Subdural Soft Electrocorticography (ECoG) Array Implantation and Long-Term Cortical Recording in Minipigs

Surgical and behavioral procedures were approved by the local ethical committee in accordance with the guidelines for the Care and Use of Laboratory Animals and approved by local (Canton of Geneva) and federal (Swiss) veterinary authorities with authorization number GE11120A. Female Göttingen minipigs (n = 7) at 2-6 months of age (5-8 kg) were used in this study.

1. Presurgical planning

1. In vitro characterization of the soft implant system
   1. Chronoamperometry: Record the voltage drop upon the injection of a biphasic current pulse evolution (i.e., the voltage transient [VT]) using an oscilloscope connected in parallel to a pulse generator. Connect the pulse generator sequentially to each electrode and a platinum counter in saline solution (phosphate-buffered saline [PBS] 1x). Refer to step 3.1 for the settings.
   2. Electrochemical impedance spectrogram: Measure the electrochemical impedance at different frequencies using a potentiometer. Connect the potentiometer sequentially to each electrode using the platinum counter and an Ag/AgCl reference electrode in saline solution (PBS 1x). Refer to step 3.2 for the settings.
   3. Inter-channel resistance: In the dry state, measure the direct current (DC) resistance between adjacent channels using a handheld multimeter.
   4. Implant selection: After the three measurements cited above, select the implant along with the following criteria: impedance at 1 kHz below 100 kΩ and no inter-channel resistance below 1 MΩ.
2. Sterilization
   1. Implant sterilization: Place the selected implants individually into sterilization bags along with a sterilization marker and seal them. Employ double packaging to ensure sterility during surgery.
      NOTE: In this case, hydrogen peroxide (H₂O₂) gas sterilization is used due to the short time cycle and low temperature (55 °C). Alternatives are ethylene oxide (ETO) gas or autoclave sterilization, but compatibility with the implant system should be ensured.
   2. Instrument sterilization: Place the cleaned instruments in double sterilization bags or sterile instrument boxes along with sterilization markers. Autoclave sterilization is most common for instruments, but H₂O₂ or ETO are possible alternatives.

2. Surgical implantation of soft ECoG arrays

1. Anesthesia
   1. Premedication: Isolate the animal and fast it overnight. Inject a mix of midazolam at 0.75 mg/kg, atropine at 0.25 µg/kg, and haldol at 0.1 mg/kg intradermally, and wait until the animal is sedated. Weigh the animal before proceeding.
   2. Installation of intravenous (IV) lead:
      1. Place the animal on the surgery table on a heating pad. Induce anesthesia by placing a face mask on the animal, using sevoflurane at 3%-3.5%.
      2. Place electrocardiogram leads on the abdomen, a blood saturation sensor on the tail, and a temperature sensor in the nostril.
      3. Place an IV lead on an ear vein and confirm blood access using a syringe filled with saline. Ensure the eyes are kept hydrated by using ointment.
   3. Intubation: Inject a bolus of atracurium at 0.5 mg/kg, ketamine at 1 mg/kg, and fentanyl at 1-2 µg/kg. Place the animal on its back for intubation. Insert a 4.5 mm tube.
   4. Medication: After intubation, stop the sevoflurane anesthesia and install a propofol infusion at 10 mg/kg/h, fentanyl at 2 µg/kg/h, atracurium at 0.2-0.5 mg/kg/h, and saline at 4-7 mg/kg/h. Start an infusion of mannitol at 1 g/kg/h to reduce brain swelling during the surgery.
      NOTE: A multimodal analgesia regimen may be used if recommended by the local animal ethics committee.
2. Preoperative x-ray
   1. Place the animal on its abdomen in the sphinx position. Temporarily remove any metallic object in the vicinity of the brain and skull of the animal (e.g., temperature lead in the nostril).
   2. Acquire an axial and sagittal plane x-ray with a contrast of the bone. Place a metallic object with known dimensions in the sagittal acquisition field to serve as a scale to measure the skull thickness at the front and back of the brain.
   3. Identify the location of the frontal sinuses (visible by voids below the skull) and mark the most posterior location on the head of the animal using a permanent marker. This will indicate the furthest point where any craniotomy or screw placement can be done in the surgical approach described below.
3. Aseptic field and skin preparation: Shave the entire surface of the head beyond the surgical field. Using a sterile pad, thoroughly scrub the head with betadine. Next, place sterile drapes on the instrumentation table and on the animal to reveal only the surgical window. Finally, scrub the head again with a sterile pad using betadine.
4. Craniotomy and durotomy
   1. Skin cut: Incise the skin with a scalpel knife along the midline. Separate the muscle and periosteum (25 mm laterally from bregma on both sides and 40 mm anterior and posterior to bregma) from the bone using a raspatory and place spreaders to get optimum access for later drilling.
   2. Measurements and marking: Identify bregma and lambda, and mark them with a sterile surgical pen (Figure 2A, B). Using a sterile ruler, define the bone flap outline centered around the implantation target on both hemispheres. In this specific case, the auditory cortex was chosen, with coordinates -5 mm to -15 mm from bregma and -4 mm to -20 mm laterally. Then, adjust the craniotomy to the size of the implant and anatomical landmarks, limiting the opening size.
   3. Craniotomy:
      1. Using a bone drill with a round cutting bit, drill the outline of the craniotomy, taking into account the thickness of the skull measured in step 2.2. Irrigate the drilling location with saline solution to avoid overheating of the bone.
      2. Carefully drill the outline homogeneously until reaching the dura mater. At the first breakthrough, finish drilling the outline until it has thinned enough to almost break through. Then, use a flat spatula (either on the midline side or the lateral side) to break away the bone flap in one piece, using the craniotomy edge as leverage. If too much resistance is encountered, continue thinning the bone.
      3. Place the bone piece in sterile saline.
      4. Once the bone flap is removed, carefully chip the edge of the craniotomy away, using a Kerison to avoid the sharp bone edge from cutting into the dura mater.
      5. If excessive bleeding is encountered on the dura mater or the bone, use Gelfoam or bone wax, respectively. Place a wet compress (standard pad in sterile saline solution) in the craniotomy and repeat this step on the other hemisphere (Figure 2B).
   4. Durotomy:
      1. Using the needle from a 6-0 suture kit, carefully pierce and lift the dura mater at the anterior or posterior end of the craniotomy halfway between the medial and lateral side and create the beginning of an incision with the stab knife.
      2. Then, using a small flat spatula inserted in the subdural space acting as a cutting base to protect the cortex, create an anteroposterior slit in the dura mater by advancing simultaneously with both tools. Ensure that the slit is slightly larger than the width of the implant (Figure 2C). If any bleeding or damage occurs at this step, cover it with gel foam and wait until it stops.
         NOTE: The slit trajectory should be adapted if large blood vessels are present in the dura mater to avoid bleeding.
5. Implantation
   1. Device insertion:
      1. Irrigate the implant (Supplementary Figure 1A) with saline on both sides so it slides into the subdural space more easily. Place the implant above the dura mater slit and, with small forceps, subdurally insert the device by sliding it sequentially on each edge.
      2. Carefully hold the pedestal end of the device and advance with the implant in order not to create tension hindering the insertion. When the connector edge is located on top of the slit, stop the insertion.
   2. Secure the implant: To secure the implant in place, place a titanium bridge over the cable after the edge of the craniotomy or in the anchoring wings and secure it with one or two titanium screws using the appropriate screwdriver (Figure 2D).
   3. Ground placement: Carefully remove 1 cm of the insulation of the ground wires and insert it epidurally at the posterior end of the craniotomy (or any epidural location far from the cortex of interest or large blood vessels) (wire in Figure 2E)
   4. Repeat steps 2.4.4. and 2.5.1.-2.5.3 on the contra-lateral hemisphere.
   5. Intraoperative x-ray to confirm placement:
      1. Place a wet pad (standard pad in sterile saline solution) over the surgery location to keep the tissue hydrated. Next, place a sterile surgery drape to cover the head of the animal.
      2. Take plane x-ray images (axial and sagittal) to ensure the implants are well-placed and are not folded, using the x-ray markers as indicators. If not, remove the drape and explant the device to insert it again (follow steps 2.4.4. and 2.5.1.-2.5.3 again).
   6. Dura mater closure: Suture the dura mater carefully around the implant cable using a 3-0 resorbable suture and a small needle holder. Bring the two dura mater edges together as much as possible without tearing through the thin membrane with the suture wire (Figure 2D, E).
   7. Bone flap placement: Fix a titanium bridge on the anterior and posterior part of each bone flap using a titanium screw. Be careful to plan the positioning of the Ti-bridges with respect to the placement of the footplate legs in the next steps. Screw the end of the titanium bridges to the skull (Figure 2F, G).
6. Pedestal and footplate placement
   1. Positioning: In this configuration, the footplate has six legs with two screw holes each (Supplementary Figure 1B). Plan the placement of the footplate on the skull to optimize the location of the screws (avoid placing them at the edge of the craniotomy or in the temporal muscle). Skip the holes in the legs if they cannot be screwed in.
   2. Footplate securing: Screw in the titanium screws of the footplate until it is firmly in place (see Figure 2H).
   3. Pedestal placement: Remove the titanium bridges over the connection cables and flip over the pedestal to land on the footplate. Screw the pedestal onto the footplate. Check that the pedestal is firmly in place (Figure 2I).
7. Suture and closure
   1. Cleaning of the wound: Clean the subcutaneous space of any bone or other debris by flushing with saline. Cut away some skin around the pedestal edges to create a round edge following the cylinder.
   2. Subcutaneous sutures: Remove the spreaders and fold the skin flaps together. Create subcutaneous sutures with a 4-0 non-resorbable suture wire, 3 mm apart using simple interrupted sutures or simple continuous sutures. Start away from the pedestal, moving toward it on both sides of the incision.
   3. Dermal sutures: Suture the skin using a 6-0 non-resorbable suture wire, with sutures 5 mm apart. Start away from the pedestal, moving toward it on both sides of the incision. Take care to achieve good tissue apposition between the two skin flaps and near the pedestal edge to avoid a void (Figure 2J).
   4. Wound dressing: Clean the wound area again with a sterile pad and betadine. Apply a self-adhesive sterile bandage over the wound.
8. In vivo measurements: For in vivo measurements, follow sections 3, 4, and 5.
9. Awakening: After all the measurements have been performed, take the animal off all anesthetics but keep under ventilation. For analgesia, apply a buprenorphine patch (25 mg/h) for 24 h. Place the animal on a heating pad covered with drapes to speed up the wake-up time. When spontaneous breathing is recovered, extubate the animal and put under an oxygen face mask until consciousness is recovered (which can take 1 to 4 h).
10. Postoperative animal care: For 5 days, keep the animal under close surveillance. Give a dose of cephalexin at 75 mg twice daily with food, separated from other animals. Carry out disinfection of the wound daily by applying copious amounts of betadine with soaked sterile pads (best done during feeding).
    NOTE: Long-term care and housing: The operated animal is kept isolated for 24 h. It is put back in its original social group if the animal is well enough to interact socially with its peers. Daily observation of the pedestal and skin opening needs to be conducted to follow the integration of the device on the head. When appropriate, clean the location around the pedestal with copious amounts of betadine.

3. In vivo characterization of the soft implant

1. Chronoamperometry: Record the voltage drop upon the injection of a biphasic current pulse evolution (i.e., the VT) using an oscilloscope connected in parallel to a pulse generator. Connect the pulse generator sequentially to each electrode and the ground wire. Perform the stimulation pulse at 100 µA with a pulse width of 300 µs at 100 Hz.
2. Electrochemical impedance spectroscopy: Measure the electrochemical impedance at different frequencies using a potentiometer. Connect the pulse generator sequentially to each electrode, using the ground wire as both the counter electrode and the ground. Set the excitation voltage to 200 mV and the frequency range from 1 Hz to 1 MHz, with three points per decade.
3. Inter-channel short measurements: Measure the DC resistance between adjacent channels using a handheld multimeter. In vivo, the inter-channel DC resistance is only measured to verify that no shortages below 1 kΩ appear, indicating total encapsulation failure.

4. Electrophysiological recording

1. Spontaneous activity: Plug the wireless recording system through the pedestal and record the baseline activity for 2-3 min. These recordings will serve as a control to analyze the auditory evoked potentials.
2. Auditory evoked potentials: In addition to the wireless system, insert speakers in a closed field in the animal ears. Play tone burst acoustic stimulation at different frequencies (ranging from 200-20,000 Hz) at around 70 dB sound pressure level (SPL) over 120 repetitions. Then, average the recordings and align them over the stimulus period for analysis.
3. Sensory evoked potentials: Place the needles in the snout at three different positions. Evoke sensory potentials by stimulating the snout for ~30 s with the pulse generator at different amplitudes to obtain the recruitment curves.

5. In vivo imaging

1. Animal transport: Keep the animal under propofol anesthesia, as described in step 2.1. Use a transport cart that houses a ventilator, syringe pump, and vital signs monitor to transport the animal from the surgery room to the imaging facilities and back.
2. Computed tomography x-ray scan: Place the animal on the scanner table and remove any metallic object around the head (e.g., the temperature sensor). Acquire a CT scan at the smallest resolution (0.4 mm slice thickness) using isometric acquisition with automatic current and voltage selection for bone contrast.
3. Magnetic resonance imaging: Remove all equipment containing metal from the animal (use MRI-compatible IV leads and intubation tube). Keep the animal ventilated and anesthetized under sevoflurane at 3%-3.5%, using a ventilator situated outside the MRI chamber and linked through a long tube to the animal. Before the first sequence, inject a bolus of fentanyl at 1-2 µg/kg. Use three isometric sequences at the smallest resolution: T1-, T2- and turbo spin echo (TSE)-weighted sequences (parameters shown in Supplementary File 1).

6. Freely moving recording

1. Follow the same procedure described in section 4 for recording awake signals from the brain. Plug the wireless headstage by either holding the animal in the experimenter's arms or feeding the animal with treats to distract it. Provide the acoustic stimulation using external speakers placed close to the animal.

7. Perfusion and tissue preparation

1. OPTIONAL: If the animal is not already under anesthesia, follow step 2.1 for the anesthesia protocol.
2. Insertion of a catheter in the carotid artery for perfusion (Supplementary Figure 2)
   1. Carotid artery and jugular vein dissection: Cut the throat at the midline using a cauterizer/cutter. Cut the skin first and then the muscle following the middle white line (no vessels underneath) (Supplementary Figure 2A).
   2. Open further, using fingers to make space around the trachea and under the muscle. Look for the carotid artery (beating and pinkish); the vagal nerve is sometimes also around (white), and the jugular vein can be underneath or on the side (red). Place the spreaders (see Supplementary Figure 2B).
   3. Start the dissection of the carotid artery using fine forceps and round scissors. Use scissors to open conjunctive tissue. If there are no blood vessels, cut; if there are blood vessels, cauterize and move forward (Supplementary Figure 2C, D). When dissected enough, use a clamp to go underneath the carotid artery so it is completely isolated (Supplementary Figure 2E).
   4. Repeat the same operation with the jugular vein (Supplementary Figure 2F).
   5. When both vessels are fully dissected and isolated, place suture wire around them. Do not close them yet. Two sutures around the carotid, one at the very base (suture 1-heart side for brain perfusion) and one on the other side (suture 2), are shown in Supplementary Figure 2G.
   6. Place one suture around the jugular vein (suture 3), not closed; mark the wires with tape for sectioning of the vein later.
   7. Close suture 1 very firmly, otherwise it will bleed (Supplementary Figure 2H). Tie three knots. Place a clamp on the wire to put weight and create tension in the carotid artery.
   8. Clamp the carotid artery using a vessel clamp on the opposite side of the carotid artery dissection (brain side for brain perfusion; Supplementary Figure 2I). Suture 2 is in the middle, not closed yet.
   9. Use black forceps to catch and pull the carotid artery. Using fine scissors, section half of the carotid artery near the base of the dissection (near suture 1, heart side; see Supplementary Figure 2J). Ensure that the section is as neat as possible and reaches the vessel itself, not only the "sheath"; otherwise, the catheter will not go through. Insert the catheter as shown in Supplementary Figure 2K.
   10. Flush and fill the catheter, first with PBS, so there is no air left in the catheter (Supplementary Figure 2K inset).
   11. Close suture 2 firmly enough so it does not go away, but not too much so that the catheter can still move a bit (Supplementary Figure 2L). Then, remove the vessel clamp, finish inserting the catheter as far as possible, and finalize the closure of suture 2 (close firmly).
   12. If desired, use suture wires at suture 1 to attach the base of the catheter to the skin at the exit of the wound as an extra security step. Then, transfer the animal to the perfusion zone and plug it to PBS/heparin. When the perfusion starts, pull suture 3 and cut the jugular vein
3. Euthanasia: Deliver pentobarbital (90 mg/kg) intravenously and flush the line with saline to ensure the full dose is administered successfully.
4. Perfusion: Using a perfusion pump (200 mL/min) plug the carotid catheter into PBS/heparin (1 L for a 15 kg pig) and then into PBS containing 4% paraformaldehyde (PFA) (5 L for a 15 kg pig). When the perfusion starts, pull suture 3 and cut the jugular vein.
5. Tissue collection
   1. Beheading: When perfusion is over, detach the animal's head from the body using a scalpel, by cutting through the skin and muscle and inserting the blade between the first and second vertebrae.
   2. Postfixation: Plunge the head in 4% PFA in PBS for another 48 h at 4 °C and then transfer to PBS before brain extraction.
   3. Brain and implant extraction:
      1. Remove the skin using a scalpel and start to cut the bone carefully using a rongeur starting from the first vertebrae, following the spinal cord to the cerebellum. Once the cerebellum is exposed, carefully remove the temporal bones and expose the parietal and frontal lobes.
      2. At this point, screw the pedestal off the footplate and cut the feet of the footplate with pliers. Remove the bone near the cable entry into the skull to free the implant system from the bone, without pulling the implants out of the dura mater exit.
      3. If the implant cables are embedded into the bone, cut the cable as close as possible to the exit. Once enough brain surface is exposed, carefully cut the dura following the midline using small scissors.
      4. Free the implant cables from the dura mater exit. Take photographs from the implant placement on the brain. Then, use a small spoon to detach the brain from the cranial nerves below. Carefully extract the brain. Remove the implants from the brain.
   4. Postfixation of the brain: Depending on the perfusion quality, postfix the extracted brain again for 24 h in 4% PFA in PBS at 4 °C. Keep in 0.1 M PBS at 4 °C until the brain is cut.
   5. Brain cut: Separate the two hemispheres using a razor blade. Then, cut the brain orthogonally in four different pieces. Cut the implanted zone in half to obtain two blocks containing the implanted zone and a control area. Store the other two blocks if more control slides are needed.
   6. Brain cryoprotection and freezing: Transfer the brain blocks to a solution of 15% of sucrose first and then 30% of sucrose at 4 °C until the brain plunges and reaches equilibrium. Then, freeze the tissue in isopentane at -55 °C in a tissue freezing system.

8. Histology

1. Tissue sectioning
   1. Cryostat: Place the frozen brain in a cryostat and trim until full sections are achieved. Then, slice the brain into 40 µm sections and immerse in groups of three in 0.1 M PBS in well plates. Carefully note the order of the plates.
   2. Section selection: Select the sections depending on the zone to analyze (implanted region or control zone). Take the sections sequentially out of the well plates, using a fine brush to inspect them for damage. Place them in new well plates filled with 0.1 M PBS for further staining.
2. Immunohistochemistry
   1. Preparation: Incubate the sections in 0.3% Triton X/PBS for 15 min, followed by 3% bovine serum albumin (BSA)/PBS for 1 h at room temperature (RT).
   2. Primary antibodies: Incubate the tissues with primary antibodies for 48 h at RT (anti-GFAP, rat, diluted at 1/300; anti Iba1, rabbit, diluted at 1/400; anti-NeuN, guineapig, diluted at 1/1,000; all in 1% BSA/PBS). Cover the well plates with aluminum foil.
   3. Washing: Wash the wells with 0.1 M PBS three times for 5 min.
   4. Secondary antibodies: Incubate the tissues with secondary antibodies for 2 h at RT (Alexa Fluor 488, Alexa Fluor 647, Alexa Fluor 555; all diluted at 1/400 in 1% BSA/PBS).
   5. DAPI (4′,6-diamidino-2-phenylindole): Incubate the tissues with DAPI for 15 min (1/1,000 in 1% BSA/PBS).
   6. Washing: Wash the wells with 0.1 M PBS five times for 15 min.
   7. Mounting: Mount the slides using mounting media and a coverslip. Keep the slides in the dark in a fridge at 4 °C.
3. Imaging
   1. Whole slide imaging: Image the slides at 10x magnification (objective working distance value = 3,100 µm) using a slide scanner microscope at three different wavelengths (640 nm, 560 nm, 485 nm). All power and gain information can be found in Supplementary File 2.
   2. Microscopic imaging: Image the region of interest at 20x magnification (apochromat 20x/0.8 M27) using a confocal microscope at four wavelengths (Alexa Fluor 647, DAPI, Cy3, EGFP). All power and gain information can be found in Supplementary File 3.