Implantation and Control of Wireless, Battery-free Systems for Peripheral Nerve Interfacing

All procedures described in this protocol are carried out in accordance with the NIH Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee (IACUC) of Northwestern University. This protocol follows the animal care guidelines of Northwestern University's Center for Comparative Medicine and IACUC. It is necessary to consult with IACUC when adapting the protocols.

1. Wireless electronic stimulator fabrication (Figure 2)

1. Use copper/polyimide/copper (18 µm thick top and bottom copper, 75 µm thick polyimide) as a substrate for the radio frequency power harvester coil (i.e., wireless receiver antenna).
2. Use direct laser ablation to pattern holes for the electrodes on the top and bottom copper layers and shape the device. Electrically connect the top and bottom layers using silver paste through the holes.
3. Attach the electronic components with commercial packaging, such as diode and capacitor, via soldering.
4. Use the bioresorbable dynamic covalent polyurethane (b-DCPU; 200 µm thick) encapsulated molybdenum (Mo; 15 µm thick; serpentine structure) as stretchable extension electrodes¹⁹.
5. Form the cuff electrode for the interface between the device and the nerve using poly(lactic-co-glycolic acid) (PLGA) film (300 µm thick).
6. After connecting the wireless receiver antenna and stretchable extension electrode, encapsulate the wireless receiver antenna and the connection with commercialized waterproof epoxy or polydimethysiloxane (PDMS). See Figure 2 (right) for the fully assembled device.
7. Confirm the wireless operation of the device, using a waveform generator to generate monophasic electrical impulses via the primary coil (i.e., transmission coil).
   ​NOTE: Examining the recruitment of peripheral axons and induction of axonal regeneration by monophasic and biphasic stimuli, prior studies reported a negligible effect due to the differences in waveform characteristic²⁰, and this group has been able to achieve therapeutic electrical stimulation enhancement with the same monophasic current parameters in mice²¹ and rats¹⁸. Furthermore, prior studies examined biocompatibility in vivo and in vitro and did not find any evidence of tissue damage from heating effects or the materials themselves. Because of these findings and the limited duration of therapeutic electrical stimulation in the present study, monophasic, rather than biphasic, stimuli were used in this protocol.
8. Measure the resulting direct current output voltage with an oscilloscope connected to the cuff electrode.

2. Device preparation for implantation

1. Place the implant devices into a sterile Petri dish and seal it with parafilm.
2. Irradiate the devices with UV light for 30 min per side.

3. Surgical procedure of rat right sciatic nerve implantation of wireless, battery-free peripheral nerve interface for electrical stimulation (Figure 3)

NOTE: Maintain sterile conditions. Perform surgeries within the designated surgical area of an animal procedure room. The surgeon will don a facemask, coat, cap, and sterile gloves during surgery. If more than one surgery is performed, change sterile gloves between animals and use clean, sterile surgical instruments for each surgery. Sterilize tools between surgeries by heat sterilization (autoclave or glass bead sterilizer). Use adult Sprague-Dawley rats that weighing 200-250 g.

1. Induce anesthesia using isoflurane gas anesthesia (3% induction, 1-3% maintenance) in oxygen (2 L/min), with subcutaneous administration of meloxicam (1-2 mg/kg). Cover the eyes of the rats with designated ophthalmic ointment to prevent drying.
2. Place the rats in a prone position on disinfected surgical tables for subsequent procedures. For the remaining surgical duration, assess breathing rate (should be ~2/s), tissue color, and depth of anesthesia no less than every 15 min, and maintain isoflurane levels accordingly. Confirm the appropriate depth of anesthesia by checking pedal reflex (lack of response to a firm toe pinch). Monitor the mucous membranes, which should remain pink and moist.
3. Shave the surgical area, including the right leg and lower half of the back. Scrub the shaved surgical area with a betadine pad, followed by a 70% medical ethanol swab, and repeat this scrub process three times for skin disinfection.
4. Make a 1.5 – 2 cm incision in the skin parallel to the right femur bone using tissue scissors, followed by blunt separation of subcutaneous connective tissue on the back (directly medial to the incision) to clear a subcutaneous pocket for the receiver coil (Figure 4A). Make a subsequent incision (1.2-1.5 cm) on the right gluteal muscle parallel to the skin incision.
5. Gently isolate the sciatic nerve with metal dissection probes with blunt ends (Figure 4B).
   NOTE: The sciatic nerve is located deep to the biceps femoris and runs parallel to the femur. A dissecting scope is suggested.
6. Implant the wireless, battery-free device on the sciatic nerve (Figure 4C) by wrapping the cuff around the isolated right sciatic nerve, without putting the nerve under tension or distorting its path^18,19,20. Mark out on the skin where the receiver coil is placed for further electrical stimulation.
7. Suture the gluteal muscle incision using absorbable sutures (Figure 4D).
   NOTE: The upper half of the receiver coil sits above the gluteal muscle and the cuff interface beneath it.
8. Close the skin incision with wound clips (or buried suture; Figure 4E). Match the skin edges.
9. Deliver 1 h of continuous postsurgical 20 Hz electrical stimulation with 200 µs pulse width under anesthesia (Figure 4F). Return animals to their home cages after they have fully recovered from anesthesia.
   NOTE: The detailed protocol is described below. The receiver coil is shown above the skin in Figure 4F.
10. Postsurgical treatment
    1. Place the rat in a recovery cage without bedding, lined with paper towels, with half the cage placed on an appropriate temperature-regulated heat source (approved heating pad).
    2. Monitor the rat carefully until it is ambulatory. Once ambulatory and assessed to be stable, return the rat to the home cage and watch for social reintegration.
    3. After acute recovery, monitor the rats for incision site infection and for symptoms of neurogenic pain, including but not limited to guarding, writhing, scratching, and self-mutilation. Monitor the rats daily for the 5-day postsurgical recovery period, and at least once every three days after that if the rats are not sacrificed at day 5.
    4. Administer meloxicam (1-2 mg/kg) subcutaneously once daily for two to three days after the surgery, depending on the animal's displayed level of pain/discomfort. If persistent pain is suspected, continue meloxicam beyond this period postoperatively, and if it proves refractory, euthanize the rat early in consultation with the veterinary team.
    5. Remove the skin sutures or wound clips 10-12 days post-surgery.

4. Surgical procedure of rat left phrenic nerve implantation of wireless stimulators (Figure 5A)

NOTE: Maintain sterile conditions, as in section 3. Use adult Sprague-Dawley rats weighing 200-250 g. Sterilize all surgical tools before use.

1. Induce anesthesia using isoflurane gas anesthesia (3% induction, 1-3% maintenance) in oxygen (2 L/min), with subcutaneous administration of meloxicam (1-2 mg/kg). Cover the rat's eyes with designated ophthalmic ointment to prevent desiccation.
2. Place the rats in a supine position on disinfected surgical tables for subsequent procedures. For the remaining surgical duration, assess breathing rate, tissue color, and depth of anesthesia no less than every 15 min, and maintain isoflurane levels accordingly. Confirm the appropriate depth of anesthesia by checking pedal reflex (lack of response to a firm toe pinch). Monitor the mucous membranes, which should remain pink and moist.
3. Shave the surgical area on the ventral aspect of the neck. Scrub the shaved surgical area with a betadine pad, followed by a 70% medical ethanol swab, and repeat this scrub process three times for skin disinfection.
4. Administer bupivacaine (2 mg/kg, diluted in saline not exceeding total volume 0.5mL) subcutaneously at midline on the neck, targeting the most superficial layer. Make a 3 cm midline incision through the skin and superficial cervical fascia to expose the sternohyoid and sternocleidomastoid muscles (Figure 5B).
   NOTE: A dissecting scope is suggested.
5. Elevate the sternocleidomastoid using gentle blunt dissection with a probe and retract it laterally using a vessel loop (Figure 5C). Gently free and retract the omohyoid. Next, gently free and medially retract the vagus nerve and the carotid bundle beneath the omohyoid muscle.
   NOTE: The major discrimination here is between the vagus nerve and the phrenic nerve. Cut the omohyoid if it is necessary to expose the beneath structures.
6. Isolate the phrenic nerve (Figure 5D).
   NOTE: The phrenic nerve runs along the surface of the anterior scalene muscle, running as a notably small longitudinal nerve crossing perpendicularly to the brachial plexus. Unlike the sciatic nerve, the anatomy around the phrenic nerve at the neck is more complex. Perform electrophysiological confirmation (step 4.7) before implantation for best results.
7. Place the recording electrode subcutaneously, just caudally to the rib cage, ipsilateral to the isolated phrenic nerve (Figure 5E). Place the stimulators on the phrenic nerve and confirm via synchronous signaling (Figure 6).
   NOTE: Evoking a maximal response with a stimulus intensity of ~3-6 mA and stimulus duration of 0.02 ms is typical.
8. Verify complete transection of the phrenic nerve by showing complete abolishment of the evoked response when an electrical stimulus is applied on the proximal nerve end relative to the transection site (Figure 6).
9. Implant a wireless, battery-free device on the phrenic nerve (Figure 5F) by placing the receiver coil of the implantable device on the sternohyoid, deep relative to the bilateral sternocleidomastoid muscles, with the cuff around the phrenic nerve and the contact electrodes positioned perpendicular to the nerve.
10. Close the superficial cervical fascia with simple running absorbable sutures (Figure 5G). Close the skin with interrupted inverted absorbable sutures in the deep dermis. Return the animals to their home cages only after they have fully recovered from anesthesia.
11. For postsurgical treatment, follow step 3.10.

5. Wireless delivery of therapeutic electrical stimulation

1. Apply electrical stimulation for 1 h to the rats under general anesthesia. For the wireless stimulation, place a waveform/function generator (voltage: 1-15 Vpp) and optional amplifier above the animal to provide electrical power to an external inductive coil (i.e., transmission coil) (two-dimensional spiral coil with 5 turns; diameter: 2 cm) to ensure good inductive coupling with the implanted receiver coil. Deliver monophasic, 200 µs pulses at 20 Hz for 1 h duration.
2. To verify and quantify electrical stimulation delivery, record CMAPs from the tibial anterior muscle, adjusting the stimulation voltage to deliver supramaximal activation of the sciatic nerve. Use concentric needle electrodes for all recordings.
   NOTE: If function-generated maximal voltage is insufficient to evoke a maximal response, use an amplifier.

6. Euthanasia

1. Primary method
   1. Place the cage under a CO₂ delivery chamber, set to a rate of 8-12 LPM (or appropriate flow rate based on the chamber size). Monitor the rats for unconsciousness and then for at least 1 min of cessation of breathing.
2. Secondary method
   1. Perform cervical dislocation or bilateral thoracotomy.