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 organs2,3. 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 gyrencephalic3,5,6. 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 settings7,8. 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.
Subscription Required. Please recommend JoVE to your librarian.
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
- 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
- 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.
- Place the animal in a supine position on a heating blanket to ensure optimal thermoregulation.
- 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. - Maintain anesthesia by inhalation of 1% to 2% sevoflurane.
- Apply ophthalmic ointment carefully to both eyes of the subject to prevent dryness during anesthesia.
- Ensure the degree of anesthesia by checking for ciliary reflexes and adjusting the sevoflurane accordingly.
- 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.
- Monitor the animal's vital signs throughout the procedure.
NOTE: Vital signs include pulse, continuous arterial blood pressure, temperature, and end-tidal CO2. - 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
- 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. - Shave the hair from the surgical site using a trimmer or razor.
4. Preparation of surgical equipment
- Prepare the surgical equipment as listed in the Table of Materials.
5. Exposing the frontal bone
- Identify the nuchal prominence and the caudal aspect of each superior orbital crest (Figure 1) to define the expected sagittal midline.
- 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.
- Wipe away the blood with surgical swabs.
- Gradually separate the galea aponeurotica from the underlying frontal bone around the incision using a 12 mm flattened rongeur.
- 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
- Identify the sagittal suture as a reference for the anatomical midline and the coronal suture (Figure 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
- 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.
- Connect each corner with a 4 mm rounded diamond burr to ensure the correct location of the opening (Figure 4B).
- 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. - Use the first point of contact with the dura to visually assess the thickness of the remaining frontal bone.
- 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. - 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. - Apply sterile saline with a syringe to clear the view.
8. Removal of the bone plate
- 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). - 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. - 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
- 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)
- 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.
- Slowly withdraw the needle while ensuring the MDC remains at the same location within the superficial cerebral cortex.
- 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. - 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).
- 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.
- Prepare a 2 mL disposable plastic syringe with an 18 G disposable needle. Fill the syringe with 0.5 mL of sterile saline.
- 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.
- Observe the syringe during each aspiration until cerebrospinal fluid (CSF) is obtained.
- 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.
- Securely fasten the MDC with surgical tape as described above.
11. Microdialysis (MD)
- Connect each microdialysis catheter (MDC) to a separate microdialysis pump.
- 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.
- Start a tissue-calibration period of 30 min of continuous microdialysis at a flow rate of 1 µL/min.
12. Euthanaziation
- Administer a bolus of intravenous pentobarbital (50 mg/kg) through the central venous catheter (following institutionally approved protocols).
- Observe the pulse, blood pressure, and end-tidal CO2 curves on the respirator for a flatline as confirmation of cardiac arrest.
Subscription Required. Please recommend JoVE to your librarian.
Representative 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 line before making the incision. After the incision and removal of the galea aponeurotica, the sagittal suture was identified to determine the true anatomical midline (Figure 2). Following exposure of the frontal bone, the three palpable landmarks (Figure 1 and Figure 3) and the sagittal suture (Figure 2) were used to define a triangle within which the borders of the craniectomy were determined within the desired location (Figure 4A). These landmarks are well-suited for this procedure as they are closely related to the pig brain (Figure 7).
After gradually drilling and thinning the frontal bone along the borders of the craniectomy (Figure 4B), a contact point for the underlying dura mater was achieved, allowing for assessment of the remaining thickness of the frontal bone.
Finally, after carefully removing the frontal bone, the underlying dura mater covering the cranial two-thirds of both cerebral hemispheres was revealed. The procedure was considered successful because (1) the revealed underlying dura mater was intact (Figure 6); (2) the opening in the frontal bone was centered, confirmed by the location of the dorsal sagittal sinus (Figure 5); and (3) only minor venous hemorrhages occurred along the exposed spongiosa bone.
The integrity of the dura mater was assessed by visually inspecting for obvious cerebrospinal fluid (CSF) leakage through smaller defects. These leaks would pulsate synchronously with the pulsation of both hemispheres. The intact dura mater was confirmed as no visual CSF leakage was observed, and both hemispheres appeared elevated and rounded synchronously with the pulsation (Figure 6). This elevated appearance of the hemispheres indicates a net positive pressure within the CSF, equivalent to intact meninges.
The correct centered location of the craniectomy was crucial for several reasons. Firstly, it ensured maximal exposure of both hemispheres for later instrumentalization. Secondly, it reduced the risk of accidentally damaging the dorsal sagittal sinus with the drill. The correct location was confirmed after the bone plate was removed, and the contours of both cerebral hemispheres were symmetrically separated by the fissure just above the dorsal sagittal sinus (Figure 4).
Minor venous bleeding is expected from both the dura mater and the exposed trabecular bone. This can be managed by applying bone dust from the craniectomy for a few minutes to achieve hemostasis. If this is not sufficient, hemostasis can be ensured by applying low-voltage cauterization to the origin of the hemorrhage using a mono- or bipolar cautery.
Figure 1: Image of the exposed frontal bone. The three anatomical landmarks are identified by manual palpation: the nuchal prominence and the caudal aspect of each superior orbital crest. Please click here to view a larger version of this figure.
Figure 2: Image of the exposed frontal bone after dissection, revealing the sagittal suture and the coronal suture. Please click here to view a larger version of this figure.
Figure 3: 3-dimensional computed tomography scan. The scan of the posterior aspect of the porcine skull visualized from a frontolateral perspective (A) and a posteriosuperior perspective (B). Anatomical landmarks of the frontal bone are highlighted in red (superior orbital crest) and blue (nuchal crest). Please click here to view a larger version of this figure.
Figure 4: Images of the exposed frontal bone. (A) Defining each corner of the rectangular craniectomy using the nuchal prominence and superior orbital crests as landmarks. (B) Drilling the initial line of the craniectomy by connecting each corner. Please click here to view a larger version of this figure.
Figure 5: Depiction of the exposed dura mater with underlying cerebral hemispheres and sagittal suture. Please click here to view a larger version of this figure.
Figure 6: Image of the dura mater with underlying cerebral hemispheres seen from a frontal perspective. If the dura mater is intact, both hemispheres are elevated and rounded synchronously to the pulsation (marked with white stipulated lines). Please click here to view a larger version of this figure.
Figure 7: Horizontal computed tomography scan. The scan of the porcine skull revealing the anatomical relationship between the posterior orbital crest (red), nuchal crest (blue), and cerebrum (yellow). Please click here to view a larger version of this figure.
Subscription Required. Please recommend JoVE to your librarian.
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 sagittal sinus. However, it is important to note that pigs exhibit significant individual variations in the composition of their cranium, and not all pigs show a clearly defined sagittal or coronal suture, unfortunately11. Therefore, the method presented in this procedure relies on more general anatomical landmarks such as the nuchal crest and orbital prominences to improve accuracy in locating the craniectomy.
While a rounded drill at high speeds is efficient, it also entails a high risk of accidental dura lesions, especially at the initial contact point. Gradual practical experience can help lower this risk. However, pneumatic high-speed drills are generally expensive compared to alternatives like regular manual drills, which is a significant limitation. One could argue for improving the procedure by using a manual cranial drill for the initial dura contact, followed by a footed craniotome. This approach might decrease the risk of dura lesions as the dura would not be exposed to high-speed moving objects. However, in our experience, the varying thickness and distribution of sinusoids within the porcine frontal bone make the use of regular craniotomes challenging.
The overall purpose of the presented method is to achieve visual access to a large proportion of the porcine brain through a practical, simple, and time-efficient procedure. We believe that the demonstrated procedure offers several advantages compared to less invasive procedures like burr-hole craniotomies. Firstly, this method exposes a significantly larger proportion (approximately 20 cm2) of the pig brain compared to single burr-hole craniotomies (0.7-1.5 cm2, depending on the diameter of the drill). This not only allows for substantial instrumentalization of the brain (e.g., insertion of ventricular drains, neuronal sensors, multiple catheters, probes, or stimulating electrodes) but also enhances navigation for specific brain areas using visible structures such as the lateral hemisphere borders, gyri contours, or the dorsal sagittal sinus. Secondly, the brain itself is not damaged during the procedure when performed correctly, which can be achieved relatively quickly by an experienced practitioner.
Although the demonstrated procedure was used to describe the intracerebral penetration of moxifloxacin in a non-survival porcine study12, we believe that this method is suitable for survival studies too. Reinserting the bone plate and closing the overlying skin tissue through sutures would allow the pig to wake up unharmed following the procedure. However, this would only be practically and ethically feasible if the underlying cerebral cortex remains relatively unharmed from possible instrumentalization, ensuring an intact neurological state. Survival studies of this nature would require strict antiseptic techniques and close post-operative monitoring of the pigs for signs of neurological deficits due to surgical trauma or hemorrhage, such as epidural hematomas.
In general, we believe the presented method is simple, reproducible, and has significant applications within various future porcine models relevant to the central nervous system (CNS), especially in cases where considerable instrumentalization and/or monitoring of the otherwise uninterrupted porcine brain is needed.
Subscription Required. Please recommend JoVE to your librarian.
Disclosures
The authors have no conflicts of interest to declare.
Acknowledgments
The authors would like to express our gratitude for the support and technical experience shared by the personnel at the Biomedical Laboratory, Aalborg University Hospital, Denmark.
Materials
| 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 |
References
- Mariager, T., Bjarkam, C., Nielsen, H., Bodilsen, J. Experimental animal models for brain abscess: a systematic review. Br J Neurosurg. , (2022).
- Bassols, A., et al. The pig as an animal model for human pathologies: A proteomics perspective. Proteomics Clin Appl. 8, 715-731 (2014).
- Meurens, F., Summerfield, A., Nauwynck, H., Saif, L., Gerdts, V. The pig: A model for human infectious diseases. Trends Microbiol. 20 (1), 50-57 (2012).
- Swindle, M. M., Makin, A., Herron, A. J., Clubb, F. J., Frazier, K. S. Swine as models in biomedical research and toxicology testing. Vet Pathol. 49 (2), 344-356 (2012).
- Lind, N. M., et al. The use of pigs in neuroscience: Modeling brain disorders. Neurosci Biobehav Rev. 31 (5), 728-751 (2007).
- Hoffe, B., Holahan, M. R. The use of pigs as a translational model for studying neurodegenerative diseases. Front Physiol. 10, 838 (2019).
- Ettrup, K. S., et al. Basic surgical techniques in the göttingen minipig: Intubation, bladder catheterization, femoral vessel catheterization, and transcardial perfusion. J Vis Exp. 52, e2652 (2011).
- Bjarkam, C. R., Glud, A. N., Orlowski, D., Sørensen, J. C. H., Palomero-Gallagher, N. The telencephalon of the Göttingen minipig, cytoarchitecture and cortical surface anatomy. Brain Struct Funct. 222 (5), 2093-2114 (2017).
- Hou, N., Du, X., Wu, S. Advances in pig models of human diseases. Animal Model Exp Med. 5 (2), 141-152 (2022).
- Munk, M., Poulsen, F. R., Larsen, L., Nordström, C. H., Nielsen, T. H. Cerebral metabolic changes related to oxidative metabolism in a model of bacterial meningitis induced by lipopolysaccharide. Neurocrit Care. 29 (3), 496-503 (2018).
- Kyllar, M., et al. Radiography, computed tomography and magnetic resonance imaging of craniofacial structures in pig. J Vet Med C: Anat Histol Embryol. 43 (6), 435-452 (2014).
- Mariager, T., et al. Continuous evaluation of single-dose moxifloxacin concentrations in brain extracellular fluid, cerebrospinal fluid, and plasma: A novel porcine model. J Antimicrobial Chemother. , (2024).
