Use of a Piglet Model for the Study of Anesthetic-induced Developmental Neurotoxicity (AIDN): A Translational Neuroscience Approach

Healthy, domestic piglets (Sus scrofa) are obtained from a farm approved by The Ohio State University Institutional Animal Care and Use Committee (IACUC). All animal experimentations are performed in accordance with The Ohio State University IACUC policy, after protocol approval.

1. Animals and Animal Handling

1. Use male piglets in this experiment to eliminate the potential confounding effects of sex. NOTE: If experimental goals include evaluation of effects of the experiment on animals while in the period of maximal brain growth, do not use piglets older than 14 days.
2. Schedule the piglets to arrive in the vivarium at least 24 h before experimentation to allow acclimation to the environment.
   NOTE: Trained veterinary technicians supervised by licensed veterinarians provide routine animal care.
   1. Keep the piglets in individual temperature-controlled cages and give a nutritionally complete, commercial piglet milk replacer ad libitum. Supply the animals with a blanket and a toy. Continuously monitor the temperature in the animal enclosures.
3. For this preliminary feasibility study, used 18 piglets for the isoflurane arm and 22 piglets for the control arm. Perform sample size calculations based on study design when possible. Randomize the available piglets to either control or exposure group for the appropriate length of exposure. From experience, even with multiple investigators, expect to be able to perform no more than 2 experiments per day (2 animals total).

2. Control Animals

1. Do not perform experimental intervention on control animals.
2. Induce deep general anesthesia via face cone mask for the perfusion and tissue collection procedure. Specifically, after the 24-h acclimation period, anesthetize the piglets with 5% isoflurane or 8% sevoflurane in 100% oxygen via face cone mask. Do not use desflurane for induction.
   NOTE: The time between induction of anesthesia and institution of cold PBS perfusion should be as short as possible. Experienced technicians can complete this process in under 5 min.
   1. Confirm adequate depth of anesthesia by lack of dewclaw-pinch reflex using a surgical clamp.
   2. To avoid hypoxic/ischemic insults to the brain, monitor the piglet using a pulse oximeter to ensure maintenance of adequate oxygenation, ventilation, and cardiac output until perfusion of cold phosphate-buffered saline (PBS) begins.
      NOTE: To provide additional protection against tissue damage, pack the animal (including the head) in ice after induction of anesthesia.
3. Perform a transcardiac perfusion.
   NOTE: because paraformaldehyde is used, the perfusion procedure should be performed under a fume hood or on a downdraft table.
   1. Make a craniocaudal incision along the length of the sternum using a scalpel. The depth of the incision should be sufficient to expose the sternum.
   2. Carefully, perform a midline sternotomy with a pair of sharp heavy scissors, avoiding damage to the heart, lungs, or diaphragm. If necessary, place a finger placed between the posterior aspect of the sternum and the intrathoracic contents to avoid injuries. Maneuver a finger into the mediastinum by making a small (finger-sized) incision in the diaphragm.
   3. After entering the thoracic cavity, keep the rib cage open using a self-retaining retractor.
   4. Incise the pericardium using forceps and a pair of scissors, exposing the beating heart. Use caution not to injure the heart.
   5. Identify the left ventricle and carefully place a cannula (such as a 14 G angiocatheter) through the apex of the ventricle. Remove the needle, leaving the catheter in place.
      NOTE: Use caution not to puncture the posterior wall of the ventricle. Pulsatile blood return from the catheter indicates that it is properly placed. Blood can easily be sampled from the animal at this point.
   6. After identifying the right atrium, perform an atriotomy by making a large incision in the atrium with scissors to allow for exsanguination and the escape of perfusate.
      NOTE: Isoflurane via inhalation should be continued until cardiac death is confirmed. Cardiac death is confirmed by direct observed lack of cardiac output.
4. Perfuse the piglet using a perfusate consisting of cold (4 °C) phosphate-buffered saline (PBS) containing heparin at a concentration of 5 units per mL. Perfuse at 300 mL per min for 5 min or until the solution runs clear.
   1. Use caution that the perfusion cannula does not become dislodged during perfusion. Use a commercially available peristaltic pump for this and all other perfusions.
5. Perform a hemicraniectomy to remove one hemisphere of the brain for fresh tissue analysis.
   NOTE: This protocol allows for retrieval of one hemisphere of fresh brain tissue. The other hemisphere is fixed. If no fresh tissue is required, skip to step 2.6.
   1. During this procedure, continue the circulation of ice-cold PBS at a rate of 50 mL per hour to ensure that the brain remains cold.
   2. Make a longitudinal incision in the scalp along the length of the sagittal suture until the foramen magnum using a scalpel. During the process, use firm pressure to create a score in the skull. Reflect the scalp to expose the entire cranium.
   3. Using rongeurs and beginning at the foramen magnum, remove the skull on one side by inserting the rongeurs between the skull and the dura mater, using caution not to injure the underlying brain tissue. Remove bone in pieces, using the rongeurs to pry it away from the brain parenchyma.
   4. Once the skull has been removed, incise and remove the dura mater using forceps and scissors, again using caution not to injure the underlying brain tissue.
   5. Place a scalpel blade between the two hemispheres to carefully divide the corpus callosum.
   6. Using a flat tool such as the broad handle end of forceps, gently retract the frontal lobe, gradually severing the cranial nerves, working anterior to posterior. At the most posterior aspect of the hemisphere, use a scalpel to cut the spinal cord. Remove the hemisphere en bloc.
      NOTE: Unfixed brain tissue is fragile. Use caution when removing the hemisphere to prevent disruption of the remaining hemisphere's blood supply.
   7. Section the removed hemisphere. If indicated, immediately flash-freeze in 2-methylbutane cooled to -160 °C in a liquid nitrogen bath to avoid tissue breakdown, and immediately stored at -80 °C for later analysis.
      NOTE: We recommend sectioning brain coronally in 2 mm increments using a matrix, but specific details of sectioning will depend upon specific experimental goals.
6. Change the perfusate to 4% paraformaldehyde (PFA). Continue the PFA perfusion at 300 mL per minute for at least 5 min.
   CAUTION! PFA is toxic, avoid contact with skin, eyes, or mucous membranes. Do not inhale PFA fumes.
7. Expect the piglet's body to stiffen due to the formation of aldehyde cross-linkages being created in muscle. After perfusion of PFA is complete, remove the remaining hemisphere in identical fashion to that described in step 2.5.5.
   NOTE: Properly perfused brain will be pale and completely exsanguinated.
   1. Place the remaining hemisphere in a small container with fresh 4% PFA at 4 °C. Keep the hemisphere in PFA for 24-48 h to complete the fixation process.
   2. After 24-48 h, move the fixed brain to a solution of PBS containing 0.1% sodium azide, as it is essential to prevent over-fixation. Over-fixation could result in masking of the epitope or strong non-specific background staining. The addition of sodium azide prevents bacterial growth.
      NOTE: The tissue can be stored for up to one month at 4 °C.

3. Isoflurane (Experimental) Animals

NOTE: Any anesthetic or intervention can be used, but we do not recommend desflurane for inhalation induction.

1. Induction and maintenance of anesthesia:
   1. Perform the anesthesia using a clinical anesthesia workstation equipped with a pediatric ventilator and monitoring devices.
   2. After the 24 h acclimation period, anesthetize the piglets with 8% sevoflurane in 100% O₂ via face cone mask.
   3. Continuously monitor pulse oximetry, non-invasive blood pressure, electrocardiography, and temperature during the induction period and at all times during the study procedure.
   4. After induction, titrate sevoflurane or isoflurane to a concentration that allows for adequate depth of anesthesia while ensuring sustained spontaneous respiration (typically at a concentration of 3-4%).
   5. Place a 24 G peripheral intravenous catheter in the marginal ear vein (Figure 1).
   6. Place the piglet in the dorsal recumbent position for tracheal intubation (Figure 2, panel A). Use a Miller #1 or #1.5 blade to facilitate instrumentation of the hypopharynx and intubation of the trachea. NOTE: An experienced operator and assistant are needed during laryngoscopy.
      1. Have an assistant displace the tongue of the animal using a dry gauze to facilitate exposure of the larynx and visualization of the vocal cords (Figure 2B).
         NOTE: The piglet epiglottis is morphologically similar to that of humans (Figure 2C). The piglet vocal cords can be difficult to visualize since they are several millimeters deep within the laryngeal inlet (Figure 2D).
      2. Displace the epiglottis: place the tip of the laryngoscope blade underneath the epiglottis and lift the blade upwards to expose the larynx.
      3. Prior to placement of the tube into the trachea, spray the vocal cords with 0.5 mL of 2% lidocaine to prevent laryngospasm during the passage of the endotracheal tube, as piglets are especially prone to laryngospasm.
   7. Place and secure a 3.0 mm cuffed tracheal tube.
      1. Ensure bilateral breath sounds and sustained end-tidal carbon dioxide using chest auscultation with a stethoscope and EtCO₂ monitoring.
      2. Inflate the piglet's lungs to a continuous airway pressure of 20 cm H₂O. Then, inflate cuff of the endotracheal tube to the minimum pressure required to prevent air leak at a pressure of 20 cm H₂O, listening over the trachea to identify air leak if present.
         NOTE: This is important to prevent mucosal ischemia during intermittent positive pressure ventilation.
      3. Normoxia and normocarbia are maintained during anesthesia.
   8. Start administration of 2% isoflurane in 50% oxygen/50% air. Titrate oxygen to maintain PaO₂ of 90 – 100 mmHg. Continue for 3 h (or desired experimental duration).
   9. Apply ophthalmic ointment on eyes to prevent dryness for the duration of anesthesia.
2. Begin femoral artery catheterization after initiation of 2% isoflurane.
   1. Administer broad-spectrum antibiotics pre-incision (cefazolin, 25 mg/kg) via the peripheral intravenous line to prevent surgical site infection.
   2. Sterilize both groins using tinted chlorhexidine to ensure proper sterile field, and place an appropriate sterile drape (Figure 3B). At a minimum, staff participating in survival surgery should wear a surgical cap, mask, sterile gloves, and eye protection.
   3. Palpate the femoral pulse at the inguinal crease using index and middle fingers.
   4. Make a superficial, 1.5 cm, craniocaudal incision using a scalpel (Figure 3C).
   5. Dissect between the two heads of the gracilis muscle using a blunt instrument, such as surgical hemostat or blunt-tipped scissors (Figure 3D).
      NOTE: The femoral neurovascular bundle is typically found between and just deep to these two muscles. (Figure 4A).
   6. Using vascular loops or silk ties, isolate the artery at the proximal and distal end. Use the loop to pull the artery up to the level of the skin distally (Figure 4B).
   7. While placing traction on the proximal tie sufficient to interrupt blood flow, make a small arteriotomy with a pair of tenotomy scissors.
      1. Take caution to not transect the artery. A small arteriotomy is sufficient. Alternatively, use a needle-and-wire approach to access the artery (Figure 4C).
      2. Gentle traction on the proximal tie will prevent excessive blood loss at any point during this portion of the procedure. If using the needle-and wire approach, pass the guidewire (supplied with the kit or obtained separately) through the needle and into the artery up to 5 cm.
      3. Take caution to not advance the wire further as it can cause ventricular ectopy. Withdraw the wire immediately 1-2 cm if this occurs.
   8. Remove the needle from the vessel, taking care to leave the guide wire in the vessel. Gently pass the catheter over the wire and into the vessel (Figure 4D).
      1. Use a 3-French, 8 centimeter catheter for femoral artery catheterization. Expect mild resistance when the tip of the catheter first enters the superficial wall of the vessel.
   9. If using the arteriotomy approach, advance the catheter or polyethylene tubing directly into the vessel. Blood return should be observed immediately.
   10. Immediately attach the catheter attached to a pressure transducer. Flush the catheter with normal saline to maintain catheter patency.
       1. Place sutures to secure the catheter in place. Cover the incision with sterile gauze to prevent contamination. Utilize a percutaneous approach to femoral artery catheterization.
          NOTE: Ensure that the ultrasound is performed by an experienced technician.
3. Intra-operative conditions and monitoring:
   1. Actively warm the piglet with a forced-air warming device and continuously monitor rectal temperature (Figure 3A).
   2. Infuse dextrose-containing, isotonic fluid (5% dextrose in Ringer's lactate or normal saline) at maintenance rate (4 times the piglet's weight in kilograms, mL/hr).
   3. Monitor vital signs for perturbations (hypotension, arrhythmia, hypo/hyperthermia, hypoxia)
      NOTE: Normal vital sign ranges and corresponding suggested management in the case of abnormalities are summarized in Table 1.
   4. Using a commercially available blood analysis system, measure arterial blood gases (arterial pH, pCO₂, pO₂), electrolytes (bicarbonate, base excess/deficit, sodium, potassium, ionized calcium), hemoglobin, and glucose at least hourly during the experimental period. Draw arterial blood sample from the femoral artery catheter.
      NOTE: Normal acid-base and electrolyte values along with recommendations for correction if abnormalities are seen are summarized in Table 1.
4. After 3 h of isoflurane exposure, remove the femoral artery catheter.
   1. Carefully tie down the proximal vascular silk to permanently occlude the femoral artery to prevent bleeding. Alternatively, use a vascular clip.
   2. Ensure complete hemostasis before closing the incision. Irrigate the incision with 10-20 mL of sterile saline solution to help prevent infection.
5. At the conclusion of the experiment, close the skin incision with simple, interrupted sutures using a 3-0 non-absorbable suture material.
   1. Infiltrate the wound using 0.5-1 mL/kg of 0.25% bupivacaine with 1:200,000 epinephrine for pain control. Coat the incision with a sterile, surgical skin adhesive.
      NOTE: No dressing is required.
6. Discontinue the anesthetic and allow the piglet to awaken.
7. Remove the endotracheal tube upon signs of awakening (opening of the eyes, attempts to stand, kicking, and opening and closing of the mouth), with signs of stable hemodynamics, adequate oxygenation, and adequate ventilation.
8. After extubation, supply supplemental oxygen via face cone until adequacy of oxygenation and ventilation are assured.
9. Administer buprenorphine 0.05 mg/kg subcutaneously for additional pain control. Alternatively, transdermal fentanyl can be used.
10. When appropriate, return the piglet to its home cage. Do not leave the piglet unattended until it has regained sufficient consciousness to maintain sternal recumbency. Do not return an animal to a cage with other animals until it has fully recovered from anesthesia.
    1. Actively warm the private home cage with a warming light.
    2. Closely monitor the animal after anesthesia by trained veterinary or research staff. Provide the piglet milk replacer.
11. Allow the piglets to recover for 48-72 h, depending on experimental goals.
    1. Inject piglets with buprenorphine subcutaneously at an appropriate dose, every three hours as needed to ensure pain control, at the discretion of animal care staff experienced to control post-surgical discomfort in animals.
    2. Monitor the piglets every hour for the first 6 h after surgery and every 4 h thereafter. Perform animal sacrifice and tissue procurement in identical fashion to control animals, described above.