Fabrication of the Composite Regenerative Peripheral Nerve Interface (C-RPNI) in the Adult Rat

Abstract

Recent advances in neuroprosthetics have enabled those living with extremity loss to reproduce many functions native to the absent extremity, and this is often accomplished through integration with the peripheral nervous system. Unfortunately, methods currently employed are often associated with significant tissue damage which prevents prolonged use. Additionally, these devices lack any meaningful degree of sensory feedback as their complex construction dampens any vibrations or other sensations a user may have previously depended on when using more simple prosthetics. The composite regenerative peripheral nerve interface (C-RPNI) was developed as a stable, biologic construct with the ability to amplify efferent motor nerve signals while providing simultaneous afferent sensory feedback. The C-RPNI consists of a segment of free dermal and muscle graft secured around a target mixed sensorimotor nerve, with preferential motor nerve reinnervation of the muscle graft and sensory nerve reinnervation of the dermal graft. In rats, this construct has demonstrated the generation of compound muscle action potentials (CMAPs), amplifying the target nerve’s signal from the micro- to milli-volt level, with signal to noise ratios averaging approximately 30-50. Stimulation of the dermal component of the construct generates compound sensory nerve action potentials (CSNAPs) at the proximal nerve. As such, this construct has promising future utility towards the realization of the ideal, intuitive prosthetic.

SUMMARY:

The following manuscript describes a novel method for developing a biologic, closed loop neural feedback system termed the composite regenerative peripheral nerve interface (C-RPNI). This construct has the ability to integrate with peripheral nerves to amplify efferent motor signals while simultaneously providing afferent sensory feedback.

INTRODUCTION:

Extremity amputations affect nearly 1 in 190 Americans¹, and their prevalence is projected to increase from 1.6 million today to over 3.6 million by 2050². Despite documented use for over a millennium, the ideal prosthetic has yet to be realized³. Currently, there exist complex prosthetics capable of multiple joint manipulations with the potential to reproduce many motor functions of the native extremity^4,5. However, these devices are not considered intuitive as the desired prosthetic motion is typically functionally separate from the input control signal. Users typically consider these “advanced prosthetics” difficult to learn and therefore not suitable for everyday use^1,6. Additionally, complex prosthetics do not provide any appreciable degree of subtle sensory feedback for adequate control. The sense of touch and proprioception are vital to carrying out daily tasks, and without these, simple acts such as picking up a cup of coffee become burdensome as it relies entirely on visual cues^7-9. For these reasons, advanced prostheses are associated with a significant degree of mental fatigue and are often described as burdensome and unsatisfactory^5,10,11.

For this population, the ideal prosthetic device would provide high fidelity motor control alongside meaningful somatosensory feedback from the individual’s environment. Critical to the design of said ideal prosthetic is the development of a stable, reliable interface that would allow for simultaneous transmission of afferent somatosensory information with efferent motor signals. The most promising of current human-machine interfaces are those that interact with the peripheral nervous system directly, and recent developments in the field of neuro-integrated prosthetics have worked towards bridging the gap between bioelectric and mechanical signals¹². Current interfaces utilized include: flexible nerve plates¹³, extra-neural cuff electrodes^14-18, tissue penetrating electrodes^19-20, and intrafascicular electrodes^21-23. However, each of these methods have demonstrated limitations with regard to nerve specificity, tissue injury, axonal degeneration, and/or scar tissue formation associated with chronic indwelling foreign body response²⁴.

To address these limitations, our laboratory developed a biologic interface termed the Regenerative Peripheral Nerve Interface (RPNI) to integrate transected peripheral nerves in a residual limb with a prosthetic device. RPNI fabrication involves surgically implanting a peripheral nerve into an autologous free muscle graft, which subsequently revascularizes and reinnervates previously denervated neuromuscular junctions. Our lab has developed this interface over the past decade, with success in amplifying and transmitting motor signals in both animal and human trials, allowing for suitable prosthetic control with multiple degrees of freedom^2,26. In addition, we have separately demonstrated sensory feedback through the use of peripheral nerves embedded in dermal grafts, termed the Dermal Sensory Interface (DSI)^3,27. In more distal amputations, using these interfaces simultaneously is feasible as motor and sensory fascicles within the target peripheral nerve can be surgically separated. However, for more proximal level amputations, this is not feasible due to intermingling of motor and sensory fibers. The Composite Regenerative Peripheral Nerve Interface (C-RPNI) was developed for more proximal amputations, and it involves implanting a mixed sensorimotor nerve into a construct consisting of free muscle graft secured to a segment of dermal graft (Figure 1). Peripheral nerves demonstrate preferential targeted reinnervation, thus sensory fibers will re-innervate the dermal graft and motor fibers, the muscle graft. This construct thus has the ability to simultaneously amplify motor signals while providing somatosensory feedback²⁸, allowing for the realization of the ideal, intuitive, complex prosthetic.

Figure 1: Illustrative schematic of the C-RPNI construct.

The common peroneal nerve can be seen secured between the top dermal layer and bottom muscle layer. This construct is secured to the femur periosteum proximally and distally via EDL’s tendinous junctions.

PROTOCOL:

All animal experiments are performed under the approval of the University of Michigan’s Committee on the Use and Care of Animals. Donor rats are allowed free access to food and water prior to skin and muscle donation procedures. Euthanasia is performed under deep anesthesia followed by intra-cardiac potassium chloride injection with a secondary method of bilateral pneumothorax. Male and female Fischer rats greater than two months of age are used with no appreciable differences between the sexes. Donor rats must be isogenic to the experimental rats.

1. Preparation of the dermal graft

1.1 Anesthetize donor rat in an induction chamber utilizing a solution of 5% isoflurane oxygen at 0.8-1L/min. Once the rat has been anesthetized, remove from induction chamber and place on a rebreathing nose cone, lowering the isoflurane to 2-2.5% for maintenance of anesthesia.

1.2 Administer a solution of 0.02-0.03 ml Carprofen (50mg/ml) in 0.2 ml sterile saline subcutaneously between the shoulder blades for analgesia.

1.3 Apply artificial tears ointment to both eyes to prevent corneal ulcers.

1.4 Using clippers, shave the entire lower hindlimb(s), ankle region, and sides of paw(s).

1.5 Cleanse chosen hindlimb and plantar surface of paw with alcohol, followed by betadine solution, ending with a final cleanse with alcohol to remove residual betadine.

1.6 Using a hand-held micro motor high speed drill with a removable round fine grit polishing stone (4000 rpms), burr the plantar surface of the paw to remove the epidermis. While burring, apply drips of saline as to not burn the skin. The underlying dermis will have a shiny appearance with pinpoint bleeding.

1.7 Apply a tourniquet to the lower extremity to slow blood flow

1.8 Remove the plantar skin sharply with a #15 scalpel and place in saline moistened gauze to prevent desiccation. Some tendinous and connective tissue will inherently be removed with the skin in this step and will be removed later.

1.9 Apply gauze wrap to the bleeding foot to slow hemorrhage.

2.0 Repeat steps 1.5-1.9 if doing two constructs.

2.1 Under a microscope, remove the tendinous and connective tissue from the deep layer of the skin graft using micro-scissors. Take care not to make holes in the graft. The thinned dermal graft should be slightly opaque containing only dermis, measuring approximately 0.5x1.0cm in size. Place in saline-moistened gauze until ready for C-RPNI construct fabrication.

2. Preparation of muscle graft

2.1 Make a longitudinal incision along the anterior aspect of the lower hindlimb from just above the ankle to just below the knee with a #15 scalpel. Dissect through subcutaneous tissue to expose the underlying musculature.

2.2 At the distal aspect of the incision, expose the tendinous insertions of the lower limb musculature. Tibialis anterior (TA) is typically the largest and most anterior of the muscles, and just underneath and posterior to this muscle lies the extensor digitorum longus (EDL). Isolate the distal EDL tendon from the other tendons in the area, taking care not to incise its insertion at this point.

2.3 Ensure you have the correct tendon by inserting both tines of a forceps underneath the tendon and exerting upward pressure with opening of the forceps to cause tendon excursion. Manipulation of this tendon should cause all of the toes to extend simultaneously.

2.4 Perform a distal tenotomy with sharp iris scissors and separate the muscle from the surrounding tissues bluntly with tenotomies (or other blunt-tipped scissor) working proximally to find the tendinous origin.

2.5 Once the proximal tendon is visualized, again perform a tenotomy utilizing sharp iris scissors. Place muscle graft in a saline-moistened gauze to prevent desiccation.

2.6 Once all desired grafts have been removed from the donor rat, euthanize according to protocol.

3. Common peroneal nerve isolation and preparation

3.1 Anesthetize and provide analgesia to the experimental rat according to protocol outlined in 1.1-1.3.

3.2 Shave the desired thigh and cleanse with alcohol, betadine, ending with alcohol to remove traces of betadine.

3.3 Move animal from surgical prep table to surgical microscope table and place on heating pad with temperature probe for body temperature maintenance. Isoflurane is maintained at 2-2.5% and oxygen at 0.8-1L/min

3.4 Mark your incision, extending from just distal to sciatic notch to the inferior portion of the knee. This marking should be inferior to, and angled away from, the femur. Make the incision with a #15 blade.

3.5 Carry your incision through the underlying biceps femoris fascia.

3.6 Carefully dissect through the biceps femoris muscle with either hemostat or blunt tipped micro-scissors to the space underlying biceps femoris.

Note: The sciatic nerve will be traveling approximately the same direction as the initial incision that was made. There will be three branches, typically with sural nerve posteriorly and common peroneal and tibial nerve traveling superficial and deep to the knee, respectively.

3.7 Following identification of the common peroneal (CP) nerve, using a pair of micro-, fine-tipped forceps and micro-scissors, carefully isolate the CP nerve from the other sciatic branches and remove any lingering connective tissue distally.

3.8 At the point where the nerve crosses the surface of the knee, sharply transect the nerve with a pair of micro-scissors.

Note: Using sharp scissors is extremely important in this step as causing significant trauma to the nerve could increase the risk of neuroma formation.

3.9 Carefully free any remaining connective tissue from the CP nerve and work proximally to free the nerve to a length of approximately 2 centimeters.

4. C-RPNI Construct Fabrication

4.1 Remove muscle graft from saline-moistened gauze and remove all central tendinous tissue as well as a small segment of epimysium. Leave the tendinous ends intact.

4.2 Using an 8-0 nylon suture, secure the epineurium of the transected end of the CP nerve to the area of the muscle graft devoid of epimysium with two interrupted stitches on either side of the nerve.

4.3 Secure the muscle graft to the femur periosteum with a single 6-0 nylon interrupted stitch both proximally and distally with the nerve-muscle junction facing away from the femur.

Note: Secure the muscle so that it is at normal relaxed length. Try not to stretch the muscle significantly or leave too much laxity when securing.

4.4 Place an 8-0 nylon stitch at the inferior, central margin of the muscle graft epimysium, securing it to the CP nerve epineurium in a way as to create laxity in the nerve within the muscle graft and help to relieve any future tension it may be exposed to with later ambulation.

4.5 Remove the skin graft from the saline-moistened gauze and arrange on the muscle graft in such a way to completely cover the nerve and the majority of the muscle. Ensure that the deep margin of the dermis is resting on the muscle. Trim any dermis that extends beyond the border of the muscle.

4.6 Secure the skin graft to the muscle graft circumferentially using 8-0 nylon interrupted sutures. Typically, 4-8 total sutures are used depending on the size of the construct.

4.7 Close the biceps femoris fascia over the construct in a running fashion with 5-0 chromic suture.

4.8 Close the overlying skin with 4-0 chromic suture in running fashion.

4.9 Swab the surgical area with an alcohol pad and apply antibiotic ointment.

5.0 Cease inhalational anesthetic and allow rat to recover with food and water sources separate from cage mates.

REPRESENTATIVE RESULTS:

Construct fabrication is considered unsuccessful if rats develop an infection or do not survive surgical anesthesia. Previous research has indicated these constructs require approximately three months to revascularize and reinnervate^2,3,12,28. Following the three-month recovery period, construct testing can be pursued to examine viability. Surgical exposure of the constructs after three months will reveal revascularized muscle and skin if successful (Figure 2). At times, the free muscle and dermal grafts will have been replaced with scar tissue, and/or the nerve will not be attached to the construct; these findings indicate an unsuccessful attempt. However, if successful, gentle squeezing of the common peroneal nerve with forceps proximal to the construct will result in visible muscle contraction (Figure 3). Histological analysis of constructs should demonstrate viable skin, nerve, and muscle (Figure 4). Immunostaining will also reveal motor and sensory nerve reinnervation to their neuromuscular junctions and sensory end organs, respectively (Figure 5). If the common peroneal nerve did not reinnervate those tissues, immunostaining will not demonstrate any individual nerve fibers within the construct with the exception of the implanted nerve itself.

Figure 2: C-RPNI in vivo.

(A) A C-RPNI immediately following fabrication and at (B) 3 months post-construction at time of electrophysiologic testing. The muscle component is the deep layer of the construct and the dermal, the superficial. Muscle tissue is marked by (M), dermis (D), and common peroneal nerve (N).

Figure 4: C-RPNI histology 6 months.

C-RPNI H&E at 6 months in (A) cross-section and (B) longitudinal section. Muscle noted by (M), dermis (D), and nerve (N).

Figure 5: Immunostaining of the C-RPNI.

(A) Representative example of a cross-section of muscle tissue, with red arrows identifying neuromuscular junctions. (B) Close-up of a neuromuscular junction noted in the sample. For (A) and (B), red staining (alpha-bungarotoxin) indicates presence of cholinergic receptors in muscle tissue; blue (neurofilament 200) specifies presence of neurofilaments within axonal tissue; and green (choline acetyltransferase) notes specifically motor neuron presence. (C) Representative example of an iDISCO image focusing on the dermal junction with red arrows marking sensory neurons (white) entering the dermis. (D) On-lay view of iDISCO demonstrating multiple sensory neurons (white, neurofilament 200) innervating the dermis with some scattered underlying neuromuscular junctions noted in red (alpha-bungarotoxin).

Electrophysiologic testing can be performed on these constructs in vivo, and previous research has been conducted at 3 and 9 months following C-RPNI fabrication²⁸ (Figure 6). Following maximal stimulation at the proximal common peroneal nerve, compound muscle action potentials (CMAPs) can be measured at the muscle component with visible muscle contraction. The average CMAP amplitude recorded at the muscle was 8.7 +/− 1.6 mV at 3 months and 10.2 +/− 2.1 mV at 9 months. The average conduction velocity was 10 +/− 1.2 m/sec at 3 months and +/− 0.6 m/sec at 9 months. In comparison, CMAPs generated by physiologic EDL muscle typically range from 10-18 mV (Sabbagh SW May 2018; Kubiak CA May 2019). Following stimulation at the dermal component of the C-RPNI, compound sensory nerve action potentials (CSNAPs) were produced at the proximal common peroneal nerve, with average CSNAP amplitude measuring 113.7 +/− 35.1 μV at 3 months and 142.9 +/− 63.7 μV at 9 months. Figure 7 illustrates single and summation CMAP and CSNAP signals obtained during electrophysiologic testing in a graphical format.

Figure 6: Electrophysiologic testing of C-RPNIs at 3 and 9 months post-construction.

To obtain CMAPs, a recording electrode was placed on the muscle with a stimulating electrode on the proximal common peroneal nerve. A series of stimulations increasing in amplitude was applied to the nerve until maximal CMAP values were obtained and results recorded. A similar methodology was applied to the dermal component but with the recording electrode placed on the nerve and stimulating electrode on the dermis. For the sensory evaluation of rat 4690 at 9 months, the dermal graft was found to be too scarred to allow for testing.

Figure 7: Typical C-RPNI electrophysiologic signaling.

(A) A single CMAP signal recorded at the muscle component following a 5.00 mA signal applied to the CP nerve. (B) 24 CMAPs generated by 5.00 mA stimulation at the nerve. (C) A single CSNAP signal recorded from the proximal CP nerve following dermal component stimulation at 900 μA. (D) A series of CSNAPs recorded from the proximal CP nerve following increasing stimulation at the dermal component from 500 μA to 1000 μA.

The C-RPNI serves to amplify a nerve’s inherent microvolt signal, and previous research has demonstrated sufficient amplification from the microvolt to millivolt level²⁹. Therefore, if a construct does not provide that level of amplification, it is not considered successful. If either the dermal, muscle, or both components of the C-RPNI fail, testing would result in recordings that mimic the stimulation signal utilized. For the muscle component specifically, a suboptimal result (but one that is still considered operational) would be one that has CMAP amplitude and conduction velocity in the range that falls between the signal stimulation value and that of physiologic EDL muscle. Additionally, these signals can become attenuated and lack the characteristic CMAP waveform (Figure 8A). Suboptimal results at the level of the dermal component can occur but are difficult to quantify given that rats cannot express the quality of sensation they experience. These suboptimal results usually involve dampening of the waveform with significant background noise (Figure 8B). However, if there is significant scarring or callusing of the skin graft, or minimal graft survived, no CSNAPs will be appreciated at the proximal common peroneal nerve regardless of stimulation value.

Figure 8: Abnormal C-RPNI signaling.

(A) A series of CMAPs obtained while ramping CP nerve stimulation from 0.2 to 4 mA. Wave-forms peak at different points and fail to return to baseline, possibly indicating defective electrodes or inadequate overall construct function. (B) Summation of CSNAPs obtained while stimulating dermal component, ramping 0.1 to 5 mA. These findings can occur for a multitude of reasons, including malfunctioning electrode(s), dermal graft scarring, and/or nerve damage.

DISCUSSION:

The C-RPNI is a novel construct that provides simultaneous amplification of a target nerve’s motor efferent signals with provision of afferent sensory feedback. In particular, the C-RPNI has unique utility for those living with proximal amputations as their motor and sensory fascicles cannot easily be mechanically separated during surgery. Instead, the C-RPNI utilizes the inherent preferential reinnervation properties of the nerve itself to encourage sensory fiber reinnervation to dermal sensory end organs and motor fibers to neuromuscular junctions.

As C-RPNI fabrication relies on the reinnervation abilities of the target nerve, careful handling of the nerve is paramount during the procedure. During dissection, avoid direct manipulation of, and trauma to, the target nerve. If the nerve must be handled, it is recommended to manipulate epineurium or surrounding connective tissue instead. Although our laboratory has not encountered neuroma formation within this construct, theoretically, significant nerve trauma could increase the risk. An additional key step in the process is the harvesting of the dermal grafts. All epidermal tissue must be removed from the hindpaw graft as retained epidermis can increase the risk of infection and inclusion cysts during the healing process. Furthermore, the dermal graft must be adequately thinned to promote imbibition and revascularization throughout the graft and avoid significant ischemia and necrosis.

Although the majority of studies conducted with the C-RPNI have been performed on the common peroneal nerve, any mixed sensorimotor nerve could be substituted. A pure motor or pure sensory nerve could be utilized, but the results are difficult to predict and would likely result in either largely muscle or dermal reinnervation, respectively. With regards to the muscle graft, as long as epimysium is removed from the portion contacting the nerve, any muscle graft similar in size could be utilized as long as it contained tendinous or fascial tissue at either end suitable for anchoring to nearby periosteum. For the dermal graft, glabrous tissue is specifically used due to the potential for hair growth following grafting. Non-glabrous skin was previously attempted, but due to the difficulty of removing individual hair follicles, all resultant constructs had significant hair growth, inflammation, and scarring following the three-month maturation period. Additionally, other rat species can be employed, but Lewis and Fischer rats are recommended for this experiment as many other rat species will self-mutilate secondary to nerve transection^30-31.

Given the delay between procedure and results, it is difficult to know ahead of time if any alterations must be made to the method. Infection is a theoretical risk rarely encountered by our laboratory, but if infection occurs, it is typically responsive to antibiotics. Occasionally, rats will chew on their incisions causing dehiscence, and this can be treated with washout, debridement, and re-closure. If, after three months at time of exposure, the construct is found to be nonfunctional and/or scarred, there are several potential causes. At times, if the nerve is not secured correctly to the construct with at least three sutures, the nerve can tear from the construct with ambulation. Additionally, the muscle and/or dermal grafts can necrose, causing failure. Typically, this is a result of either repeated infection, the dermal graft being too thick, or the muscle too damaged to recover properly. Additionally, if the muscle is not secured to periosteum at resting length, contraction can be impaired causing inadequate signals during testing. At times, the construct will appear viable but fail to produce CMAPs/CSNAPs on testing. This could be secondary to failure in equipment, elevated electrode impedance, or significant skin callusing. Skin callusing can dampen and completely block signal transduction if the dermis is not thinned properly during fabrication. If any of the preceding described events are seen frequently during the testing process, one must return to the protocol and make appropriate alterations.

Other methods commonly employed to amplify or record nerve signals include flexible nerve plates¹³, extra-neural cuff electrodes^14-18, tissue penetrating electrodes^19-20, and intrafascicular electrodes^21-23, all of which are associated with tissue injury, axonal degeneration, and/or scar tissue formation associated with chronic indwelling foreign body response²⁴. The C-RPNI, however, is a biologic construct and thus does not induce foreign body response. Histologic analysis of these samples has not demonstrated any significant degree of scar tissue formation in the nerve with chronic use, thus allowing the C-RPNI to interface with the nerve for extended periods in comparison to the methods listed above. Although this method is highly effective at amplification of efferent motor signals, it is limited with regards to sensory afferent signal production. We have measured and characterized signal transduction produced with mechanical and electrical stimulation of the dermal component of the C-RPNI²⁸; however, these rat subjects cannot qualify the type or degree of sensations elicited from stimulation of this construct. As such, at this time it is impossible to know what kind of effect the C-RPNI is producing with regards to sensation and will be the target of future experiments.

The predecessor to the C-RPNI, the RPNI (regenerative peripheral nerve interface), consists of a free muscle graft attached to a transected nerve, with motor fibers reinnervating previously denervated neuromuscular junctions. The RPNI has demonstrated utility in human subjects, with several patients controlling advanced prosthetics from signals amplified by—and recorded from—these RPNIs²⁶. A common complaint for those utilizing these advanced prosthetics, however, is the need to visualize the prosthetic during use as these prosthetics lack proprioception and provide minimal sensory feedback. The C-RPNI could be a solution to this common criticism by providing a way to deliver sensory feedback via the dermal component, leading to the realization of the much-desired, ideal prosthetic.

Supplementary Material

Figure 3

Figure 3: Muscle contraction within a C-RPNI. A pair of forceps can be seen to the left of the video gently squeezing the proximal common peroneal nerve. This results in contraction of the muscle component of a 3-month-old C-RPNI that is visible to the viewer.

Table 1.

Table of Materials

Name of Material/ Equipment	Company	Catalog Number	Comments/Description
Rimadyl (Caprofen)	Zoetis, Inc.	NADA# 141-199	Injectable form, 50 mg/mL
VaporStick 3	Surgivet	V7015	Anesthesia tower with space for isofluorane and oxygen canister
Fluriso (Isofluorane)	VetOne	13985-528-40	Inhalational Anesthetic
Oxygen	Cryogenic Gases	UN1072	Standard medical grade oxygen canisters
Stereo Microscope	Leica	Model M60	User can adjust magnification to their preference
Puralube Vet Opthalmic Ointment	Dechra	17033-211-38	Corneal protective ointment for use during procedure
Surgical Instruments	Fine Science Tools	Various	User can choose instruments according to personal preference or from what is currently available in their lab
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
6-0 Ethilon Suture	Ethicon	SKU# 697G	P-1 Reverse Cutting Needle (Nylon suture)
Micro Motor High Speed Drill with Stone	Master Mechanic	Model 151369	Handheld rotary tool; kit comes with multiple fine grit stones
#15 Scalpel	Aspen Surgical, Inc	Ref 371115	Rib-Back Carbon Steel Surgical Blades (#15)
Triple Antibiotic Ointment	MediChoice	39892-0830-2	Ointment comes in sterile, disposable packets
Povidone Iodine USP	MediChoice	65517-0009-1	10% Topical Solution, can use one bottle for multiple surgical preps
Webcol Alcohol Prep	Coviden	Ref 6818	Alcohol prep wipes; use a new wipe for each prep
Experimental Rats	Envigo	F344-NH-sd	Rats are Fischer F344 Strain
Potassium Chloride	APP Pharmaceuticals	63323-965-20	Injectable form, 2 mEq/mL