, to achieve a final volume of CSF injected tracer of 5 µL or 10 µL, thus compensating the aCSF remaining in the implanted cannula. At the end of the injection, allow the CSF tracer to circulate throughout the brain for 30 min with the cannula undisturbed. Ensure that the tubing remains attached during injection and circulation periods.
5. At the end of CSF tracer circulation time, proceed to euthanasia by decapitation, assuring that the animal is deeply anesthetized. Quickly dissect the brain and fix the tissue by immersion in 4% paraformaldehyde (PFA) diluted in phosphate buffered saline (PBS; 0.01M; pH 7.4) overnight (o/n) at 4 °C.

Results

Upon fixation of mice or rats in a stereotaxic frame, the neck muscles around the occipital crest region are bluntly dissected to expose the cisterna magna (CM). The triangular structure of the CM is readily recognized between the caudal portion of the cerebellum and the medulla (Figure 1A-1C). The cannula is inserted 1 - 2 mm into the CM by gently piercing the atlanto-occipital membrane (Figure 1D). The dura mem...

Discussion

We have presented a protocol that describes a detailed procedure for cisterna magna cannulation (CMc), which offers a straightforward method to deliver labeled molecules to the CSF compartment. CMc allows the subsequent visualization of CSF dynamics, both in vivo and ex vivo, using different imaging modalities or histology.

One of the main advantages of the CMc technique lies in its direct access to the subarachnoid space without the need to expose the brain by craniotomy. By...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
SOPIRA Carpule 30G 0.3 x 12mm	Kulzer	AA001
Polyethylene Tubing 0.024” OD x 0.011” ID	Scandidact	PE10-CL-500
30G x ½” 0.3 x 12 mm Luer-Lock	Chirana T. Injecta	CHINS01
Chlorhexidine 0.5% (chlorhexidine digluconate)	Meda AS	no catalogue number, see link in comments	http://www.meda.dk/behandlingsomrader/desinfektion/desinfektion-af-hud/klorhexidin-sprit-medic-05/
Alcohol Swab 70% Isopropyl Alcohol 30 x 60mm	Vitrex Medical A/S	520213
Viskoese Oejendraeber Ophtha	Ophtha	145250
Wooden applicator, Double cotton bud (Ø appr. 4 - 5 mm, length appr. 12 mm)	Heinz Herenz	1032018
Eye spears	Medicom	A18005
Ferric chloride 10% solution	Algeos	NV0382
Kimtech Science Precision Wipes Tissue Wipers	Kimberly Clark Professional	05511
Loctite Super Glue Precision 5g	Loctite	no catalogue number, see link in comments	http://www.loctite-consumer.dk/da/produkter/superglue-liquid.html
Insta-Set CA Accelerator	Bob Smith Industries	BSI-152
Dental Cement Powder	A-M Systems	525000
Surgical weld	Kent Scientific Corporation	INS750391
Hamilton syringe GASTIGHT® , 1700 series, 1710TLL, volume 100 μL, PTFE Luer lock	Hamilton syringes	1710TLL
LEGATO 130 Syringe pump	KD Scientific	788130
Paraformaldehyde powder, 95%	Sigma Aldrich	158127
Phosphate buffered saline (PBS; 0.01M; pH 7.4)	Sigma Aldrich	P3813
Ovalbumin, Alexa Fluor 647 Conjugate	Thermo Fisher Scientific	O34784
DAPI (diamidino-2-phenylindole) Solution (1 mg/mL)	Thermo Fisher Scientific	62248
Dextran, Fluorescein, 3000 MW, Anionic	Thermo Fisher Scientific	D3305
E-Z Anesthesia EZ-7000 Classic System	E-Z Systems	EZ-7000
Attane Isofluran 1000 mg/g	ScanVet	55226
Euthanimal 200mg/mL (sodium pentobarbital)	ScanVet	545349
Ketaminol Vet 100 mg/mL (ketamine)	Intervet International BV	511519
Rompin Vet 20 mg/mL (xylazin)	KVP Pharma + Veterinär Produkte GmbH	148999
Xylocain 20 mg/mL (lidocain)	AstraZeneca	158543
Marcain 2.5 mg/mL (bupivacain)	AstraZeneca	123918
Bupaq Vet 0.3 mg/mL (buprenorphine)	Richter Pharma AG	185159