The Muscle Cuff Regenerative Peripheral Nerve Interface for the Amplification of Intact Peripheral Nerve Signals

Summary

This manuscript provides an innovative method for developing a biologic peripheral nerve interface termed the Muscle Cuff Regenerative Peripheral Nerve Interface (MC-RPNI). This surgical construct can amplify its associated peripheral nerve's motor efferent signals to facilitate accurate detection of motor intent and the potential control of exoskeleton devices.

Abstract

Robotic exoskeletons have gained recent acclaim within the field of rehabilitative medicine as a promising modality for functional restoration for those individuals with extremity weakness. However, their use remains largely confined to research institutions, frequently operating as a means of static extremity support as motor detection methods remain unreliable. Peripheral nerve interfaces have arisen as a potential solution to this shortcoming; however, due to their inherently small amplitudes, these signals can be difficult to differentiate from background noise, lowering their overall motor detection accuracy. As current interfaces rely on abiotic materials, inherent material breakdown can occur alongside foreign body tissue reaction over time, further impacting their accuracy. The Muscle Cuff Regenerative Peripheral Nerve Interface (MC-RPNI) was designed to overcome these noted complications. Consisting of a segment of free muscle graft secured circumferentially to an intact peripheral nerve, the construct regenerates and becomes reinnervated by the contained nerve over time. In rats, this construct has demonstrated the ability to amplify a peripheral nerve's motor efferent action potentials up to 100 times the normal value through the generation of compound muscle action potentials (CMAPs). This signal amplification facilitates high accuracy detection of motor intent, potentially enabling reliable utilization of exoskeleton devices.

Introduction

In the United States alone, approximately 130 million people are affected by neuromuscular and musculoskeletal disorders, resulting in over $800 billion in annual economic impact^1,2. This group of disorders is typically secondary to pathology within the nervous systems, at the neuromuscular junction, or within the muscle itself³. Despite the variety of pathologic origins, the majority share some degree of extremity weakness^1,3. Unfortunately, this weakness is often permanent given the limitations in neural and muscle tissue regeneration, especially in the setting of severe trauma^4,5,6.

Extremity weakness treatment algorithms have classically focused on rehabilitative and supportive measures, often relying on harnessing the capabilities of the remaining intact limbs (canes, wheelchairs, etc.)⁷. This strategy falls short, however, for those whose weakness is not limited to a single extremity. With recent innovations in robotic technologies, advanced exoskeleton devices have been developed that restore extremity functionality to those living with extremity weakness^{8,9,10,11,12,13}. These robotic exoskeletons are often powered, wearable devices that can assist with initiation and termination of movement or maintenance of limb position, providing a varying amount of force that can be individually tailored for the user^{8,9,10,11,12,13}. These devices are classified as either passive or active depending on how they provide motor assistance to the user: active devices contain electrical actuators that augment power to the user, whereas passive devices store energy from the user's motions in order to release it back to the user when necessary¹⁴. As active devices have the ability to increase a user's power capabilities, these devices are utilized far more frequently in the setting of extremity weakness^[14].

In order to determine motor intent in this population, modern exoskeletons commonly rely on pattern recognition algorithms generated from either electromyography (EMG) of distal limb muscles^8,15,16,17 or surface electroencephalography (sEEG) of the brain^18,19,20. Despite the promise of these detection modalities, both options have significant limitations that preclude widespread utilization of these devices. As sEEG detects microvolt-level signals transcranially^18,19,20, criticisms frequently focus on the inability to differentiate these signals from background noise²¹. When background noise is similar to the desired recording signal, this produces low signal-to-noise ratios (SNRs), resulting in inaccurate motor detection and classification^22,23. Accurate signal detection additionally relies on stable, low-impedance scalp contact²¹, which can be significantly affected by the presence of coarse/thick hair, user activity, and even sweating^22,24. In contrast, EMG signals are several magnitudes larger in amplitude, facilitating greater motor signal detection accuracy^15,18,25. This comes at a cost, however, as nearby muscles can contaminate the signal, decreasing the degrees of freedom able to be controlled by the device^16,17,25 and an inability to detect deep muscle motion^25,26,27,28. Most importantly, EMG cannot be used as a control method when there is significant muscle compromise and complete absence of tissue²⁹.

In order to advance the development of robotic exoskeletons, consistent and accurate detection of motor intent of the intended user is required. Interfaces that utilize the peripheral nervous system have arisen as a promising interface technique, given their relatively simple access and functional selectivity. Current peripheral nerve interfacing methods can be invasive or non-invasive and typically fall within one of three categories: extraneural electrodes^30,31,32,33, intrafascicular electrodes^34,35,36 and penetrating electrodes^37,38,39,40. As peripheral nerve signals are generally on the level of microvolts, it can be difficult to differentiate these signals from similar amplitude background noise^41,42, which reduces the overall motor detection accuracy capabilities of the interface. These low signal-to-noise (SNR) ratios often worsen over time secondary to worsening electrode impedance⁴³ produced from either degradation of the device^39,43, or local foreign body reaction producing scar tissue around the device and/or local axonal degeneration^37,44. Although these shortcomings can generally be resolved with reoperation and implantation of a new peripheral nerve interface, this is not a viable long-term solution as foreign-body-associated reactions would continue to occur.

To avoid these local tissue reactions generated from peripheral nerves' interaction with abiotic interfaces, an interface incorporating a biologic component is necessary. To address this shortcoming, the Regenerative Peripheral Nerve Interface (RPNI) was developed to integrate transected peripheral nerves in the residual limbs of those with amputations with prosthetic devices^45,46,47,48. Fabrication of the RPNI involves surgical implantation of a transected peripheral nerve into a segment of autologous free muscle graft, with revascularization, regeneration, and reinnervation occurring over time. Through the generation of milli-volt level compound muscle action potentials (CMAPs), the RPNI is able to amplify its contained nerve's micro-volt level signal by several magnitudes, facilitating accurate detection of motor intent^45,48,49. There has been considerable development of the RPNI over the past decade, with notable success in amplifying and transmitting efferent motor nerve signals in both animal^50,51 and human⁴⁷ trials, facilitating high accuracy prosthetic device control with multiple degrees of freedom.

Individuals with extremity weakness but intact peripheral nerves would similarly benefit from high accuracy detection of motor intent through peripheral nerve interfaces in order to control exoskeleton devices. As the RPNI was developed for integration with transected peripheral nerves, such as in persons with amputations, surgical modifications were necessary. Building from experience with the RPNI, the Muscle Cuff Regenerative Peripheral Nerve Interface (MC-RPNI) was developed. Consisting of a similar segment of free muscle graft as in the RPNI, it is instead secured circumferentially to an intact peripheral nerve (Figure 1). Over time, it regenerates and becomes reinnervated through collateral axonal sprouting, amplifying and translating these efferent motor nerve signals to EMG signals that are several orders of magnitude larger⁵². As the MC-RPNI is biologic in origin, it avoids the inevitable foreign body reaction that occurs with peripheral nerve interfaces currently in use⁵². Furthermore, the MC-RPNI confers the ability to control multiple degrees of freedom simultaneously as they can be placed on distally dissected nerves to individual muscles without significant cross-talk, as has been previously demonstrated in RPNIs⁴⁹. Finally, the MC-RPNI can operate independent of distal muscle function as it is placed on the proximal nerve. Given its advantages over current peripheral nerve interfaces, the MC-RPNI holds substantial promise for providing a safe, accurate, and reliable method of exoskeleton control.

Protocol

All animal procedures and experiments were carried out with the approval of the University of Michigan's Institutional Care and Use of Animals Committee (IACUC). Male and female Fischer F344 and Lewis rats (~200-300 g) at 3-6 months of age are most frequently utilized in experiments, but any strain can theoretically be utilized. If utilizing donor rats instead of autologous muscle grafts, donor rats must be isogenic to the experimental strain. Rats are allowed free access to food and water both pre- and post-operatively. Following terminal endpoint evaluations, euthanasia is performed under deep anesthesia with intra-cardiac potassium chloride injection followed by a secondary method of bilateral pneumothorax.

1. Experimental preparation of the rat

1. Anesthetize the experimental rat utilizing a solution of 5% isoflurane in oxygen at 0.8-1.0 L/min in an induction chamber. Once adequate anesthesia is achieved and confirmed with the absence of corneal reflex, place the rat on a rebreather nose cone with isoflurane lowered to 1.75%-2.25% for maintenance of anesthesia.
2. Inject a solution of 0.02-0.03 mL Carprofen (50 mg/mL) in 0.2 mL of sterile saline with 27 G needle in the subcutaneous plane between the shoulder blades for peri- and post-operative analgesia.
3. Apply sterile eye ointment to both eyes to prevent corneal ulcers while anesthetized.
4. Using an electric razor, shave the lateral portion of the bilateral lower limbs, extending from the hip joint, over the thigh, and to the dorsal surface of the paw.
5. Sterilize the surgical site by first wiping with an alcohol prep pad, followed by povidone-iodine solution application, ending with a final cleanse with a new alcohol prep pad to remove the residual povidone-iodine solution. Repeat this alternating cleansing process three times to maintain sterility.
   NOTE: This can be a dermatological irritant; ensure the majority of the solution is removed.

2. Preparation of the muscle graft

1. Place the rat on a heating pad underneath a surgical microscope with an intraoral body temperature probe of choice for body temperature monitoring. Maintain isoflurane at 1.75%-2.25% and oxygen at 0.8-1.0 L/min.
2. Make a longitudinal incision along the anterior aspect of the desired donor hindlimb extending from just above the ankle to just below the knee with a #15 scalpel.
3. Dissect through the underlying subcutaneous tissue using sharp iris scissors to expose the underlying musculature and distal tendons just proximal to the ankle joint. Tibialis anterior (TA) is the largest and most anterior of the muscles; the extensor digitorum longus (EDL) muscle can be found just deep and posterior to this muscle. Isolate the EDL muscle and its distal tendon from the surrounding musculature.
4. Ensure isolation of the correct tendon by inserting both tines of a forceps or iris scissor underneath the distal tendon just proximal to the ankle joint. Exert upward pressure on the tendon by opening either the forceps or iris scissors. This motion should produce a simultaneous extension of all of the toes simultaneously. If isolated ankle dorsiflexion, ankle eversion, or single toe dorsiflexion occurs, the wrong tendon has been isolated.
5. Perform a distal tenotomy of the EDL muscle at the level of the ankle with sharp iris scissors and dissect the muscle free from surrounding tissues working proximally towards its tendinous origin.
6. Once the proximal tendon is visualized, perform a proximal tenotomy utilizing sharp iris scissors to free the graft.
7. Trim both tendinous ends of the muscle graft and cut to the desired length with sharp iris scissors.
   NOTE: Grafts measuring 8-13 mm have been utilized with success; however, the most common length utilized is 10 mm.
8. On one side of the muscle graft, make a longitudinal incision along the entire trimmed length to facilitate placement of the nerve within the muscle graft and provide contact of the nerve with endomysium.
9. Place the prepared muscle graft in a saline-moistened gauze to prevent tissue desiccation.
10. Close the skin overlying the donor site with 4-0 chromic suture in a running fashion.

3. Common peroneal nerve isolation and preparation

1. Mark the surgical incision, which will extend from a line ~5 mm from the sciatic notch, extending to just inferior to the knee joint. Ensure that this marking is inferior to, and angled away from, the femur that can be palpated below.
2. Incise through the skin and subcutaneous tissues along the marked incision line with a #15 blade. Carefully incise through the underlying biceps femoris fascia, taking care not to extend through the entire depth of the muscle as the sciatic nerve lies just below.
3. Utilizing blunt-tipped small scissors or a hemostat, dissect carefully through the biceps femoris muscle.
   NOTE: The sciatic nerve travels in this space underlying the biceps, oriented in approximately the same direction as the incision marked on the skin. There are three notable sciatic nerve branches: sural (most posterior and smallest of the nerves), tibial (typically most anterior, but this nerve always dives deep to the knee joint), and common peroneal (typically located between tibial and sural, always travels above the knee joint).
4. Identify the common peroneal (CP) nerve and carefully isolate it from the surrounding nerves using a pair of micro-forceps and micro-scissors. Remove any surrounding connective tissue from the middle 2 cm of the nerve. Take care not to crush the CP nerve with forceps in this process, as crush injury can alter endpoint results.
5. Over the central-most portion of the freed CP nerve, perform an epineurial window by removing 25% of the epineurium along the length of the nerve that matches the desired length of the muscle graft.
6. To perform this hold the proximal epineurium with micro-forceps, cut into the epineurium immediately underlying with micro-dissection scissors, and remove ~25% of the epineurium traveling distally along the nerve. Take care to remove this segment in one piece, as multiple attempts can cause irregular epineurial removal, increasing the risk of nerve injury. 
   NOTE: The nerve tissue underlying the epineurium will have a goo-like texture; noting this quality of nerve ensures the correct tissue plane has been removed.

4. MC-RPNI construct fabrication

1. Remove the muscle graft from the saline-moistened gauze and place it under the central portion of the CP nerve where the epineurial window was created. Rotate the nerve 180° so the epineurial window section contacts intact muscle and does not underlie the eventual suture line.
2. Using an 8-0 nylon suture, suture the epineurium of the CP nerve both proximally and distally to the muscle graft within the groove created in step 2.8 using simple interrupted sutures to secure epineurium to endomysium.
   NOTE: Place these stitches, ensuring the muscle is at normal resting length. Stretching or compressing the muscle too much can impact regeneration and signaling capabilities later on.
3. Circumferentially wrap the edges of the muscle graft surrounding the now-secured nerve and suture in place utilizing simple interrupted 8-0 nylon stitches (~4-6 depending on length).
4. Once hemostasis is achieved, close the biceps femoris fascia over the construct with 5-0 chromic suture in running fashion.
5. Close the overlying skin in running fashion with a 4-0 chromic suture.
6. Cleanse the surgical area with an alcohol prep pad and apply antibiotic ointment.
7. Terminate inhalational anesthetic and place the rat in a clean cage isolated from cage mates and allow to recover with food and water ad lib.
8. Once the rat has suitably recovered, place it back with cage mates in a clean cage.
   NOTE: These constructs require maturation of three months at the minimum to produce adequate nerve signal amplification.

Results

MC-RPNI surgical fabrication is considered a peri-operative failure if rats do not survive emergence from surgical anesthesia or develop an infection within a week of the operation. Previous research has indicated a 3 month maturation period will result in reliable signal amplification from this constructs^42,45,48,49. At that time or thereafter, surgical exposure of the constructs and evaluatio...

Discussion

The MC-RPNI is a novel construct that allows for amplification of an intact, peripheral motor nerve's efferent action potentials in order to accurately control an exoskeleton device. Specifically, the MC-RPNI confers a particular benefit to those individuals with extremity weakness caused by significant muscle disease and/or absence of muscle where EMG signals cannot be recorded. Reducing already compromised muscle function would be devastating in this population; however, the MC-RPNI has the ability to provide this ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
#15 Scalpel	Aspen Surgical, Inc	Ref 371115	Rib-Back Carbon Steel Surgical Blades (#15)
2-N-thin film load cell (S100)	Strain Measurement Devices, Inc	SMD100-0002	Measures force generated by the attached muscle
4-0 Chromic Suture	Ethicon	SKU# 1654G	P-3 Reverse Cutting Needle
5-0 Chromic Suture	Ethicon	SKU# 687G	P-3 Reverse Cutting Needle
8-0 Monofilament Suture	AROSurgical	T06A08N14-13	Black polyamide monofilament suture on a threaded tapered needle
Experimental Rats	Envigo	F344-NH-sd	Rats are Fischer F344 Strain
Fine Forceps - mirror finish	Fine Science Tools	11413-11	Fine tipped forceps with mirror finish ideal for handling delicate structures like nerves
Fluriso (Isofluorane)	VetOne	13985-528-40	Inhalational Anesthetic
Force Measurement Jig	Red Rock	n/a	Custom designed force measurement jig that allows for immobilization of hindlimb to allow for accurate muscle force recording
MATLAB software	Mathworks, Inc	PR-MATLAB-MU-MW-707-NNU	Calculates active force for each recorded force trace from passive and total force measurements
Nicolet Viasys EMG EP System	Nicolet	MFI-NCL-VIKING-SELECT-2CH-EMG	Portable EMG and nerve signal recording system capable of simultaneous 2 channel recordings from nerve and/or muscle
Oxygen	Cryogenic Gases	UN1072	Standard medical grade oxygen canisters
Potassium Chloride	APP Pharmaceuticals	63323-965-20	Injectable form, 2 mEq/mL
Povidone Iodine USP	MediChoice	65517-0009-1	10% Topical Solution, can use one bottle for multiple surgical preps
Puralube Vet Opthalmic Ointment	Dechra	17033-211-38	Corneal protective ointment for use during procedure
Rimadyl (Caprofen)	Zoetis, Inc.	NADA# 141-199	Injectable form, 50 mg/mL
Stereo Microscope	Leica	Model M60	User can adjust magnification to their preference
Surgical Instruments	Fine Science Tools	Various	User can choose instruments according to personal preference or from what is currently available in their lab
Triple Antibiotic Ointment	MediChoice	39892-0830-2	Ointment comes in sterile, disposable packets
Vannas Spring Scissors - 2mm cutting edge	Fine Science Tools	15000-04	Curved micro-dissection scissors used to perform the epineurial window
VaporStick 3	Surgivet	V7015	Anesthesia tower with space for isofluorane and oxygen canister
Webcol Alcohol Prep	Coviden	Ref 6818	Alcohol prep wipes; use a new wipe for each prep