Accessing the Porcine Brain via High-Speed Pneumatic Drill Craniectomy

Summary

This protocol describes performing a craniectomy using a high-speed pneumatic drill on a 3-month-old Danish Landrace pig. The access is made through the frontal bone and reveals the ventral dura mater and underlying cerebral hemispheres. This procedure allows for access to a large portion of the pig brain.

Abstract

The use of pigs as an experimental animal model is especially relevant in neuroscience research, as the porcine and human central nervous systems (CNS) share many important functional and architectural properties. Consequently, pigs are expected to have an increasingly important role in future research on various neurological diseases. Here, a method to perform an anterior craniectomy through the porcine frontal bone is described. After a midline incision and subsequent exposure of the porcine frontal bone, anatomical landmarks are used to ensure the optimal location of the craniectomy. By careful and gradual thinning of the frontal bone with a rounded drill, a rectangular opening to the dura mater and underlying cerebral hemispheres is achieved. The presented method requires certain surgical materials, including a pneumatic high-speed drill, and some degree of surgical experience. Potential complications include unintended lesions of the dura mater or dorsal sagittal sinus. However, the method is simple, time-efficient, and offers a high degree of reproducibility for researchers. If performed correctly, the technique exposes a large portion of the unaffected pig brain for various neuromonitoring or analyses.

Introduction

In general, animal models are used when practical and/or ethical limitations prohibit the use of human patients to examine diseases or test surgical methods. Novel animal models are generally established to provide new knowledge with translational value to human conditions. Rodents are often utilized due to practical and financial considerations, but they have limited translational value to humans, especially due to substantial anatomical differences¹. Pigs, however, offer several advantages compared to rodents. Not only do pigs share several key anatomical, physiological, metabolic, and genetic features with humans, but the size of the porcine organ systems can be weight-matched to resemble human organs^2,3. This gives pigs a unique role among surgical animal models and in procedural training⁴. Although the use of porcine models requires certain practical and financial capabilities compared to the use of rodents, pigs offer both a financially and ethically more acceptable option compared to the use of non-human primates.

The porcine brain is of particular interest in translational neuroscience research. Firstly, the architecture of the pig brain is similar to that of the human brain, as both are white matter-predominant and gyrencephalic^3,5,6. Secondly, the larger brain size in pigs compared to rodents permits the use of surgical equipment and various imaging modalities equivalent to those used in clinical settings^7,8. Consequently, various porcine models have been used extensively in neuroscience research over recent decades⁹. The majority of these porcine CNS models, however, require direct analysis of brain tissue, which can be obtained in various ways (e.g., implantation of catheters or electrodes, tissue biopsies, etc.)¹⁰. Since most of these modalities require some degree of instrumentalization and direct access to the brain, different approaches for surgical access must be considered.

This method involves performing an anterior craniectomy through the frontal bone on a sedated 3-month-old female Danish Landrace pig. The overall purpose of this manuscript is to describe a method for exposing a large proportion of the ventral porcine brain through a craniectomy using a pneumatic high-speed drill. The first step is to place the subject in a suitable position with an elevated head. Since the porcine cranium is quite different from that of humans, the second step involves planning the placement of the craniectomy using various anatomical landmarks. The third step is to access the underlying dura mater covering both hemispheres without damaging it.

Protocol

All animal experiments described were performed at Aalborg University Hospital, Denmark, in accordance with existing laws and under the approval of the Danish Animal Experiments Inspectorate (license no. 2020-15-0201-00401). Domestic swine, female, approximately 40 kg and 3 months of age, were used for this study. The details regarding the reagents and equipment used are listed in the Table of Materials.

1. Subject housing

1. House subjects in groups with 12 h light/dark cycles in approved pens for at least 7 days before the surgical procedure to ensure acclimation.

2. Anesthesia and monitoring

1. Sedate the animal with an intramuscular injection of 2 mL/10 kg of a mixture containing Tiletamine 6.25 mg/mL, Ketamine 6.25 mg/mL, Zolazepam 6.25 mg/mL, Butorphanol 1.25 mg/mL, and Xylazine 6.25 mg/mL.
2. Place the animal in a supine position on a heating blanket to ensure optimal thermoregulation.
3. Intubate the subject with a size 6.5 tube⁷ and ventilate it mechanically with non-humidified air, a tidal volume of 8 mL/kg, and a respiratory rate of 16-20 breaths/min according to the expiratory end-tidal CO₂ concentrations <6.0 kPa.
   NOTE: This also ensures correct placement of the tube within the trachea.
4. Maintain anesthesia by inhalation of 1% to 2% sevoflurane.
5. Apply ophthalmic ointment carefully to both eyes of the subject to prevent dryness during anesthesia.
6. Ensure the degree of anesthesia by checking for ciliary reflexes and adjusting the sevoflurane accordingly.
7. Insert a bladder catheter with a thermometer into the subject's bladder through the urethra to monitor temperature and collect urine in a catheter bag.
8. Monitor the animal's vital signs throughout the procedure.
   NOTE: Vital signs include pulse, continuous arterial blood pressure, temperature, and end-tidal CO₂.
9. Insert a central venous catheter (6 Fr sheath) in the right jugular vein by percutaneous puncture and use it for continuous saline (NaCl, 0.9%) infusion, drug infusion, and euthanasia at the end of the study.

3. Animal positioning

1. Place the subject in a prone position with the head raised and stabilized with sandbags until the frontal bone is nearly horizontal to ensure optimal positioning.
   NOTE: Ensure the head is somewhat immobilized to prevent unwanted movement.
2. Shave the hair from the surgical site using a trimmer or razor.

4. Preparation of surgical equipment

1. Prepare the surgical equipment as listed in the Table of Materials.

5. Exposing the frontal bone

1. Identify the nuchal prominence and the caudal aspect of each superior orbital crest (Figure 1) to define the expected sagittal midline.
2. Begin with a midline incision using scalpel no. 24 to cut through both the skin and the galea aponeurotica onto the periosteum of the frontal bone.
3. Wipe away the blood with surgical swabs.
4. Gradually separate the galea aponeurotica from the underlying frontal bone around the incision using a 12 mm flattened rongeur.
5. Use a surgical retractor to separate the galea aponeurotica and expose the underlying frontal bone (Figure 2).

6. Identifying anatomical landmarks of the exposed frontal bone

1. Identify the sagittal suture as a reference for the anatomical midline and the coronal suture (Figure 2).
2. Identify the three bone structures through manual palpation: the nuchal prominence and the caudal aspect of both superior orbital crests (Figure 1 and Figure 3).
   NOTE: These landmarks form a triangle within which the craniectomy is performed (Figure 1 and Figure 4A).

7. Access to the dura mater

1. Using a high-speed pneumatic drill with a rounded diamond-coated burr (Figure 3A), define each corner of a rectangle within the borders of the previously defined triangle.
2. Connect each corner with a 4 mm rounded diamond burr to ensure the correct location of the opening (Figure 4B).
3. Gradually thin the frontal bone with the 4 mm rounded diamond burr until the dura mater is exposed.
   NOTE: Avoid making the first point of contact with the dura mater in the midline, corresponding to the sagittal suture (Figure 1), as the large dorsal sagittal sinus is located here. Perforation at this point can cause significant venous hemorrhage.
4. Use the first point of contact with the dura to visually assess the thickness of the remaining frontal bone.
5. Continue carefully thinning the frontal bone in the defined rectangle using the 4mm rounded diamond burr.
   NOTE: Save the bone dust for later hemostasis of bleeding from veins or exposed spongiosa of the frontal bone.
6. Slide a 3 mm dissector under the sufficiently thinned bone around the bone plate and chip it off with gentle manual pressure to expand the opening to the dura mater.
   NOTE: Alternate between drilling and chipping off the bone until multiple points of the dura mater around the bone plate are exposed.
7. Apply sterile saline with a syringe to clear the view.

8. Removal of the bone plate

1. Insert the 3 mm dissector under the bone plate and apply gradual downward pressure to its handle to break off the bone plate.
   NOTE: The underlying dura mater should now be exposed, revealing the contours of both cerebral hemispheres (Figure 5 and Figure 6).
2. Assess the integrity of the dura mater by visually inspecting for cerebrospinal fluid (CSF) leakages.
   NOTE: Both hemispheres will elevate in synchronous pulsation if there are no significant defects in the dura mater.
3. Stop minor venous bleedings using saved bone dust or by carefully applying monopolar or bipolar coagulation with a cautery at low voltage.
   NOTE: Minor venous bleedings from emissary or diploic veins are expected.

9. Protecting exposed dura mater

1. Cover the exposed dura mater with a sterile surgical swab soaked in sterile saline to prevent drying out of the underlying tissue.

10. Insertion of microdialysis catheters (MDC)

1. Use the dorsal sagittal sinus as a reference for the midline. Place a microdialysis catheter (MDC) within the lumen of a disposable 18 G needle. Penetrate the dura mater 10 mm laterally from the dorsal sagittal sinus with the needle at a 10°-15° angle in a cranial-rostral direction until 20 mm of the needle is within the tissue.
2. Slowly withdraw the needle while ensuring the MDC remains at the same location within the superficial cerebral cortex.
3. Securely fasten the catheter to nearby skin to prevent dislocation of the MDC.
   NOTE: Ensure the tip of the MDC does not contact the sharp tip of the needle to avoid damage.
4. Repeat the procedure using the dorsal sagittal sinus as a midline reference. Place an MDC within a needle as described above and penetrate the dura mater on the contralateral hemisphere 15 mm from the midline with the needle perpendicular (90° angle).
   1. Insert the needle 20 mm before slowly withdrawing it as described, leaving the MDC within the subcortical cerebral tissue. Secure the MDC as described above.
5. Prepare a 2 mL disposable plastic syringe with an 18 G disposable needle. Fill the syringe with 0.5 mL of sterile saline.
6. Penetrate the dura mater 20 mm laterally from the midline while keeping the needle at a 45° angle aimed towards the midline. Slowly introduce the needle 1 mm at a time while gently aspirating with each step of insertion.
7. Observe the syringe during each aspiration until cerebrospinal fluid (CSF) is obtained.
8. Separate the needle from the syringe while keeping the needle in the same anatomical location. Insert an MDC through the needle into the lateral ventricle until resistance is felt.
9. Securely fasten the MDC with surgical tape as described above.

11. Microdialysis (MD)

1. Connect each microdialysis catheter (MDC) to a separate microdialysis pump.
2. Initiate the microdialysis process by pumping sterile saline through the MDC and collecting it in a suitable sample. Ensure flow through each MDC by observing saline in each sample.
3. Start a tissue-calibration period of 30 min of continuous microdialysis at a flow rate of 1 µL/min.

12. Euthanaziation

1. Administer a bolus of intravenous pentobarbital (50 mg/kg) through the central venous catheter (following institutionally approved protocols).
2. Observe the pulse, blood pressure, and end-tidal CO₂ curves on the respirator for a flatline as confirmation of cardiac arrest.

Results

The prone position of the pig's head provides optimal access for the surgeon during the procedure, and the use of stabilizing sandbags reduces the risk of unintended shifts in the pig's head position while drilling.

During this demonstration, the superficial anatomical landmarks of the pig's superior skull (both superior orbital crests and the nuchal crest) (Figure 1 and Figure 3) were used to precisely identify the centered sagittal ...

Discussion

The demonstrated procedure involves several critical steps. Firstly, the accurate planning of the craniectomy's location is crucial due to the composition of the porcine skull. Since the thickness of the porcine frontal bone increases at the lateral edges, placing the opening too laterally¹¹ can make it difficult to reach the dura mater during drilling. Additionally, locating the opening correctly within the midline is important to reduce the risk of unintended damage to the underlying dorsal ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
10 mL plastic syrringes	Becton, Dickinson and Company	303219
107 Microdialysis pump	M Dialysis	P000127	107 Microdialysis Pump
2 mL plastic syrringes	Becton, Dickinson and Company	300928
25 mm, 18 G needles	Becton, Dickinson and Company	304100
Bair Hugger heater	3M	B5005241003
Bair Hugger heating blanket	3M	B5005241003
Batery for microdialysis pump	M Dialysis	8001788	Battery 6V, 106 & MD Pump
Dissector	Karl Storz	223535	Flattended 3 mm dissector
Endotracheal tube size 6.5	DVMed	DVM-107860	Cuffed endotracheal tube
Euthasol Vet	Dechra Veterinary Products A/S	380019	phentobarbital for euthanazia, 400 mg/mL
Farabeuf Rougine	Mahr Surgical		Flat headed rougine (12 mm)
Foley Catheter 12 F	Becton, Dickinson and Company	D175812E	Catherter with in-built thermosensor
Intravenous sheath	Coris Avanti		Avanti Cordis Femoral Sheath 6 F
Microdialysis brain catheters	M Dialysis	P000050	membrane length 10 mm -shaft 100 mm 4/pkg
Microdialysis syringe	M Dialysis	8010191	106 Pump Syringe 20/pkg
Microvials for microdialysis sampling	M Dialysis	P000001	Microvials 250/pkg
Operating table
Pneumatic high-speed drill	Medtronic		Medtronic Midas Rex 7 drill
Primus respirator	Dräger		Respirator with in-built vaporiser for supplementary Sevofluran anesthesia
Rounded diamond drill	Medtronic	7BA40D-MN
Self-retaining retractor	World Precission Instruments	501722	Weitlander retractor, self-retaining, 14 cm blunt
Sterile Saline	Fresnius Kabi	805541	1000 mL
Sterile surgical swaps
Surgical scalpel no 24	Swann Morton	5.03396E+12	Swann Morton Sterile Disposable Scalpel No. 24
Zoletil Vet	Virbac		Medical mixture for induction of anesthesia