Journal of Neuroscience Methods 332 (2020) 108523

Contents lists available at ScienceDirect

Journal of Neuroscience Methods
journal homepage: www.elsevier.com/locate/jneumethA review for the peripheral nerve interface designeraChristopher E. Larson , Ellis Meng
a
b

a,b,

T

*

Department of Biomedical Engineering, University of Southern California, Los Angeles, CA, 90089, USA
Ming Hsieh Department of Electrical and Computer Engineering, University of Southern California, Los Angeles, CA, 90089, USA

G R A P H I C A L A B S T R A C T

A R T I C LE I N FO

A B S T R A C T

Keywords:
Peripheral nerve interface
Neural prosthesis
Bioelectronic medicine
Vagus nerve stimulation
Functional electrical stimulation
Brain machine interface

Informational density and relative accessibility of the peripheral nervous system make it an attractive site fortherapeutic intervention. Electrode-based electrophysiological interfaces with peripheral nerves have been
under development since the 1960s and, for several applications, have seen widespread clinical implementation.
However, many applications require a combination of neural target resolution and stability which has thus far
eluded existing peripheral nerve interfaces (PNIs). With the goal of aiding PNI designers in development ofdevices that meet the demands of next-generation applications, this review seeks to collect and present practicalconsiderations and best practices which emerge from the literature, including both lessons learned during early
PNI development and recent ideas. Fundamental and practical principles guiding PNI design are reviewed,
followed by an updated and critical account of existing PNI designs and strategies. Finally, a brief survey of invitro and in vivo PNI characterization methods is presented.

Abbreviations: AP, action potential; BMI, brain machine interface; CAP, compound action potential; CE, counter electrode; CNS, central nervous system; CSCc,
cathodal charge storage capacity; CV, cyclic voltammetry; ECH, electrically-conductive hydrogel; EIS, electrochemical impedance spectroscopy; EMG, electromyography; ENG, electroneurography; ESA, electrochemical surface area; FES, functional electrical stimulation; FFC, ﬂat ﬂexible cable; FINE, ﬂat interface nerve
electrode; GSA, geometric surface area; hd-USEA, high-density Utah slanted electrode array; LIFE, longitudinal intrafascicular electrode; MPM, multiphoton microscopy; OCT, optical coherence tomography; PBS, phosphate-buﬀered saline; PCB, printed circuit board; PEEK, polyether ether ketone; PNI, peripheral nerve
interface; PNS, peripheral nervous system; PTFE, polytetraﬂuoroethylene; RE, reference electrode; REMI, regenerative multielectrode interface; RSE, regenerative
scaﬀold electrode; SCE, saturated calomel electrode; SHG, second harmonic generation; SI, selectivity index; SNR, signal-to-noise ratio; TEENI, tissue-engineered
electronic nerve interface; TIME, transverse intrafascicular multichannel electrode; UEA, Utah electrode array; USEA, Utah slanted electrode array; VNS, vagus nerve
stimulation; WE, working electrode; ZIF, zero-insertion-force connector⁎
Corresponding author.
E-mail addresses: larsonce@usc.edu (C.E. Larson), ellis.meng@usc.edu (E. Meng).
https://doi.org/10.1016/j.jneumeth.2019.108523
Received 7 June 2019; Received in revised form 14 November 2019; Accepted 15 November 2019
Available online 16 November 20190165-0270/ © 2019 Elsevier B.V. All rights reserved.

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

1. Introduction

autonomously maintain healthy organ function (Ganzer and Sharma,
2019; Plachta et al., 2014; Sun and Morrell, 2014; Zanos, 2019).
Despite ﬁve decades of development and multiple instances ofclinical translation, each of the PNI application areas described above
remain highly active areas of research. Successful clinical applications
have largely been limited to those that rely only on non-speciﬁc stimulation. Chronically stable, high-resolution PNIs for ambitious applications such as neurally-integrated prosthetic limbs and closed-loop
organ modulation have not yet been realized. From the late 1990s topresent, the variety of PNI designs being investigated at the researchlevel has expanded dramatically thanks to new technologies from ﬁelds
such as biomedical microelectromechanical systems (bioMEMS) andmicroelectronics. Development continues to expand as PNI designers
strive for devices that can record and stimulate with increased resolution and maintain performance in the body for decades.
Previous reviews have covered, in varying detail, particular PNI
applications, fundamental concepts of PNI function, and anecdotal examples from the most prominent types of PNI designs (del Valle andNavarro, 2013; Giagka and Serdijn, 2018; Grill et al., 2009; Heiduschka
and Thanos, 1998; Kim and Romero-Ortega, 2012; Micera and Navarro,
2009; Mortimer and Bhadra, 2004; Navarro et al., 2005; Ortiz-Catalan
et al., 2012; Russell et al., 2019; Rutten, 2002; Saal and Bensmaia,
2015; Schultz and Kuiken, 2011; Tyler, 2018). This review presents an
updated, comprehensive survey that closely examines critical design
choices, strategies, and best practices that can inform next generationPNI designs. This review purposefully encompasses a broad range ofwork to counteract symptoms of recency-bias. Much of the foundational
knowledge and practical wisdom regarding PNI design and use wasascertained in the 1960s to 1990s. Lastly, there is a need to collect, in
one place, the methods and best practices to guide in vitro and in vivo
characterization of a PNI towards the goal of clinical impact. In sum,
the present review aims to aid the PNI designer in development of nextgeneration devices by reviewing principles guiding PNI design, examining diﬀerent PNI design strategies that have been explored, andsurveying methods of PNI characterization.

The peripheral nervous system (PNS) carries control (eﬀerent) andinformational (aﬀerent) signals between the central nervous system andother parts of the body. In cases of neurological or other dysfunction
due to injury or disease, the informational density and relative accessibility of the peripheral nervous system presents an attractive target fortherapeutic intervention. To this end, a wide variety of implantable
peripheral nerve interfaces (PNIs) have been developed to enable
monitoring and modulation of peripheral nerve signals.
Clinical implementation of implanted PNIs was ﬁrst achieved in the
1960s. The ﬁrst applications included diaphragmatic pacing (Judson,1968) and treatment of urinary incontinence (Alexander and Rowan,1968). Though these early devices were merely looped wires, they were
suﬃcient to produce the desired muscle contractions. The 1970s and80s saw reﬁnement of devices and methods for these same applications(Brindley, 1977; Brindley et al., 1986, 1982; Glenn and Phelps, 1985;
Schmidt et al., 1978) and further exploration of functional electricalstimulation (FES) via PNIs, including chronic pain management (Avery
and Wepsic, 1973; Picaza et al., 1977) and lower limb activation forwalking and standing assistance in cases of paralysis (Brindley et al.,
1979; McNeal and Bowman, 1985; Waters et al., 1975). Similar
methods were later applied to upper body control in spinal cord injury
patients (Popovic et al., 2002). FES applications of PNIs continue to be
an important area of development. For example, a recently approvedcommercial FES system stimulates the hypoglossal nerve overnight,
maintaining a slight contraction of the tongue muscle to treat obstructive sleep apnea (Eastwood et al., 2011; FDA Approves Inspire
Upper Airway Stimulation Therapy for Obstructive Sleep Apnea, 2014;
Schwartz et al., 2001)
Concurrent with advances in robotics and computing, a new focus of
PNI application beginning in the late 1990s, though anticipated muchearlier (Hoﬀer and Loeb, 1980; Stein et al., 1975), was that of hybrid
bionic systems, or brain-machine interfaces (BMI) (Micera et al., 2006;
Navarro et al., 2005; Wright et al., 2016). BMIs aim to directly link the
nervous system with a computer or robotic prosthesis, ideally utilizing
two-way communication of sensory and control signals and thereby
enabling possibilities such as neurally-controlled prosthetic limbs for
amputees (Ciancio et al., 2017; Lebedev and Nicolelis, 2006; Lu et al.,2012; Micera et al., 2010; Pasluosta et al., 2018; Schultz and Kuiken,2011). While many BMIs directly interface with the cerebral cortex, thePNS may be an attractive alternative in cases where the target nerve is
intact due to lower surgical risk and greater isolation of the desired
neural targets.
There are also a range of non-motor applications of PNIs. The vagus
nerve, which carries autonomic signals between visceral organs and thebrain stem, was targeted for therapeutic stimulation as early as the late
1800s (Guiraud et al., 2016). Vagus nerve stimulation (VNS) was ﬁrst
shown to oﬀer reduction of epileptic seizures in 1985 (Zabara, 1985),
and since 1988, VNS systems have been implanted in more than
168,000 patients around the world (VNS Therapy, 2019). Because of
the vagus nerve’s innervation of visceral organs, it is a potential site ofintervention for many disorders that would otherwise be treated withlimited success by medications, including depression (Rush et al.,2000), hypertension (Plachta et al., 2014), obesity (Banni et al., 2012;
Cork et al., 2018; Guiraud et al., 2016), heart failure (Guiraud et al.,2016), and autoimmune or inﬂammatory diseases such as rheumatoid
arthritis (Koopman et al., 2016; Payne et al., 2019; Zitnik, 2011). The
use of peripheral nerve recording and stimulation, especially of thevagus nerve and its branches, for such applications is often calledbioelectronic medicine, or “electroceuticals” (Birmingham et al., 2014;
Giagka and Serdijn, 2018). While bioelectronic medicine has largely
focused on use of pre-programmed or “open-loop” stimulation schedules, recent approaches not only modulate organ function via stimulation, but also monitor organ status via recording of vagus nerve signals with the aim of eventually closing the feedback loop to

2. Guiding principles for peripheral nerve interface design
2.1. Peripheral nerve anatomy
PN ﬁbers are extensions of neurons which have their cell body in ornear the central nervous system (CNS). Their ability to function asneural conductors is dependent on the support of complex, composite
tissue structures which provide nutrition and protection (Lundborg,2004). Coarsely described, the anatomical structure of a PN trunk is “abundle of bundles” (Fig. 1). Myelinated and unmyelinated nerve ﬁbers
of varying diameter and function (e.g. motor and sensory) are tightly
packed together in fascicles. A specialized sheath-like structure called
the perineurium forms the outer boundary of each fascicle. The perineurium is the primary source of mechanical strength for the nerve
trunk and is comprised of up to 15 dense layers of cells and collagen(Lundborg, 2004). Analogous to the crucial blood-brain barrier of the
CNS, the perineurium additionally serves as a blood-nerve barrier tomaintain the privileged microenvironment of the enclosed endoneurial
space. In addition to nerve ﬁbers, the endoneurium consists of Schwann
cells, immune cells, capillaries, and a matrix of collagen and otherconnective tissue ﬁbers (Grill et al., 2009; Lundborg, 2004). Finally,
holding multiple fascicles together to form the outer bundle of a compound nerve trunk is the epineurium. The epineurium is a relatively
loose matrix of collagen, fat, elastin ﬁbers and blood vessels, which,besides holding the bundle together, also serves to cushion the fascicles
from compressive forces (Sunderland, 1965). Blood vessels, being crucial to nerve ﬁber health and function, form a dense intraneural network along the entire length of the nerve trunk, as shown in Fig. 2b(Rydevik et al., 1990; Sunderland, 1978). “Extrinsic” vessels runningalongside the nerve are connected to intraneural vessels via frequent2

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 1. Illustration of nerve trunk anatomy, highlighting the epineurium (epi), fascicles (arrow), perineurium (p), and endoneurium (end). Also shown is the networkof extrinsic (exv) and regional (rv) blood vessels and internal vascular structures (*). Reprinted from (Lundborg, 2004) with permission from Elsevier.

“regional” vessels which obliquely penetrate into the nerve trunk’sconnective tissue layers (Fig. 1, Lundborg, 2004). The smallest endoneurial capillaries are 6–10 μm in diameter and possess looping andcoiling forms which help accommodate motion (Lundborg, 2004).
Nerves vary widely in terms of size, fascicle count, and ﬁber countacross species, nerve types, and proximal to distal positions within thesame nerve. The number of fascicles in a nerve trunk ranges from 1 to100, and the number of individual ﬁbers ranges from hundreds to tensof thousands (Sunderland, 1990). To give a sense of the numbers in acommon model for PNI study, the rat sciatic nerve has four fascicles
comprised of approximately 8,000 myelinated ﬁbers and 19,000 unmyelinated ﬁbers in total. The number of myelinated and unmyelinatedﬁbers in each fascicle ranges from 300 to 5,000 and 2,000 to 9,000,
respectively (Schmalbruch, 1986). The cross-section of a PN trunk is

generally presumed to be circular or ovicular, though the gradual separation and eventual branching of fascicles at more distal locations
produces a more ﬂattened or irregular shape, as evident in Fig. 3(Schmalbruch, 1986).
A remarkable feature of PN anatomy, especially in the extremities, is
its accommodation of relative motion among nerve components andsurrounding tissues. Peripheral nerves in high-motion areas such as
limbs, and especially joints, are served by a loose, fatty, lubricating
layer of paraneural connective tissue to facilitate gliding in the nerve
groove (Millesi et al., 1995). Within the nerve, the epineurium is only
loosely connected to perineurium so that individual fascicles can slide
past one another and rearrange under transverse compression(Lundborg, 2004; Millesi et al., 1995; Sunderland, 1990). Undulated
patterns are found throughout the microanatomy of the nerve in its

Fig. 2. A) A photograph of an unstretched nerve shows bands of Fontana caused by the undulating pattern of the tissue structure. These bands reversibly disappear
when the nerve is stretched. B) An injected and cleared sample of a peripheral nerve trunk shows the dense microvascular network. Reprinted from (Sunderland,1978) with permission from Elsevier.
3

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 3. A) Surgically exposed rat sciatic nerve, with the trochanter (t) and knee (k) marked by pins. B), C), and D) show cross-sections of the nerve at positions 1, 2,
and 3, respectively. The cross-section becomes ﬂatter as the sural (s), tibial (t), peroneal (p), and cutaneous (c) fascicles separate and eventually branch oﬀ. Reprinted
from (Schmalbruch, 1986) with permission from Wiley.

unstretched state, such as the wavy shape of nerve ﬁbers within a fascicle and the arrangement of collagen ﬁbers in the epi- and endoneurium (Millesi et al., 1995; Stolinski, 1995; Sunderland, 1978).
The nerve trunk and fascicles default to a wavy form when unstretched(Sunderland, 1978). Visible striations, known as bands of Fontana(Fig. 2a), reversibly disappear when the nerve is stretched, attesting totheir role in allowing dynamic elongation and shortening (Sunderland,1978). During length changes, volume of a PN trunk is conserved; thediameter decreases when elongated and increases when shortened(Millesi et al., 1995).
In summary, the structural and dynamic features of PN anatomy
have profound implications for PNI design.

which an electrode operates depends on a variety of factors such as
electrode size, location, and stimulation protocol. Methods for characterizing PNI electrodes and ﬁnding the limits between charge injection regimes are discussed in Section 4.1.1.
Several surface modiﬁcation strategies exist to increase the amountof charge that an electrode can capacitively and pseudocapacitively
deliver. The charge injection required by PNIs is usually beyond thecapacitive limit of common noble metal electrodes (namely, platinum
and platinum-iridium alloys) so these electrodes predominantly operate
in the pseudocapacitive regime (Robblee and Rose, 1990). However, anelectrode’s capacitive limit can be increased to a usable range by
coating it with a thin dielectric layer which imparts a higher dielectricstrength and prevents transfer of charge carriers; these are known ascapacitive electrodes (Howell and Grill, 2015). Assuming the electrodedielectric-electrolyte system exhibits parallel plate capacitor behavior,
the electrode’s capacitance is C = εoεrAe/d where εo is the permittivityof free space, εr is the relative permittivity of the dielectric, Ae is theelectrode’s electrochemical surface area, and d is the thickness of thedielectric. An eﬀective dielectric coating for an electrode provides increased εr while minimizing d (Howell and Grill, 2015). Electrode capacitance can be further enhanced by increasing the electrochemical or“real” surface area (ESA). Surface roughness increases ESA without
increase of the geometric surface area (GSA). Titanium nitride is often
selected for its electrical conductivity, amenability to thin ﬁlm deposition, and surface roughness when deposited by sputtering (Cogan,2008).
In some cases, such as with small electrodes (tens of μm), not evenpseudocapacitive mechanisms may provide enough charge transfer
with unmodiﬁed Pt or Pt-Ir electrodes (Cogan, 2008; Robblee and Rose,1990). One method to increase the reversible charge transfer limit isapplication of faradaic electric coatings, also called valence change oxides(Howell and Grill, 2015; Robblee and Rose, 1990). Iridium oxide ﬁlms
are the most common example; these possess two oxidative states andreadily accept or donate electrons during pulsing of the electrode.
Another method is to roughen the surface by etching, abrading, or
coating for increased ESA (Cogan, 2008; Geddes and Roeder, 2003; Leeet al., 2016a; Ordonez et al., 2014; Robblee and Rose, 1990; Weremfo
et al., 2015). The advantages of surface roughening may reach a limit
when signiﬁcant roughness or porosity increases the so-called pore resistance. As the conductive path through solution reaches deeper into
the roughened electrode surface, a time constant may be added to theelectrode’s response, and the deep surfaces of the electrode may not beutilized (Cogan, 2008; Robblee and Rose, 1990). Rough electrode surfaces may also be less mechanically robust and more prone to biofouling by protein adhesion (Cogan, 2008; Straka et al., 2018; Wellman

2.2. Electrodes
From an extracellular perspective, the inward rush of sodium ions
during the rising phase of an action potential is a strong current sink,accompanied by adjacent current sources: ahead, a weak source arisingfrom the initial discharging of the membrane; behind, a strong source
arising from the eﬄux of potassium ions. Given the current sources andsinks occurring at any instant during an action potential, there arenecessarily currents passing between them in the extracellular space. It
is these extracellular currents that may be measured by the electrodesof a PNI (Loeb and Gans, 1986; Stein and Oğuztöreli, 1978).
While charge in neural signalling is carried by ions, in metallic
conductors it is carried by electrons. Thus, at the metal-saline interface
of an implanted PNI electrode, there is necessarily some translation
between the two. The ﬂow of charge across this interface can occur by
two types of mechanisms: capacitive and faradaic. Water molecules
orient themselves on the electrode surface, forming what is known as
the double layer and collectively act as a dielectric between ioniccharges in the saline and electronic charges in the metal. Up to about 20
μC/cm2, the transfer of charge occurs only by Columbic attraction andrepulsion across this layer, with no charge-carrying species crossing theinterface (capacitive, Robblee and Rose, 1990). Above this limit, thedouble layer breaks down and both reversible and irreversible faradaic
reactions can occur. In reversible faradaic processes, also called pseudocapacitive, charge carriers cross the boundary and new chemical
species are created in oxidation and reduction (redox) reactions, however they do not leave the electrode surface and are readily reversed bya ﬂow of charge in the opposite direction. Irreversible faradaic processes
produce new chemical species that are not bound to the electrodesurface. They lead to corrosion or dissolution of the electrode, release of
toxic products into the surrounding ﬂuid and tissue, potentially harmful
shifts in pH, and gas formation from electrolysis of water. The regime in4

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

decreases sharply with distance. However, this tactic involves othercrucial compromises in PNI performance. Other strategies that havebeen explored include current steering and diﬀerential activation basedon ﬁber size. These and other examples of design strategies for improved stimulation performance will be discussed in Section 3.
Stimulation with varied parameters can also be used for the purposeof blocking, rather than activating neural activity, as discussed by
Bhadra and Kilgore (2018). Several reviews provide further discussion
regarding strategic parameters and waveforms for enhanced stimulation (Cogan et al., 2016; Grill, 2018; Mortimer, 1990; Mortimer andBhadra, 2004; Pasluosta et al., 2018).

et al., 2018).
Low electrode impedance yields improved stimulation eﬃciency
and noise reduction. Depending on the design, placement, and application of a PNI, geometric dimensions of the electrodes on the order ofhundreds of μm or even single mm may be appropriate and can achieve
low impedance (tens of kΩ at 1 kHz) without any further modiﬁcation
to a platinum surface. For smaller dimensions, faradaic coatings andincreased ESA also decrease electrode impedance (Howell and Grill,2015).
2.2.1. Stimulation
PN stimulation initiates an action potential by depolarization of theaxonal membrane to the threshold potential. This is accomplished byinjecting negative charge (i.e. a cathodic pulse) into the tissue, which,as it approaches the membrane, locally cancels the net positive charge
outside the cell and repels negative charges on the inside of the cell.
Though without any physical transfer of ions in or out of the cell, thespatial shifting of charges produces a local outward current and discharging of the membrane, thereby activating the sodium channels ofthat region and initiating the action potential (Mortimer and Bhadra,
2018). The goals of PN stimulation are to accomplish this activation
safely and eﬃciently while ideally aﬀecting only the targeted nerve
ﬁbers.
As for safety, general recommendations are to avoid irreversible
faradaic reactions and combinations of high charge density and charge
per phase. A long-established technique for the former is to use chargebalanced biphasic pulses. The second phase of opposite polarity reverses any surface-bound faradaic products that result from the initial
pulse, thereby avoiding charge accumulation on the electrode or in thetissue after repeated cycles and keeping the electrode potential within asafe range (Cogan, 2008; Donaldson and Donaldson, 1986). A general
guide for safe charge density, though it does not account for all relevant
factors (Cogan et al., 2016), is the Shannon limit, given bylog(D) = k − log(Q) where Q is the charge per phase or time integral of
the current, D = Q/A is the charge density or charge per phase divided
by the geometric area of the electrode, and k is a constant usually set at
1.85, or 1.5 for a more conservative guideline (Shannon, 1992). Any
combination of parameters above this limit is expected to induce tissue
damage. While these guidelines may serve as a starting point, tissue
damage resulting from stimulation is a complex topic and the reader isreferred to a number of other reviews and studies (Agnew et al., 1999,
1990; Agnew and McCreery, 1990; Cogan et al., 2018; Günter et al.,2019; McCreery et al., 1995, 1990; Pasluosta et al., 2018).
Stimulation eﬃciency is concerned with the amount of electricalenergy needed to achieve the desired neural activation, or, equivalently, the time integral of electrical power, which is the product of
current and voltage. Therefore, eﬃciency is increased by minimizing
the current, voltage, or duration of stimulation waveforms. Low electrode impedance contributes to eﬃciency by decreasing the voltage
required to deliver a given current. Another tactic is to shape the extracellular potential such that greater activation is achieved for thesame amount of current. Activation is proportional to the second spatial
diﬀerence in the longitudinal direction of the potential on the surface ofan axon (Howell and Grill, 2015; Rattay, 1986). Positioning two return
electrodes longitudinally on either side of a central cathode, known as a
tripolar conﬁguration, can achieve a greater second spatial diﬀerence
for the same amount of current as compared with a single (monopolar)
electrode with distant return (Howell and Grill, 2015). PNIs may also
take advantage of insulative materials and other electrode arrangements to shape the electric ﬁeld and “steer” the current for more eﬃciency and localization to the targeted nerve ﬁbers (Tarler andMortimer, 2004, 2003).
How best to limit activation to only the targeted nerve ﬁbers is oneof the prevailing considerations of PNI development. The moststraightforward approach is to position small electrodes as close as
possible to the nerve ﬁbers, taking advantage of activation that

2.2.2. Recording
A crucial consideration for peripheral nerve recording is the relative
amplitudes of extracellular neural signals and noise sources. Electricﬁelds produced by nerve ﬁbers are about three orders of magnitude
smaller than the electomyographic (EMG) ﬁelds produced by nearby
muscles (i.e. microvolts versus millivolts). Hoﬀer and Kallesøe (2001)
draw an analogy by equating the signal amplitudes to audio decibels, inwhich case recording a PN signal in the body is like listening to a quiet
voice while a loud horn is sounding one meter away.
The simplest recording arrangement is a monopolar electrodeplaced near the nerve and a distant electrode used as reference.
However, this arrangement is highly susceptible to EMG noise sources.
Diﬀerential or bipolar recording is eﬀective in cancelling some of thisnoise by subtracting the signals of two electrodes placed near the nerve
and outputting their diﬀerence. Bipolar electrodes should be oriented
along the length of the nerve, ideally spaced to match the distance
between source and sink of a propagating action potential which will
maximize the amplitude of the measured diﬀerential signal. This distance can be approximated by multiplying the nerve ﬁber’s conduction
velocity and the time between maximum inward and outward currents,
or about half the period of an action potential. As a rule of thumb,myelinated ﬁber conduction velocity, in m/s, is approximately six times
the ﬁber diameter in μm; for unmyelinated ﬁbers it is a factor of two(Kandel, 2013). A “medium” diameter myelinated nerve ﬁber of 10 μm,
for example, has a conduction velocity of ∼60 m/s. Combined with thenominal 1 ms duration of an action potential, we can estimate an ideal(though not necessarily practical) electrode spacing of 30 mm. The
most eﬀective electrode arrangement for rejecting noise is the tripolar
conﬁguration in which a central electrode is measured diﬀerentiallywith respect to electrodes on either side. The tripolar electrode conﬁguration and its variations, along with other strategies for enhanced
PN recording, are discussed in Section 3.1.1.1.
A general principle for optimal recording of neural signals is toposition the electrode(s) as close as possible to the signal source, which
for PNIs is the axonal membrane exposed at nodes of Ranvier. As distance between the signal source and electrode increases, more current isshunted away through the tissue and the signal is attenuated. Anotherfactor aﬀecting recording quality is tissue impedance. Diﬀerential recording of a nerve, perhaps counterintuitively, can gain increased
signal amplitude from increased tissue impedance between the electrodes, assuming the source-to-electrode distance remains the same. To
ﬂow from current source to sink, extracellular currents must ﬂow parallel to nerve ﬁber and the bipolar electrode alignment. Thus, increased
impedance of this path yields an increased voltage diﬀerence between
the electrodes.
2.3. Invasiveness versus selectivity
As described above, for optimal stimulation and recording performance it is desirable to position electrodes close to the targeted nerve
ﬁbers. However, the layered, bundled, densely-vascularized anatomical
structure of PN conﬂicts with this goal. Directly inserting electrodes
into nerve ﬁbers may allow high stimulation and recording selectivity,
but breaching the blood-nerve barrier and inducing other injuries to5

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

implanted at diﬀerent locations (e.g. extremity versus core) may incite
vastly diﬀerent host responses due to diﬀerent degrees of nerve
movement (Naples et al., 1990).
Geometry also plays an important role in biocompatibility. Whilemacroscale geometry of PNIs will be discussed in Section 3, a generalrecommendation provided here is to avoid sharp edges and bulky
masses. Surface geometry may be adjusted to achieve a desired eﬀect
depending on the application. A smooth surface minimizes tissue adhesion and facilitates device mobility. A porous or perforated surface
will encourage tissue ingrowth, which may be desirable for anchoring
to the nerve and in some cases for increasing recorded signal amplitude,
but will make future removal of the device diﬃcult (Hoﬀer andKallesøe, 2001; Marks and Loeb, 1976; Naples et al., 1990; Stein andOğuztöreli, 1978).
In addition to the generalized host response discussed here, otherpotential agents of neural damage must be considered in all aspects of
PNI design, including surgical implantation and connection methods(Sections 2.6 and 2.7). Furthermore, diﬀerent types of PNI designs
come with their own considerations of implicit nerve damage, as discussed in Section 3.

Fig. 4. Plot illustrating the strategic tradeoﬀ between invasiveness to the nerveand selectivity. Existing PNI types generally lie on the diagonal, while a variety
of strategies attempt to enhance performance toward the “ideal” region of theplot. Categories omitted from the plot are optical PNIs (Sec. 3.5) and hybrid
approaches (Sec. 3.6).

2.5. Material selection

nerve tissues may increase likelihood of interface failure in the longterm. On the other hand, positioning electrodes outside of the nerve ismore likely to maintain long-term interface stability but suﬀers frompoor selectivity and signal discrimination. Such trade-oﬀs can be described in multiple ways: invasiveness versus selectivity; stability versus
resolution; safety versus eﬃcacy; chronic viability versus acute functionality (Fig. 4). Section 3 will highlight approaches that balance or
circumvent trade-oﬀs towards the combined goal of selectivity andlong-term performance.

Based on the multiple facets of biocompatibility discussed above,
there exists no list of universally-accepted materials for use in PNIs.
Rather, material selection must account for interplay of multiple aspects of the design and application. Materials commonly used in PNIs
include silicone, polyimide, and Parylene C as insulation and platinum
and iridium as conductors. Among these, only silicone, platinum, andplatinum-iridium have been clinically implemented in PNIs on a large
scale. Parylene is known for its biological inertness and use in many
approved medical devices and has seen success in neural interfaces usedin research (Hara et al., 2016; Lecomte et al., 2017; Scholten and Meng,
2015; Sohal et al., 2016). Polyimide has seen similar success in research
applications but with limited historical use in approved medical devices(Hassler et al., 2011; Stieglitz et al., 2011, 2005). However, thesepolymer materials suﬀer from moisture permeation, delamination, andfabrication diﬃculties and limitations (Hassler et al., 2011; OrtigozaDiaz et al., 2018; Rubehn and Stieglitz, 2010; Srinivasan et al., 2016).
When inserted into the endoneurium, Parylene and polyimide canprovoke growth of a signiﬁcant encapsulation layer (de la Oliva et al.,2018a,c; del Valle et al., 2015; Wurth et al., 2017). Silicone elastomer
may eventually degrade in the body (Garrido et al., 1993). According to
the physician’s manual for a commercially distributed PNI, the integrityof silicone is not guaranteed after about 25 years (Finetech-Brindley
Sacral Anterior Root Stimulator (CPC2): Notes for Surgeons andPhysicians, 2016). Even platinum, with its reputation for robustness
and excellent characteristics, cannot completely avoid dissolution
under chronic stimulation (Robblee and Rose, 1990; Wellman et al.,2018).
New materials being explored for PNIs include hydrogels, liquid
crystal polymer, SU-8, conductive polymers, carbon ﬁber, graphene,carbon nanotubes, and even collagen (Fattahi et al., 2014; Green and
Abidian, 2015; Kim and Romero-Ortega, 2012; Tian et al., 2018;
Wellman et al., 2018; Woloszko et al., 1998). If clinical translation ofdevices using exploratory materials is desired, the associated regulatory
obstacles should be evaluated. Materials selected may impact a device’s
safety proﬁle which is determined following a rigorous battery of biocompatibility tests, such as those outlined by the widely accepted
standard ISO 10993-1, “Biological evaluation of medical devices”(Standards and Guidances for Neurological Devices, 2018). As Wellman
et al. (2018) pointed out, beyond in vitro and animal demonstrations ofsafety, any “new” material will also need substantial historical data to
warrant the ﬁnancial risk of a clinical trial (which typically exceeds$100 million). Overall strategy should not be limited to optimization of
one or several material variables such as softness or conductivity, but

2.4. Host response
Implantation of any object triggers an immune reaction which isintended to facilitate healing by destroying, removing, or containing
the intruding object, clearing debris from the area, and increasing local
ﬂuid for migration of repair cells (Coleman et al., 1974). This “foreign
body response” generally involves, at a minimum, immediate bloodclotting around the implantation site, acute inﬂammation, and ensuingencapsulation of the implant in ﬁbrotic tissue (Stroncek and Reichert,2008). If the implant continues to have some injurious eﬀect other thanits presence, such as chemical or mechanical irritation, the response
may intensify and include worse eﬀects such as extreme inﬂammation,
edema, hemmorage, or necrosis (Naples et al., 1990). Therefore, a PNI
implementation should minimize or tailor host response to maximize its
operational lifetime.
Naples et al. (1990) provide a useful framework to summarize themultiple aspects of biocompatibility that the PNI designer should consider in order to minimize the host response: chemical, mechanical, andgeometric. It is important to note that in and among these properties,
which are mostly intrinsic to the device, there are also a number of
extrinsic considerations such as implant site and stimulation characteristics.
Chemical biocompatibility is chieﬂy a material issue. The interface
between tissue and the implant surface is the target of chemical reactions that will progress until an equilibrium is reached. In the process,the implant may degrade and release noxious products into the body.
Therefore, inert or near-inert materials are preferred (Naples et al.,1990). Furthermore, the materials should remain inert and intact whensubjected to prolonged electrical stimulation (White and Gross, 1974).
Mechanically, an implant should ideally possess similar properties
to the nerve and also accommodate relative motion, either by moving
with the tissue or avoiding friction at the interface (Naples et al., 1990).
Accordingly, the chosen implantation site may play a large role in theapparent mechanical biocompatibility of a device; the same device
6

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 5. Illustrations of common forms (blue) representing the main PNI categories. The PNI forms contain both the electrode, insulation, and any additional
supporting structure.

2009; Ramachandran et al., 2006; Schuettler et al., 2000; Wodlinger
et al., 2015) and others have replaced the cable entirely with wirelessapproaches (Larson and Towe, 2011; Lee et al., 2018a,b; Park et al.,2015a,b; SetPoint Medical, 2019; Sharma et al., 2011).
A recently proposed packaging solution involves routing the remaining nerve ending of an amputated limb into the interior of thebone, which oﬀers a protected space for housing the interface and othercomponents, and also a convenient site for direct connection to a robotic prosthesis (Israel et al., 2018).
Section 4.2.1.2 will cover packaging in more detail with speciﬁc
considerations for implementation in animal studies. Though some ofthe same principles apply to clinical use of PNIs in humans, packaging
of implantable devices for human use is a highly specialized topic in the
medical device industry and will not be speciﬁcally covered.

entail a careful balance of numerous interdependent trade-oﬀs acrossall material-related design factors.
2.6. Surgical implantationPNI design should incorporate the entire scope of a device’s use,
including handling and surgical implantation (Mortimer et al., 1995).
Devices should be robust and the handling instructions straightforward.
Some designs may beneﬁt from auxiliary features, such as tabs, loops,
or handles that solely serve as a grasping point during device preparation and installation (Payne et al., 2019; VNS Therapy System
Epilepsy Physician’s Manual (US), 2018). Such features can be designed
for easy in situ removal after installation.
The implantation process itself should be made as eﬃcient andfailproof as possible. Something as simple as an “up” label on anotherwise transparent polymer device can help avoid the easy but
terminal mistake of installing it inside-out (Cobo et al., 2019). Chancesof accidental damage to the nerve during the installation process should
be minimized, such as pinching trauma resulting from premature closure of a nerve cuﬀ. Several groups have devised special tools to facilitate installation of the PNI, such as a syringe-like electrode injector(Yim et al., 2018), or purpose-made forceps for opening, positioning,and closing a helical or spiral electrode (Bakula and Mortimer, 2000;
Bullara, 1990; Rise and Klepinski, 1992). It is a prudent choice to makethese tools in forms that are already familiar to surgeons. Others haveincorporated installation mechanisms into the device itself, such as aneyelet for leading the device around or through the nerve with a suture(Boretius et al., 2010; Cobo et al., 2019; Yoshida et al., 2000).

3. Types of peripheral nerve interfaces
This section surveys the diﬀerent PNI approaches that have been
explored. The categorized approaches (Fig. 5) are presented along withdesign considerations and best practices that have emerged. Particularattention is given to strategies for optimizing eﬃcacy and safety, and,
where available, in vivo longevity of the interface is reported.
3.1. Extraneural
Extraneural PNIs are situated outside the nerve; this placement canpreserve nerve health at the cost of greater distance to individual ﬁbers.
However, extraneural PNIs may injure the nerve by constriction which
prevents crucial nutritional and metabolic transport through neuralblood vessels and the axons (Ju et al., 2006; Krarup et al., 1989). Ju
et al. (2006) observed a decrease in blood prefusion beginning at 30.5mmHg of nerve constriction. Constriction may inadvertently arise whena circumneural electrode is ﬁtted too closely around a nerve trunk. The
ensuing acute inﬂammation that follows implantation temporarily increases the nerve diameter and after the swelling subsides, a layer ofﬁbrotic encapsulation invariably forms between the nerve and the interface, eﬀectively increasing its diameter (Hoﬀer and Kallesøe, 2001).
Nerve diameter will also change with motion since volume is conserved
during changes in length, though the literature does not specify if thischange is signiﬁcant enough to be aﬀected by PNIs.
The anatomical groove in which a nerve trunk resides may induce
lateral compression as it attempts to accommodate the extra bulk of an
extraneural PNI. Mechanical pressure from adjacent tissues can betransmitted through the compliant material of an extraneural PNI to thenerve. Rydevik et al. (1981) observed blood ﬂow in a nerve to be affected starting at 20 mmHg of lateral compression and complete occlusion at 60 mmHg.
Relative motion between the PNI and the nerve can also be a sourceof damage. Cabling connected to the PNI can apply tension, creating
irregular pressure points on the nerve via lateral pulling or torquing(Hoﬀer and Kallesøe, 2001; Restaino et al., 2014). Before ﬁbrotic encapsulation anchors a PNI to the nerve, there may be motion between

2.7. Packaging
Another crucial aspect of any PNI is the series of supporting components which facilitate connection to the interface from outside thebody, frequently referred to as packaging. In practice, packaging has
many more potential points of failure than the interface itself (Mortimer
et al., 1995). Most PNI packaging is concerned with electrical connections, but newer devices may also include ﬂuidic connections for drugdelivery.
Usually, the minimum arrangement of components consists of theinterface connected to either a telemetry and control module for clinicaland long-term use, or a percutaneous connector for shorter-term studies. The main principles of successful packaging for PNIs are to avoid
any transfer of tension from the leads to the interface, protect the leads
from breakage, and ensure sealing of any components that should remain isolated from tissues and ﬂuids (Naples et al., 1990).
The standard practice of accessing each of a PNI’s electrodes via its
own lead wire with a percutaneous connection is not scalable to highchannel counts (dozens or more); increased channel counts are inevitable as advanced PNIs seek more degrees of freedom. To reduce thenumber of electronic connections, a number of groups have integrated
amplifying and multiplexing electronics into the device or the cable(Ballini et al., 2017; Lancashire et al., 2018, 2019; Lertmanorat et al.,7

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Mengthem, causing abrasion and an aggravated response (Naples et al.,1990). If ﬁbrotic tissue on the outside of the PNI anchors it in place
relative to the surrounding tissues, the nerve may also become anchored and experience damaging levels of tension (Hoﬀer and Kallesøe,
2001; Sunderland, 1990).
The following explores extraneural PNI strategies, highlighting bestpractices for mitigating nerve damage and enhancing recording andstimulation.
3.1.1. Cuﬀ
Extraneural cuﬀs are the most widely used and studied of PNIs.
Their general form is an insulative tube wrapped around the nerve
containing exposed metal sites on its inner wall. In 1948, Sarnoﬀ et al.
described a method of looping wires around the phrenic nerve andwrapping the assembly with a piece of polyethylene sheeting. Sarnoﬀ
explains: “In this way the phrenic nerve could not escape contact withthe electrode but was not compressed by it and the latter could notdeliver an impulse to any other structure” (Sarnoﬀ et al., 1948). A more
standardized cuﬀ design emerged consisting of wires sewn into the wallof a section of silicone tubing with a longitudinal slit across its length(Loeb and Peck, 1996; Stein et al., 1977). New fabrication technologies
enabled greater dimensional precision, design ﬂexibility, and a wider
choice of material properties. Hoﬀer and Kallesøe (2001) provide athorough and practical review on the design and use of nerve cuﬀsusing traditional fabrication techniques.

Fig. 6. Demonstration of EMG rejection using a tripolar electrode conﬁguration. In monopolar and bipolar recordings, the neural signal is dominated by
EMG and only evident after muscle denervation. In the tripolar recording theneural signal is clearly seen in the presence of EMG. Reprinted from (Steinet al., 1977) with permission from Elsevier.

other. The ﬁrst four requirements are readily met by leveraging the
precision and layout ﬂexibility of microfabrication techniques. The
latter requirement has bearing on design of the cuﬀ’s closure mechanism which will be discussed in more detail below.
Several improvements to this historical quasi-tripolar conﬁguration
have been explored to further enhance its noise cancellation abilities(Fig. 7). Fibrotic tissue growth over time may aﬀect the impedance
balance between the two end electrodes or the linearity of the voltage
gradient inside the cuﬀ. To compensate for this, Struijk and Thomsen(1995) connected the two end electrodes with a potentiometer, allowing for adaptive balancing of the asymmetric tissue eﬀects. This
adaptive tripolar conﬁguration decreased the EMG artifact by 30 %while only decreasing the ENG by 3 %. Pﬂaum et al. (1997) ampliﬁed
the central electrode with respect to each end electrode separately thenadded the two outputs in a conﬁguration called true tripolar. This resolved the dissimilar impedance between one and two-electrode inputs
and yielded a 10 % improvement in signal-to-noise ratio (SNR). Plachta
et al. (2014) later used the true tripolar conﬁguration to perform
adaptive balancing by making adjustments to the ﬁrst ampliﬁcation
stage. In a revised quasi-tripolar conﬁguration, Chu et al. (2012)
achieved an 11 % improvement in SNR by replacing the central electrode with a pair of shorted electrodes separated by a strategic distance.
The mechanism by which this improvement is achieved is subtle: as theseparation distance between the center electrodes increases from zero(i.e. a single electrode), the EMG signal decreases linearly while the
ENG decreases non-linearly, remaining near its maximum value at short
separation distances. Therefore, the ratio ENG/EMG is maximized witha short separation between a pair of central electrodes (Chu et al., 2012;
Lee et al., 2016a).
Other strategies for noise rejection include passive shunting or
shielding features. The screened tripole devised by Rahal et al. consists of
two additional electrodes which are shorted together and positioned
outside the end electrodes. They serve to provide a low-resistance pathfor shunting the external signal around the lumen of the cuﬀ rather than
through it. The eﬀect is a shallower voltage gradient through the cuﬀ sothat the recording is less aﬀected by asymmetries in the tripole, similar
to what would be achieved by a longer cuﬀ (Rahal et al., 2000b). Another approach is to include a conductive shield layer covering theentire outside of the cuﬀ. Modelling by Sadeghlo (2013) estimated a 70
% reduction in noise using this technique, and Sabetian et al. (2017)
found it reduced stimulus artifacts by 63 %.

3.1.1.1. Strategies for enhanced recording with extraneural cuﬀs. The
goals of neural recording are rejection of noise from extraneuralsources (namely, EMG) and discrimination of signals from diﬀerent
nerve ﬁbers.
3.1.1.1.1. Noise rejection. An eﬀective and widely used method toreject large extraneural noise sources in cuﬀs is the tripolar electrodeconﬁguration: a central electrode is connected to one input of adiﬀerential ampliﬁer and two equidistant surrounding electrodes areshorted together and connected to the other input (Stein et al., 1975).
The external EMG source can only access the cuﬀ electrodes by entering
one end and leaving the other; the resultant voltage gradient is linearly
dependant on distance. The two shorted electrodes eﬀectively measure
the average potential between them, and therefore any EMG voltage
picked up by them is equivalent to the potential at their midpoint.
Consequently, the EMG signal measured by the ampliﬁer is thediﬀerence between the central electrode and a virtual electrode atthat same point, which is zero. Meanwhile, signals generated within thecuﬀ, i.e. the electroneurogram (ENG), are not spatially linear and are
not cancelled. Mathematically, the measured ENG is the second spatial
diﬀerence of the voltage across the length of the cuﬀ (Stein et al.,1977).
Stein et al. (1977) provided a demonstration of EMG cancellation
with a tripolar cuﬀ, shown in Fig. 6. Monopolar, bipolar, and tripolar
cuﬀ recordings were recorded in the presence of EMG from nearby
muscles, after which the muscles were denervated and the recordings
were repeated. In the monopolar and bipolar recordings, the neuralsignal is dominated by EMG and only revealed following denervation,
while the tripolar recording captures the major features albeit withsome EMG contamination.
Several conditions must be met in order for tripolar cancellation of
extraneural noise to be eﬀective (Loeb and Gans, 1986). First, the impedance of the two end electrodes must be as similar as possible.
Second, the middle electrode must be equidistant between the twoends. Third, the impedance of the ampliﬁer inputs should be similar,
accounting for the fact that one input is coming from a single electrodeand the other input is coming from two electrodes. Fourth, the voltage
gradient is slightly nonlinear at the entrance and exit of the cuﬀ, so theend electrodes should be positioned away from the cuﬀ ends (generally
by a few mm) (Rahal et al., 2000a). Finally, the cuﬀ must be well-sealed
and insulated to ensure a constant voltage gradient from one end to the8Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 7. Electrode conﬁgurations for recording with extraneural cuﬀs.

Finally, ﬁltering can further improve noise rejection. ENG generally
occupies the range of 100–5000 Hz with most of its power between 1
and 3 kHz, while EMG is 1–500 Hz with most of its power between 100
and 300 Hz (Tyler, 2018). A notch ﬁlter around 60 Hz is also usuallynecessary to reject room noise.
3.1.1.1.2. Signal discrimination. The signals recorded by a cuﬀ are asummation of all the signals from the nerve ﬁbers it encircles, which
often number in the thousands. The use of small electrodes may
enhance the ability to localize signals coming from fascicles near thesurface of the nerve. However, arraying small electrodes around theinner wall of a cuﬀ may only yield modest improvements due toextracellular currents’ spreading throughout the cuﬀ lumen (Hoﬀer andKallesøe, 2001; Lichtenberg and De Luca, 1979; Sahin and Durand,
1997; Struijk et al., 1997). Such a strategy is most eﬀective whencombined with distal placement on the nerve where there is greater
separation among fascicles as they near branching points (Silveira et al.,2018).
Discrimination of signals based on location in the nerve may behelped by modiﬁcation of the cuﬀ’s inner wall, as proposed by Hoﬀer
et al. (Hoﬀer et al., 1998; Hoﬀer and Kallesøe, 2001). The multichambered cuﬀ has parallel ridges on the inside wall which are intended to contact the surface of the nerve and form isolated extraneuralchambers, each with its own tripolar set of electrodes. This would potentially limit the spreading of extracellular currents that would
otherwise be picked up on multiple electrodes around the circumference.
One long-established method for isolating a signal is coherent
averaging, also called spike-triggered averaging (Hoﬀer et al., 1981;
Mendell and Henneman, 1971). A signal which is too small to distinguish in one recording may emerge after tens or hundreds of the recordings are averaged. Coherent averaging requires a repeated signalthat is time-coupled to some other detectable signal such as a stimulus
or biological signal. Plachta et al. (2014), for example, averaged recordings of baroreceptor signals in the vagus nerve using the rising edgeof the electrocardiogram as a trigger.
Discrimination between eﬀerent and aﬀerent signals may be possible given the reversal of the recorded signal shape. However, a

tripolar conﬁguration may make it more diﬃcult to distinguish becauseof the symmetric electrode arrangement. It is possible to maintain directionality with the use of multiple tripoles in a single cuﬀ, perhaps
interleaved or sharing electrodes in each set (Hoﬀer et al., 1981).
Extending the idea of using multiple electrodes to distinguish signal
direction, Taylor and co-workers demonstrated the ability to discriminate signal sources by conduction velocity (Al-Shueli et al., 2014;
Donaldson et al., 2008; Metcalfe et al., 2018, 2014; Schuettler et al.,2013; Taylor et al., 2004). Conduction velocity of a ﬁber correlates to
its size, and diﬀerent sized ﬁbers generally correlate to diﬀerent typesof signals within the nerve. For example, pain signals are carried bysmall, slow ﬁbers while proprioceptive signals are carried by large, fast
ﬁbers. Velocity-selective recording is based on a “delay-and-add” process using a cuﬀ with about ten electrodes longitudinally arranged witheven spacing. A delay, corresponding to position in the row of electrodes, is added to the output of each overlapping tripole before thesignals of all tripoles are summed. The result is that a neural signal
propagating at a particular velocity will end up being summed constructively with itself in all of the outputs. The resulting peak reveals
the propagation velocity of the signal and its amplitude is multiplied by
the number of electrodes (Taylor et al., 2015). An extension of thismethod may be used to extract the amount of activity occurring in anerve at each velocity, called the “velocity spectral density” (Metcalfe
et al., 2014). These methods have recently been demonstrated to capture distinct physiological neural signals from the vagus nerve of a pig,
discriminated by direction and velocity (Metcalfe et al., 2018).
An emerging technique for recordings of higher spatial resolution isfast neural electrical impedance tomography (EIT) (Aristovich et al.,2018). In this technique, the impedance changes among an array of
extraneural electrodes are measured while a constant current is passed
between any two of them. The measurements are used to reconstructtomographic “images” of the nerve cross-section using a process similar
to inverse source analysis for electroencephalography. The changes inmembrane conductivity as ion channels open and close are captured in
the image and thereby give an indication of neural activity. Fast neural
EIT has been demonstrated to localize compound action potentials to aparticular fascicle in rat sciatic nerve with temporal resolution of 0.3 ms
9

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Mengand spatial resolution of 100 μm. This technique currently requires
averaging over tens of seconds or minutes (Aristovich et al., 2018).
Relatedly, a more direct adaptation of inverse source analysis techniques was investigated by Zariﬀa et al. (2011) though with limited
success. Wodlinger and Durand (2010, 2009) also present a source-localization technique utilizing spatial ﬁlters determined by the interface
geometry and demonstrated localization of activity in two fascicles.

side eﬀects.
Fiber size selectivity is based on the fact that larger diameter ﬁbers
have a lower stimulation threshold. Gorman and Mortimer (1983)
found that shortened pulse durations exaggerate the eﬀect as they dofor spatial selectivity, by increasing the diﬀerences in threshold for
diﬀerent ﬁber sizes. Notably, the tendency to recruit large ﬁbers before
small ﬁbers is opposite to the physiological recruitment order for voluntary muscle contractions which begins with low-force, fatigue-resistant motor units and progresses to high-force, quickly-fatiguing
motor units. Consequently, graded stimulation levels intended to reproduce natural-like motor control will instead produce poor gradationof force and quick onset of fatigue. This is evident in the steep slope ofmost recruitment curves (plots of a muscle’s contraction force versus
stimulation amplitude) found in the literature. Sub-threshold depolarizing pre-pulses before the main pulse, as with the distance-threshold
relationship, have been found to invert the diameter-threshold relationship, which can be used to achieve a more natural recruitment
order (Sassen and Zimmermann, 1973).
Lertmanorat and Durand (2006; 2004) proposed an extraneural
method to recruit small myelinated ﬁbers before large ones that does
not rely on use of a long pre-pulse and large amounts of charge injection, but instead strategically shapes the extracellular voltage proﬁleusing an array of electrodes. This technique is based on the proportionality of ﬁber excitation to the second spatial diﬀerence of thevoltage proﬁle along the nerve, and internodal spacing to ﬁber diameter. The electrode conﬁguration is an evenly spaced array of 5–11
alternating anodes and cathodes. A ﬁber whose internodal spacing
approximately matches the spacing between cathodes will see a more
uniform voltage proﬁle and thus a drastically diminished second spatial
diﬀerence. Meanwhile, ﬁbers with smaller internodal spacing, especially those matching the cathode-anode spacing, will have a larger
second spatial diﬀerence and thus can be activated independently of thelarger ﬁbers. This technique was demonstrated to be eﬀective at preferably recruiting smaller myelinated ﬁbers in both simulations andexperiments and decreased the slope of recruitment curves by at least afactor of four as compared to a tripolar cuﬀ (Lertmanorat et al., 2006).

3.1.1.2. Strategies for enhanced stimulation with extraneural cuﬀs. Goalsof neural stimulation are eﬃciency and selectivity. Extraneural cuﬀsderive their eﬃciency from the insulative wall which conﬁnes
stimulation currents. A well-sealed cuﬀ is not as crucial for
stimulation as it is for recording, but prevents wasteful spill-over ofcurrent which may cause unwanted side eﬀects (Hoﬀer and Kallesøe,
2001). A tripolar electrode conﬁguration with return electrodes near
the cuﬀ ends helps conﬁne current and also shapes the generated ﬁeld
to have a higher second spatial diﬀerence for the same amount of
current (as compared with a monopole) thereby more eﬃciently
initiating action potentials (Howell and Grill, 2015).
To achieve selectivity, it is straightforward to install cuﬀs at distal
locations after branch points (Payne et al., 2019). However, this approach is often undesirable because the branched nerves are smaller,
more fragile, and less accessible. It also requires multiple implantations,cables, and connectors if multiple degrees of freedom are required. It ispreferable to implant a single PNI on a larger compound nerve trunk.
The primary mechanisms for diﬀerential activation of nerve ﬁbers
are spatial selectivity and ﬁber size selectivity. Grill (1995) provided anin-depth review of selective stimulation strategies using cuﬀs. A summary is provided here along with recent advancements.
Spatial selectivity is based on the increase in a ﬁber’s stimulus
threshold as distance from the electrode increases. A given stimulus
pulse can activate ﬁbers within a certain radius. Spatial selectivity may
be improved by arraying and positioning electrodes around the cuﬀ
with reference to the fascicle positions in the nerve trunk. Furthermore,
the spatial pattern of activated ﬁbers can be shaped via electrode arrangement. A tripolar arrangement is generally used with the cathodeat center and anodes at the ends, though Tarler and Mortimer found amonopolar arrangement to be nearly as selective (Tarler et al., 1995;
Tarler and Mortimer, 2003). A transverse anode, i.e. on the other side of
the cuﬀ, may also be used to “steer” the current depending on theamplitude of the anodal pulse (Grill et al., 1991; Sweeney et al., 1990;
Veraart et al., 1993). Grill and Mortimer (1996a) found the use of
shortened pulses can exaggerate the eﬀect of spatial selectivity, increasing the diﬀerence in stimulation threshold for ﬁbers at diﬀerent
distances from the electrode. An inversion of the distance-threshold
relationship can be achieved by use of sub-threshold depolarizing prepulses before the main stimulation pulse. These pre-pulses deactivate
the sodium channels in nearby ﬁbers while those farther away are
unaﬀected. In such a manner, ﬁbers at the center can be activatedwithout activating those near the surface (Grill and Mortimer, 1997).
Other waveforms for blocking can also be used for this purpose(Vuckovic et al., 2008).
Beyond these guidelines, spatially selective activation of fascicles
with cuﬀs is a case-by-case practice. A period of characterization and“tuning” after implantation will generally be required to ﬁnd theelectrode combinations that achieve the desired eﬀects. Tarler andMortimer (2004) demonstrated selective activation of each of four
fascicles in cat sciatic nerve using only four monopolar electrodes incombination with anodic and cathodic steering currents. Veraart et al.
(1998) implanted a cuﬀ on a human optic nerve and with four electrodes elicited a range of percepts roughly corresponding to four
quadrants in the visual ﬁeld. Good selectivity was achieved in humanupper extremity using a combination of monopolar electrodes andcurrent steering (Polasek et al., 2009). Plachta et al. (2014), with atripolar conﬁguration, achieved activation of baroreceptors in rat vagus
nerve that was at least selective enough to avoid cardiac and respiratory

3.1.1.3. Cuﬀ closure mechanisms. Cuﬀs are largely diﬀerentiated fromeach other by the mechanisms by which they close around a nerve.
Closure mechanism design should consider sealing, sizing, and ease of
installation. As previously mentioned, complete sealing along the entire
length of the cuﬀ is crucial for maximal recording quality. The seal
should also deny the aggressive ingrowth of ﬁbrotic tissue at any point
other than the ends of the cuﬀ. The host immune response will breachand expand even the tiniest gap or crack in the seal by packing it withﬁbrotic tissue, which then provides a path for internal signals to leakout and for external noise to invade (Hoﬀer and Kallesøe, 2001; Loeband Gans, 1986; Loeb and Peck, 1996; Naples et al., 1990).
Sizing of a cuﬀ’s diameter must balance several factors. A ﬁt that is
too tight will compromise nerve health by constriction during postoperative swelling, tissue encapsulation, and motion. If it is too loose,the recording and stimulation performance will be impaired. The prevailing convention for cuﬀ sizing is to use an inner diameter that is
1.2–1.5 times the nerve diameter. Hoﬀer and Kallesøe (2001) note thata looser ﬁt may increase nerve damage because it has a tendency toallow misalignment of the cuﬀ’s and nerve’s axes, resulting in sharppressure points at either end of the cuﬀ. Likewise, Naples et al. (1990)
suggested that a closer ﬁt may improve stabilization of the cuﬀ on thenerve, and that the greater cause of damage is incongruous motion
between the two. Naples’ review found a variety of confounding factors
in reports of health or harm caused by various cuﬀ diameters, including
various implantation sites subject to diﬀerent amounts of motion anddiﬀerent cabling qualities that may serve to either aggravate or stabilize
the interface. From these discussions, when selecting a ﬁxed cuﬀ diameter, it seems prudent to aim for 1.2–1.3 times the nerve diameter.
3.1.1.3.1. Split cuﬀs. Split cuﬀs possess a simple design and were10

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 8. Extraneural electrodes. A) Cuﬀ with “piano hinge” closure mechanism (Kallesøe et al., 1996). B) Silicone spiral cuﬀ. Image reproduced courtesy of Ardiem
Medical, Inc (Neural Cuﬀ | Ardiem Medical, 2019). C) Two-electrode helical array (McCreery, 2004).

preferred throughout the ﬁrst two decades of PNI research. They consistof a sheet of insulation folded around the nerve or a piece of tubing witha longitudinal end-to-end slit. To close and seal split cuﬀs, sutures and/or silicone can be applied and the silicone cured in situ. This method isstill widely used for its simplicity (Avery and Wepsic, 1973; Caravaca
et al., 2017; González-González et al., 2018; Hoﬀer and Kallesøe, 2001;
Loeb and Peck, 1996; Stein et al., 1977; Tian et al., 2018). However, theobvious drawbacks are imprecision and inconsistency. Sutures on theexterior of a cuﬀ may cause problematic adhesion to surrounding
tissues (Hoﬀer and Kallesøe, 2001).
Alternatively, an outer shell can be applied which gives form to the
ﬂexible inner cuﬀ, holding it closed via spring force (Micro-Leads
Research Products, 2019). A double-layered cuﬀ design used an outersplit cuﬀ wrapped around the inner cuﬀ with its longitudinal slit positioned on the opposite side of the nerve, which was then sealed withsilicone (Loeb and Peck, 1996). This design is particularly resistant totissue ingrowth but at the expense of a bulkier proﬁle.
Kallesøe et al. presented a split cuﬀ having inter-locking teeth thatare aligned and linked by inserting a single nylon suture (Fig. 8a), much
like the hinge of a piano lid (Hoﬀer and Kallesøe, 2001; Kallesøe et al.,1996).
A few examples in literature have bendable metal skeletons, or armatures, embedded inside the cuﬀ wall, giving it a formable structure.
An armature cuﬀ intended for acute experiments by Foldes et al. (2011)
is simply pinched closed to hold it in place around the nerve. A designby Crampon et al. (1999) used a temperature sensitive shape-memory
alloy which can maintain an open state for easy installation around thenerve, followed by automatic closing once it warms to body temperature.
3.1.1.3.2. Spiral cuﬀs. Unlike split cuﬀs that require additional
closure mechanisms to achieve appropriate ﬁtting around a nerve,
spiral cuﬀs were introduced that could “self-size” (Naples et al., 1988,
1986). Spiral cuﬀs (Fig. 8b) have a dynamically adapting diameter andcan be installed with a snug ﬁt (a resting diameter of about 0.9 times
the nerve diameter), keeping electrodes in contact with the surface ofthe epineurium while accommodating swelling, ﬁbrosis, and motion.
The accommodating ﬁt can minimize abrasion of the nerve before tissue
encapsulation can anchor it (Naples et al., 1990).
Models showed that a spiral cuﬀ could accommodate a 21 % expansion of the nerve diameter before reaching the 20 mmHg pressure
safety threshold Rydevik observed as the starting point of eﬀects on
blood ﬂow (Naples et al., 1988). Later, in vitro measurements by Cuoco
and Durand (2000) revised this limit to at least 33 % expansion of thenerve. A seven-month implantation on cat sciatic nerves showed morphological changes only attributed to cabling issues while a subsequent
four-month implantation showed no signiﬁcant morphological diﬀerences as compared with controls (Grill and Mortimer, 2000; Romero
et al., 2001). The majority of investigations on cuﬀ-based stimulation
by Mortimer and co-workers at Case Western Reserve University in the
1990s and 2000s were also performed using spiral cuﬀs (Grill et al.,
1991; Grill and Mortimer, 1998, 1997, 1996b, 1996a; Tarler et al.,

1995; Tarler and Mortimer, 2004, 2003; Veraart et al., 1998, 1993).
Spiral cuﬀ implantations in humans have seen stable function without
adverse eﬀects up to 11 years in limbs (Christie et al., 2017; Fisher
et al., 2009; Polasek et al., 2009), and up to 7 years on an optic nerve(Brelén et al., 2005).
Most spiral cuﬀs have been made from silicone and platinum foil,
like the example shown in Fig. 8b. It is desirable to minimize thethickness of the cuﬀ in order to avoid a bulky discontinuity in the lumen
where the inner edge of the spiral ends. For this purpose, and to takeadvantage of the other beneﬁts of microfabrication, several groups havemicrofabricated spiral cuﬀs from the thin-ﬁlm polymers polyimide andParylene (Kang et al., 2015; Lee et al., 2016a; Plachta et al., 2014;
Rodríguez et al., 2000; Schuettler et al., 2000).
3.1.1.3.3. Other closure mechanisms. Several other cuﬀ closure
mechanisms can be found in literature. Ratchet or zip-tie inspired
mechanisms presented by Yu et al. (2014) and Cobo et al. (2019)
involve microfabricated teeth on the edge of a tab which is passed
around the nerve and through a slit, the edges of which lock the teeth
into place. The tooth spacing oﬀers a set of discrete, though ﬁxed, sizes
from which a precise ﬁt can be selected. Chronic experiments have notyet shown if the longitudinal slit in these designs may compromise theintegrity of the cuﬀ seal.
Another interesting closure mechanism is the hook-and-loop fastener proposed by Seki et al. (2017). Made from Parylene, one end of
this cuﬀ is patterned with three-dimensional microscale loops thatproject from the surface, while the other is patterned with hooks. After
wrapping the device around the nerve, the two ends are pressed together and the arrays of hooks and loops become entangled. It is unclear how the design will fare in chronic studies as the cuﬀ does not seal
and the microstructures may irritate the nerve.
3.1.1.4. Cuﬀ length. To optimize a tripolar cuﬀ for recording purposes,
its length would ideally match the wavelength of action potentials in
the ﬁbers of interest. For a medium-sized myelinated ﬁber of 10 μm
diameter, this can be estimated to be about 60 mm, and for the largest
ﬁbers it would be twice that. This excessive cuﬀ length is not practical,
however making the cuﬀ as long as possible given anatomical andsurgical constraints can aid in maximizing the recorded signal (Marksand Loeb, 1976; Rahal et al., 2000a).
Andreasen and Struijk (2002) estimate that single-ﬁber action potential amplitude begins to saturate when about 22 nodes of Ranvier are
included within the ends of the tripole. Internodal spacing is 100 times
the ﬁber diameter (Hursh, 1939), so the tripole length to maximallyrecord from a 10 μm ﬁber is about 22 mm. Short of maximal recordings,
Tyler (2018) gives the rule of thumb that the tripole should cover at
least 10 nodes of Ranvier, which, by the same calculation, would yield
about 10 mm. In other words, a minimum cuﬀ length for recording
might be estimated by simply transliterating the targeted ﬁber diameterfrom μm to mm and adding a few mm to account for the non-linearities
at the ends of the cuﬀ, as discussed in Section 3.1.1.1.
Hoﬀer and Kallesøe (2001) state that adequate recording during
11

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

consists of a silicone substrate formed into an open-matrix helix withplatinum foil electrodes. Like the spiral cuﬀ, helical electrodes oﬀer anadaptive ﬁt to the nerve, accommodating changes in diameter. They
have been widely successful for clinical non-selective stimulation of thevagus nerve, however the open-matrix form is not suited for sensitive
ENG recording. Multiple design iterations have included a bidirectional
helix with separate sections turning in opposite directions to prevent
travel along the nerve, and “ﬁnger-like” gripping forms that resemble
the open matrix of the helix but allow for easier implantation by obviating the need to completely wrap behind the nerve (Baker, 1993;
Bullara, 1990; Rise and Klepinski, 1992; Weinberg, 1993).
At least one thin-ﬁlm polyimide version of a helical electrode wasproposed, though it has not been speciﬁcally compared with the traditional silicone helix (Xiang et al., 2016).

behavioural tasks in animals is achieved using a cuﬀ length that is 10
times its diameter. However, longer cuﬀs yield better recordings. Ifeven a few mm can be added, Andreasen and Struijk (2002) state thatthe extra length may have a signiﬁcant impact on the RMS value of the
ENG.
For the purposes of stimulation, cuﬀ length is much less importantand has not received special treatment in literature. A longer cuﬀ will
improve eﬃciency and current containment, and otherwise it is thedesired electrode arrangements and ﬁeld shaping capabilities that will
dictate length (Hoﬀer and Kallesøe, 2001). For stimulation with a tripolar conﬁguration, length comparable to a short recording cuﬀ, asdescribed above, should be suﬃcient.
3.1.1.5. Cuﬀs for small nerves. Sizing extraneural cuﬀs for use on small,fragile nerves 500 μm in diameter and smaller introduces additional
challenges. Cuﬀs having thin walls and constructed from soft,compliant materials are necessary. Soft, thin-ﬁlm polymer substrates
may oﬀer signiﬁcant advantages in this case. A 30 μm-thick cuﬀ was
microfabricated from shape memory polymer and used to acutely
record from the pelvic nerve of rat, which has a diameter of 200 μm(González-González et al., 2018). The shape memory polymer can forma very thin cuﬀ wall that also softens upon implantation, in contrast toother commonly used materials like silicone, polyimide, and Parylenethat do not have this combined feature set.
Smaller nerves require cuﬀs capable of tighter bending radius. The
metal conductors used in larger cuﬀs may not accommodate small
bending radii without signiﬁcant stress. One strategy to address this isuse of microfabrication methods to pattern thin metal layers (approximately 200 nm thick) as opposed to methods that involve metal foils(25 μm thick). Ordonez et al. decreased the stress in a circumneural
metal electrode for a 350 μm diameter cuﬀ by longitudinally dividing
the electrode into two sections and reconnecting them with a serpentine
metal trace (2014).

3.1.4. Flat interface neural electrode (FINE)
The ﬂat interface neural electrode (FINE), introduced by Tyler andDurand (1997a), oﬀers a powerful strategy for increasing extraneuralaccess to individual fascicles in a nerve trunk. The concept is that thenerve will tolerate atraumatic rearrangement of the fascicles underlimited transverse compression. The FINE aims to gently ﬂatten thenerve, thereby bringing all fascicles closer to the nerve’s surface andincreasing the cross-sectional perimeter while maintaining cross-sectional area. In addition to spreading out the fascicles to improve accessto them, the increased perimeter allows for a greater number of electrode sites to contact the nerve (Tyler, 2018).
Individual fascicle selectivity and independent addressability of the
FINE on cat sciatic nerve was demonstrated in acute studies (Tyler andDurand, 2002). A study of the four-week response to varying degrees ofreshaping forces (via varying wall thickness) on rat sciatic nerve foundthat the low forces exerted by a 0.4 mm silicone wall were best-tolerated and still capable of achieving the desired reshaping (Tyler andDurand, 2003). Three-month studies of FINE on cat sciatic nerve demonstrated stable selectivity and no signiﬁcant histological evidence ofnerve damage in FINEs with low or medium degrees of reshaping(Leventhal et al., 2006; Leventhal and Durand, 2004). Modelling by
Leventhal and Durand (2003) suggests that selectivity is maximized bydesigning FINEs to particularly match the dimensions of the targeted
nerve, and that doing so may yield sub-fascicle selectivity.
Later, a FINE was designed, modelled, and intraoperatively tested
on human femoral nerve with the aim of eventual use in a neuroprosthesis for standing and walking. At least four of the six targeted leg
muscles were selectively and independently activated in each of theseven subjects tested, and all six muscles in one subject (Schiefer et al.,2010, 2008). Similar studies were performed on other lower-limb
nerves (Schiefer et al., 2013, 2012). Stable and selective sensory percepts were stimulated in upper arm amputees for up to two years using
FINE (Tan et al., 2015).
Several variations of the initial FINE have been produced. A liquid
crystal polymer version was designed to take advantage of microfabrication (Hess et al., 2007). A FINE was designed to close more
slowly for more gradual reshaping of the nerve by integration of abiodegradable polymer, allowing a tuneable reshaping period ranging
from 12 h to 16 days (Caparso et al., 2009). The latest generation of
FINE is designed with regionally speciﬁc stiﬀness such that the ﬂattening eﬀect is still possible but with greater ﬂexibility in the longitudinal direction to accommodate bending of the nerve. Stiﬀness was
modulated by a strip of PEEK embedded in the center of the silicone
wall (Freeberg et al., 2017). This composite-FINE was implanted inhuman trans-tibial amputees and stably elicited multiple sensory percepts in the missing limb over a period of seven months (Charkhkar
et al., 2018).

3.1.2. Book
Book electrodes, introduced by Brindley (1972), consist of deep
grooves in a silicone block with platinum foil electrodes in each groove,
where nerves are to be placed (Brindley, 1977, 1972). The ﬂaps separating each groove are reminiscent of the pages of a book, thus thename. After placing the nerves, a silicone cover is placed and sealedwith silicone adhesive. This device was designed for implantation on
the sacral nerve roots near the spine for bladder control in spinal cord
injury patients. Book electrodes were ﬁrst commercially introduced in1982 by Finetech Medical and have been implanted in thousands ofpatients (About us – Finetech Medical, 2019). The manufacturer statesthat longevity of the device is expected to be 20–25 years, limited bydegradation of the silicone rubber, though it has not been observed(Finetech-Brindley Sacral Anterior Root Stimulator (CPC2): Notes forSurgeons and Physicians, 2016). In a 2018 survey of 33 patients who
were implanted with the Brindley stimulator at one hospital, theaverage time since implant was 16 years and overall satisfaction withthe device was 87.5 % (Deberge et al., 2018). A large advantage of thedevice is its stable implant location, which avoids complications of therelatively bulky geometry. Owing to the selectivity problem, installation of the device unfortunately includes cutting some of the aﬀerent
dorsal sacral roots so that reﬂex circuits do not interrupt function,
which also eliminates other reﬂexes such as gastric emptying and sexualfunctions (Navarro et al., 2005).
3.1.3. Helical
The helical or helicoidal PNI, shown in Fig. 8c, is another extraneural design that has been successfully commercialized. It was originally developed at Huntington Medical Research Institute by Agnew
and co-workers in the 1980s, and nowadays is found in stimulation
systems by LivaNova (previously Cyberonics) (Agnew et al., 1989;
Bullara, 1986; Naples et al., 1990; VNS Therapy, 2019). The design

3.1.5. Other extraneural PNIs
Other notable extraneural PNI designs are introduced here. A microscale, printable “nanoclip” was designed for nerves as small as 50 μm
12Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

more pronounced for electrodes placed directly adjacent to the perineurium (Tyler et al., 2011). Intrafascicular electrodes discussed in thenext section are situated within a fascicle. When comparing their stimulation windows, intrafascicular electrodes have a narrower windowof stimulation amplitudes that achieve 100 % activation before spillingover into other fascicles.
According to Sunderland (1978), disruption or removal of the epineurium, as in the medical removal of neural scar tissue known as
neurolysis, has no harmful eﬀects as long as the perineurium is notdamaged. Unfortunately, there are not many examples of interfascicular
PNIs in literature from which to learn. One example is the multigroove
interfascicular PNI, proposed by Koole et al. (1997). This PNI takes
inspiration from Brindley’s book electrodes, except instead of placing
entire nerves between the pages of a “book,” the epineurium is surgically dissected and each fascicle is placed in its own groove. It is notclear whether in vivo studies were ever performed with this device.
Another example is the slowly-penetrating interfascicular nerve
electrode (SPINE) (Tyler and Durand, 1997b). This device consists of asilicone cuﬀ that has four partial longitudinal slits at one end, producing “beams” around the circumference which project ﬁn-like elements
toward the center of the nerve. As the name suggests, these elements
are intended to slowly penetrate into the epineurium, allowing gradual
rearrangement of the nerve trunk’s organization as the electrodes settle
into position between the fascicles. 24 -h implantations on cat sciatic
nerve yielded promising results, demonstrating improved selectivity ascompared to extraneural cuﬀs and no acute injury to the perineurium(Tyler and Durand, 1997b). Nielsen et al. (2014, 2012) introduced aroughly-constructed ﬁn-like device with two electrodes on each side,
led by a suture to be placed in between the tibial and peroneal fascicles
of sciatic nerve in rabbits. Although acute results were promising forboth interfascicular PNIs, no follow-up chronic experiments have beenreported.
A device intended for intrafascicular interfacing was determinedfollowing histology to have instead an interfascicular interface withelectrodes positioned between fascicles. Even so, the device could selectively stimulate ﬁve muscles (Kundu et al., 2014).
The few examples of interfascicular PNIs in literature suggest thatthe primary design concerns are the placement of electrodes among
fascicles of irregular shape and size, electrical isolation between the
fascicles, and the severity of host response under chronic use.

(Lissandrello et al., 2017). The clip itself does not oﬀer insulation but
serves to hold the electrodes near the nerve. It features a “trap-door”
closure for ease of installation. Non-active clips were installed on zebra
ﬁnch tracheal syringeal nerve for 74 days and changes in bird song
were monitored as an indicator of nerve health. Bird song deviated similarly to a nerve crush injury after implantation, but recovered tobaseline after about 10–20 days while the clip was still present. Whilefurther characterization and optimization of this design are likely
needed, the 200 nm-resolution 3D printing technique may provide a
simple solution for interfacing with small nerves.
The split-ring polyimide PNI proposed by Lee et al. (2017) prioritizes ease of fabrication and installation. No subsequent fabrication
steps are required after removing the microfabricated device from its
carrier wafer, and the ﬂexible split ring shape is intended to easily slip
around the nerve and apply minimal pressure while keeping the electrodes in contact with the nerve. A drawback of this design is thetransverse positioning of electrodes around the perimeter which is not
optimal for either recording or stimulation. Furthermore, the sharpedges of the thin ﬁlm are oriented normal to the nerve and the adjacenttissues and are likely to cause injury when implanted chronically.
A paperclip-inspired polyimide PNI is similarly ﬂat and requires
little manipulation after microfabrication (Lee et al., 2018a,b). This
design is intended for use on small nerves, as it was implanted on rat
pelvic nerves of 250−300 μm diameter and used to initiate bladdervoiding in acute experiments. The ﬁxture mechanism depends on applying constant pressure to slightly kink the nerve, and the long-term
safety of this remains to be seen in chronic experiments.
Ong et al. (2018) presented a sticky hydrogel-based PNI with mechanical compliance more similar to nerve tissue than most PNI materials. Embedded in the hydrogel substrate, platinum electrodes are
insulated by Parylene, and SU-8 structures provide a gradual change inmechanical stiﬀness between the connection end and the interface end.
Another intriguing hydrogel-based PNI presented by Liu et al.
(2019) uses micropatterned electrically-conductive hydrogel (ECH) inplace of the metal electrodes and traces, insulated by a layer ofstretchable insulative polymer and embedded in a hydrogel sheet. The
ECH possesses excellent electrical characteristics and showed minimal
host response when implanted around sciatic nerves of mice for six
weeks. While the mechanical characteristics of hydrogels are near optimal for PNIs, a hydrogel PNI in a cuﬀ-like form will not provide thesame noise rejection and current containment as a cuﬀ made fromtraditional insulative materials, and so a method to combine the meritsof both material types remains to be explored.

3.3. Intrafascicular
Despite the unavoidable damage to the nerve, much research has
explored the use of intrafascicular PNIs (Fig. 9) which beneﬁt from theinsulating properties of the perineurium and may oﬀer more eﬃcient
stimulation, sub-fascicular selectivity, and higher SNR and signal discrimination than other PNIs (Jung et al., 2018a,b).
The primary concern with intrafascicular PNIs is risk of damage tothe nerve ﬁbers and intraneural blood vessels and thus poor chronic

3.2. InterfascicularInterfascicular PNIs penetrate into the epineurium, however they do
not breach the blood-nerve barrier. Modelling has shown that electrodes placed closer to fascicles may preferentially activate nerve ﬁbers
within the fascicle but not other ﬁbers in the nerve trunk; this eﬀect is

Fig. 9. Intrafascicular PNIs. A) Structural diagram of the poly-LIFE (not to scale). Reprinted from (Lawrence et al., 2003) with permission from Elsevier. B) Transverse
intrafascicular multichannel electrode (TIME) implanted in a rat sciatic nerve. Scale bar = 1 mm. Reprinted from (Boretius et al., 2010) with permission from
Elsevier. C) Right: Utah slant electrode array (USEA). Left: high-deﬁnition Utah slanted electrode array (hd-USEA). Reprinted from (Wark et al., 2013). Scale bar =3mm.
13

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

sheaths, leads, and connectors to facilitate handling and implantation ofmultiple LIFEs, which they call a distributed intrafascicular multielectrode (DIME). LIFEs and other intrafascicular PNIs must address the
problem of interfering EMG signals (Mortimer et al., 1995; Yoshida
et al., 2010). Some groups have placed Faraday cages around thenerves, either in the form of a carbon ﬁber mesh, or a metal-and-insulator cuﬀ (Djilas et al., 2007; Mortimer et al., 1995).

performance. The host response to an object in the sensitive endoneurial space may be especially consequential as there is no barrier
to safeguard the nerve ﬁbers. Expected reactions are endoneurial edema
and ﬁbrosis, potentially leading to increased endoneurial pressure,compression of nerve ﬁbers, and consequent neural damage including
demyelination, decreased axon size, and degeneration of the ﬁbers’
distal portions (known as Wallerian degeneration) (Grill et al., 2009;
Rydevik et al., 1990).
Despite these concerns, a variety of studies have shown fairly stable
responses to intrafascicular electrodes, as reviewed by Yoshida et al.
(2010) and Jung et al. (2018a,b). After a recovery period from theimplantation (about one to four weeks), any functional deﬁcits areoften subtle and morphological changes in the nerve are limited, besides the usual signiﬁcant ﬁbrotic capsule surrounding the device. The
intrafascicular PNIs that fare best tend to minimize the encapsulationand potential for re-injury by prioritizing small size and ﬂexibility.

3.3.2. Transverse intrafascicular multichannel electrode (TIME)
The transverse intrafascicular multichannel electrode (TIME) takes
a form similar to the tf-LIFE, except it is implanted perpendicularly
through one or more fascicles so that multiple active sites gain access toa greater number of nerve ﬁbers (Boretius et al., 2012, 2010). A 90degree bend is positioned at the point where the TIME enters the nerve
so that the insulated traces leading to the connector run longitudinally(Badia et al., 2011b).
Following two-month implantation through the three main fasciclesof rat sciatic nerves, slight electrophysiological and functional deﬁcits
at one week, but recovery by one and two months were observed. There
was also 30–60 μm thick encapsulation and an 8–25 % decrease in
myelinated ﬁbers of the smaller fascicles. (Badia et al., 2011b). Though
limited to monopolar electrode conﬁgurations by having all contacts at
one longitudinal position, good selectivity in stimulation and recording
were demonstrated in acute experiments in rats (Badia et al., 2016,2011a).
TIMEs have also been implanted into human amputees. Ranging
from 4 to 12 months post-implantation, studies have demonstrated
stimulation of sensory percepts, usually perceived as tingling, electricity, or vibration in a small region of the phantom limb (Rognini
et al., 2018; Valle et al., 2018b, 2018a). Another study implanted the
TIME for one month in the median nerve and, using an artiﬁcial ﬁngertip with integrated microelectromechanical systems-based “tactile”
sensors for mechano-transduction, the subject was able to distinguishbetween textures with 96 % accuracy (Oddo et al., 2016).
TIME is inserted through multiple fascicles, which anchors them
together and disallows relative motion between them. This may explain
the higher loss of myelinated ﬁbers observed in the smaller fascicles, as
the larger fascicle has an increased “hold” on the TIME due to increased
area of interaction and the smaller fascicles must therefore endure moretrauma to accommodate motion of the device. Also, the size of thedevice and its encapsulation are proportionally larger in the smallerfascicles, which may lead to greater impact on the nerve ﬁbers there(Badia et al., 2011b). Another concern is that slight repositioning of the
TIME by approximately 0.1 mm was found to signiﬁcantly change its
selectivity and its population of interfaced nerve ﬁbers, which may beseen as an advantage for versatility but also a concern for the stabilityof input-output characteristics over the long term (Badia et al., 2016,2011a).
A variation on the TIME design is the self-opening intrafascicularneural interface (SELINE) (Cutrone et al., 2015, 2013, 2011). Upon
insertion into the nerve, SELINE is pulled back a short distance, whichcauses a set of “wings” on each side to spread from the plane of thedevice, helping to anchor it in place. Each of the four wings has a pair ofelectrodes on it, so another beneﬁt is expanded access to reach additional nerve ﬁbers which are not near the original insertion track.
Three-month implantations in rats demonstrated selectivity and nerve
health with the SELINE and also enhanced stability over the TIME
(Cutrone et al., 2015). A thorough six-month characterization of the
SELINE in rats found increasing stimulation thresholds and impedances
during the ﬁrst month corresponding to formation of a signiﬁcant encapsulation layer, followed by stable thresholds and impedances overthe ﬁnal ﬁve months (Wurth et al., 2017). 60 % of the total channels
were functional at three months post-implantation and 25 % at sixmonths, though the cause of lost channels was not speciﬁed. At six
months, cross-sectional area of the nerve trunk near the implant wasapproximately doubled due to the ﬁbrotic capsule. Despite the3.3.1. Longitudinal intrafascicular electrode (LIFE)
The longitudinal intrafascicular electrode (LIFE) is thread-like andimplanted parallel to the nerve ﬁbers. Installation is accomplished byattachment to an electro-sharpened tungsten needle which, after careful
dissection of the epineurium, pierces into the perineurium, exits about 1
cm away, and is then pulled to position the electrode site inside thefascicle (Mortimer et al., 1995).
The ﬁrst LIFE consisted of a 25 μm diameter Pt-Ir wire insulated by
Teﬂon, with about 1 mm of insulation removed at its midpoint to form
the electrode site (Malagodi et al., 1989). The length of the site wasintended to ensure positioning near a node of Ranvier. A bipolar version
used a twisted pair of 5 μm diameter carbon ﬁbers instead of the Pt-Ir
wire (Veltink et al., 1989). In another iteration, the polyLIFE utilized a
12 μm diameter metallized Kevlar thread and silicone insulation for
greater ﬂexibility (Lawrence et al., 2003; McNaughton and Horch,
1996). An 11 μm-thick thin-ﬁlm version (tf-LIFE) was microfabricatedfrom polyimide, enhancing precision, ﬂexibility, and number of electrode sites (Lago et al., 2007b; Yoshida et al., 2000, 2006). The tf-LIFE
is folded in half to position electrodes on opposite sides and also toprovide a loop that can be led by suture into the fascicle. A further
enhancement of the tf-LIFE was the integration of a shape-memory
alloy that could be actuated in situ to achieve ﬁner positioning andcounteract positional drift (Bossi et al., 2007).
Early experiments demonstrated chronic stability of LIFE over six
months in cats and the ability to record distinct signals from about 10
single ﬁbers at a time, though the Pt-Ir wire was prone to drift within
the fascicle and the population of ﬁbers gradually changed (Goodall
et al., 1991). Histology from a similar six-month experiment foundsigniﬁcant encapsulation and a 40 % decrease in axon diameters at the
implant site, but sections about 1 cm distal and proximal from the site
showed no apparent changes (Lefurge et al., 1991). Implantation of themore ﬂexible polyLIFE in rabbits for six months still showed encapsulation but no signiﬁcant eﬀects on axon count or diameter(Lawrence et al., 2002). A three-month evaluation of tf-LIFE in rats
revealed a slight functional decline after one month which recovered at
the two and three-month points and was attributed to recovery after the
implantation procedure. Histology in this case showed encapsulation
but no signiﬁcant changes in axon count or diameter (Lago et al.,2007b; Navarro et al., 2007). In a two-week implantation in humanamputees, polyLIFEs were able to stimulate sensory percepts and record
volitional motor signals (Dhillon et al., 2005). In another human study,
tf-LIFEs recorded output signals for four weeks and stimulated sensory
percepts for 10 days, though stimulation ability decreased after that
(Rossini et al., 2010).
A signiﬁcant limitation of the LIFE design is low throughput interms of the number of ﬁbers accessed per implantation. The implantation process is relatively tedious and time consuming, and if
multiple degrees of freedom are desired, multiple devices will need to
be implanted. Thota et al. (2015) proposed a supporting system of14

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

activation of inﬂammatory cells at the implant site and a downward
shift in distribution of ﬁber diameters.
Early on, an array similar to the UEA was implanted in the median
nerve of a healthy human volunteer for 96 days (Gasson et al., 2004).
Anecdotal stimulation and recordings were broadcast to and from arobotic hand, though only in a non-speciﬁc threshold-detection scheme.
The subject experienced no deﬁcits in sensation or motor control while
the array was implanted, nor up to nine months after removal. Morerecently, USEAs were implanted in median and ulnar nerves of two
human amputees for 30 days and achieved promising results (Daviset al., 2016). Recordings of volitional motor signals from the USEAs
allowed the subjects to control a virtual robotic hand, performing 15
diﬀerent movements including proportional control of individual ﬁngers. In one of the subjects, stimulation via various combinations ofelectrodes produced more than 80 diﬀerent sensations in the phantom
limb. SNR and stimulation thresholds did not signiﬁcantly change
throughout the implantation period. In another study, USEAs were
implanted in the median and ulnar nerves of an amputee for 14 months(Page et al., 2018). They were used for stimulation of sensory percepts
from a sensor-equipped prosthetic hand in combination with intramuscular recording electrodes for motor control. This study quantiﬁed an increase of prosthesis embodiment and a decrease in phantom
limb pain arising from integration with the nervous system.
Christensen et al. (2016) performed a study in which USEAs were
implanted in two human amputees for 30 days and subsequent histology of the nerve was performed. While electrophysiological tests had
demonstrated success in both recording multiple units and stimulating
sensory percepts, histology revealed that the electrode tips had only
slightly penetrated into the outermost fascicles in the best cases, and inother cases had not penetrated the perineurium at all, as shown in
Fig. 11. Still, in the penetrated cases, axons and myelin were found inclose proximity to the electrode. Activated macrophages were present
on the explanted device and at the implant site, though the administration of medication to the subjects likely aﬀected the inﬂammatory
response.
As the evidence from multiple studies suggests, the densely spaced
silicon shanks have diﬃculty penetrating the tough perineurium.
Attempting insertion by simply pressing the array into the nerve islikely result in a crush injury before penetration into the endoneurium.
Therefore, implantation is assisted by a pneumatically powered toolthat rapidly accelerates the array into the tissue (Branner and Normann,
2000). As illustrated by the examples above, this technique is not always eﬀective. A study designed to assess the safety of pneumatic implantation of the hd-USEA in rat sciatic nerve found the amplitude ofsignals propagating across the implantation site before and after implantation dropped by 13–38% on average. Furthermore, signal propagation across the site ceased completely in 1 of 8 implantations(Mathews et al., 2014a).
After implantation, Utah arrays intended for chronic use are enclosed with the nerve inside of a silicone cuﬀ or other wrap to hold it inplace. This may be an additional source of nerve injury to consider, as
discussed in Section 3.1.1. If the PNI is implanted near active muscles,
EMG will be a signiﬁcant problem for recording, and so a cuﬀ/wrap
may also serve a shielding role (Branner et al., 2004). Recent implantations in human amputees with favorable results report wrapping
the nerve and array in a sheet of manufactured collagen or other decellularized biological material (Christensen et al., 2016; Davis et al.,2016; Page et al., 2018).
The literature on rigid penetrating arrays such as the Utah arrayseems to suggest use in sites that experience little motion, have minimal
connective tissue, and have less fascicular division. The nerve roots
match this description, and, indeed, several studies have shown rigid
penetrating arrays to be highly eﬀective when implanted there, thoughchronic stability is yet to be investigated (Aoyagi et al., 2003; Gaunt
et al., 2009; Holinski et al., 2013; Stein et al., 2004; Weber et al., 2007).

Fig. 10. Cross-section of the implant site at one shank of the Utah electrodearray. Fibers were compressed under the tip and sheared adjacent to the insertion track. Reprinted from (Branner and Normann, 2000) with permission
from Elsevier. Scale bar = 100 μm.

signiﬁcant encapsulation, selectivity over the experiment duration remained stable, as did the number of axons distal and proximal to theimplant.
3.3.3. Utah slanted electrode array (USEA)The Utah electrode array (UEA) was originally designed as an intracortical neural interface (Nordhausen et al., 1996). Its use in peripheral nerves was ﬁrst explored by Branner and Normann (2000), whodemonstrated its ability to record single-unit responses from mechanoreceptors and eﬃciently evoke twitches in individual digits during 36h studies in felines. Histology of the implantation site showed a dramatic “compression wave” surrounding each shank, consisting of axonal cross-sections which were compressed under the tip and sheared
adjacent to the insertion track, as shown in Fig. 10.
An adaptation of the intracortical UEA better suited to peripheral
nerves is the Utah slanted electrode array (USEA) which has variedshank lengths so that a greater number of nerve ﬁbers may be accessed,
rather than only those that lie in a single plane (Branner et al., 2001).
The shanks are spaced 400 μm apart and range from 0.5 to 1.5 mm inlength. Acute implantations of USEA in cats demonstrated four times
broader recruitment curves and one tenth the required stimulus current
as compared with extraneural cuﬀs (Branner et al., 2001).
A high-density version of the USEA (hd-USEA) was introduced by
Wark et al. (2013) with a shank spacing of 200 μm, yielding a density of
25 electrodes/mm2 (compared with 6.25 electrodes/mm2 for the
USEA). Shank lengths of the hd-USEA range from 300 to 800 μm. Acuteexperiments in rats and felines demonstrated single unit recording,selective stimulation of motor units, and histological conﬁrmation ofelectrode placement in the fascicles (Wark et al., 2013). In anotheracute study, the hd-USEA was implanted in the feline pudendal nerve
and was able to selectively record and distinguish among neural signalsthat corresponded to diﬀerent genitourinary functions including
bladder ﬁlling and distension-evoked contractions (Mathews et al.,2014b).
In contrast to the successful intrafascicular array demonstrations inacute studies, chronic experiments have yielded mixed results. USEAs
implanted in felines for up to seven months resulted in limited functional deﬁcits which recovered in most cases, inconsistent stimulation
capabilities which seemed to depend on surgical technique, and poor
recording capabilities which completely receded by one month(Branner et al., 2004). In some cases, histology showed that the shankseither did not penetrate the perineurium or were pushed out by connective tissue. In another study, USEAs were implanted in feline sciatic
nerves for up to one year to study the foreign body reaction(Christensen et al., 2014). The reaction was poor, including persistent15

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 11. Cross-sections of human nerve fascicles under the USEA implant at the locations of two rows of diﬀerent-length shanks. It can be seen that neither row ofshanks penetrated into the fascicle (red). Reprinted from (Christensen et al., 2016) with permission from Elsevier. Scale bar =500 μm.

sieve, usually supported by a section of tubing, so that the axons growthrough the holes to reconnect the two ends. Mannard et al., in 1974,
fabricated a sieve electrode by drilling through silver wires embedded
in an epoxy wafer, and successfully recorded sensory and motor signals
in amphibians from 12 to 25 weeks after implantation (Mannard et al.,1974). Most sieve designs over the next few decades were made fromsilicon to take advantage of microfabrication techniques (Akin et al.,
1994; Bradley et al., 1997; Della Santina et al., 1997; Edell, 1986;
Kovacs et al., 1992; Mensinger et al., 2000), and since the late 1990s,
several polyimide-based designs have been introduced (Jeong et al.,2016; Park et al., 2018; Stieglitz et al., 2002, 1997). In 1997, Bradley
et al. (1997) brieﬂy summarized the short history of sieve electrodes up
to that time, stating that many had been designed but not publishedexcept in abstract form because chronic recordings were rarely successful. In one successful example, Bradley et al. (1997) were able torecord spontaneous and mechanically evoked signals in rat with a silicon sieve electrode from week 7 post-implant up to week 20.
Ramachandran et al. (2006) demonstrated axon conduction through apolyimide sieve by proximal nerve stimulation and measurement of
resulting EMG distal to the sieve, and also captured recordings of mechanically and electrically stimulated aﬀerent signals, though not reliably.
A few design considerations for sieve electrodes include “porosity”
or “transparency,” referring to the ratio of the summed hole areas to thetotal substrate area in the path of the nerve (Jeong et al., 2016; Stieglitz
et al., 2002). Higher porosity generally achieves greater axon growth
but has adverse implications for mechanical strength. Hole size is also a
trade-oﬀ. Spearman et al. (2018) state that there tends to be a sharp
tipping point between holes that are too small (discouraging axon
growth and constricting those that do grow), and too large (allowing
too many axons to grow through a single hole and thereby diminishingselectivity). The hole size recommended in the literature is 40–65 μm(Lago et al., 2007a; Navarro et al., 2005). Some notable design enhancements include coating the sieve with bioactive proteins to

3.4. Regenerative
Regenerative PNIs aim to take advantage of the long-known abilityof peripheral nerves to regenerate and reconnect after transection(Guth, 1956). Instead of inserting electrodes into the nerve, the nerve
ﬁbers can integrate with electrodes in the PNI. If successful, regenerative PNIs could potentially oﬀer far greater resolution and stability than any other of the PNI types discussed thus far. However, thedrawback of the approach is the requirement for nerve transection sothat the interface may be inserted between the proximal and distal
portions. Because of this, any applications that require the interfaced
nerve to remain intact, such as bioelectronic medicine, are excluded.
This approach may be best suited to applications where distal innervation is unnecessary or not crucial, as in neural prosthesis integration with amputated limbs.
A number of diﬃcult challenges face the implementation of regenerative PNIs. Foremost is the formation of healthy, mature nerve
ﬁbers, suﬃciently reconnected after transection through and notaround the interface. According to Sunderland (1990), axons have noparticular compulsion to reconnect with the original distal axon, oreven to grow into the same fascicle; they readily grow into other fascicles, the interfascicular space, or even out of the nerve. Furthermore,
the foreign body response to the implanted device may interfere byblocking the path with ﬁbrotic tissue before axons can grow through it.
This is especially the case for mammals, in which ﬁbrosis is more extensive as compared with amphibians or ﬁsh (Stein et al., 1975). Another challenge is interconnection when a large number of electrodesites are desired. Several regenerative PNIs were developed to addressthese challenges.
3.4.1. Sieve
Sieve electrodes (Fig. 12) consist of a planar structure with a dense
pattern of holes through it, some or all of which are ringed by a metalelectrode site. Each end of the transected nerve is positioned against the16

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 12. Sieve electrodes. A) Diagram showing arrangement of transected nerve stumps, guide tube, and sieve. B) SEM of a silicon sieve. A) and B) reprinted from
(Bradley et al., 1997) with permission from Elsevier. Scale bar = 100 μm. C) Polyimide sieve. Holes are 40 μm in diameter. Reprinted from (Stieglitz et al., 2002)
with permission from Elsevier.

microfabricating a ﬂat, ﬂexible array of open-ended microchannels
with integrated electrodes, comparable to a sheet of corrugated cardboard. This structure is then rolled up, creating a cylinder of multiple
layers of microchannels (Fig. 13b). Virtues of this design include its
amenability to standard microfabrication techniques, simple assembly
into a 3D structure, and natural cylindrical form to match the crosssection of the nerve. A variety of polymers were explored as the substrate for rolled microchannel arrays, including Parylene, polyimide,
silicone, and SU-8 (FitzGerald et al., 2012; Lacour et al., 2008;
Srinivasan et al., 2011; Suzuki et al., 2006).
Similarly, the stacked microchannel array is formed by stacking
multiple ﬂat arrays. This design was proposed by Loeb et al. in 1977
and the concept was recently revisted (Lancashire et al., 2016; Loebet al., 1977; Musick et al., 2015). Advantages of the stacked array are
better control of sealing separate layers of microchannels and greater
amenability to dense electrical connections, though the overall shape isblock-like and potentially incongruous with the nerve and the nerve
groove.
Several regenerative microchannel PNIs achieved promising in vivo
results. One study found axon growth into more than 95 % of a device’s
microchannels and recording of spontaneous and evoked single- andmultiple-unit action potentials from anesthetized rats (Srinivasan et al.,2015). Other studies achieved similar recording success in ambulatory
rats (Gore et al., 2015; Musick et al., 2015; Srinivasan et al., 2016).
As with sieve electrodes, open area ratio and channel size are key
design considerations. In a study of in vitro neuron cultures, Lacour
et al. (2008) found optimal growth of axons into microchannels whentransparency was at least 50 %. FitzGerald et al. (2012) found greateraxon growth in vivo with 62 % transparency as opposed to 21 %, though
the two devices compared had other diﬀerences. Smaller microchannels

encourage growth (Mensinger et al., 2000), and a multi-layered designthat helps address the trace-routing problem of many electrode sites ona planar structure (Jeong et al., 2016).
3.4.2. Microchannel
The microchannel electrode array, or tubular electrode array, developed alongside the sieve electrode. The design may be considered asa sieve that is extruded in the longitudinal direction, so that, instead ofgrowing through holes in a plane, the axons grow through tubes in a
volume. The microchannel electrode array has advantages akin to thoseof an extraneural cuﬀ except applied to multiple smaller groups of ﬁbers. These include conﬁnement and ampliﬁcation of the extracellularsignals, and ability to incorporate multiple electrodes in a single lumen
for noise rejection and selectivity techniques (FitzGerald et al., 2008).
After early demonstrations of nerve regeneration through various
plastic conduits and microscale gold cylinders in the late 1960s, anumber of designs were explored to use electrodes inside of microchannels, though none of the early designs demonstrated successful
regeneration and recording (Loeb et al., 1977; Marks, 1969; Spearman
et al., 2018). In the last two decades, there has been a relative proliferation of regenerative microchannel PNIs.
One method to form conduits is casting of a silicone substratearound a bundle of microwires or ﬁlaments. The microwires are removed after casting, leaving behind an array of densely packed microchannels (Fig. 13a). A limitation is that electrodes are introduced
after casting, obscuring part of the channel; microwires with diameters
smaller than the conduit diameter can be placed in several of thechannels (FitzGerald et al., 2012; Gore et al., 2015; Kim et al., 2015).
Alternatively, a bundle of conduits can be assembled from a planar
structure. The rolled microchannel array is made by ﬁrst

Fig. 13. Microchannel PNIs. A) Microchannel array formed by casting silicone around microwires. Reprinted from (Kim et al., 2015) with permission from Springer.
B) Rolled microchannel array made of PDMS and SU-8. Reprinted from (Srinivasan et al., 2015) with permission from Elsevier.
17

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

may increase selectivity by isolating groups of fewer nerve ﬁbers,
however the channel must also accommodate vasculature and othersupportive tissues if the regenerated ﬁbers are to be healthy and robust.
Several studies reported growth of “mini-fascicles” into channels whosecross sections were 75–200 μm in diameter (FitzGerald et al., 2012;
Gore et al., 2015; Srinivasan et al., 2015). These mini-fascicles included
tens of nerve ﬁbers, Schwann cells, a few blood vessels, and an outer
sheath of connective tissue. While far from the ideal isolation of only afew ﬁbers in each microchannel, this still represents a potentiallyhigher electrode density than other PNI types and may be a productive
compromise to achieve healthy integration. Inclusion of selectivityenhancing techniques already used in cuﬀs could further improve thecapabilities of interacting with each smaller group of regenerated nerve
ﬁbers.
The conduits need not be limited to a straight path; bifurcating ordiverging microchannel arrays have also been explored. In this design,
the microchannels bifurcate and grow smaller as they progress distally,
with the purpose of having larger groups of regenerating ﬁbers progressively divide until small numbers of ﬁbers are isolated in eachmicrochannel each with an electrode for increased selectivity(Stoyanova et al., 2013). In vitro and in vivo studies demonstrated feasibility of inducing the growth and separation of nerve ﬁbers into suchmicrochannels (Stoyanova et al., 2013; Wieringa et al., 2010a, 2010b).
Further studies have shown that the bifurcating approach could possibly be used to induce separation of based on ﬁber type (e.g. sensory
and motor) by preferential attraction to diﬀerent neurotrophic factors(Lotﬁ et al., 2011).

weeks (Cho et al., 2008).
Several designs orient thin-ﬁlm planar microfabricated electrodesubstrates parallel to the nerve axis (Clements et al., 2013, 2007;
Delgado-Martínez et al., 2017; Judy et al., 2018; Kuliasha et al., 2018).
Planar electrodes like these are maximally transparent in terms of crosssection, and potentially more scalable than others since they could bestacked to access a greater volume of the nerve (Spearman et al., 2018).
The regenerative scaﬀold electrode (RSE) places a thin-ﬁlm SU-8 electrode array onto a sheet of aligned nanoﬁbers which serve as the
scaﬀold (Clements et al., 2007). The RSE demonstrated recording of
evoked potentials during an 8-week implantation (Clements et al.,2013). A similar design to the RSE is the double-aisle regenerativeelectrode, which instead uses a polyimide substrate with electrodes onboth sides (Delgado-Martínez et al., 2017). This design explores the useof the electrode as a septum to divide the regenerating ﬁbers into two
fascicles. The tissue-engineered electronic nerve interface (TEENI),
shown in Fig. 14b, consists of multiple thread-like arms suspended in afull-volume, carefully-designed, biodegradable composite hydrogel
scaﬀold (Judy et al., 2018; Spearman et al., 2018). A 6-week implantation of the TEENI yielded single-unit recordings with good SNR(Kuliasha et al., 2018).

3.5. Optical
The present discussion of PNIs has focused solely on electricalmeans of stimulation and recording. However, recently-devised optogenetic methods have raised the possibility of interfacing with the nerve
via light. While the majority of optogenetics research has focused on thebrain, several groups have begun to adapt the methods for use in PNIs,
as reviewed by Anderson and Weir (2019). Incorporation of light-sensitive ion channels into the axonal membrane can enable optical activation or inhibition of action potentials. Likewise, incorporation ofvoltage- or calcium-sensitive ﬂuorescing proteins can enable opticalread-out of neural activity. So, to implement an optical PNI, the necessary components are these light-transducing proteins, a method for
introducing the proteins into the targeted nerve ﬁbers, a light source,
and a microscope.
A variety of optogenetic proteins were developed to provide spectral
variety and speciﬁcity and optimize their temporal response (Anderson
and Weir, 2019; Yizhar et al., 2011). Incorporation of these proteins
into target cells is accomplished by deployment of the genetic material(transgenes) for transcription and translation by the cell. Use of transgenic animal models is common for optogenetic research purposes andyields the most stable long-term expression of the transgenes, thoughpractical implementation of optical PNIs will require delivery by othermeans, namely viral or non-viral vectors (Anderson and Weir, 2019;
Yizhar et al., 2011).

3.4.3. Scaﬀold
A variety of synthetic supportive structures have been used to facilitate the repair of injured nerves. Several regenerative PNI designs
position electrodes within these scaﬀolds to reduce distance to the regenerated nerve, as recently reviewed by Spearman et al. (2018). Distinct from the regenerative microchannel arrays described above, thisapproach incorporates techniques from the ﬁeld of tissue engineering,using materials such as hydrogels and collagen to mimic properties of
the nerve’s extracellular matrix. Various biomolecules and growth
factors can also be incorporated to further enhance nerve regeneration.
A design called the regenerative multielectrode interface (REMI)
places an array of microwires, similar in form to the Utah array, projecting transversely into a scaﬀold of collagen inside a section ofpolyurethane tubing, as shown in Fig. 14a (Garde et al., 2009; Seifert
et al., 2012). REMI implantations of up to 120 days were able to record
single units starting at 7 days post-implantation and remained stablethroughout the study duration (Desai et al., 2014). Another design
places individual SU-8 and gold probes longitudinally into a biodegradable agarose scaﬀold and achieved robust recordings for up to 51

Fig. 14. Regenerative scaﬀold-type PNIs. A) Regenerative multielectrode interface (REMI) consisting of a microwire array projecting into a scaﬀold tube to be ﬁlledwith collagen. Scale bar = 2 mm. Reprinted from (Garde et al., 2009). B) Tissue-engineered electronic nerve interface (TEENI) and cross section diagrams showing
the potential for scaling by adding additional arrays. C is a reprint of Fig. 2 in: Kuliasha et al., “Robust and Scalable Tissue-Engineered Electronic Nerve Interfaces(TEENI),” 2018 Hilton Head Workshop on Sensors and Actuators, reprinted with the permission of the author and the Transducer Research Foundation.
18

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

sprouting of the axons and growth into the microﬂuidic channels where
the electrodes are located. Acute in vivo experiments with the LACE
have demonstrated successful lysing of the epineurium, though chronic
experiments are yet to be performed to demonstrate axonal sprouting
into the channels (Larson et al., 2018).
Another proposed regenerative design without nerve transection is adirect adaptation of Kennedy’s neurotrophic cone cortical probe, whichaims to encourage axonal growth into a polyimide cone where several
wires are housed (Jung et al., 2019, 2018a,b; Kennedy, 1989). After 19
weeks of implantation in rat, naturally-evoked potentials were capturedby the electrode, and histology revealed axonal generation and minimal
scar tissue inside the cone (Jung et al., 2019).
Other design enhancements which do not necessarily belong to aparticular PNI category include integration of pH sensing as means offeedback towards closed-loop vagus nerve stimulation (Cork et al.,2018) and glucose sensing as a means of in situ inﬂammation monitoring around the PNI (Lee et al., 2016b).
Drug delivery is a PNI enhancement that can be used for several
purposes and takes on several forms. Multiple studies have incorporated
catheters into cuﬀs in order to modulate nerve activity with pharmacological agents (Hoﬀer and Kallesøe, 2001; Pohlmeyer et al., 2009)and several designs use microfabrication techniques to directly integrate microﬂuidic channels (Cobo et al., 2019; Schuettler et al.,2011). PNI studies have also explored the use of drug-eluting coatings(Heo et al., 2016; Mensinger et al., 2000; Park et al., 2015a,b).
Aside from modulatory and regenerative goals, a common aim ofdrug delivery is mitigation of the host response. An early demonstrationof dexamethasone elution from pacemaker leads showed a thinner ﬁbrotic capsule, fewer activated immune cells, lower stimulationthreshold, and lower interface impedance as compared with controls(Radovsky and Van Vleet, 1989). Following these results, dexamethasone coatings and various vehicles for controlled dexamethasone release from neural probes were investigated, including
polypyrrole, hydrogels, and carbon nanotubes (Abidian and Martin,
2009; Luo et al., 2011; Wadhwa et al., 2006; Zhong and Bellamkonda,
2007). Park et al. (2015a,b) patterned dexamethasone-loaded coatings
onto a polyimide cuﬀ. Heo et al. (2016) developed a polyimide cuﬀ
with surface microwells and a hydrogel coating loaded with cyclosporine A, another anti-inﬂammatory agent. Five-week in vivo assessment of the microwell design showed increased ENG amplitude, decreased ﬁbrotic thickness, and increased myelination as compared with
controls (Kim et al., 2017a, b). Improved host response to PNIs has also
been achieved with systemic delivery of dexamethasone (de la Oliva

While studies in anesthetized animals have successfully demonstrated and reﬁned optical PNI concepts (Fontaine et al., 2017; Kapuret al., 2014; Liske et al., 2013; Sharp and Fromherz, 2011), hardware
implementation of the light source and microscope in awake, freelymoving animals requires further development. For optical stimulation,
extraneural silicone cuﬀs incorporating LEDs (Llewellyn et al., 2010;
Park et al., 2015a,b; Song et al., 2018) and ﬁber optic connections(Michoud et al., 2018; Towne et al., 2013) were demonstrated. Opticalread-out of peripheral nerve activity with an implanted device has not
yet been demonstrated. A multi-modal approach may provide a stopgap
until miniaturized optical PNI read-out methods are developed, such as
the cuﬀ presented by Song et al. (2018) which includes an LED for
optical stimulation and platinum electrodes to record the response.
Other issues remaining to be addressed in optical PNI development
include signal attenuation and distortion by the nerve tissue (Fontaine
et al., 2018; Futia et al., 2018), management of heat generation during
optical stimulation (Park et al., 2015a,b; Yizhar et al., 2011), and immune response to vector delivery (Anderson and Weir, 2019). Thoughmuch development remains, optical PNIs could eventually oﬀer singleﬁber resolution with low invasiveness to the nerve by combining spatial
localization, optogenetic proteins that operate in distinct bandwidths,
and targeted delivery of those proteins to particular sets of ﬁbers
3.6. Hybrid approaches and other enhancements
Several devices combine design concepts and as such do not ﬁtneatly into the above categories. A polyimide-based helical PNI wasdeveloped having probe shanks periodically projecting into the nerve
along its length (Kim et al., 2017a,b; Park et al., 2018). The intraneuralshanks potentially provide a thorough sampling of ﬁbers throughout
the nerve’s cross section while the extraneural helical arrangement
avoids both circumneural constriction and concentrated displacementof the nerve tissue.
A design that combines elements of extraneural cuﬀs and regenerative PNIs is the lyse-and-attract cuﬀ electrode (LACE) shown in
Fig. 15 (Cobo et al., 2019). The LACE is a Parylene-based extraneuralcuﬀ with integrated microﬂuidic channels inside which electrodes areembedded. Like a regenerative PNI, the LACE aims to induce growth ofnerve ﬁbers among its electrodes. However, it aims to do this fromoutside the nerve by inducing collateral sprouting of axons rather thantransection and regeneration. Collagenase is delivered through themicroﬂuidic channels to focally lyse an opening in the collagen-richconnective tissues, followed by a gradient of growth factors to induce

Fig. 15. The lyse-and-attract cuﬀ electrode (LACE) has integrated microﬂuidic channels with embedded electrodes, combining extraneural and regenerative approaches to grow axons near the electrodes without nerve transection.
19

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

sweep (Fig. 16). At any point on the voltammogram, the equivalentcapacitance is given by C = i/(dE/dt) where dE/dt is the scan rate (V/s)(Robblee and Rose, 1990). The negative and positive peaks areassociated with reduction-oxidation reactions on the surface of theelectrode, such as oxide formation and reduction and hydrogen
adsorption/desorption on platinum, as illustrated in Fig. 16. At therelatively ﬂat region (for platinum, approximately –0.2 V versus Ag/
AgCl in PBS), C is the capacitance of the double layer. The CV of apurely capacitive electrode will be smoother, lacking the peaks
associated with redox reactions. The cathodal charge storage capacity
is calculated as the negative area enclosed within the CV “loop” and isoften divided by GSA to focus on the electrode material’s surface
quality. CSCc is roughly interpreted as the amount of charge available
for a stimulation pulse. However, in practice, the equilibrium potentialof the electrode from which a stimulation pulse starts limits the usable
stored charge to a fraction of the total CSCc (Cogan, 2008). Strategies
exist to increase use of the available CSCc, such as modiﬁed waveforms
and positively biasing the electrode prior to pulse delivery (Cogan,
2008; Robblee and Rose, 1990).
4.1.1.1.2. Water window. The potential range swept during CV isdesigned to remain just inside the “water window,” beyond which H2(at negative potentials) and O2 (at positive potentials) gas evolve―thisgas evolution is an irreversible process since the newly formed gas
molecules diﬀuse away from the electrode surface (Hudak et al., 2010;
Robblee and Rose, 1990). Because this diﬀusion prevents the immediate
saturation and nucleation of gas bubbles, visual detection of
microscopic bubble formation is not a reliable technique to estimate
safe operating potentials (Robblee and Rose, 1990). Instead, excursions
beyond the water window during CV may show as sharp increases incurrent at the extremes of the voltammogram (Hudak et al., 2010;
Robblee and Rose, 1990). The water window of platinum and iridium in
1× phosphate-buﬀered saline (PBS, pH ∼ 7.2) is taken to beapproximately –0.6 to 0.8 V versus Ag/AgCl, and in sulfuric acid
(0.05 M H2SO4, pH ∼ 1) it is approximately –0.2 to 1.2 V (Cogan,
2008; Hudak et al., 2010).
4.1.1.1.3. Factors inﬂuencing CV. Factors inﬂuencing the CV include
scan rate, solution composition and pH, temperature, and GSA of theelectrode; any report of CV results should also state these parameters. A
typical scan rate is 100 mV/s. Faster scan rates generally yield higher
currents and may better reﬂect the frequencies involved in stimulation
waveforms, though lower scan rates are often used to determine an
estimate of CSCc (Cogan, 2008; Elgrishi et al., 2018; Robblee and Rose,1990). Two standard solutions used for CV are sulfuric acid (typically
0.05 to 0.15 M) and PBS. Within their respective water windows, thetwo solutions will yield similar voltammograms. However, PBS is a
closer approximation of the in vivo ﬂuid environment and, outside theet al., 2019, 2018b) and anti-TNF-α (Vince et al., 2005).
4. Characterization of peripheral nerve interfaces
Fundamental principles guiding PNI design and a survey of approaches and strategies used in PNI design were presented. Here wecomplement the prior discussion by highlighting concepts, guidelines,
and best practices that emerged from literature and can be employed in
evaluation of a PNI design both in vitro and in vivo. Many of thesetechniques also extend to evaluating certain aspects of other neuralinterface types.
4.1. In vitro characterization
After successful PNI fabrication, a variety of in vitro tests evaluate adevice’s performance-related characteristics, either informing reﬁnement of the design and fabrication methods or proving readiness for invivo studies. The main categories of in vitro evaluations are electrochemical characterization, longevity testing, simulated implantations,and other tests particular to the unique features of a device.
4.1.1. Electrochemical characterization
Electrochemical characterization is performed to evaluate theability of an electrode site to deliver suﬃcient charge for stimulation
and record signals with suﬃciently low noise and estimate safe operating parameters. Common tests performed include cyclic voltammetry(CV), electrochemical impedance spectroscopy (EIS), and pulse testing.
The metrics usually reported from these tests are cathodal charge storage capacity (CSCc), impedance, and reversible charge injection limit.
4.1.1.1. Cyclic voltammetry. CV characterizes the modes and quantities
of charge transfer that occur on the surface of an electrode. A
potentiostat sweeps the potential of an electrode (in solution) througha triangle wave proﬁle between two potentials while it measures thecurrent between the electrode (set as the working electrode, WE) and alarge counter electrode (CE, typically platinum for inertness) (Elgrishi
et al., 2018). A standard, specially-packaged Ag/AgCl electrodetypically serves as the reference electrode (RE), though hydrogenelectrodes and saturated calomel electrodes have also been used
(Robblee and Rose, 1990). The applied potential drives chargetransfer at the WE by mechanisms such as double layer charging andoxidation/reduction (redox) reactions, and the resulting current i is
proportional to the rate at which these mechanisms occur (Cogan,2008).
4.1.1.1.1. Voltammograminterpretation. The
resultingvoltammogram shows current plotted against potential over the entire

Fig. 16. Illustration of a typical cyclic voltammogram of a platinum electrode in PBS. Electrochemical processes corresponding to particular features are labelled.
Sharp increases in current magnitude just outside the water window correspond to gas evolution.
20

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 17. Illustration showing removal of the access voltage (iR error) from pulse testing. A) Voltage measured during delivery of a biphasic current pulse includes
oﬀsets from the access voltage Va. B) Same data with Va removed, giving an accurate measure of the maximum cathodic and anodic potentials (Ec and Ea). Most
commercial electrochemical instruments oﬀer automatic correction of this error.

intact electrodes have relatively high impedance (Straka et al., 2018;
Takmakov et al., 2015). Another confounding factor in interpretation of
EIS is that the reported impedance is contributed by the entire
conductive path including the traces and insulation; compromised
insulation may be diﬃcult to isolate among other factors. Since most
PNIs involve some polymer construction and polymers are nothermetic, EIS contributions arising from the polymer must also be
considered. The impedance and phase data gained from EIS are perhaps
most powerful when used to derive equivalent circuit models, yielding
insight into electrode surface characteristics and insulation quality(Franks et al., 2005; Ortigoza-Diaz et al., 2018; Straka et al., 2018).

water window, shows additional electrochemical activity involving
chloride and phosphate which may be relevant to some electrodes
and stimulation protocols (Hudak et al., 2010). Solution conductivity
and redox reaction rates increase with temperature, so it is ideal toperform CV (and other electrochemical characterizations) at bodytemperature rather than room temperature (Cogan, 2008).
O2 dissolved in solution is not inert and can interfere with the CV, so
standard practice is to prepare the solution by purging with nitrogengas, for example by bubbling pure nitrogen directly into the solution
through a submerged section of polyetheretherketone (PEEK) or polytetraﬂuoroethylene (PTFE) tubing before and during the CV. However,when simulating the in vivo electrochemical environment, it should beconsidered that dissolved oxygen will be present. Hudak et al. (2010)
compared CVs performed in PBS with nitrogen and oxygen purging, thelatter of which yielded a cathodic current due to oxygen reduction that
dominates at potentials below about 0.25 V versus Ag/AgCl, lowering
the entire left half of the CV into the negative current range. It is suggested that the high amount of oxygen reduction in this range may noteasily be reversed and may be a mechanism of stimulation-induced
tissue damage; further investigation is warranted. Another possible
source of distortion in CV is inaccurate potential measurement due to“ohmic drop,” also called iR error, which arises from the solution resistance between the WE and RE. To minimize this eﬀect withoutchanging the solution conductivity, the WE and RE should be positioned close together (Elgrishi et al., 2018). Many commercial electrochemical instruments oﬀer automatic compensation of this error (iRCompensation: Potentiostat Fundamentals / Basics of EIS, 2019).
4.1.1.1.4. Electrode cleaning. Besides characterization, CV is also
used to electrochemically clean electrodes and is a recommended step
before other electrochemical characterizations (Rose and Robblee,1990). During fabrication, by-products of machining processes orother contaminants from handling may deposit on the electrodes,
decreasing the eﬀective surface area. Cleaning is usually performed insulfuric acid. As the cycles progress, cleaning of the surface is evidenced
by increased currents. Depending on the initial electrode cleanliness,
30–50 cycles is usually suﬃcient to reach stabilized currents.

4.1.1.3. Pulse testing. The goal of pulse testing is to directly estimate
the range of parameters for stimulation that can be delivered without
excursion of the electrode potential beyond the water window. Current
pulses are delivered to the WE, submerged in PBS with a RE and large
return electrode, while the resulting potential transients are measured
and compared to the water window. The ohmic drop (sometimes called“access voltage” to include other sources of oﬀset) between the WE andRE can add a signiﬁcant oﬀset to the measured potential duringcathodic and anodic pulses. Since this test relies on accurate
measurement of the electrode potential, simply placing the WE andRE close together will not suﬃce as with CV. If the electrochemical
instrument being used does not oﬀer automatic iR compensation, theoﬀset should be manually subtracted from each phase as demonstrated
by Rose and Robblee (1990) and illustrated in Fig. 17. As noted by
Cogan (2008), the value of this oﬀset may be made clearer by insertinga brief interphase period during which i = 0; the nearly instantaneouschange in potential at the start of this period is the oﬀset to be
subtracted.
Another possible indicator of unsuitable stimulation parameters to
be found during pulse testing is drift of the electrode’s equilibrium
potential. For example, a long interphase period (> 1 ms, i = 0) willallow the potential to settle to an equilibrium. If the potential after
cathodic pulses of the same charge per phase settles to diﬀerent equilibria, irreversible charge transfer is likely occurring (Cogan, 2008).
As previously stated, solution conductivity and reaction rates increase with temperature, so pulse testing at room rather than bodytemperature may underestimate the range of suitable stimulation
parameters. This may be especially true for porous electrodes which
rely temperature-sensitive transport mechanisms occurring within theirpore structure (Cogan, 2008).

4.1.1.2. Electrochemical impedance spectroscopy. In electrochemical
impedance spectroscopy (EIS), the impedance and phase of anelectrode in PBS is measured in response to a small (∼10 mV)
sinusoidal signal across a range of frequencies (typically 1 to 105 Hz)(Cogan, 2008). The impedance at 1 kHz is typically reported to serve as
an approximate predictor of an electrode’s recording merits, with 1 kHz
presumably chosen to represent the frequency range of neural activity.
Low impedance is generally desirable, however this is traded oﬀ withfactors such as electrode size and placement. Consequently, a wide
range of 1 kHz impedances is found in literature, ranging from singlekΩ to tens of MΩ. The 1 kHz impedance is limited in its usefulness as it
may not even indicate broken electrode leads, especially when the

4.1.2. Longevity testing
The warm, saline, chemically active in vivo environment is inhospitable toward devices and over time can induce multiple failure modes
including electrode damage and insulation breakdown. However, current benchtop models to simulate this environment are limited. Even so,
there is utility in conducting exposure studies to determine
21

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

between the exposed electrode sites will likely obscure the smallerchanges of interest which arise from insulation defects. To avoid thisproblem, test structures with insulated electrodes may be included, orinterdigitated electrodes insulated by the same material can be used.
These alternatives do not accurately represent the devices to be implanted, however, and may not reproduce the same failure modes. As ofyet, no standard technique has emerged to monitor the development ofcross-talk among electrodes in complete, unmodiﬁed devices.
Interpretation of data from the above techniques is helped immensely by visual inspection of the device via microscopy. Simple optical inspection can be performed repeatedly throughout the trial andmay reveal the progression of structural damage. However, some feature sizes and damage indications may require the higher resolution of
scanning electron microscopy (SEM) which, for many device types, isdestructive and thus can only be performed at the end of the testing
period.

improvements to device construction and performance that will maximize success of future animal studies.
4.1.2.1. Simulating the in vivo environment. A basic simulated in vivo
environment is a PBS bath set at 37 °C. In accordance with the
Arrhenius equation, the chemical reactions involved in device aging
can be exponentially accelerated by increasing temperature (Takmakov
et al., 2015). To approximate the equivalent rate of aging imparted byelevated temperature, the “ten degree rule” states that every increase of
10 °C above the reference temperature doubles the rate of aging
reactions in polymers (Hukins et al., 2008). Mathematically
expressed, the ten degree rule’s accelerated aging factor faa is given by

faa = 2(T − Tref )/10
where Tref is body temperature. 57–87 °C is a common temperature
range for accelerated aging as it provides a signiﬁcant increase intesting throughput (faa = 4–32) while being relatively safe for handlingand avoiding initiation of other chemical processes that may invalidate
the approximation, such as boiling. Selection of temperature shouldtake into account the suggested operating temperature range for thematerials and whether thermally induced changes to the materials canresult that are not representative of expected in vivo failure modes.
Takmakov et al. proposed an enhancement to the simulation of invivo device aging by including reactive oxygen species (ROS) in thesolution—namely, hydrogen peroxide (Street et al., 2018; Takmakov
et al., 2015). ROS are released by activated immune cells and are likely
one of the agents responsible for in vivo device failure, so raising theperformance standard by evaluating devices against harsh simulated
environments including ROS should yield improved chronic performance.

4.1.3. Simulated implantation
Since durability during handling and simplicity in surgical implantation are key determinants of PNI success (Section 2.6), a recommended benchtop study is to simulate the installation process.
While a section of compliant tubing may provide the simplest geometrically-similar nerve phantom, fabrication of phantoms that mechanically approximate peripheral nerve is non-trivial due to inhomogeneous
tissue composition and anisotropic mechanical properties. For example,
agarose gel, which is often used as a phantom for other neural tissues,
can be made to have similar transverse compressibility but lacks similar
longitudinal tensile strength. A nerve phantom used by Cobo et al.
(2019) to simulate dye absorption on the nerve surface consisted of
paper tissue wrapped around a silicone core. In the ﬁeld of anesthesia, acreative variety of nerve-in-tissue phantoms are used for training of
local anesthetic delivery including gelatin, tofu, pork loin, and processed meats (Pollard, 2008; Rathbun et al., 2018; Sparks et al., 2014;
Wells and Goldstein, 2010). 3D printing of hydrogels and composite
structures may also oﬀer a new method to fabricate more accurate
nerve-like models. Until more reﬁned nerve phantoms are developed,
the most accurate implantation simulations are performed on cadavernerve.

4.1.2.2. Evaluation techniques. Several techniques are used to evaluate
device performance and function during longevity testing. The ﬁrst is
EIS. Impedance tends to decrease over time for many devices whensoaked. This decrease is thought to be attributed to insulation
compromise that eﬀectively increases the area of the electrode, but
EIS alone does not provide enough information to diﬀerentiate among
delamination, cracks, or moisture absorption through the bulk of thematerial, nor does it give information about the location of insulation
defects (Takmakov, 2017). Matching the Bode plot of an electrode tothat of an equivalent circuit model may lend some insight (OrtigozaDiaz et al., 2018; Straka et al., 2018). As previously noted, monitoring
impedance only at 1 kHz is insuﬃcient to capture signiﬁcant changes inelectrode condition (Straka et al., 2018; Takmakov et al., 2015).
Takmakov et al. (2015) found impedance at frequencies < 10 Hz tobe most sensitive to aging processes.
Comparison of slow and fast CVs can also provide insight into
electrode failure over time. In an example given by Cogan (2008), a
scan rate of 50,000 mV/s provides information only on the exposed
electrode site and should change minimally over time if the electrodesurface is stable, or show decreased currents in the case of deterioration. On the other hand, a scan rate of 50 mV/s will access areas of the
conductor where the insulation has been compromised by delamination
or solution ingress, showing increased current magnitudes as the insulation deteriorates.
While EIS and CV may capture decreased impedance suggestive ofcompromised insulation, these data do not necessarily indicate afunctional failure—an electrode with imperfect insulation may still beable to suﬃciently stimulate and record. The functional deﬁcit ofconcern with compromised insulation is loss of electrical isolation
among electrodes, or cross-talk. Cross-talk degrades stimulation andrecording selectivity by spreading the signals from one electricalchannel to others, and so any attempts in the PNI design to increase
selectivity will be negated. EIS performed between two electrodes may
quantify changes in their mutual isolation, however the solution path

4.1.4. Other in vitro tests
In addition to the generalized in vitro tests described above, the PNI
designer should carefully consider what other tests should be performed
to validate idiosyncratic features of a device. Examples from literatureinclude neuron cultures to study regenerative approaches (Wieringa
et al., 2010b), ﬂow and pressure characterizations for microﬂuidic
channels (Cobo et al., 2019), deformation tests for delicate structures(Ordonez et al., 2014), and force tests for insertion and anchoringmechanisms (Cutrone et al., 2015). Packaging components such as
cables and connectors should also be subjected to a battery of bench top
tests, as exempliﬁed by Pena et al. (2017). It is noted that none of the invitro tests mentioned can adequately simulate potential host response
issues or disruption of devices by behaving animals.
4.2. In vivo characterization
Borrowing the FDA’s framework, the goals of in vivo characterization are to discern a PNI’s merits in terms of safety and eﬃcacy. Morespeciﬁcally, this includes the PNI’s eﬀects on nerve health and function,and its ability to initiate and measure nerve activity. Presented here isan introductory set of considerations and best practices which emerge
from literature regarding in vivo characterization. Successful animalstudies will depend on decisions made early in the development process, such as animal model selection. The reader is advised to consult
examples in literature for more detailed guidance regarding methods ofanimal studies. Additionally, works by Loeb and Gans (1986) and
Hoﬀer (1990) are recommended for foundational techniques and22

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

consideration of packaging is critical to gaining useful in vivo results,
especially in chronic studies, and deserves signiﬁcant attention from thePNI designer (Mortimer et al., 1995). Since PNI outcomes are so
sensitive to packaging, successful packaging designs and experiences
should be reported in detail to advance the ﬁeld.
4.2.1.2.1. Device-level connection. In early PNI designs consisting ofwires stitched into the wall of a silicone cuﬀ, the lead wire andelectrode were a single continuous conductor. In most other PNI
designs, the electrodes and leads are separate conductors which mustbe electrically and mechanically connected. Since most contemporary
and next-generation devices are microfabricated, “breaking out”
microscale electrodes to interface with macroscale components is
non-trivial. Most thin-ﬁlm polymer devices include a short ribbon
cable which carries the leads away and scales up the conductive
traces to facilitate connection to macroscale leads.
Traditional ultrasonic and thermal wirebonding techniques used in
the semiconductor industry are a well-established option for PNIs
which have a rigid substrate, such as the Utah array. A similar methodof connection is resistance welding, which involves application ofpressure and a high current pulse to heat up the contact point between awire and the electrode (Loeb and Gans, 1986; Ordonez et al., 2014;
Rozman et al., 2018). However, the thin metals and ﬂexible substratesof thin-ﬁlm polymer devices may be prone to failure during theseprocesses (Ortigoza-Diaz et al., 2018).
The “MicroFlex” technique introduced by Meyer et al. is an adaptation of wirebonding for ﬂexible devices. A ball bond acts as a rivet
through a hole in the thin-ﬁlm conductor, connecting it electrically andmechanically to a rigid substrate (Meyer et al., 2001; Schuettler et al.,2008). This substrate may be an integrated circuit or an intermediateconnector between the device and larger lead wires, such as the ceramicinterconnect plate with screen-printed conductive tracks described by
Yoshida et al. (Boretius et al., 2012; Yoshida et al., 2006).
Another tactic for device-level connection is to design the PNI forcompatibility with oﬀ-the-shelf electrical connectors. In particular, theback end of thin-ﬁlm polymer PNIs can easily be designed to mimic
standard ﬂat ﬂexible cables (FFCs), and thus can be inserted into FFC
connectors such as zero insertion force (ZIF) connectors (Gutierrez
et al., 2011; Ortigoza-Diaz et al., 2018). Currently-available ZIFs are
best mounted on printed circuit boards (PCBs) which can be used tofacilitate connection to lead wires.
For all connection methods, the connection sites must be thoroughly
potted with biocompatible epoxy and/or silicone to insulate the exposed conductors from ﬂuids and tissues. Potting also lends crucial
mechanical support to the connection and provides a smooth surface tominimize tissue irritation, though bulky potting should be avoided.
Based on the literature on packaging of neural interfaces, a few bestpractices are suggested. First, the materials used at the connection siteshould be biocompatible, which excludes many standard non-medical
materials, such as solder and conductive epoxy. Second, junctions between dissimilar metals should be avoided. If such a junction comes
into contact with saline, it will produce a battery eﬀect, leading to a
low-but-steady DC current and thus metal corrosion and possible tissue
damage (Loeb and Gans, 1986). Finally, mechanical discontinuities
should be avoided, as these present preferential bending sites. For thispurpose, a tapered layer of silicone may be used to support a gradualtransition between stiﬀ and ﬂexible regions. If including non-biocompatible materials or bi-metallic junctions, as would be the case witha PCB-mounted ZIF connector, these delicate connections must be
thoroughly potted and insulated to prevent contact with body ﬂuid.
Potting may be suﬃcient for studies of limited duration but are a likely
site of failure in long term studies. Overall, we note that the limited
methods and materials available to the PNI designer make chronicallyreliable packaging a challenge and there is a need to develop more
options for device-level connections.
4.2.1.2.2. Cabling. Electrode leads conduct signals from the PNI toa percutaneous connector and their design and placement is crucial topractical tips.
4.2.1. General considerations for in vivo studies
Some general considerations for in vivo studies include animalmodel selection, packaging, and device sterilization.
4.2.1.1. Animal model selection. Several factors should be consideredwhen deciding whether to adopt one animal model or select another.
Rat sciatic nerve has become the default model for PNI studies, oﬀering
a well-characterized, inexpensive, trainable mammalian model withestablished, consistent strains, a workable size-scale, and compliant
behavior (Varejão et al., 2001). Concerning size and morphology, cat
and rabbit models oﬀer a larger size scale compared with rats, allowing
implantation of multiple PNIs or interface with more distal,
diﬀerentiated branches. Prior to the 1990s, cat was the model ofchoice for PN studies as its size accommodated the bulkier technologiesof the time and the model was supported by a substantial body ofresearch (Hoﬀer, 1990). Pig median nerve has been used to better
approximate the large diameter and high fascicle count of human
median nerve (Kundu et al., 2014). On the other hand, smaller animals(e.g. mice and song birds) may oﬀer other desirable traits which must be
weighed against the challenges that accompany smaller nerves andbodies, including microsurgical techniques and strict packaging weight
limitations.
For long term experiments, it is important to select an animal modelthat will not grow substantially throughout the implantation period, as
this will require length accommodation from the cabling and possibly
other dimensions of the device. For this reason, it is advisable to implant in fully- or near-fully grown animals. Both male and female rats
exhibit asymptotic growth trajectories, though female rats may be
preferred over males since they remain smaller and their size stabilizes
more quickly (Reichling and German, 2000). A controlled diet may
slow the rate of growth, but ﬁnal size will remain the same (Reichlingand German, 2000).
Another factor in animal and nerve selection is the set of techniquesthat will be used to evaluate nerve health and device performance.
Some useful characterization techniques found in literature (Sections
4.2.2 and 4.2.3) depend on particular behaviors such as ﬁne motor
tasks, walking patterns, or birdsongs (Lissandrello et al., 2017; Varejão
et al., 2001; Wurth et al., 2017). Others depend on particular limb sizes
and nerve targets in order to use specially-built measurement apparatuses (Grill and Mortimer, 1996b; Riso et al., 2000). PNIs intended forparticular bioelectronic medicine applications must select from a subset
of animals that oﬀer an appropriate disease model.
Regenerative PNI studies are especially sensitive to animal model
selection. Early regenerative studies experienced diﬃculties in mammals due to connective tissue proliferation while more success wasachieved in amphibians (Stein et al., 1975). However, mammal models
are preferable, assuming a goal of eventual use in humans. Tos et al.
(2009) gave an account of multiple animal models used in nerve regeneration studies. While Lewis, Sprague Dawley, and Wistar rat strains
are all used in PNI studies, regenerative PNI studies almost exclusively
use Lewis rats, reportedly because they exhibit the least-problematic
behavioral response to the limb numbness which follows nerve transection.
In summary, several animal models exist and selection can be
guided by examples in literature that share similar goals. The animalmodel will dictate device dimensions and packaging design and should
be decided upon at the beginning as designs may not always be interchangeable across models.
4.2.1.2. Packaging for in vivo studies. It is common for long-term in vivo
studies to terminate not because of interface failure, but instead
because of a packaging failure such as lead breakage (Bradley et al.,
1997; Desai et al., 2014; Grill and Mortimer, 2000; Hoﬀer, 1990;
Kuliasha et al., 2018; Stein et al., 1978). Therefore, careful
23

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

connect and disconnect from an external cable should be avoided, as
this increases the risk of injuring the animal or breaking the mount.
Loeb and Gans (1986) recommend that the mechanical and electricalroles of an external connector should be carried by separate elements inorder to avoid wear and motion artifacts. Furthermore, the mechanicalanchoring should be easily overcome in order to avoid damage or injury
in the process of connection/disconnection, or in the case that the animal applies tension (Loeb and Gans, 1986). One possible solution is touse magnetic mating between the connector mount and the cable, easily
achieved by placing magnets into the face of the connector mount andthe mating face of the external cable (Yaghouby et al., 2018).
4.2.1.2.4. Fluidic packaging. Though there are fewer examples inliterature, connections for ﬂuidic access to a PNI presumably follow
many of the same guidelines as electrical packaging. In contrast to theout-of-plane connections often used to connect to chip-type
microﬂuidic devices, Cobo et al. (2019) made an in-plane connection
to the Parylene microﬂuidic channel of an extraneural cuﬀ by insertinga length of catheter and sealing it with a ﬂexible, biocompatible
cyanoacrylate glue (Loctite 4902). At the other end of the catheter, anumber of diﬀerent methods exist to drive infusion, including
implantable active or osmotic pumps, subcutaneous injection ports(Pohlmeyer et al., 2009), or standard percutaneous ports for connection
to an external pump. For rats, a backpack-like harness is conventionallyused to support connection between a percutaneous catheter and an
external catheter/tether system for continuous infusion. Harnesses may
require supervision and regular human intervention to ﬁx problems
such as skin irritation and entanglement of limbs or teeth in the straps(Bellinger, 2015). An alternative to the infusion harness is a backmounted pedestal-like infusion button, which omits the problematic
straps and may be protected and kept clean with a magnetic metal cap(Instech Laboratories Inc., 2019).
An additional consideration for ﬂuidic devices is maintaining patency. For this purpose, Hoﬀer and Kallesøe (2001) recommend periodic saline ﬂushes to prevent connective tissue ingrowth from clogging
the outlet.

PNI success (Naples et al., 1990). Multi-stranded lead wires, as opposed
to solid core, exhibit best ﬂexibility and dependability. A long-timestandard wire for this purpose has been the stainless steel, multistranded, 40 AWG Cooner Wire AS 631. From the 1970s to the present,
the most successful multi-conductor cable formation has been helicallywound (individually insulated) wires embedded in silicone, thanks to
its accommodation of bending, twisting, stretching, and, to a lesser
degree, crushing (Boretius et al., 2012; Brindley, 1977; Donaldson,1983). A modular connector for this type of cable is described by
Donaldson (1985).
Mechanical force transmitted from the cable to the PNI and therefore the nerve has been suspected as a primary cause of nerve damage
by PNI (Grill and Mortimer, 2000; Mortimer et al., 1995). Besides
maximizing cable ﬂexibility, careful cable placement can also help
minimize this force transmission. The cable should be routed to leave
the PNI longitudinally, though it also should not contact the nerve(Hoﬀer and Kallesøe, 2001; Naples et al., 1990). Moreover, increased
strain relief has been achieved by routing the cable to leave the PNI’s
distal end and then form a 180° loop for routing in the proximal direction, anchored with a suture to the muscle (Grill and Mortimer,
2000; Larsen et al., 1998; Romero et al., 2001). Commercial VNS systems similarly advise installation of several strain-relief anchor points
and formation of a strain-relief loop in the cable (VNS Therapy System
Epilepsy Physician’s Manual (US), 2018). Letechipia et al. devised a“spring-sleeve” cable connector which includes a strain relief mechanism and was used successfully in human studies (Letechipia et al.,
1991; Tan et al., 2015).
4.2.1.2.3. Percutaneousconnection. Whilesomemodularimplantable wireless systems for electrophysiology are becoming
available (Micro-Leads Research Products, 2019; Triangle BioSystems
International, 2019; Pederson et al., 2019), percutaneous connection
currently remains the most straightforward approach for most animalstudies. In a rat model, percutaneous connector mounts or pedestals
may be placed on the head or the back. While head-mounted connectors
may limit tampering and damage by the animal, back-mounted
connectors avoid repetitive cable bending at the neck, and the shorter
cable is less susceptible to “antenna” noise. In general, a percutaneousconnector should be located at a point where skin movement is minimal(i.e. the midline), require as small a skin opening as possible, not haveany sharp or irritating edges, and be easy to clean. Commercial
pedestals are available which are generally designed for screw- or
dental cement-mounting onto solid bone—typically the skull, though
the pelvis may also be suitable (Barrese et al., 2013; Seifert et al., 2012).
Recently, high-resolution 3D printing services oﬀer a convenient means
to produce custom plastic and/or or metal connector mounts (Straka
et al., 2018; Vasudevan et al., 2016; Wurth et al., 2017). For mounting
to the lumbar fascia instead of bone, a sheet of surgical mesh cloth may
be attached to the bottom of the pedestal which is placed under the skinand encourages tissue ingrowth for stabilization (Straka et al., 2018). A
back-mounted pedestal, or at least its top, is best made of metal toprotect the connector from gnawing by the animal (Straka et al., 2018;
Yaghouby et al., 2018). It is good practice to apply a coating of silicone
to any surface of the pedestal that will contact soft tissues (Straka et al.,2018; Yaghouby et al., 2018). If a pedestal is not suited to the site, aswith percutaneous connections located on a limb, Tan et al. (2015)
reported use of individually-passed open-matrix helical leads which
minimize the opening size and allow skin to better anchor theconnection, thereby avoiding piston-like motion of the lead anddecreasing the risk of infection.
The same considerations also apply to the connection between thecable and the external connector. If possible, the electrical junction
between the two should be positioned outside the skin so that compromised potting poses less of a risk (Loeb and Gans, 1986). The customizability of 3D-printed pedestals makes it possible to mount any oﬀthe-shelf external connectors for connection to electrophysiology
equipment. In general, connectors which require signiﬁcant force to

4.2.1.3. Device sterilization. In vivo studies typically require thoroughsterilization of the components to be implanted. Selection of an
appropriate sterilization technique depends on multiple factors.
Autoclaving is a widely available method which has been used tosterilize silicone and platinum foil devices, however the hightemperature and moisture of autoclaving may damage other PNIs.
Lower-temperature, dry methods include ethylene oxide and hydrogen
peroxide plasma processes, with hydrogen peroxide plasma becoming
the preferred choice for environmental and safety reasons.
Electromagnetic radiation methods are often not ideal for PNIs as
they tend to degrade many polymers (Ortigoza-Diaz et al., 2018).
Whichever method is chosen, characterization should be performed
before and after sterilization to conﬁrm that the integrity of the device(and packaging components) remains intact.
4.2.2. Characterizing PNI safety
A primary aim of in vivo studies is to characterize a PNI’s eﬀects on
nerve health and function. The methods for accomplishing this can begrouped into functional tests, electrophysiological tests, and histology.
In all of these characterizations, experimental controls should be
carefully conceived and executed in order to elucidate the cause of any
damage to the nerve. For further reading, an exceptionally thoroughand authoritative reference on the topic of nerve injuries is provided by
Sunderland (1978).
4.2.2.1. Functional tests. Of the methods used to evaluate PNI safety,
functional tests are the most indirect, aiming to infer information about
nerve health by observing the end results of nerve activity, i.e. sensory
responses and motor tasks.
The general form of sensory tests is to apply a stimulus to the distal
24

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

information (Krarup and Loeb, 1988). For a longitudinal study, themost eﬀective option may be implantation of simple, well-characterized
extraneural cuﬀs proximal and distal to the PNI under test, if space
allows. Krarup et al. provided a rigorous example of this approach in aseries of studies observing the conduction of cat nerve through nerve
injury and recovery using multiple implanted cuﬀs and patch electrodes(Krarup et al., 1989, 1988; Krarup and Loeb, 1988). Experimental
methods and controls should be ﬁrmly established to minimize any
confounding nerve damage from the implanted electrodes other than
the PNI under test.

part of the limb, and either quantify the degree of response or vary thestimulus intensity to determine the threshold at which a certain response is achieved. For example, the traditional von Frey test consists ofpoking the foot with a series of monoﬁlaments of increasing stiﬀness
and noting the stiﬀness which provokes a response (Chaplan et al.,
1994; Nedic et al., 2013). A similar “foot ﬂick” test uses electrical shockas the stimulus (Sta et al., 2014). Lago et al. (2007b) describe a pinprick
test in which the foot is pricked at ﬁve diﬀerent points, each response isscored on a simple 0–2 scale (absent, decreased, or normal), and scores
are summed. Algesiometry tests follow a similar process but aim to
separate pain ﬁber activation from mechanical stimulation, for example
by irradiating the foot with heat and measuring the time to response(Lago et al., 2007b).
The primary functional motor test for the rat model is walking track
analysis, in which video of the rat’s gait viewed from below is analysedfor parameters such as foot spreading and toe spacing (Varejão et al.,
2001; Vasudevan et al., 2017). These parameters are fed into a calculation which yields a value called the sciatic function index (SFI).
Manifestation of nerve health in ﬁner motor control tasks may be observed by kinematic tests. For example, Wurth et al. (2017) reported
ﬁlming the rat’s leg motion (marked with infrared markers) as it ran
over a ladder of unevenly-spaced rungs. The percentage of missed steps
was taken as an indicator of motor control.
Overall, existing functional tests oﬀer more qualitative value thanquantitative as they are susceptible to a number of behavioral variables
which cannot be tightly controlled. They also tend to quantify responses
into broad ranges (i.e. absent, diminished, or normal function) and thus
have low sensitivity. Furthermore, while each variety of functional test
seems to emphasize one modality of nerve ﬁber over others, the output
is truly a measure of entire-circuit function. For example, a diminishedresponse to a pain stimulus does not necessarily correlate to damaged
pain ﬁbers; rather, it may be the motor ﬁbers which are damaged andcannot conduct reﬂexive signals to the limb. Similarly, diminishedmotor control may be an indication of damaged motor neurons, or it
may actually be a consequence of absent sensory feedback.

4.2.2.3. Histology. The most direct and precise method of measuring a
PNI’s eﬀects on nerve health is visual observation of the nerve’s
microanatomy. Histological methods for peripheral nerve vary
widely, and once again the reader is advised to follow speciﬁc
examples in literature which possess similar histological goals, and to
consult further resources on the topic (Cold Spring Harbor Protocols,
2019; Ghnenis et al., 2018; Highley and Sullivan, 2018; Raimondo
et al., 2009; Yuste, 2011). In general, excised nerve is prepared througha multi-step process of ﬁxation, dehydration, embedding, sectioning,and staining. An osmium tetroxide pre-stain prior to embedding is
recommended in most cases (Raimondo et al., 2009). Embedding ismost often performed in paraﬃn, resin, or optimal cutting temperature
medium (Highley and Sullivan, 2018; Raimondo et al., 2009).
Transverse sections should be taken at three positions: proximal to,
distal to, and at the location of the implant (Grill and Mortimer, 2000;
Larsen et al., 1998; Naples et al., 1990). A longitudinal section at the
implant site may also be valuable (Grill and Mortimer, 2000;
Vasudevan et al., 2017). Silver-based preparations have been largely
replaced by a number of simpler and more reliable stains, the mostcommon of which are hematoxylin and eosin (H&E), Masson’s
trichrome, and toluidine or methylene blue, though H&E does notstain myelin (Highley and Sullivan, 2018; Raimondo et al., 2009).
Immunohistochemical labelling has become increasingly favored for its
ability to diﬀerentially visualize speciﬁc cellular features andbiomarkers, providing more detailed information about the nerve’s
structural condition and immune response to an implant (Christensen
et al., 2014; Highley and Sullivan, 2018; Raimondo et al., 2009; Wurthet al., 2017). In addition to post-sacriﬁcial labelling, Somann et al.
(2018) demonstrated administration of ﬂuorogold tracer several days
prior to sacriﬁce, revealing axonal transport as an indicator of ﬁber
integrity across the implant site.
Variety is also found regarding microscopy techniques. In addition
to light microscopy, transmission electron microscopy has also long
been used to provide the highest resolution and is most eﬀective to
visualize small, unmyelinated ﬁbers (Raimondo et al., 2009; Thomas,
1963). Confocal microscopy, in combination with immunoﬂuorescentlabelling, oﬀers three-dimensional imaging and diﬀerentiation of
structures and cells. Multiphoton microscopy (MPM) also oﬀers imaging at depth up to several hundred microns. MPM additionally enables
imaging by second harmonic generation (SHG) which acts as a nonphotobleaching, label-free contrast (Vijayaraghavan et al., 2014). In
SHG, emission from molecules of non-centrosymmetric structure isexactly half the excitation wavelength. Collagen has such a molecular
structure and thus the collagen-rich connective tissue of peripheral
nerve is well-suited for SHG imaging (Vijayaraghavan et al., 2014).
Jung et al. (2014) applied tissue-clearing techniques to enhance imaging depth of whole-mount nerve after transection and regeneration.
As an alternative to the necessarily destructive techniques of nerve
excision and sectioning, Vasudevan et al. (2019) presented a method forreal-time in vivo imaging of nerve using optical coherence tomography(OCT) and a mechanically-stabilizing ﬁxture. This method provides
another mode of nerve health characterization by using OCT angiography to measure changes in microvasculature and blood ﬂow.
After cross-sectional images of the nerve are acquired, morphometric (also called stereologic) analysis should be performed. The ﬁber

4.2.2.2. Electrophysiological tests. Electrophysiological tests possibly
oﬀer a more direct and quantitative measure of nerve health. The
essence of nerve damage is degeneration and demyelination of nerve
ﬁbers, and these changes in the nerve’s ﬁber population can potentiallybe detected in electrophysiological measurements (Somann et al.,2018). The general form of these tests is to proximally/distally evoke
action potentials in the nerve trunk and distally/proximally record thesignals which are conducted across the implant site. Alternatively, asignal evoked at a third site may be recorded both proximally anddistally to the implant site, providing a more direct comparison. A
compound action potential (CAP) resulting from evocation is arelatively large signal which is the summation of the extracellularsignals of all recruited ﬁbers. Peak amplitude of the CAP depends inpart on the synchronized arrival of individual APs at the recording
electrode. Thus, variation among ﬁbers’ conduction velocities gives rise
to desynchronization which increases with distance from thestimulation site, accompanied by temporal spreading of the CAP and
decrease of its peak amplitude. Alternatively, the integrated rectiﬁed
value of a recorded CAP provides a more reliable reﬂection of thequantity of nerve ﬁbers which are carrying a signal. Thus, a decrease in
the CAP area as it crosses the implant may indicate ﬁber degeneration.
Demyelination of ﬁbers without complete degeneration manifests as adecrease in conduction velocity, which may be directly measured bymultiple recording sites or observed as an increased latency in a portionof signals contributing to the CAP.
A limitation of using electrophysiological tests to evaluate nerve
health is diﬃculty in achieving reproducible recording and stimulationon either side of the PNI. Acute insertion of needle probes is highly
sensitive to changes in placement between tests. Surface EMG may be amore reproducibly recordable distal signal but yields limited25

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Fig. 18. In this example of software-aided morphometry, tiled images were stitched together to form an image of the whole nerve, followed by isolation of individual
ﬁbers and sorting into rejected (red), suspect (blue), or accepted (green) categories based on parameters that indicate distortion from sectioning or imaging. The
suspect category is manually sorted by the researcher and the other categories are conﬁrmed before morphometric analysis is performed. Reprinted from (Christensen
and Tresco, 2018) with permission from Elsevier.

A shallow slope in the recruitment curve is generally desirable as itcorresponds to more ﬁnely graded control of ﬁber recruitment. The
recruitment curve also provides an indication of stimulus eﬃciency interms of threshold and the amount of stimulus needed to achieve signiﬁcant activation. Another desirable characteristic is to have one recruitment curve left-shifted from all other curves which arise from aparticular stimulation condition (e.g. a given electrode or combinationof electrodes). This indicates an ability to selectively achieve that
output.
Quantiﬁcation of selectivity varies based on the mode of output
measurement. A common calculation in cases which involve multiple
simultaneous measurements (e.g. EMGs from multiple patch electrodes)
is the selectivity index (SI). Deﬁned by Veraart et al. (1993), SI is thenormalized output of the muscle of interest divided by the sum of thenormalized outputs of all measured muscles for a given set of stimulus
conditions. For example, for forces F measured from N muscles, the SI of
activating muscle j is given by

population is quantiﬁed in terms of number, density, size distribution,and myelination. A commonly reported measurement is the g-ratio
which is the axon diameter divided by total ﬁber diameter which includes the myelin sheath (Raimondo et al., 2009). Previously, morphometry was a tedious, manual process prone to error and bias. Algorithms for sampling and counting were developed to standardize theprocess and remove bias, and readily-available software tools for image
analysis have since improved eﬃciency (Christensen and Tresco, 2018;
Jung et al., 2014; Larsen, 1998; Raimondo et al., 2009; Silva et al.,2007; Wurth et al., 2017). Shown in Fig. 18, Christensen and Tresco(2018) demonstrated an image processing method to help the researcher isolate and sort ﬁbers prior to morphometric measurements.
4.2.3. Characterizing PNI eﬃcacy
For most PNIs, evaluation of eﬃcacy includes stimulation and/or
recording performance, plus long-term stability.
4.2.3.1. Stimulation performance. A common gauge of stimulation
performance in vivo is the recruitment curve, which is an s-shaped
plot of output signal as a function of the stimulation charge (or otherstimulus parameter such as amplitude, duration, or frequency). The
measured output may be neural CAP area, EMG amplitude, limb
motion, muscle force, or joint torque, however these should not betaken to be equivalent or reliably correlated (Loeb and Gans, 1986).
EMG for stimulation characterization can be recorded by multiple patchor wire electrodes implanted on/in the main muscles innervated by anerve (Kundu et al., 2014; Loeb and Gans, 1986; Wurth et al., 2017).
Limb motion can be recorded by ﬁlm or multi-camera motion-capturesystems for subsequent analysis and quantiﬁcation (González-González
et al., 2018; Wurth et al., 2017). Muscle force has been measured byattaching the tendons to load cells or by implanting tendon force
transducers, though success with these methods was limited to acute or
sub-chronic studies (Grill and Mortimer, 1996b; Hoﬀer, 1990; Loeb andGans, 1986; Veraart et al., 1993). Grill and Mortimer (1996b)
introduced an apparatus for non-invasive three-dimensional isometricjoint torque measurements of the cat foot consisting of an aluminium
boot coupled to a 3D force transducer. They found that the peak of thetorque measurement correlated well with its time integral and thus
served as a suﬃcient representation of ﬁber activation. Stieglitz et al.
(2003) developed a similar apparatus for rat.

SI (stimulusconditions) =FjN∑i = 1 Fi

SI yields a range of values from 0 to 1. To summarize a device’s
ability to selectively activate multiple targets, Badia et al. (2011a) introduced a device selectivity index which is a product of the highest SIs
achieved by each measured muscle. However, as Kundu et al. (2014)
pointed out, this deﬁnition is biased towards smaller muscle counts andstrongly penalizes a device for one low SI though its others may be high;instead, a simple mean of the SIs was proposed.
Leventhal and Durand (2003) oﬀered another quantiﬁcation of selectivity based on 3D joint torque measurements. Two electrodes of the
PNI were independently stimulated, yielding diﬀerent torque vectors.
Then they were stimulated simultaneously. If the simultaneous stimulation yielded a torque vector that did not match the sum of the two
independent vectors, it was presumed that there was some overlap inthe ﬁber populations activated by each electrode. This measure of selectivity S is summarized by comparison of the experimental (simultaneous) and theoretical (summed) vectors, and calculated byS=(τs − τ h ) ∙τ l
|τ l |2

where τs is the simultaneous torque vector, and τh and τl are the are the26

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

histology should be performed to illuminate causes of change or toreveal detrimental changes which were not previously detected.

higher- and lower-magnitude independent vectors, respectively.
Non-motor applications of PN stimulation may have a wide variety
of other physiological biomarkers to serve as the performance indicator.
For example, with vagus nerve stimulation the physiological outputs ofinterest may be changes in heart rate, blood pressure, or concentrationof blood analytes (Plachta et al., 2014; Somann et al., 2018). In cases
like these, selectivity is presumed to manifest as the absence of unwanted side eﬀects, though such studies will beneﬁt from standardization and a more thorough accounting of all the potentially aﬀected
targets (Birmingham et al., 2014).

5. Conclusion
This review has covered guiding principles for PNI design, PNI types
and strategies for enhancing performance, and methods of PNI characterization. These topics may serve as a starting point for a new entrant to the ﬁeld who wishes to design a PNI. They may also serve as astarting point for further discussion in the PNI community and a call to
conduct further comparative studies of diﬀerent approaches so that thestrengths of each approach might be more deﬁnitively elucidated.
While an attempt has been made to provide an initial collection of bestpractices in PNI design and development, there is a strong need for PNI
designers to further consider and converge on the most eﬀective paths
to further the ﬁeld toward highly functional, long-lasting PNIs.

4.2.3.2. Recording performance. Fewer examples in literaturedemonstrate rigorous in vivo characterization of PNI recording
performance. The signals recorded may be electrically evoked,
naturally evoked, or spontaneous. Recording of large, electricallyevoked CAPs may serve as a rudimentary demonstration of a PNI’s
basic recording functionality but provide little value as a performanceindicator since such a signal would not naturally arise during clinicaluse. More meaningful signals can be evoked by natural mechanisms,
such as mechanical stimulation via brushing or pinching of the foot.
Spontaneous nerve activity can also be recorded, but without
correlative observations provides little information. Wodlinger andDurand (2010) elicited “pseudo-spontaneous” nerve activity bystimulating with sine waves at 5 or 10 kHz which resulted in arandom ﬁring pattern.
The end goal of PNI recordings is to meaningfully distinguish signals(or functional ensembles of signals) from one another and correlatethem to speciﬁc origins or functions, such as volitional motor control
signals or proprioceptive muscle aﬀerents. Riso et al. (2000) used anapparatus for precise rotation of the ankle of a rabbit which allowed
detailed study of muscle aﬀerent signals recorded via cuﬀ. Many of thesame output measurements described for stimulation performance canpotentially serve as correlative signals to help interpret recorded signals, as can a variety of naturally evoked stimuli. To characterize a
PNI’s ability to localize signals within the nerve trunk, separate distal
branches of the nerve may be stimulated so that the aﬀerent CAPs are
conﬁned to a known fascicle (Aristovich et al., 2018; Wodlinger andDurand, 2010; Zariﬀa et al., 2011).
Signal-to-noise ratio (SNR) is frequently mentioned in PNI literature
but not often clearly deﬁned. In many cases, SNR is reported simply as
the ratio of peak-to-peak amplitudes of a spike and the steady background noise. However, it should be noted that this deﬁnition may not
be consistent with adjacent disciplines which use power rather thanamplitude (Schultz, 2007). Furthermore, what is interpreted as noise
may in some cases be ongoing spontaneous activity, and so the ratio
might be better described as evoked-to-spontaneous.
Since PNI experimental setups can be prone to multiple interference
signals such as EMG, motion artifacts, and stimulus artifacts, the neuralnature of recorded signals should be veriﬁed by subsequent applicationof local anesthetic to the nerve, such as lidocaine or bupivacaine, whichblock action potential propagation and should suppress the previously
recorded signal (Elyahoodayan et al., 2019; Lissandrello et al., 2017;
Riso et al., 2000). Temporary placement of paraﬁlm or plastic wrap
under or around the nerve can also help reject EMG interference inacute setups, and also limits dehydration of the nerve during extended
surgical exposures (Vasudevan et al., 2017, 2016).

Declaration of Competing Interest
None.
Acknowledgements
This work was sponsored in part by the Defense Advanced Research
Projects Agency (DARPA) BTO under the auspices of Drs. Doug Weberand Eric Van Gieson through the DARPA Contracts Management Oﬃce
Cooperative Agreement HR0011-15-2-0006. C.L. was partially supported by the University of Southern California Provost Fellowship.
References
Abidian, M.R., Martin, D.C., 2009. Multifunctional nanobiomaterials for neural interfaces.
Adv. Funct. Mater. 19, 573–585. https://doi.org/10.1002/adfm.200801473.
About us – Finetech Medical, n.d. URL https://ﬁnetech-medical.co.uk/about-us/(Accessed 4.26.19).
Agnew, W.F., McCreery, D.B., 1990. Considerations for safety with chronically implanted
nerve electrodes. Epilepsia 31, S27–S32. https://doi.org/10.1111/j.1528-1157.1990.
tb05845.x.
Agnew, W.F., McCreery, D.B., Bullara, L.A., Yuen, T.G.H., 1990. Eﬀects of prolonged
electrical stimulation of peripheral nerve. In: Agnew, W.F., McCreery, D.B. (Eds.),
Neural Prostheses: Fundamental Studies, Prentice Hall Biophysics and Bioengineering
Series. Prentice Hall, Englewood Cliﬀs, N.J, pp. 147–168.
Agnew, W.F., McCreery, D.B., Yuen, T.G.H., Bullara, L.A., 1999. Evolution and resolutionof stimulation-induced axonal injury in peripheral nerve. Muscle Nerve 22,1393–1402. https://doi.org/10.1002/(SICI)1097-4598(199910)22:10<1393::AIDMUS9>3.0.CO;2-E.
Agnew, W.F., McCreery, D.B., Yuen, T.G.H., Bullara, L.A., 1989. Histologic and physiologic evaluation of electrically stimulated peripheral nerve: considerations for theselection of parameters. Ann. Biomed. Eng. 17, 39–60. https://doi.org/10.1007/
BF02364272.
Akin, T., Najaﬁ, K., Smoke, R.H., Bradley, R.M., 1994. A micromachined silicon sieve
electrode for nerve regeneration applications. IEEE Trans. Biomed. Eng. 41, 305–313.
https://doi.org/10.1109/10.284958.
Alexander, S., Rowan, D., 1968. Electrical control of urinary incontinence by radio implant. A report of 14 patients. Br. J. Surg. 55, 358–364. https://doi.org/10.1002/bjs.
1800550508.
Al-Shueli, A.I.K., Clarke, C.T., Donaldson, N., Taylor, J., 2014. Improved signal processing
methods for velocity selective neural recording using multi-electrode cuﬀs. IEEE
Trans. Biomed. Circuits Syst. 8, 401–410. https://doi.org/10.1109/TBCAS.2013.
2277561.
Anderson, H.E., Weir, R.F.F., 2019. On the development of optical peripheral nerve interfaces. Neural Regen. Res. 14, 425. https://doi.org/10.4103/1673-5374.245461.
Andreasen, L.N.S., Struijk, J.J., 2002. Signal strength versus cuﬀ length in nerve cuﬀ
electrode recordings. IEEE Trans. Biomed. Eng. 49, 1045–1050. https://doi.org/10.
1109/TBME.2002.800785.
Aoyagi, Y., Stein, R.B., Branner, A., Pearson, K.G., Normann, R.A., 2003. Capabilities of apenetrating microelectrode array for recording single units in dorsal root ganglia of
the cat. J. Neurosci. Methods 128, 9–20. https://doi.org/10.1016/S0165-0270(03)00143-2.
Aristovich, K., Donegá, M., Blochet, C., Avery, J., Hannan, S., Chew, D.J., Holder, D.,2018. Imaging fast neural traﬃc at fascicular level with electrical impedance tomography: proof of principle in rat sciatic nerve. J. Neural Eng. 15, 056025. https://doi.
org/10.1088/1741-2552/aad78e.
Avery, R.E., Wepsic, J.S., 1973. Implantable Electrodes for the Stimulation of the SCIATIC
Nerve. US3738368.
Badia, J., Boretius, T., Andreu, D., Azevedo-Coste, C., Stieglitz, T., Navarro, X., 2011a.

4.2.3.3. Stability. In both stimulation and recording characterizations,a crucial dimension of PNI performance is long-term stability. Clinicalusefulness of a PNI will depend on predictable operation throughout the
implant duration. Host response, tissue rearrangement, and device
deterioration may cause changes in a PNI’s input-output properties
and signal capturing. Throughout a long-term study, the researcher
should monitor for changes in such benchmarks as electrode
impedance, stimulation threshold, and stimulated or recorded targets.
After a long-term trial, close examination of the explanted device and27

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

ﬂexible cuﬀ-like microelectrode for dual purpose, acute and chronic electrical interfacing with the mouse cervical vagus nerve. J. Neural Eng. 14, 066005. https://doi.org/10.1088/1741-2552/aa7a42.
Chaplan, S.R., Bach, F.W., Pogrel, J.W., Chung, J.M., Yaksh, T.L., 1994. Quantitative
assessment of tactile allodynia in the rat paw. J. Neurosci. Methods 53, 55–63.
https://doi.org/10.1016/0165-0270(94)90144-9.
Charkhkar, H., Shell, C.E., Marasco, P.D., Pinault, G.J., Tyler, D.J., Triolo, R.J., 2018.
High-density peripheral nerve cuﬀs restore natural sensation to individuals withlower-limb amputations. J. Neural Eng. 15, 056002. https://doi.org/10.1088/17412552/aac964.
Cho, S.-H., Lu, H.M., Cauller, L., Romero-Ortega, M.I., Lee, J.-B., Hughes, G.A., 2008.
Biocompatible SU-8-based microprobes for recording neural spike signals from regenerated peripheral nerve ﬁbers. IEEE Sens. J. 8, 1830–1836. https://doi.org/10.
1109/JSEN.2008.2006261.
Christensen, M.B., Pearce, S.M., Ledbetter, N.M., Warren, D.J., Clark, G.A., Tresco, P.A.,
2014. The foreign body response to the Utah Slant Electrode Array in the cat sciatic
nerve. Acta Biomater. 10, 4650–4660. https://doi.org/10.1016/j.actbio.2014.07.
010.
Christensen, M.B., Tresco, P.A., 2018. The foreign body response and morphometricchanges associated with mesh-style peripheral nerve cuﬀs. Acta Biomater. 67, 79–86.
https://doi.org/10.1016/j.actbio.2017.11.059.
Christensen, M.B., Wark, H.A.C., Hutchinson, D.T., 2016. A histological analysis of human
median and ulnar nerves following implantation of Utah slanted electrode arrays.
Biomaterials 77, 235–242. https://doi.org/10.1016/j.biomaterials.2015.11.012.
Christie, B.P., Freeberg, M., Memberg, W.D., Pinault, G.J.C., Hoyen, H.A., Tyler, D.J.,
Triolo, R.J., 2017. Long-term stability of stimulating spiral nerve cuﬀ electrodes onhuman peripheral nerves. J. NeuroEng. Rehabil. 14. https://doi.org/10.1186/
s12984-017-0285-3.
Chu, J.-U., Song, K.-I., Han, S., Lee, S.H., Kim, J., Kang, J.Y., Hwang, D., Suh, J.-K.F.,
Choi, K., Youn, I., 2012. Improvement of signal-to-interference ratio and signal-tonoise ratio in nerve cuﬀ electrode systems. Physiol. Meas. 33, 943–967. https://doi.
org/10.1088/0967-3334/33/6/943.
Ciancio, A.L., Cordella, F., Hoﬀmann, K.-P., Schneider, A., Guglielmelli, E., Zollo, L.,2017. Current achievements and future directions of hand prostheses controlled via
peripheral nervous system. In: Bertolaso, M., Di Stefano, N. (Eds.), The Hand.
Springer International Publishing, Cham, pp. 75–95. https://doi.org/10.1007/978-3319-66881-9_5.
Clements, I.P., Kim, Y., Andreasen, D., Bellamkonda, R.V., 2007. A regenerative electrodescaﬀold for peripheral nerve interfacing. In: 2007 3rd International IEEE/EMBS
Conference on Neural Engineering. Presented at the 2007 3rd International IEEE/
EMBS Conference on Neural Engineering. IEEE, Kohala Coast, HI, USA. pp. 390–393.
https://doi.org/10.1109/CNE.2007.369691.
Clements, I.P., Mukhatyar, V.J., Srinivasan, A., Bentley, J.T., Andreasen, D.S.,
Bellamkonda, R.V., 2013. Regenerative scaﬀold electrodes for peripheral nerve interfacing. IEEE Trans. Neural Syst. Rehabil. Eng. 21, 554–566. https://doi.org/10.
1109/TNSRE.2012.2217352.
Cobo, A.M., Larson, C.E., Scholten, K., Miranda, J.A., Elyahoodayan, S., Song, D., Pikov,
V., Meng, E., 2019. Parylene-based cuﬀ electrode with integrated microﬂuidics forperipheral nerve recording, stimulation, and drug delivery. J. Microelectromech.
Syst. 28, 36–49. https://doi.org/10.1109/JMEMS.2018.2881908.
Cogan, S.F., 2008. Neural stimulation and recording electrodes. Annu. Rev. Biomed. Eng.
10, 275–309. https://doi.org/10.1146/annurev.bioeng.10.061807.160518.
Cogan, S.F., Hara, S., Ludwig, K.A., 2018. The safe delivery of electrical currents andneuromodulation. Neuromodulation. Elsevier, pp. 83–94. https://doi.org/10.1016/
B978-0-12-805353-9.00007-3.
Cogan, S.F., Ludwig, K.A., Welle, C.G., Takmakov, P., 2016. Tissue damage thresholds
during therapeutic electrical stimulation. J. Neural Eng. 13, 021001. https://doi.org/
10.1088/1741-2560/13/2/021001.
Cold Spring Harbor Protocols [WWW Document], n.d. URL http://cshprotocols.cshlp.
org/ (Accessed 4.21.19).
Coleman, D.L., King, R.N., Andrade, J.D., 1974. The foreign body reaction: a chronic
inﬂammatory response. J. Biomed. Mater. Res. 8, 199–211. https://doi.org/10.1002/
jbm.820080503.
Cork, S.C., Eftekhar, A., Mirza, K.B., Zuliani, C., Nikolic, K., Gardiner, J.V., Bloom, S.R.,
Toumazou, C., 2018. Extracellular pH monitoring for use in closed-loop vagus nerve
stimulation. J. Neural Eng. 15, 016001. https://doi.org/10.1088/1741-2552/
aa8239.
Crampon, M.-A., Sawan, M., Brailovski, V., Trochu, F., 1999. New easy to install nerve
cuﬀ electrode using shape memory alloy armature. Artif. Organs 23, 392–395.
https://doi.org/10.1046/j.1525-1594.1999.06357.x.
Cuoco, F.A., Durand, D.M., 2000. Measurement of external pressures generated by nerve
cuﬀ electrodes. IEEE Trans. Rehabil. Eng. 8, 35–41. https://doi.org/10.1109/86.
830947.
Cutrone, A., Bossi, S., Micera, S., 2013. Development of a self-opening neural interface.
Am. J. Respir. Med. 7, 020938. https://doi.org/10.1115/1.4024379.
Cutrone, A., Sergi, P.N., Bossi, S., Micera, S., 2011. Modelization of a self-opening peripheral neural interface: a feasibility study. Med. Eng. Phys. 33, 1254–1261. https://doi.org/10.1016/j.medengphy.2011.06.001.
Cutrone, A., Valle, J.D., Santos, D., Badia, J., Filippeschi, C., Micera, S., Navarro, X., Bossi,
S., 2015. A three-dimensional self-opening intraneural peripheral interface (SELINE).
J. Neural Eng. 12, 016016. https://doi.org/10.1088/1741-2560/12/1/016016.
Davis, T.S., Wark, H.A.C., Hutchinson, D.T., Warren, D.J., O’Neill, K., Scheinblum, T.,
Clark, G.A., Normann, R.A., Greger, B., 2016. Restoring motor control and sensory
feedback in people with upper extremity amputations using arrays of 96 microelectrodes implanted in the median and ulnar nerves. J. Neural Eng. 13, 036001. https://doi.org/10.1088/1741-2560/13/3/036001.

Comparative analysis of transverse intrafascicular multichannel, longitudinal intrafascicular and multipolar cuﬀ electrodes for the selective stimulation of nerve
fascicles. J. Neural Eng. 8, 036023. https://doi.org/10.1088/1741-2560/8/3/
036023.
Badia, J., Boretius, T., Pascual-Font, A., Udina, E., Stieglitz, T., Navarro, X., 2011b.
Biocompatibility of chronically implanted transverse intrafascicular multichannel
electrode (TIME) in the rat sciatic nerve. IEEE Trans. Biomed. Eng. 58, 2324–2332.
https://doi.org/10.1109/TBME.2011.2153850.
Badia, J., Raspopovic, S., Carpaneto, J., Micera, S., Navarro, X., 2016. Spatial and functional selectivity of peripheral nerve signal recording with the transversal intrafascicular multichannel electrode (TIME). IEEE Trans. Neural Syst. Rehabil. Eng.
24, 20–27. https://doi.org/10.1109/TNSRE.2015.2440768.
Baker, R.G., 1993. Electrode Assembly for Nerve Stimulation. US5215089.
Bakula, T., Mortimer, J.T., 2000. Spiral Nerve Cuﬀ Electrode Implantation Tool.
US6093197.
Ballini, M., Bae, J., Marrocco, N., Verplancke, R., Schaubroeck, D., Cuypers, D., Cauwe,
M., O’Callaghan, J., Fahmy, A., Maghari, N., Bashirullah, R., Van Hoof, C., Van
Helleputte, N., de Beeck, M.O., Braeken, D., Mitra, S., 2017. Intraneural active probe
for bidirectional peripheral nerve interface. In: 2017 Symposium on VLSI Circuits.
Presented at the 2017 Symposium on VLSI Circuits. IEEE, Kyoto, Japan. pp. C50–C51.
https://doi.org/10.23919/VLSIC.2017.8008544.
Banni, S., Carta, G., Murru, E., Cordeddu, L., Giordano, E., Marrosu, F., Puligheddu, M.,
Floris, G., Asuni, G.P., Cappai, A.L., Deriu, S., Follesa, P., 2012. Vagus nerve stimulation reduces body weight and fat mass in rats. PLoS One 7, e44813. https://doi.org/
10.1371/journal.pone.0044813.
Barrese, J.C., Rao, N., Paroo, K., Triebwasser, C., Vargas-Irwin, C., Franquemont, L.,
Donoghue, J.P., 2013. Failure mode analysis of silicon-based intracortical microelectrode arrays in non-human primates. J. Neural Eng. 10, 066014. https://doi.org/
10.1088/1741-2560/10/6/066014.
Bellinger, D., 2015. Harness Versus Button Device With Automated Blood Sampler.
Bhadra, N., Kilgore, K.L., 2018. Fundamentals of kilohertz frequency alternating current
nerve conduction block of the peripheral nervous system. Neuromodulation. Elsevier,pp. 111–120. https://doi.org/10.1016/B978-0-12-805353-9.00010-3.
Birmingham, K., Gradinaru, V., Anikeeva, P., Grill, W.M., Pikov, V., McLaughlin, B.,
Pasricha, P., Weber, D., Ludwig, K., Famm, K., 2014. Bioelectronic medicines: a research roadmap. Nat. Rev. Drug Discov. 13, 399–400. https://doi.org/10.1038/
nrd4351.
Boretius, T., Badia, J., Pascual-Font, A., Schuettler, M., Navarro, X., Yoshida, K., Stieglitz,
T., 2010. A transverse intrafascicular multichannel electrode (TIME) to interface withthe peripheral nerve. Biosens. Bioelectron. 26, 62–69. https://doi.org/10.1016/j.
bios.2010.05.010.
Boretius, T., Yoshida, K., Badia, J., Harreby, K., Kundu, A., Navarro, X., Jensen, W.,
Stieglitz, T., 2012. A transverse intrafascicular multichannel electrode (TIME) to treat
phantom limb pain: towards human clinical trials. In: 2012 4th IEEE RAS &amp;
EMBS International Conference on Biomedical Robotics and Biomechatronics(BioRob). Presented at the 2012 4th IEEE RAS &amp; EMBS International Conference
on Biomedical Robotics and Biomechatronics (BioRob 2012). IEEE, Rome, Italy. pp.
282–287. https://doi.org/10.1109/BioRob.2012.6290787.
Bossi, S., Menciassi, A., Koch, K.P., Hoﬀmann, K.-P., Yoshida, K., Dario, P., Micera, S.,
2007. Shape memory alloy microactuation of tf-LIFEs: preliminary results. IEEE
Trans. Biomed. Eng. 54, 1115–1120. https://doi.org/10.1109/TBME.2007.895186.
Bradley, R.M., Cao, X., Akin, T., Najaﬁ, K., 1997. Long term chronic recordings fromperipheral sensory ﬁbers using a sieve electrode array. J. Neurosci. Methods 73,
177–186. https://doi.org/10.1016/S0165-0270(97)02225-5.
Branner, A., Normann, R.A., 2000. A multielectrode array for intrafascicular recording
and stimulation in sciatic nerve of cats. Brain Res. Bull. 51, 293–306. https://doi.org/
10.1016/S0361-9230(99)00231-2.
Branner, A., Stein, R.B., Fernandez, E., Aoyagi, Y., Normann, R.A., 2004. Long-term stimulation and recording with a penetrating microelectrode array in cat sciatic nerve.
IEEE Trans. Biomed. Eng. 51, 146–157. https://doi.org/10.1109/TBME.2003.
820321.
Branner, A., Stein, R.B., Normann, R.A., 2001. Selective stimulation of cat sciatic nerve
using an array of varying-length microelectrodes. J. Neurophysiol. 85, 1585–1594.
https://doi.org/10.1152/jn.2001.85.4.1585.
Brelén, M.E., Duret, F., Gérard, B., Delbeke, J., Veraart, C., 2005. Creating a meaningful
visual perception in blind volunteers by optic nerve stimulation. J. Neural Eng. 2,
S22–S28. https://doi.org/10.1088/1741-2560/2/1/004.
Brindley, G.S., 1977. An implant to empty the bladder or close the urethra. J. Neurol.
Neurosurg. Psychiatry 40, 358–369. https://doi.org/10.1136/jnnp.40.4.358.
Brindley, G.S., 1972. Electrode-arrays for making long-lasting electrical connexion tospinal roots. J. Physiol. 222, 135P–136P.
Brindley, G.S., Polkey, C.E., Rushton, D.N., 1982. Sacral anterior root stimulators for
bladder control in paraplegia. Spinal Cord 20, 365–381. https://doi.org/10.1038/sc.
1982.65.
Brindley, G.S., Polkey, C.E., Rushton, D.N., 1979. Electrical splinting of the knee inparaplegia. Spinal Cord 16, 428–435. https://doi.org/10.1038/sc.1978.78.
Brindley, G.S., Polkey, C.E., Rushton, D.N., Cardozo, L., 1986. Sacral anterior root stimulators for bladder control in paraplegia: the ﬁrst 50 cases. J. Neurol. Neurosurg.
Psychiatry 49, 1104–1114. https://doi.org/10.1136/jnnp.49.10.1104.
Bullara, L.A., 1990. Bidirectional Helical Electrode for Nerve Stimulation. US4920979.
Bullara, L.A., 1986. Implantable Electrode Array. US4573481.
Caparso, A.V., Durand, D.M., Mansour, J.M., 2009. A nerve cuﬀ electrode for controlledreshaping of nerve geometry. J. Biomater. Appl. 24, 247–273. https://doi.org/10.
1177/0885328208097426.
Caravaca, A.S., Tsaava, T., Goldman, L., Silverman, H., Riggott, G., Chavan, S.S., Bouton,
C., Tracey, K.J., Desimone, R., Boyden, E.S., Sohal, H.S., Olofsson, P.S., 2017. A novel

28

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Mengof four-contact spiral nerve-cuﬀ electrodes in stimulating the human femoral nerve. J.
Neural Eng. 6, 046010. https://doi.org/10.1088/1741-2560/6/4/046010.
FitzGerald, J.J., Lacour, S.P., McMahon, S.B., Fawcett, J.W., 2008. Microchannels asaxonal ampliﬁers. IEEE Trans. Biomed. Eng. 55, 1136–1146. https://doi.org/10.
1109/TBME.2007.909533.
FitzGerald, J.J., Lago, N., Benmerah, S., Serra, J., Watling, C.P., Cameron, R.E., Tarte, E.,
Lacour, S.P., McMahon, S.B., Fawcett, J.W., 2012. A regenerative microchannel
neural interface for recording from and stimulating peripheral axons in vivo. J.
Neural Eng. 9, 016010. https://doi.org/10.1088/1741-2560/9/1/016010.
Foldes, E.L., Ackermann, D.M., Bhadra, Niloy, Kilgore, K.L., Bhadra, Narendra, 2011.
Design, fabrication and evaluation of a conforming circumpolar peripheral nerve cuﬀ
electrode for acute experimental use. J. Neurosci. Methods 196, 31–37. https://doi.
org/10.1016/j.jneumeth.2010.12.020.
Fontaine, A.K., Gibson, E.A., Caldwell, J.H., Weir, R.F., 2017. Optical read-out of neuralactivity in mammalian peripheral axons: calcium signaling at nodes of ranvier. Sci.
Rep. 7, 4744. https://doi.org/10.1038/s41598-017-03541-y.
Fontaine, A.K., Kirchner, M.S., Caldwell, J.H., Weir, R.F., Gibson, E.A., 2018. deep-tissue
two-photon imaging in brain and peripheral nerve with a compact high-pulse energy
ytterbium ﬁber laser. In: Optical Interactions with Tissue and Cells XXIX. Presented at
the Optical Interactions with Tissue and Cells XXIX. International Society for Optics
and Photonics. pp. 1049217. https://doi.org/10.1117/12.2309490.
Franks, W., Schenker, I., Schmutz, P., Hierlemann, A., 2005. Impedance characterizationand modeling of electrodes for biomedical applications. IEEE Trans. Biomed. Eng. 52,
1295–1302. https://doi.org/10.1109/TBME.2005.847523.
Freeberg, M.J., Stone, M.A., Triolo, R.J., Tyler, D.J., 2017. The design of and chronic
tissue response to a composite nerve electrode with patterned stiﬀness. J. Neural Eng.
14, 036022. https://doi.org/10.1088/1741-2552/aa6632.
Futia, G.L., Fontaine, A., McCullough, C., Caldwell, J., Restrepo, D., Weir, R., Gibson,
E.A., Ozbay, B.N., George, N.M., 2018. Measurement of wavefront aberrations incortex and peripheral nerve using a two-photon excitation guidestar. In: Bifano, T.G.,
Gigan, S., Kubby, J. (Eds.), Adaptive Optics and Wavefront Control for Biological
Systems IV. Presented at the Adaptive Optics and Wavefront Control for Biological
Systems IV. SPIE, San Francisco, United States, pp. 58. https://doi.org/10.1117/12.
2309492.
Ganzer, P., Sharma, G., 2019. Opportunities and challenges for developing closed-loopbioelectronic medicines. Neural Regen. Res. 14, 46. https://doi.org/10.4103/16735374.243697.
Garde, K., Keefer, E., Botterman, B., Galvan, P., Romero-Ortega, M.I., 2009. Early interfaced neural activity from chronic amputated nerves. Front. Neuroeng. 2. https://doi.
org/10.3389/neuro.16.005.2009.
Garrido, L., Pﬂeiderer, B., Papisov, M., Ackerman, J.L., 1993. In vivo degradation of silicones. Magn. Reson. Med. 29, 839–843. https://doi.org/10.1002/mrm.
1910290620.
Gasson, M., Hutt, B., Goodhew, I., Kyberd, P., Warwick, K., 2004. Invasive neural prosthesis for neural signal detection and nerve stimulation. Int. J. Adapt. Control Signal
Process. https://doi.org/10.1002/acs.854.
Gaunt, R.A., Hokanson, J.A., Weber, D.J., 2009. Microstimulation of primary aﬀerent
neurons in the L7 dorsal root ganglia using multielectrode arrays in anesthetized cats:
thresholds and recruitment properties. J. Neural Eng. 6, 055009. https://doi.org/10.
1088/1741-2560/6/5/055009.
Geddes, L.A., Roeder, R., 2003. Criteria for the selection of materials for implanted
electrodes. Ann. Biomed. Eng. 31, 879–890. https://doi.org/10.1114/1.1581292.
Ghnenis, A.B., Czaikowski, R.E., Zhang, Z.J., Bushman, J.S., 2018. Toluidine blue stainingof resin-embedded sections for evaluation of peripheral nerve morphology. J. Vis.
Exp. 58031. https://doi.org/10.3791/58031.
Giagka, V., Serdijn, W.A., 2018. Realizing ﬂexible bioelectronic medicines for accessing
the peripheral nerves – technology considerations. Bioelectron. Med. 4. https://doi.
org/10.1186/s42234-018-0010-y.
Glenn, W.W.L., Phelps, M.L., 1985. Diaphragm pacing by electrical stimulation of thephrenic nerve. Neurosurgery 17, 974–984. https://doi.org/10.1227/00006123198512000-00021.
González-González, M.A., Kanneganti, A., Joshi-Imre, A., Hernandez-Reynoso, A.G.,
Bendale, G., Modi, R., Ecker, M., Khurram, A., Cogan, S.F., Voit, W.E., RomeroOrtega, M.I., 2018. Thin ﬁlm multi-electrode softening cuﬀs for selective neuromodulation. Sci. Rep. 8. https://doi.org/10.1038/s41598-018-34566-6.
Goodall, E.V., Lefurge, T.M., Horch, K.W., 1991. Information contained in sensory nerve
recordings made with intrafascicular electrodes. IEEE Trans. Biomed. Eng. 38,
846–850. https://doi.org/10.1109/10.83604.
Gore, R.K., Choi, Y., Bellamkonda, R., English, A., 2015. Functional recordings fromawake, behaving rodents through a microchannel based regenerative neural interface. J. Neural Eng. 12, 016017. https://doi.org/10.1088/1741-2560/12/1/016017.
Gorman, P.H., Mortimer, J.T., 1983. The eﬀect of stimulus parameters on the recruitment
characteristics of direct nerve stimulation. IEEE Trans. Biomed. Eng. BME-30,
407–414. https://doi.org/10.1109/TBME.1983.325041.
Green, R., Abidian, M.R., 2015. Conducting polymers for neural prosthetic and neuralinterface applications. Adv. Mater. 27, 7620–7637. https://doi.org/10.1002/adma.
201501810.
Grill, W.M., 2018. Waveforms for neural stimulation. Neuromodulation. Elsevier, pp.
95–102. https://doi.org/10.1016/B978-0-12-805353-9.00008-5.
Grill, W.M., 1995. Spatially Selective Activation of Peripheral Nerve for Neuroprosthetic
Applications. Case Western Reserve University.
Grill, W.M., Mortimer, J.T., 2000. Neural and connective tissue response to long-term
implantation of multiple contact nerve cuﬀ electrodes. J. Biomed. Mater. Res. 50,
215–226. https://doi.org/10.1002/(SICI)1097-4636(200005)50:2<215::AIDJBM17>3.0.CO;2-A.
Grill, W.M., Mortimer, J.T., 1998. Stability of the input-output properties of chronicallyde la Oliva, N., del Valle, J., Delgado-Martinez, I., Mueller, M., Stieglitz, T., Navarro, X.,
2019. Long-term functionality of transversal intraneural electrodes is improved bydexamethasone treatment. IEEE Trans. Neural Syst. Rehabil. Eng. https://doi.org/10.
1109/TNSRE.2019.2897256. 1–1.
de la Oliva, N., Mueller, M., Stieglitz, T., Navarro, X., del Valle, J., 2018a. On the use of
Parylene C polymer as substrate for peripheral nerve electrodes. Sci. Rep. 8. https://doi.org/10.1038/s41598-018-24502-z.
de la Oliva, N., Navarro, X., del Valle, J., 2018b. Dexamethasone reduces the foreign bodyreaction to intraneural electrode implants in the peripheral nerve of the rat. Anat.
Rec. 301, 1722–1733. https://doi.org/10.1002/ar.23920.
de la Oliva, N., Navarro, X., Valle, Jdel, 2018c. Time course study of long-term biocompatibility and foreign body reaction to intraneural polyimide-based implants. J.
Biomed. Mater. Res. A 106, 746–757. https://doi.org/10.1002/jbm.a.36274.
Deberge, L., Delleci, C., Joseph, P.A., Cassoudesalle, H., Glize, B., 2018. Longitudinallong-term evaluation of patients with a Brindley’s neurostimulator and followed in
Bordeaux university hospital: return on 30 years of use. Ann. Phys. Rehabil. Med. 61,
e380. https://doi.org/10.1016/j.rehab.2018.05.884.
del Valle, J., de la Oliva, N., Muller, M., Stieglitz, T., Navarro, X., 2015. Biocompatibility
evaluation of parylene C and polyimide as substrates for peripheral nerve interfaces.
In: 2015 7th International IEEE/EMBS Conference on Neural Engineering (NER).
Presented at the 2015 7th International IEEE/EMBS Conference on NeuralEngineering (NER). IEEE, Montpellier, France. pp. 442–445. https://doi.org/10.
1109/NER.2015.7146654.
del Valle, J., Navarro, X., 2013. Interfaces with the peripheral nerve for the control ofneuroprostheses. International Review of Neurobiology. Elsevier, pp. 63–83. https://doi.org/10.1016/B978-0-12-420045-6.00002-X.
Delgado-Martínez, I., Righi, M., Santos, D., Cutrone, A., Bossi, S., D’Amico, S., Del Valle,J., Micera, S., Navarro, X., 2017. Fascicular nerve stimulation and recording using anovel double-aisle regenerative electrode. J. Neural Eng. 14, 046003. https://doi.
org/10.1088/1741-2552/aa6bac.
Della Santina, C.C., Kovacs, G.T.A., Lewis, E.R., 1997. Multi-unit recording from regenerated bullfrog eighth nerve using implantable silicon-substrate microelectrodes.
J. Neurosci. Methods 72, 71–86. https://doi.org/10.1016/S0165-0270(96)00159-8.
Desai, V.H., Anand, S., Tran, M., Kanneganti, A., Vasudevan, S., Seifert, J.L., Cheng, J.,
Keefer, E.W., Romero-Ortega, M.I., 2014. Chronic sensory-motor activity in behaving
animals using regenerative multi-electrode interfaces. 2014 36th Annual
International Conference of the IEEE Engineering in Medicine and Biology Society.
Presented at the 2014 36th Annual International Conference of the IEEE Engineeringin Medicine and Biology Society 1973–1976. https://doi.org/10.1109/EMBC.2014.
6944000.
Dhillon, G.S., Krüger, T.B., Sandhu, J.S., Horch, K.W., 2005. Eﬀects of short-term training
on sensory and motor function in severed nerves of long-term human amputees. J.
Neurophysiol. 93, 2625–2633. https://doi.org/10.1152/jn.00937.2004.
Djilas, M., Yoshida, K., Kurstjens, M., Azevedo-Coste, C., 2007. Improving the signal-tonoise ratio in recordings with thin-ﬁlm longitudinal intra-fascicular electrodes using
shielding cuﬀs. In: 2007 3rd International IEEE/EMBS Conference on NeuralEngineering. Presented at the 2007 3rd International IEEE/EMBS Conference on
Neural Engineering. IEEE, Kohala Coast, HI, USA. pp. 167–170. https://doi.org/10.
1109/CNE.2007.369638.
Donaldson, Nde N., Donaldson, P.E.K., 1986. When are actively balanced biphasic (‘Lilly’)
stimulating pulses necessary in a neurological prosthesis? I Historical background; Pt
resting potential;Q studies. Med. Biol. Eng. Comput. 24, 41–49. https://doi.org/10.
1007/BF02441604.
Donaldson, N., Rieger, R., Schuettler, M., Taylor, J., 2008. Noise and selectivity of velocity-selective multi-electrode nerve cuﬀs. Med. Biol. Eng. Comput. 46, 1005–1018.
https://doi.org/10.1007/s11517-008-0365-4.
Donaldson, P.E.K., 1985. The craggs connector; a termination for cooper cable. Med. Biol.
Eng. Comput. 23, 195–196. https://doi.org/10.1007/BF02456759.
Donaldson, P.E.K., 1983. The Cooper cable: an implantable multiconductor cable forneurological prostheses. Med. Biol. Eng. Comput. 21, 371–374. https://doi.org/10.
1007/BF02478508.
Eastwood, P.R., Barnes, M., Walsh, J.H., Maddison, K.J., Hee, G., Schwartz, A.R., Smith,P.L., Malhotra, A., McEvoy, R.D., Wheatley, J.R., O’Donoghue, F.J., Rochford, P.D.,
Churchward, T., Campbell, M.C., Palme, C.E., Robinson, S., Goding, G.S., Eckert, D.J.,
Jordan, A.S., Catcheside, P.G., Tyler, L., Antic, N.A., Worsnop, C.J., Kezirian, E.J.,
Hillman, D.R., 2011. Treating obstructive sleep apnea with hypoglossal nerve stimulation. Sleep 34, 1479–1486. https://doi.org/10.5665/sleep.1380.
Edell, D.J., 1986. A peripheral nerve information transducer for amputees: long-term
multichannel recordings from rabbit peripheral nerves. IEEE Trans. Biomed. Eng.
BME-33, 203–214. https://doi.org/10.1109/TBME.1986.325892.
Elgrishi, N., Rountree, K.J., McCarthy, B.D., Rountree, E.S., Eisenhart, T.T., Dempsey,
J.L., 2018. A practical beginner’s guide to cyclic voltammetry. J. Chem. Educ. 95,
197–206. https://doi.org/10.1021/acs.jchemed.7b00361.
Elyahoodayan, S., Larson, C., Cobo, A., Meng, E., Song, D., 2019. Acute in vivo testing of apolymer cuﬀ electrode with integrated microﬂuidic channels for stimulation, recording, and drug delivery on rat sciatic nerve. J. Neurosci. Methods.
Fattahi, P., Yang, G., Kim, G., Abidian, M.R., 2014. A review of organic and inorganic
biomaterials for neural interfaces. Adv. Mater. 26, 1846–1885. https://doi.org/10.
1002/adma.201304496.
FDA Approves Inspire Upper Airway Stimulation Therapy for Obstructive Sleep Apnea,
2014. Press Releases Inspire Med. Syst. [WWW Document] URL https://www.inspiresleep.com/wp-content/uploads/2015/10/FDA-Approves-Inspire-Upper-AirwayStimulation-UAS-Therapy-for-Obstructive-Sleep-Apnea.pdf (Accessed 2.1.19). .
Finetech-Brindley Sacral Anterior Root Stimulator (CPC2): Notes for Surgeons andPhysicians.
Fisher, L.E., Tyler, D.J., Anderson, J.S., Triolo, R.J., 2009. Chronic stability and selectivity

29

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Instech Laboratories Inc., n.d. Instech Laboratories [WWW Document]. URL https://www.instechlabs.com/products/buttons-tethers/vascular-access (Accessed 3.26.19).
R Compensation: Potentiostat Fundamentals / Basics of EIS [WWW Document], n.d.
Gamry Instrum. URL https://www.gamry.com/application-notes/instrumentation/
understanding-ir-compensation/ (Accessed 4.24.19).
Israel, J.S., Dingle, A.M., Sanchez, R.J., Kapur, S.K., Brodnick, S., Richner, T.J., Ness, J.P.,
Novello, J., Williams, J.C., Poore, S.O., 2018. Neuroma Implantation Into Long
Bones: Clinical Foundation for a Novel Osseointegrated Peripheral Nerve Interface 4.
Jeong, J., Jung, W., Kim, O., Chu, J.-U., Youn, I., Kim, K., Sang Rok, O., Park, J.W., Kim,J., 2016. 64-channel double-layered sieve electrode with increased porosity for improved axon regeneration and high spatial resolution. In: 2016 6th IEEE International
Conference on Biomedical Robotics and Biomechatronics (BioRob). Presented at the2016 6th IEEE International Conference on Biomedical Robotics and Biomechatronics(BioRob). IEEE, Singapore, Singapore. pp. 1148–1153. https://doi.org/10.1109/
BIOROB.2016.7523786.
Ju, M.-S., Lin, C.-C.K., Fan, J.-L., Chen, R.-J., 2006. Transverse elasticity and blood perfusion of sciatic nerves under in situ circular compression. J. Biomech. 39, 97–102.
https://doi.org/10.1016/j.jbiomech.2004.10.026.
Judson, J.P., 1968. Radio-frequency electrophrenic respiration: long-term application to apatient with primary hypoventilation. JAMA 203, 1033. https://doi.org/10.1001/
jama.1968.03140120031007.
Judy, J., Schmidt, C.E., Otto, K., Rinaldi, C., KULIASHA, C.A., 2018. Tissue-engineered
Electronic Peripheral Nerve Interface. US20180338765A1. .
Jung, R., Abbas, J.J., Kuntaegowdanahalli, S., Thota, A.K., 2018a. Bionic intrafascicularinterfaces for recording and stimulating peripheral nerve ﬁbers. Bioelectron. Med. 1,
55–69. https://doi.org/10.2217/bem-2017-0009.
Jung, W., Jung, S., Kim, O., Park, H., Choi, W., Son, D., Chung, S., Kim, J., 2019. Wireelectrodes embedded in artiﬁcial conduit for long-term monitoring of the peripheral
nerve signal. Micromachines 10, 184. https://doi.org/10.3390/mi10030184.
Jung, W., Jung, S., Kim, O., Park, H., Choi, W., Son, D., Kim, J., 2018b. Neural electrodewith artiﬁcial conduit for chronic signal recording. Presented at the Society for
Biomaterials 2018 Annual Meeting & Exposition.
Jung, Y., Ng, J.H., Keating, C.P., Senthil-Kumar, P., Zhao, J., Randolph, M.A., Winograd,
J.M., Evans, C.L., 2014. Comprehensive evaluation of peripheral nerve regenerationin the acute healing phase using tissue clearing and optical microscopy in a rodent
model. PLoS One 9, e94054. https://doi.org/10.1371/journal.pone.0094054.
Kallesøe, K., Hoﬀer, J., Strange, K., Valenzuela, I., 1996. Implantable Cuﬀ Having
Improved Closure. US5487756. .
Kandel, E.R. (Ed.), 2013. Principles of Neural Science, 5th ed ed. McGraw-Hill, New York.
Kang, X., Liu, J.-Q., Tian, H., Yang, B., Nuli, Y., Yang, C., 2015. Self-closed parylene cuﬀ
electrode for peripheral nerve recording. J. Microelectromech. Syst. 24, 319–332.
https://doi.org/10.1109/JMEMS.2014.2381634.
Kapur, S.K., Richner, T., Brodnick, S., Williams, J.C., Poore, S.O., 2014. Abstract 62:
development of an optogenetic sensory peripheral nerve interface. Plast. Reconstr.
Surg. 133, 10–99. https://doi.org/10.1097/01.prs.0000445095.84222.cc.
Kennedy, P.R., 1989. The cone electrode: a long-term electrode that records from neurites
grown onto its recording surface. J. Neurosci. Methods 29, 181–193. https://doi.org/
10.1016/0165-0270(89)90142-8.
Kim, B., Reyes, A., Garza, B., Choi, Y., 2015. A microchannel neural interface with embedded microwires targeting the peripheral nervous system. Microsyst. Technol. 21,
1551–1557. https://doi.org/10.1007/s00542-014-2340-3.
Kim, H.-J., Heo, D.N., Lee, Y.J., Lee, S.J., Kang, J.Y., Lee, S.H., Kwon, I.K., Do, S.H.,
2017a. Biological assessments of multifunctional hydrogel-decorated implantableneural cuﬀ electrode for clinical neurology application. Sci. Rep. 7, 15245. https://doi.org/10.1038/s41598-017-15551-x.
Kim, O., Choi, W., Jung, W., Jung, S., Park, H., Park, J.W., Kim, J., 2017b. Novel neuralinterface electrode array for the peripheral nerve. In: 2017 International Conference
on Rehabilitation Robotics (ICORR). Presented at the 2017 International Conference
on Rehabilitation Robotics (ICORR). IEEE, London. pp. 1067–1072. https://doi.org/
10.1109/ICORR.2017.8009391.
Kim, Y., Romero-Ortega, M.I., 2012. Material considerations for peripheral nerve interfacing. MRS Bull. 37, 573–580. https://doi.org/10.1557/mrs.2012.99.
Koole, P., Holsheimer, J., Struijk, J.J., Verloop, A.J., 1997. Recruitment characteristics ofnerve fascicles stimulated by a multigroove electrode. IEEE Trans. Rehabil. Eng. 5,
40–50. https://doi.org/10.1109/86.559348.
Koopman, F.A., Chavan, S.S., Miljko, S., Grazio, S., Sokolovic, S., Schuurman, P.R., Mehta,
A.D., Levine, Y.A., Faltys, M., Zitnik, R., Tracey, K.J., Tak, P.P., 2016. Vagus nerve
stimulation inhibits cytokine production and attenuates disease severity in rheumatoid arthritis. Proc. Natl. Acad. Sci. 113, 8284–8289. https://doi.org/10.1073/pnas.
1605635113.
Kovacs, G.T.A., Storment, C.W., Rosen, J.M., 1992. Regeneration microelectrode array for
peripheral nerve recording and stimulation. IEEE Trans. Biomed. Eng. 39, 893–902.
https://doi.org/10.1109/10.256422.
Krarup, C., Loeb, G.E., 1988. Conduction studies in peripheral cat nerve using implanted
electrodes: I. Methods and ﬁndings in controls. Muscle Nerve 11, 922–932. https://doi.org/10.1002/mus.880110905.
Krarup, C., Loeb, G.E., Pezeshkpour, G.H., 1989. Conduction studies in peripheral cat
nerve using implanted electrodes: III. The eﬀects of prolonged constriction on the
distal nerve segment. Muscle Nerve 12, 915–928. https://doi.org/10.1002/mus.
880121108.
Krarup, C., Loeb, G.E., Pezeshkpour, G.H., 1988. Conduction studies in peripheral cat
nerve using implanted electrodes: II. The eﬀects of prolonged constriction on regeneration of crushed nerve ﬁbers. Muscle Nerve 11, 933–944. https://doi.org/10.
1002/mus.880110906.
Kuliasha, C.A., Spearman, B.S., Atkinson, E.W., Rustogi, P., Furniturewalla, A.S.,
Nunamaker, E.A., Otto, K.J., Schmidt, C.E., Judy, J.W., 2018. Robust and scalable

implanted multiple contact nerve cuﬀ stimulating electrodes. IEEE Trans. Rehabil.
Eng. 6, 364–373. https://doi.org/10.1109/86.736150.
Grill, W.M., Mortimer, J.T., 1997. Inversion of the current-distance relationship bytransient depolarization. IEEE Trans. Biomed. Eng. 44, 1–9. https://doi.org/10.1109/
10.553708.
Grill, W.M., Mortimer, J.T., 1996a. The eﬀect of stimulus pulse duration on selectivity ofneural stimulation. IEEE Trans. Biomed. Eng. 43, 161–166. https://doi.org/10.1109/
10.481985.
Grill, W.M., Mortimer, J.T., 1996b. Non-invasive measurement of the input-output
properties of peripheral nerve stimulating electrodes. J. Neurosci. Methods 65,
43–50. https://doi.org/10.1016/0165-0270(95)00143-3.
Grill, W.M., Norman, S.E., Bellamkonda, R.V., 2009. Implanted neural interfaces: biochallenges and engineered solutions. Annu. Rev. Biomed. Eng. 11, 1–24. https://doi.
org/10.1146/annurev-bioeng-061008-124927.
Grill, W.M., Veraart, C., Mortimer, J.T., 1991. Selective activation of peripheral nerve
fascicles: use of ﬁeld steering currents. In: 1991, IEEE, Orlando, FL, USA. Proceedingsof the Annual International Conference of the IEEE Engineering in Medicine andBiology Society Volume 13: 1991. Presented at the Annual International Conferenceof the IEEE Engineering in Medicine and Biology Society 13. pp. 904–905. https://doi.org/10.1109/IEMBS.1991.684254.
Guiraud, D., Andreu, D., Bonnet, S., Carrault, G., Couderc, P., Hagège, A., Henry, C.,
Hernandez, A., Karam, N., Le Rolle, V., Mabo, P., Maciejasz, P., Malbert, C.-H.,
Marijon, E., Maubert, S., Picq, C., Rossel, O., Bonnet, J.-L., 2016. Vagus nerve stimulation: state of the art of stimulation and recording strategies to address autonomic
function neuromodulation. J. Neural Eng. 13, 041002. https://doi.org/10.1088/
1741-2560/13/4/041002.
Günter, C., Delbeke, J., Ortiz-Catalan, M., 2019. Safety of long-term electrical peripheral
nerve stimulation: review of the state of the art. J. NeuroEng. Rehabil. 16. https://doi.org/10.1186/s12984-018-0474-8.
Guth, L., 1956. Regeneration in the mammalian peripheral nervous system. Physiol. Rev.
36, 441–478. https://doi.org/10.1152/physrev.1956.36.4.441.
Gutierrez, C.A., Lee, C., Kim, B., Meng, E., 2011. Epoxy-less packaging methods forelectrical contact to parylene-based ﬂat ﬂexible cables. In: Presented at the 16th
International Solid-State Sensors, Actuators and Microsystems Conference(TRANSDUCERS). IEEE. pp. 2299–2302.
Hara, S.A., Kim, B.J., Kuo, J.T.W., Lee, C.D., Meng, E., Pikov, V., 2016. Long-term stability of intracortical recordings using perforated and arrayed Parylene sheath electrodes. J. Neural Eng. 13, 066020. https://doi.org/10.1088/1741-2560/13/6/
066020.
Hassler, C., Boretius, T., Stieglitz, T., 2011. Polymers for neural implants. J. Polym. Sci.
Part B: Polym. Phys. 49, 18–33. https://doi.org/10.1002/polb.22169.
Heiduschka, P., Thanos, S., 1998. Implantable bioelectronic interfaces for lost nerve
functions. Prog. Neurobiol. 55, 433–461. https://doi.org/10.1016/S0301-0082(98)00013-6.
Heo, D.N., Song, S.-J., Kim, H.-J., Lee, Y.J., Ko, W.-K., Lee, S.J., Lee, D., Park, S.J., Zhang,
L.G., Kang, J.Y., Do, S.H., Lee, S.H., Kwon, I.K., 2016. Multifunctional hydrogel
coatings on the surface of neural cuﬀ electrode for improving electrode-nerve tissue
interfaces. Acta Biomater. 39, 25–33. https://doi.org/10.1016/j.actbio.2016.05.009.
Hess, A.E., Dunning, J., Tyler, D., Zorman, C.A., 2007. Development of a microfabricated
ﬂat interface nerve electrode based on liquid crystal polymer and polynorbornene
multilayered structures. In: 2007 3rd International IEEE/EMBS Conference on NeuralEngineering. Presented at the 2007 3rd International IEEE/EMBS Conference on
Neural Engineering. IEEE, Kohala Coast, HI. pp. 32–35. https://doi.org/10.1109/
CNE.2007.369604.
Highley, J.R., Sullivan, N., 2018. 18 - Neuropathology and muscle biopsy techniques. In:
Suvarna, S.K., Layton, C., Bancroft, J.D. (Eds.), Bancroft’s Theory and Practice ofHistological Techniques. Elsevier, Amsterdam, pp. 306–336.
Hoﬀer, J.A., 1990. Techniques to study spinal-cord, peripheral nerve, and muscle activity
in freely moving animals. In: Boulton, A.A., Baker, G.B., Vanderwolf, C.H. (Eds.),
Neurophysiological Techniques, Neuromethods. Humana Press, Clifton, N.J.
Hoﬀer, J.A., Chen, Y., Strange, K.D., Christensen, P.R., 1998. Nerve Cuﬀ Having One orMore Isolated Chambers. US5824027A. .
Hoﬀer, J.A., Kallesøe, K., 2001. How to use nerve cuﬀs to stimulate, record or modulate
neural activity. In: Moxon, K.A., Chapin, J.K. (Eds.), Neural Prostheses for
Restoration of Sensory and Motor Function. CRC Press, pp. 139–175.
Hoﬀer, J.A., Loeb, G.E., 1980. Implantable electrical and mechanical interfaces withnerve and muscle. Ann. Biomed. Eng. 8, 351–360. https://doi.org/10.1007/
BF02363438.
Hoﬀer, J.A., Loeb, G.E., Pratt, C.A., 1981. Single unit conduction velocities from averaged
nerve cuﬀ electrode records in freely moving cats. J. Neurosci. Methods 4, 211–225.
https://doi.org/10.1016/0165-0270(81)90033-9.
Holinski, B.J., Everaert, D.G., Mushahwar, V.K., Stein, R.B., 2013. Real-time control ofwalking using recordings from dorsal root ganglia. J. Neural Eng. 10, 056008.
https://doi.org/10.1088/1741-2560/10/5/056008.
Howell, B., Grill, W.M., 2015. Design of electrodes for stimulation and recording.
Implantable Neuroprostheses for Restoring Function. Elsevier, pp. 59–93. https://doi.
org/10.1016/B978-1-78242-101-6.00004-5.
Hudak, E.M., Mortimer, J.T., Martin, H.B., 2010. Platinum for neural stimulation: voltammetry considerations. J. Neural Eng. 7, 026005. https://doi.org/10.1088/17412560/7/2/026005.
Hukins, D.W.L., Mahomed, A., Kukureka, S.N., 2008. Accelerated aging for testing
polymeric biomaterials and medical devices. Med. Eng. Phys. 30, 1270–1274.
https://doi.org/10.1016/j.medengphy.2008.06.001. Special issue to commemorate
the 30th anniversary of Medical Engineering & Physics.
Hursh, J.B., 1939. Conduction velocity and diameter of nerve ﬁbers. Am. J. Physiol.-Leg.
Content 127, 131–139. https://doi.org/10.1152/ajplegacy.1939.127.1.131.

30

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Letechipia, J.E., Peckham, P.H., Gazdik, M., Smith, B., 1991. In-line lead connector for
use with implanted neuroprosthesis. IEEE Trans. Biomed. Eng. 38, 707–709.
Leventhal, D.K., Cohen, M., Durand, D.M., 2006. Chronic histological eﬀects of the ﬂatinterface nerve electrode. J. Neural Eng. 3, 102–113. https://doi.org/10.1088/17412560/3/2/004.
Leventhal, D.K., Durand, D.M., 2004. Chronic measurement of the stimulation selectivityof the ﬂat interface nerve electrode. IEEE Trans. Biomed. Eng. 51, 1649–1658.
https://doi.org/10.1109/TBME.2004.827535.
Leventhal, D.K., Durand, D.M., 2003. Subfascicle stimulation selectivity with the ﬂatinterface nerve electrode. Ann. Biomed. Eng. 31, 643–652. https://doi.org/10.1114/
1.1569266.
Lichtenberg, B.K., De Luca, C.J., 1979. Distinguishability of functionally distinct evokedneuroelectric signals on the surface of a nerve. IEEE Trans. Biomed. Eng. BME-26,
228–237. https://doi.org/10.1109/TBME.1979.326507.
Liske, H., Qian, X., Anikeeva, P., Deisseroth, K., Delp, S., 2013. Optical control of neuronal excitation and inhibition using a single opsin protein, ChR2. Sci. Rep. 3, 3110.
https://doi.org/10.1038/srep03110.
Lissandrello, C.A., Gillis, W.F., Shen, J., Pearre, B.W., Vitale, F., Pasquali, M., Holinski,
B.J., Chew, D.J., White, A.E., Gardner, T.J., 2017. A micro-scale printable nanoclip
for electrical stimulation and recording in small nerves. J. Neural Eng. 14, 036006.
https://doi.org/10.1088/1741-2552/aa5a5b.
Liu, Y., Liu, J., Chen, S., Lei, T., Kim, Y., Niu, S., Wang, H., Wang, X., Foudeh, A.M., Tok,
J.B.-H., Bao, Z., 2019. Soft and elastic hydrogel-based microelectronics for localized
low-voltage neuromodulation. Nat. Biomed. Eng. 3, 58–68. https://doi.org/10.1038/
s41551-018-0335-6.
Llewellyn, M.E., Thompson, K.R., Deisseroth, K., Delp, S.L., 2010. Orderly recruitment ofmotor units under optical control in vivo. Nat. Med. 16, 1161–1165. https://doi.org/
10.1038/nm.2228.
Loeb, G.E., Gans, C., 1986. Electromyography for Experimentalists. University of Chicago
Press, Chicago.
Loeb, G.E., Marks, W.B., Beatty, P.G., 1977. Analysis and microelectronic design of tubular electrode arrays intended for chronic, multiple singleunit recording from captured nerve ﬁbres. Med. Biol. Eng. Comput. 15, 195–201. https://doi.org/10.1007/
BF02442964.
Loeb, G.E., Peck, R.A., 1996. Cuﬀ electrodes for chronic stimulation and recording of
peripheral nerve activity. J. Neurosci. Methods 64, 95–103. https://doi.org/10.1016/
0165-0270(95)00123-9.
Lotﬁ, P., Garde, K., Chouhan, A.K., Bengali, E., Romero-Ortega, M.I., 2011. Modalityspeciﬁc axonal regeneration: toward selective regenerative neural interfaces. Front.
Neuroeng. 4. https://doi.org/10.3389/fneng.2011.00011.
Lu, C.W., Patil, P.G., Chestek, C.A., 2012. Current challenges to the clinical translation ofbrain machine interface technology. International Review of Neurobiology. Elsevier,pp. 137–160. https://doi.org/10.1016/B978-0-12-404706-8.00008-5.
Chapter 3 - The nerve trunk. In: Lundborg, G. (Ed.), Nerve Injury and Repair, Second
Edition. Churchill Livingstone, Philadelphia. https://doi.org/10.1016/B978-0-44306711-2.50006-8. 27-cp2.
Luo, X., Matranga, C., Tan, S., Alba, N., Cui, X.T., 2011. Carbon nanotube nanoreservior
for controlled release of anti-inﬂammatory dexamethasone. Biomaterials 32,
6316–6323. https://doi.org/10.1016/j.biomaterials.2011.05.020.
Malagodi, M.S., Horch, K.W., Schoenberg, A.A., 1989. An intrafascicular electrode forrecording of action potentials in peripheral nerves. Ann. Biomed. Eng. 17, 397–410.
https://doi.org/10.1007/BF02368058.
Mannard, A., Stein, R.B., Charles, D., 1974. Regeneration electrode units: implants forrecording from single peripheral nerve ﬁbers in freely moving animals. Science 183,
547–549. https://doi.org/10.1126/science.183.4124.547.
Marks, A., 1969. Bullfrog nerve regeneration into porous implants. In: Presented at theAnatomical Record. Wiley-Liss Div John Wiley & Sons Inc, 605 Third Ave, New York,NY 10158-0012. pp. 226.
Marks, W.B., Loeb, G.E., 1976. Action currents, internodal potentials, and extracellular
records of myelinated mammalian nerve ﬁbers derived from node potentials.
Biophys. J. 16, 655–668. https://doi.org/10.1016/S0006-3495(76)85719-0.
Mathews, K.S., Wark, H.A.C., Normann, R.A., 2014a. Assessment of rat sciatic nerve
function following acute implantation of high density utah slanted electrode array(25 electrodes/mm 2) based on neural recordings and evoked muscle activity: nerve
Function with Multielectrode Array. Muscle Nerve 50, 417–424. https://doi.org/10.
1002/mus.24171.
Mathews, K.S., Wark, H.A.C., Warren, D.J., Christensen, M.B., Nolta, N.F., Cartwright,
P.C., Normann, R.A., 2014b. Acute monitoring of genitourinary function using intrafascicular electrodes: selective pudendal nerve activity corresponding to bladderﬁlling, bladder fullness, and genital stimulation. Urology 84, 722–729. https://doi.
org/10.1016/j.urology.2014.05.021.
McCreery, D., 2004. Apparatus and Methods for Diﬀerential Stimulation of Nerve Fibers.
US20040111139A1. .
McCreery, D.B., Agnew, W.F., Yuen, T.G.H., Bullara, L., 1990. Charge density and charge
per phase as cofactors in neural injury induced by electrical stimulation. IEEE Trans.
Biomed. Eng. 37, 996–1001. https://doi.org/10.1109/10.102812.
McCreery, D.B., Agnew, W.F., Yuen, T.G.H., Bullara, L.A., 1995. Relationship between
stimulus amplitude, stimulus frequency and neural damage during electrical stimulation of sciatic nerve of cat. Med. Biol. Eng. Comput. 33, 426–429. https://doi.org/
10.1007/BF02510526.
McNaughton, T.G., Horch, K.W., 1996. Metallized polymer ﬁbers as leadwires and intrafascicular microelectrodes. J. Neurosci. Methods 70, 103–107. https://doi.org/10.
1016/S0165-0270(96)00111-2.
McNeal, D.R., Bowman, B.R., 1985. Selective activation of muscles using peripheral nerve
electrodes. Med. Biol. Eng. Comput. 23, 249–253. https://doi.org/10.1007/
BF02446866.

tissue-engineerined electronic nerve interfaces (Teeni). In: Presented at the SolidState Sensors, Actuators and Microsystems Workshop. Hilton Head. pp. 4. https://doi.org/10.31438/trf.hh2018.13.
Kundu, A., Harreby, K.R., Yoshida, K., Boretius, T., Stieglitz, T., Jensen, W., 2014.
Stimulation selectivity of the “Thin-ﬁlm longitudinal intrafascicular electrode”(tfLIFE) and the “Transverse intrafascicular multi-channel electrode” (TIME) in thelarge nerve animal model. IEEE Trans. Neural Syst. Rehabil. Eng. 22, 400–410.
https://doi.org/10.1109/TNSRE.2013.2267936.
Lacour, S.P., Atta, R., FitzGerald, J.J., Blamire, M., Tarte, E., Fawcett, J., 2008. Polyimide
micro-channel arrays for peripheral nerve regenerative implants. Sens. Actuators
Phys. 147, 456–463. https://doi.org/10.1016/j.sna.2008.05.031.
Lago, N., Udina, E., Ramachandran, A., Navarro, X., 2007a. Neurobiological assessment ofregenerative electrodes for bidirectional interfacing injured peripheral nerves. IEEE
Trans. Biomed. Eng. 54, 1129–1137. https://doi.org/10.1109/TBME.2007.891168.
Lago, N., Yoshida, K., Koch, K.P., Navarro, X., 2007b. Assessment of biocompatibility ofchronically implanted polyimide and platinum intrafascicular electrodes. IEEE Trans.
Biomed. Eng. 54, 281–290. https://doi.org/10.1109/TBME.2006.886617.
Lancashire, H., Jiang, D., Demosthenous, A., Donaldson, N., 2018. A MicroChannel neuralinterface ASIC. In: 2018 IEEE International Symposium on Circuits and Systems(ISCAS). Presented at the 2018 IEEE International Symposium on Circuits andSystems (ISCAS). IEEE, Florence. pp. 1–4. https://doi.org/10.1109/ISCAS.2018.
8351380.
Lancashire, H.T., Jiang, D., Demosthenous, A., Donaldson, N., 2019. An ASIC for recording and stimulation in stacked microchannel neural interfaces. IEEE Trans.
Biomed. Circuits Syst. https://doi.org/10.1109/TBCAS.2019.2891284. 1–1.
Lancashire, H.T., Vanhoestenberghe, A., Pendegrass, C.J., Ajam, Y.A., Magee, E.,
Donaldson, N., Blunn, G.W., 2016. Microchannel neural interface manufacture bystacking silicone and metal foil laminae. J. Neural Eng. 13, 034001. https://doi.org/
10.1088/1741-2560/13/3/034001.
Larsen, J.O., 1998. Stereology of nerve cross sections. J. Neurosci. Methods 85, 107–118.
https://doi.org/10.1016/S0165-0270(98)00129-0.
Larsen, J.O., Thomsen, M., Haugland, M., Sinkjær, T., 1998. Degeneration and regeneration in rabbit peripheral nerve with long-term nerve cuﬀ electrode implant: a
stereological study of myelinated and unmyelinated axons. Acta Neuropathol. (Berl.)
96, 365–378. https://doi.org/10.1007/s004010050907.
Larson, C., Elyahoodayan, S., Cobo, A., Scholten, K., Song, D., Meng, E., 2018. Reliability
and acute in vivo testing of the lyse-and-attract cuﬀ electrode. In: Neuroscience
Meeting Planner. Presented at the Society for Neuroscience. San Diego, CA. pp.
700.15.
Larson, P.J., Towe, B.C., 2011. Miniature ultrasonically powered wireless nerve cuﬀ stimulator. 2011 5th International IEEE/EMBS Conference on Neural Engineering.
Presented at the 2011 5th International IEEE/EMBS Conference on NeuralEngineering 265–268. https://doi.org/10.1109/NER.2011.5910538.
Lawrence, S.M., Dhillon, G.S., Horch, K.W., 2003. Fabrication and characteristics of an
implantable, polymer-based, intrafascicular electrode. J. Neurosci. Methods 131,
9–26. https://doi.org/10.1016/S0165-0270(03)00231-0.
Lawrence, S.M., Larsen, J.O., Horch, K.W., Riso, R., Sinkjaer, T., 2002. Long-term biocompatibility of implanted polymer-based intrafascicular electrodes: implanted
Polymer-Based Intrafascicular Electrodes. J. Biomed. Mater. Res. 63, 501–506.
https://doi.org/10.1002/jbm.10303.
Lebedev, M.A., Nicolelis, M.A.L., 2006. Brain–machine interfaces: past, present and future. Trends Neurosci. 29, 536–546. https://doi.org/10.1016/j.tins.2006.07.004.
Lecomte, A., Degache, A., Descamps, E., Dahan, L., Bergaud, C., 2017. In vitro and in vivo
biostability assessment of chronically-implanted Parylene C neural sensors. Sens.
Actuators B Chem. 251, 1001–1008. https://doi.org/10.1016/j.snb.2017.05.057.
Lee, B., Koripalli, M.K., Jia, Y., Acosta, J., Sendi, M.S.E., Choi, Y., Ghovanloo, M., 2018a.
An implantable peripheral nerve recording and stimulation system for experiments
on freely moving animal subjects. Sci. Rep. 8. https://doi.org/10.1038/s41598-01824465-1.
Lee, S., Peh, W.Y.X., Ho, J.S., Thakor, N.V., Yen, S.-C., Lee, C., 2018b. Batteryless pelvic
nerve direct modulation for bladder voding using an active neural clip. In:
Proceedings of the 13th Annual IEEE International Conference on Nano/MicroEngineered and Molecular Systems. Singapore. pp. 4.
Lee, S., Sheshadri, S., Xiang, Z., Delgado-Martinez, I., Xue, N., Sun, T., Thakor, N.V., Yen,
S.-C., Lee, C., 2017. Selective stimulation and neural recording on peripheral nervesusing ﬂexible split ring electrodes. Sens. Actuators B Chem. 242, 1165–1170. https://doi.org/10.1016/j.snb.2016.09.127.
Lee, Y.J., Kim, H.-J., Do, S.H., Kang, J.Y., Lee, S.H., 2016a. Characterization of nerve-cuﬀ
electrode interface for biocompatible and chronic stimulating application. Sens.
Actuators B Chem. 237, 924–934. https://doi.org/10.1016/j.snb.2016.06.169.
Lee, Y.J., Park, S.J., Yun, K.-S., Kang, J.Y., Lee, S.H., 2016b. Enzymeless glucose sensor
integrated with chronically implantable nerve cuﬀ electrode for in-situ inﬂammation
monitoring. Sens. Actuators B Chem. 222, 425–432. https://doi.org/10.1016/j.snb.
2015.08.091.
Lefurge, T., Goodall, E., Horch, K., Stensaas, L., Schoenberg, A., 1991. Chronically implanted intrafascicular recording electrodes. Ann. Biomed. Eng. 19, 197–207. https://doi.org/10.1007/BF02368469.
Lertmanorat, Z., Durand, D.M., 2004. A novel electrode array for diameter-dependent
control of axonal excitability: a simulation study. IEEE Trans. Biomed. Eng. 51,
1242–1250. https://doi.org/10.1109/TBME.2004.827347.
Lertmanorat, Z., Gustafson, K.J., Durand, D.M., 2006. Electrode array for reversing therecruitment order of peripheral nerve stimulation: experimental studies. Ann.
Biomed. Eng. 34, 152–160. https://doi.org/10.1007/s10439-005-9012-5.
Lertmanorat, Z., Montague, F.W., Durand, D.M., 2009. A ﬂat interface nerve electrodewith integrated multiplexer. IEEE Trans. Neural Syst. Rehabil. Eng. 17, 176–182.
https://doi.org/10.1109/TNSRE.2008.2009307.

31

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng
Mendell, L.M., Henneman, E., 1971. Terminals of single Ia ﬁbers: location, density, anddistribution within a pool of 300 homonymous motoneurons. J. Neurophysiol. 34,
171–187. https://doi.org/10.1152/jn.1971.34.1.171.
Mensinger, A.F., Anderson, D.J., Buchko, C.J., Johnson, M.A., Martin, D.C., Tresco, P.A.,
Silver, R.B., Highstein, S.M., 2000. Chronic recording of regenerating VIIIth nerveaxons with a sieve electrode. J. Neurophysiol. 83, 611–615. https://doi.org/10.1152/
jn.2000.83.1.611.
Metcalfe, B., Chew, D., Clarke, C., Donaldson, N., Taylor, J., 2014. An enhancement tovelocity selective discrimination of neural recordings: extraction of neuronal ﬁring
rates. In: 2014 36th Annual International Conference of the IEEE Engineering in
Medicine and Biology Society. Presented at the 2014 36th Annual International
Conference of the IEEE Engineering in Medicine and Biology Society (EMBC). IEEE,
Chicago, IL. pp. 4111–4114. https://doi.org/10.1109/EMBC.2014.6944528.
Metcalfe, B., Nielsen, T.N., Donaldson, N., de, N., Hunter, A.J., Taylor, J.T., 2018. First
demonstration of velocity selective recording from the pig vagus using a nerve cuﬀ
shows respiration aﬀerents. Biomed. Eng. Lett. 8, 127–136. https://doi.org/10.1007/
s13534-017-0054-z.
Meyer, J.-, Stieglitz, T., Scholz, O., Haberer, W., Beutel, H., 2001. High density interconnects and ﬂexible hybrid assemblies for active biomedical implants. IEEE Trans.
Adv. Packag. 24, 366–374. https://doi.org/10.1109/6040.938305.
Micera, S., Carrozza, M.C., Beccai, L., Vecchi, F., Dario, P., 2006. Hybrid bionic systems
for the replacement of hand function. Proc. IEEE 94, 1752–1762. https://doi.org/10.
1109/JPROC.2006.881294.
Micera, S., Citi, L., Rigosa, J., Carpaneto, J., Raspopovic, S., Di Pino, G., Rossini, L.,
Yoshida, K., Denaro, L., Dario, P., Rossini, P.M., 2010. Decoding information fromneural signals recorded using intraneural electrodes: toward the development of aneurocontrolled hand prosthesis. Proc. IEEE 98, 407–417. https://doi.org/10.1109/
JPROC.2009.2038726.
Micera, S., Navarro, X., 2009. Chapter 2 bidirectional interfaces with the peripheral
nervous system. International Review of Neurobiology. Elsevier, pp. 23–38. https://doi.org/10.1016/S0074-7742(09)86002-9.
Michoud, F., Sottas, L., Browne, L.E., Asboth, L., Latremoliere, A., Sakuma, M., Courtine,G., Woolf, C.J., Lacour, S.P., 2018. Optical cuﬀ for optogenetic control of the peripheral nervous system. J. Neural Eng. 15, 015002. https://doi.org/10.1088/17412552/aa9126.
Micro-Leads Research Products [WWW Document], n.d. Wirel. Nerve Cuﬀ Electrodes U.
S. Micro-Leads. URL https://www.micro-leads.com/research (Accessed 4.21.19).
Millesi, H., Zöch, G., Reihsner, R., 1995. Mechanical properties of peripheral nerves. Clin.
Orthop. 76–83.
Mortimer, J.T., 1990. Electrical excitation of nerve. In: Agnew, W.F., McCreery, D.B.
(Eds.), Neural Prostheses: Fundamental Studies, Prentice Hall Biophysics andBioengineering Series. Prentice Hall, Englewood Cliﬀs, N.J, pp. 67–84.
Mortimer, J.T., Bhadra, N., 2018. Fundamentals of electrical stimulation.
Neuromodulation. Elsevier, pp. 71–82. https://doi.org/10.1016/B978-0-12-8053539.00006-1.
Mortimer, J.T., Bhadra, N., 2004. Peripheral nerve and muscle stimulation, in: neuroprosthetics: theory and practice, series on bioengineering and biomedical engineering. World Sci. 638–682. https://doi.org/10.1142/9789812561763_0020.
Mortimer, T., Agnew, W.F., Horch, K., Citron, P., Creasey, G., Kantor, C., 1995.
Perspectives on new electrode technology for stimulating peripheral nerves withimplantable motor prostheses. IEEE Trans. Rehabil. Eng. 3, 145–154. https://doi.org/
10.1109/86.392373.
Musick, K.M., Rigosa, J., Narasimhan, S., Wurth, S., Capogrosso, M., Chew, D.J., Fawcett,
J.W., Micera, S., Lacour, S.P., 2015. Chronic multichannel neural recordings from soft
regenerative microchannel electrodes during gait. Sci. Rep. 5. https://doi.org/10.
1038/srep14363.
Naples, G.G., Mortimer, J.T., Scheiner, A., Sweeney, J.D., 1988. A spiral nerve cuﬀ
electrode for peripheral nerve stimulation. IEEE Trans. Biomed. Eng. 35, 905–916.
https://doi.org/10.1109/10.8670.
Naples, G.G., Mortimer, J.T., Yuen, T.G.H., 1990. Overview of peripheral nerve electrodedesign and implantation. In: Agnew, W.F., McCreery, D.B. (Eds.), Neural Prostheses:
Fundamental Studies, Prentice Hall Biophysics and Bioengineering Series. Prentice
Hall, Englewood Cliﬀs, N.J, pp. 85–106.
Naples, G.G., Sweeney, J.D., Mortimer, J.T., 1986. Implantable Cuﬀ, Method of
Manufacture, and Method of Installation. US4602624. .
Navarro, X., Krueger, T.B., Lago, N., Micera, S., Stieglitz, T., Dario, P., 2005. A criticalreview of interfaces with the peripheral nervous system for the control of neuroprostheses and hybrid bionic systems. J. Peripher. Nerv. Syst. 10, 229–258. https://doi.org/10.1111/j.1085-9489.2005.10303.x.
Navarro, X., Lago, N., Vivo, M., Yoshida, K., Koch, K.P., Poppendieck, W., Micera, S.,
2007. Neurobiological evaluation of thin-ﬁlm longitudinal intrafascicular electrodes
as a peripheral nerve interface. In: 2007 IEEE 10th International Conference on
Rehabilitation Robotics. Presented at the 2007 IEEE 10th International Conference on
Rehabilitation Robotics. IEEE, Noordwijk, Netherlands. pp. 643–649. https://doi.
org/10.1109/ICORR.2007.4428492.
Nedic, A., Moon, J.D., Kung, T.A., Langhals, N.B., Cederna, P.S., Urbanchek, M.G., 2013.
Von frey monoﬁlament testing successfully discriminates between sensory function of
mixed nerve and sensory nerve regenerative peripheral nerve interfaces. In: 2013 6thInternational IEEE/EMBS Conference on Neural Engineering (NER). Presented at the2013 6th International IEEE/EMBS Conference on Neural Engineering (NER). IEEE,
San Diego, CA, USA. pp. 255–258. https://doi.org/10.1109/NER.2013.6695920.
Neural Cuﬀ | Ardiem Medical, (2019) n.d. URL http://www.ardiemmedical.com/neuralcuﬀ/ (Accessed 4.25.19).
Nielsen, T.N., Sevcencu, C., Struijk, J.J., 2014. Comparison of mono-, Bi-, and tripolar
conﬁgurations for stimulation and recording with an interfascicular interface. IEEE
Trans. Neural Syst. Rehabil. Eng. 22, 88–95. https://doi.org/10.1109/TNSRE.2013.

2278479.
Nielsen, T.N., Sevcencu, C., Struijk, J.J., 2012. Fascicle-selectivity of an intraneural stimulation electrode in the rabbit sciatic nerve. IEEE Trans. Biomed. Eng. 59, 192–197.
https://doi.org/10.1109/TBME.2011.2169671.
Nordhausen, C.T., Maynard, E.M., Normann, R.A., 1996. Single unit recording capabilities of a 100 microelectrode array. Brain Res. 12.
Oddo, C.M., Raspopovic, S., Artoni, F., Mazzoni, A., Spigler, G., Petrini, F.,
Giambattistelli, F., Vecchio, F., Miraglia, F., Zollo, L., Di Pino, G., Camboni, D.,
Carrozza, M.C., Guglielmelli, E., Rossini, P.M., Faraguna, U., Micera, S., 2016.
Intraneural stimulation elicits discrimination of textural features by artiﬁcial ﬁngertip in intact and amputee humans. eLife 5. https://doi.org/10.7554/eLife.09148.
Ong, X.C., Huang, W.-C., Kwon, I.S., Gopinath, C., Wu, H., Fisher, L.E., Gaunt, R.A.,
Bettinger, C.J., Fedder, G.K., 2018. Ultra-compliant peripheral nerve cuﬀ electrodewith hydrogel adhesion. In: 2018 IEEE Micro Electro Mechanical Systems (MEMS).
Presented at the 2018 IEEE Micro Electro Mechanical Systems (MEMS). IEEE, Belfast.
pp. 376–379. https://doi.org/10.1109/MEMSYS.2018.8346566.
Ordonez, J.S., Pikov, V., Wiggins, H., Patten, C., Stieglitz, T., Rickert, J., Schuettler, M.,
2014. Cuﬀ electrodes for very small diameter nerves — Prototyping and ﬁrst recordings in vivo. IEEE 6846–6849. https://doi.org/10.1109/EMBC.2014.6945201.
Ortigoza-Diaz, J., Scholten, K., Larson, C., Cobo, A., Hudson, T., Yoo, J., Baldwin, A.,
Weltman Hirschberg, A., Meng, E., 2018. Techniques and considerations in the microfabrication of parylene C microelectromechanical systems. Micromachines 9, 422.
https://doi.org/10.3390/mi9090422.
Ortiz-Catalan, M., Brånemark, R., Håkansson, B., Delbeke, J., 2012. On the viability of
implantable electrodes for the natural control of artiﬁcial limbs: review and discussion. Biomed. Eng. Online 11, 33. https://doi.org/10.1186/1475-925X-11-33.
Page, D.M., George, J.A., Kluger, D.T., Duncan, C., Wendelken, S., Davis, T., Hutchinson,
D.T., Clark, G.A., 2018. Motor control and sensory feedback enhance prosthesis
embodiment and reduce phantom pain after long-term hand amputation. Front. Hum.
Neurosci. 12. https://doi.org/10.3389/fnhum.2018.00352.
Park, H., Choi, W., Jung, W., Kim, O., Chu, J.-U., Son, D., Park, W., Kim, J., 2018.
Towards the development of bidirectional peripheral nerve interface for long-term
implantation. In: Proceedings of the 2018 IEEE International Conference on Cyborg
and Bionic Systems. Presented at the International Conference on Cyborg and Bionic
Systems. Shenzhen, China. pp. 4.
Park, S.I., Brenner, D.S., Shin, G., Morgan, C.D., Copits, B.A., Chung, H.U., Pullen, M.Y.,
Noh, K.N., Davidson, S., Oh, S.J., Yoon, J., Jang, K.-I., Samineni, V.K., Norman, M.,
Grajales-Reyes, J.G., Vogt, S.K., Sundaram, S.S., Wilson, K.M., Ha, J.S., Xu, R., Pan,
T., Kim, T., Huang, Y., Montana, M.C., Golden, J.P., Bruchas, M.R., Gereau Iv, R.W.,
Rogers, J.A., 2015a. Soft, stretchable, fully implantable miniaturized optoelectronicsystems for wireless optogenetics. Nat. Biotechnol. 33, 1280–1286. https://doi.org/
10.1038/nbt.3415.
Park, S.J., Lee, Y.J., Heo, D.N., Kwon, I.K., Yun, K.-S., Kang, J.Y., Lee, S.H., 2015b.
Functional nerve cuﬀ electrode with controllable anti-inﬂammatory drug loading andrelease by biodegradable nanoﬁbers and hydrogel deposition. Sens. Actuators B
Chem. 215, 133–141. https://doi.org/10.1016/j.snb.2015.03.036.
Pasluosta, C., Kiele, P., Stieglitz, T., 2018. Paradigms for restoration of somatosensory
feedback via stimulation of the peripheral nervous system. Clin. Neurophysiol. 129,
851–862. https://doi.org/10.1016/j.clinph.2017.12.027.
Payne, S.C., Burns, O., Stebbing, M., Thomas, R., Silva, Ade, Sedo, A., Weissenborn, F.,
Hyakumura, T., Huynh, M., May, C.N., Williams, R.A., Furness, J.B., Fallon, J.B.,
Shepherd, R.K., 2019. Vagus nerve stimulation to treat inﬂammatory bowel disease: achronic, preclinical safety study in sheep. Bioelectron. Med. https://doi.org/10.
2217/bem-2018-0011.
Pederson, D.J., Quinkert, C.J., Arafat, M.A., Somann, J.P., Williams, J.D., Bercich, R.A.,
Wang, Z., Albors, G.O., Jeﬀerys, J.G.R., Irazoqui, P.P., 2019. The bionode: a closedloop neuromodulation implant. ACM Trans Embed Comput Syst 18 (9), 1–9. https://doi.org/10.1145/3301310. 20.
Pena, A.E., Kuntaegowdanahalli, S.S., Abbas, J.J., Patrick, J., Horch, K.W., Jung, R., 2017.
Mechanical fatigue resistance of an implantable branched lead system for a distributed set of longitudinal intrafascicular electrodes. J. Neural Eng. 14, 066014.
https://doi.org/10.1088/1741-2552/aa814d.
Pﬂaum, C., Riso, R.R., Wiesspeiner, G., 1997. Performance of alternative ampliﬁer conﬁgurations for tripolar nerve cuﬀ recorded ENG. In: Proceedings of 18th Annual
International Conference of the IEEE Engineering in Medicine and Biology Society.
Presented at the 18th Annual International Conference of the IEEE Engineering in
Medicine and Biology Society. IEEE, Amsterdam, Netherlands. pp. 375–376. https://doi.org/10.1109/IEMBS.1996.657000.
Picaza, J.A., Hunter, S.E., Cannon, B.W., 1977. Pain suppression by peripheral nerve
stimulation. Stereotact. Funct. Neurosurg. 40, 223–234. https://doi.org/10.1159/
000102446.
Plachta, D.T.T., Gierthmuehlen, M., Cota, O., Espinosa, N., Boeser, F., Herrera, T.C.,
Stieglitz, T., Zentner, J., 2014. Blood pressure control with selective vagal nerve
stimulation and minimal side eﬀects. J. Neural Eng. 11, 036011. https://doi.org/10.
1088/1741-2560/11/3/036011.
Pohlmeyer, E.A., Jordon, L.R., Kim, P., Miller, L.E., 2009. A fully implanted drug delivery
system for peripheral nerve blocks in behaving animals. J. Neurosci. Methods 182,
165–171. https://doi.org/10.1016/j.jneumeth.2009.06.006.
Polasek, K.H., Hoyen, H.A., Keith, M.W., Kirsch, R.F., Tyler, D.J., 2009. Stimulation
stability and selectivity of chronically implanted multicontact nerve cuﬀ electrodes in
the human upper extremity. IEEE Trans. Neural Syst. Rehabil. Eng. 17, 428–437.
https://doi.org/10.1109/TNSRE.2009.2032603.
Pollard, B.A., 2008. New model for learning ultrasound-guided needle to target localization. Reg. Anesth. Pain Med. 33, 360–362. https://doi.org/10.1016/j.rapm.2008.
02.006.
Popovic, M.R., Popovic, D.B., Keller, T., 2002. Neuroprostheses for grasping. Neurol. Res.

32

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

stimulation artifact noise in a tripolar nerve cuﬀ electrode by application of a conductive shield layer. Med. Eng. Phys. 40, 39–46. https://doi.org/10.1016/j.
medengphy.2016.11.010.
Sadeghlo, B., 2013. Design of a Peripheral Nerve Electrode for Improved Neural
Recording of the Cervical Vagus Nerve. University of Toronto.
Sahin, M., Durand, D.M., 1997. Selective recording with a multi-contact nerve cuﬀ
electrode. In: Proceedings of 18th Annual International Conference of the IEEE
Engineering in Medicine and Biology Society. Presented at the 18th Annual
International Conference of the IEEE Engineering in Medicine and Biology Society.
IEEE, Amsterdam, Netherlands. pp. 369–370. https://doi.org/10.1109/IEMBS.1996.
656997.
Sarnoﬀ, S.J., Hardenbergh, E., Whittenberger, J.L., 1948. Electrophrenic respiration. Am.
J. Physiol. 155, 9.
Sassen, M., Zimmermann, M., 1973. Diﬀerential blocking of myelinated nerve ﬁbres bytransient depolarization. Pﬂugers Arch. Eur. J. Physiol. 341, 179–195. https://doi.
org/10.1007/BF00592788.
Schiefer, M.A., Freeberg, M., Pinault, G.J.C., Anderson, J., Hoyen, H., Tyler, D.J., Triolo,
R.J., 2013. Selective activation of the human tibial and common peroneal nerves witha ﬂat interface nerve electrode. J. Neural Eng. 10, 056006. https://doi.org/10.1088/
1741-2560/10/5/056006.
Schiefer, M.A., Polasek, K.H., Triolo, R.J., Pinault, G.C.J., Tyler, D.J., 2010. Selective
stimulation of the human femoral nerve with a ﬂat interface nerve electrode. J.
Neural Eng. 7, 026006. https://doi.org/10.1088/1741-2560/7/2/026006.
Schiefer, M.A., Triolo, R.J., Tyler, D.J., 2008. A model of selective activation of the femoral nerve with a ﬂat interface nerve electrode for a lower extremity neuroprosthesis. IEEE Trans. Neural Syst. Rehabil. Eng. 16, 195–204. https://doi.org/10.
1109/TNSRE.2008.918425.
Schiefer, M.A., Tyler, D.J., Triolo, R.J., 2012. Probabilistic modeling of selective stimulation of the human sciatic nerve with a ﬂat interface nerve electrode. J. Comput.
Neurosci. 33, 179–190. https://doi.org/10.1007/s10827-011-0381-5.
Schmalbruch, H., 1986. Fiber composition of the rat sciatic nerve. Anat. Rec. 215, 71–81.
https://doi.org/10.1002/ar.1092150111.
Schmidt, R.A., Bruschini, H., Tanagho, E.A., 1978. Feasibility of inducing micturition
through chronic stimulation of sacral roots. Urology 12, 471–477. https://doi.org/
10.1016/0090-4295(78)90309-6.
Scholten, K., Meng, E., 2015. Materials for microfabricated implantable devices: a review.
Lab Chip 15, 4256–4272. https://doi.org/10.1039/C5LC00809C.
Schuettler, M., Donaldson, N., Seetohul, V., Taylor, J., 2013. Fibre-selective recordingfrom the peripheral nerves of frogs using a multi-electrode cuﬀ. J. Neural Eng. 10,036016. https://doi.org/10.1088/1741-2560/10/3/036016.
Schuettler, M., Henle, C., Ordonez, J.S., Meier, W., Guenther, T., Stieglitz, T., 2008.
Interconnection technologies for laser-patterned electrode arrays. In: 2008 30th
Annual International Conference of the IEEE Engineering in Medicine and Biology
Society. Presented at the 2008 30th Annual International Conference of the IEEE
Engineering in Medicine and Biology Society. IEEE, Vancouver, BC. pp. 3212–3215.
https://doi.org/10.1109/IEMBS.2008.4649887.
Schuettler, M., Koch, K.P., Stieglitz, T., Scholz, O., Haberer, W., Keller, R., Meyer, J.-U.,
2000. Multichannel neural cuﬀ electrodes with integrated multiplexer circuit. In: 1st
Annual International IEEE-EMBS Special Topic Conference on Microtechnologies in
Medicine and Biology. Proceedings (Cat. No.00EX451). Presented at the 1st Annual
International IEEE-EMBS Special Topic Conference on Microtechnologies in Medicine
and Biology. Proceedings, IEEE, Lyon, France. pp. 624–629. https://doi.org/10.
1109/MMB.2000.893861.
Schuettler, M., Schroeer, S., Ordonez, J.S., Stieglitz, T., 2011. Laser-fabrication of peripheral nerve cuﬀ electrodes with integrated microﬂuidic channels. In: 2011 5thInternational IEEE/EMBS Conference on Neural Engineering. Presented at the 5thInternational IEEE/EMBS Conference on Neural Engineering (NER 2011). IEEE,
Cancun. pp. 245–248. https://doi.org/10.1109/NER.2011.5910533.
Schultz, A.E., Kuiken, T.A., 2011. Neural interfaces for control of upper limb prostheses:
the state of the art and future possibilities. PMR 3, 55–67. https://doi.org/10.1016/j.
pmrj.2010.06.016.
Schultz, S., 2007. Signal-to-noise ratio in neuroscience. Scholarpedia 2, 2046. https://doi.
org/10.4249/scholarpedia.2046.
Schwartz, A.R., Bennett, M.L., Smith, P.L., Backer, W.D., Hedner, J., Boudewyns, A., de
Heyning, P.V., Ejnell, H., Hochban, W., Knaack, L., Podszus, T., Penzel, T., Peter, J.H.,
Goding, G.S., Erickson, D.J., Testerman, R., Ottenhoﬀ, F., Eisele, D.W., 2001.
Therapeutic electrical stimulation of the hypoglossal nerve in obstructive sleep
apnea. Arch. Otolaryngol. Head Neck Surg. 127, 8.
Seifert, J.L., Desai, V., Watson, R.C., Musa, T., Kim, Y., Keefer, E.W., Romero, M.I., 2012.
Normal molecular repair mechanisms in regenerative peripheral nerve interfaces
allow recording of early spike activity despite immature myelination. IEEE Trans.
Neural Syst. Rehabil. Eng. 20, 220–227. https://doi.org/10.1109/TNSRE.2011.
2179811.
Seki, Y., Yamagiwa, S., Morikawa, Y., Sawahata, H., Numano, R., Ishida, M., Kawano, T.,
2017. Hook and loop microfastener: ﬂexible microelectrodes tied to a nerve. In: 2017
IEEE 30th International Conference on Micro Electro Mechanical Systems (MEMS).
Presented at the 2017 IEEE 30th International Conference on Micro Electro
Mechanical Systems (MEMS). IEEE, Las Vegas, NV, USA. pp. 117–120. https://doi.
org/10.1109/MEMSYS.2017.7863354.
SetPoint Medical [WWW Document], n.d. SetPoint Med. URL https://setpointmedical.
com/ (Accessed 4.21.19).
Shannon, R.V., 1992. A model of safe levels for electrical stimulation. IEEE Trans.
Biomed. Eng. 39, 424–426. https://doi.org/10.1109/10.126616.
Sharma, A., Rieth, L., Tathireddy, P., Harrison, R., Oppermann, H., Klein, M., Töpper, M.,
Jung, E., Normann, R., Clark, G., Solzbacher, F., 2011. Long term in vitro functional
stability and recording longevity of fully integrated wireless neural interfaces based

24, 443–452. https://doi.org/10.1179/016164102101200311.
Radovsky, A.S., Van Vleet, J.F., 1989. Eﬀects of dexamethasone elution on tissue reaction
around stimulating electrodes of endocardial pacing leads in dogs. Am. Heart J. 117,
1288–1298. https://doi.org/10.1016/0002-8703(89)90408-0.
Rahal, M., Taylor, J., Donaldson, N., 2000a. The eﬀect of nerve cuﬀ geometry on interference reduction: a study by computer modeling. IEEE Trans. Biomed. Eng. 47,
136–138. https://doi.org/10.1109/10.817629.
Rahal, M., Winter, J., Taylor, J., Donaldson, N., 2000b. An improved conﬁguration for the
reduction of EMG in electrode cuﬀ recordings: a theoretical approach. IEEE Trans.
Biomed. Eng. 47, 1281–1284. https://doi.org/10.1109/10.867963.
Raimondo, S., Fornaro, M., Di Scipio, F., Ronchi, G., Giacobini‐Robecchi, M.G., Geuna, S.,
2009. Chapter 5 Methods and protocols in peripheral nerve regeneration experimental research: part ii—morphological techniques. International Review of
Neurobiology. Academic Press, pp. 81–103. https://doi.org/10.1016/S00747742(09)87005-0.
Ramachandran, A., Schuettler, M., Lago, N., Doerge, T., Koch, K.P., Navarro, X.,
Hoﬀmann, K.-P., Stieglitz, T., 2006. Design, in vitro and in vivo assessment of a multichannel sieve electrode with integrated multiplexer. J. Neural Eng. 3, 114–124.
https://doi.org/10.1088/1741-2560/3/2/005.
Rathbun, K.M., Brader, W.T., Norbury, J.W., 2018. A simple, realistic, inexpensive nerve
phantom. J. Ultrasound Med. https://doi.org/10.1002/jum.14905.
Rattay, F., 1986. Analysis of models for external stimulation of axons. IEEE Trans.
Biomed. Eng. BME-33, 974–977. https://doi.org/10.1109/TBME.1986.325670.
Reichling, T.D., German, R.Z., 2000. Bones, muscles and visceral organs of protein-malnourished rats (Rattus norvegicus) grow more slowly but for longer durations toreach normal ﬁnal size. J. Nutr. 130, 2326–2332. https://doi.org/10.1093/jn/130.9.
2326.
Restaino, S.M., Abliz, E., Wachrathit, K., Krauthamer, V., Shah, S.B., 2014. Biomechanicaland functional variation in rat sciatic nerve following cuﬀ electrode implantation. J.
NeuroEng. Rehabil. 11, 73. https://doi.org/10.1186/1743-0003-11-73.
Rise, M., Klepinski, R.J., 1992. Tool for Implantable Neural Electrode. US5143067. .
Riso, R.R., Mosallaie, F.K., Jensen, W., Sinkjaer, T., 2000. Nerve cuﬀ recordings of muscle
aﬀerent activity from tibial and peroneal nerves in rabbit during passive ankle motion. IEEE Trans. Rehabil. Eng. 8, 244–258. https://doi.org/10.1109/86.847826.
Robblee, L.S., Rose, T.L., 1990. Electrochemical guidelines for selection of protocols andelectrode materials for neural stimulation. In: Agnew, W.F., McCreery, D.B. (Eds.),
Neural Prostheses: Fundamental Studies, Prentice Hall Biophysics and Bioengineering
Series. Prentice Hall, Englewood Cliﬀs, N.J, pp. 25–66.
Rodríguez, F.J., Ceballos, D., Schu¨ttler, M., Valero, A., Valderrama, E., Stieglitz, T.,
Navarro, X., 2000. Polyimide cuﬀ electrodes for peripheral nerve stimulation. J.
Neurosci. Methods 98, 105–118. https://doi.org/10.1016/S0165-0270(00)00192-8.
Rognini, G., Petrini, F.M., Raspopovic, S., Valle, G., Granata, G., Strauss, I., Solcà, M.,
Bello-Ruiz, J., Herbelin, B., Mange, R., D’Anna, E., Di Iorio, R., Di Pino, G., Andreu,D., Guiraud, D., Stieglitz, T., Rossini, P.M., Serino, A., Micera, S., Blanke, O., 2018.
Multisensory bionic limb to achieve prosthesis embodiment and reduce distorted
phantom limb perceptions. J. Neurol. Neurosurg. Psychiatry. https://doi.org/10.
1136/jnnp-2018-318570. jnnp-2018-318570.
Romero, E., Denef, J.F., Delbeke, J., Robert, A., Veraart, C., 2001. Neural morphologicaleﬀects of long-term implantation of the self-sizing spiral cuﬀ nerve electrode. Med.
Biol. Eng. Comput. 39, 90–100. https://doi.org/10.1007/BF02345271.
Rose, T.L., Robblee, L.S., 1990. Electrical stimulation with Pt electrodes. VIII.
Electrochemically safe charge injection limits with 0.2 ms pulses (neuronal application). IEEE Trans. Biomed. Eng. 37, 1118–1120. https://doi.org/10.1109/10.61038.
Rossini, P.M., Micera, S., Benvenuto, A., Carpaneto, J., Cavallo, G., Citi, L., Cipriani, C.,
Denaro, L., Denaro, V., Di Pino, G., Ferreri, F., Guglielmelli, E., Hoﬀmann, K.-P.,
Raspopovic, S., Rigosa, J., Rossini, L., Tombini, M., Dario, P., 2010. Double nerve
intraneural interface implant on a human amputee for robotic hand control. Clin.
Neurophysiol. 121, 777–783. https://doi.org/10.1016/j.clinph.2010.01.001.
Rozman, J., Pečlin, P., Ribarič, S., Godec, M., Burja, J., 2018. An improved method ofcrafting a multi-electrode spiral cuﬀ for the selective stimulation of peripheral nerves.
Sci. Rep. 8. https://doi.org/10.1038/s41598-018-19318-w.
Rubehn, B., Stieglitz, T., 2010. In vitro evaluation of the long-term stability of polyimide
as a material for neural implants. Biomaterials 31, 3449–3458. https://doi.org/10.
1016/j.biomaterials.2010.01.053.
Rush, A.J., George, M.S., Sackeim, H.A., Marangell, L.B., Husain, M.M., Giller, C., Nahas,Z., Haines, S., Simpson, R.K., Goodman, R., 2000. Vagus nerve stimulation (VNS) for
treatment-resistant depressions: a multicenter study. Biol. Psychiatry 47, 276–286.
https://doi.org/10.1016/S0006-3223(99)00304-2.
Russell, C., Roche, A.D., Chakrabarty, S., 2019. Peripheral nerve bionic interface: a review of electrodes. Int. J. Intell. Robot. Appl. https://doi.org/10.1007/s41315-01900086-3.
Rutten, W.L.C., 2002. Selective electrical interfaces with the nervous system. Annu. Rev.
Biomed. Eng. 4, 407–452. https://doi.org/10.1146/annurev.bioeng.4.020702.
153427.
Rydevik, B., Lundborg, G., Bagge, U., 1981. Eﬀects of graded compression on intraneuralblood ﬂow. J. Hand Surg. 6, 3–12. https://doi.org/10.1016/S0363-5023(81)80003-2.
Rydevik, B.L., Danielsen, N., Dahlin, L.B., Lundborg, G., 1990. Pathophysiology of peripheral nerve injury with special reference to electrode implantation. In: Agnew,
W.F., McCreery, D.B. (Eds.), Neural Prostheses: Fundamental Studies, Prentice Hall
Biophysics and Bioengineering Series. Prentice Hall, Englewood Cliﬀs, N.J, pp.
85–106.
Saal, H.P., Bensmaia, S.J., 2015. Biomimetic approaches to bionic touch through a peripheral nerve interface. Neuropsychologia 79, 344–353. https://doi.org/10.1016/j.
neuropsychologia.2015.06.010.
Sabetian, P., Sadeghlo, B., Zhang, C.H., Yoo, P.B., 2017. Characterizing the reduction of33

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

Trans. Neural Syst. Rehabil. Eng. 11, 427–437. https://doi.org/10.1109/TNSRE.
2003.819793.
Stolinski, C., 1995. Structure and composition of the outer connective tissue sheaths ofperipheral nerve. J. Anat. 186 (Pt. 1), 123–130.
Stoyanova, I.I., van Wezel, R.J.A., Rutten, W.L.C., 2013. In vivo testing of a 3D bifurcating
microchannel scaﬀold inducing separation of regenerating axon bundles in peripheral
nerves. J. Neural Eng. 10, 066018. https://doi.org/10.1088/1741-2560/10/6/
066018.
Straka, M., Shafer, B., Vasudevan, S., Welle, C., Rieth, L., 2018. Characterizing longitudinal changes in the impedance spectra of in-vivo peripheral nerve electrodes.
Micromachines 9, 587. https://doi.org/10.3390/mi9110587.
Street, M.G., Welle, C.G., Takmakov, P.A., 2018. Automated reactive accelerated aging
for rapid in vitro evaluation of neural implant performance. Rev. Sci. Instrum. 89,
094301. https://doi.org/10.1063/1.5024686.
Stroncek, J.D., Reichert, W.M., 2008. Overview of wound healing in diﬀerent tissue types.
In: Reichert, W.M. (Ed.), Indwelling Neural Implants: Strategies for Contending with
the in Vivo Environment, Frontiers in Neuroengineering. CRC Press, Boca Raton.
Struijk, J.J., Haugland, M.K., Thomsen, M., 1997. Fascicle selective recording with anerve cuﬀ electrode. In: Proceedings of 18th Annual International Conference of theIEEE Engineering in Medicine and Biology Society. Presented at the 18th Annual
International Conference of the IEEE Engineering in Medicine and Biology Society.
IEEE, Amsterdam, Netherlands. pp. 361–362. https://doi.org/10.1109/IEMBS.1996.
656993.
Struijk, J.J., Thomsen, M., 1995. Tripolar nerve cuﬀ recording: stimulus artifact, EMG and
the recorded nerve signal. In: Proceedings of 17th International Conference of theEngineering in Medicine and Biology Society. Presented at the 17th International
Conference of the Engineering in Medicine and Biology Society. IEEE, Montreal,Que., Canada. pp. 1105–1106. https://doi.org/10.1109/IEMBS.1995.579534.
Sun, F.T., Morrell, M.J., 2014. Closed-loop neurostimulation: the clinical experience.
Neurotherapeutics 11, 553–563. https://doi.org/10.1007/s13311-014-0280-3.
Sunderland, S., 1978. Nerves and Nerve Injuries, 2d ed. Churchill Livingstone; distributed
by Longman, Edinburgh; New York: New York.
Sunderland, S., 1965. The connective tissues of peripheral nerves. Brain 88, 841–854.
https://doi.org/10.1093/brain/88.4.841.
Sunderland, S.S., 1990. The anatomy and physiology of nerve injury. Muscle Nerve 13,
771–784. https://doi.org/10.1002/mus.880130903.
Suzuki, T., Kotake, N., Mabuchi, K., Takeuchi, S., 2006. Flexible regeneration-type nerve
electrode with integrated microﬂuidic channels. In: 2006 International Conference on
Microtechnologies in Medicine and Biology. Presented at the 2006 International
Conference on Microtechnologies in Medicine and Biology. IEEE, Okinawa. pp.
303–305. https://doi.org/10.1109/MMB.2006.251557.
Sweeney, J.D., Ksienski, D.A., Mortimer, J.T., 1990. A nerve cuﬀ technique for selective
excitation of peripheral nerve trunk regions. IEEE Trans. Biomed. Eng. 37, 706–715.
https://doi.org/10.1109/10.55681.
Takmakov, P., Ruda, K., Scott Phillips, K., Isayeva, I.S., Krauthamer, V., Welle, C.G., 2015.
Rapid evaluation of the durability of cortical neural implants using accelerated aging
with reactive oxygen species. J. Neural Eng. 12, 026003. https://doi.org/10.1088/
1741-2560/12/2/026003.
Takmakov, P.A., 2017. Electrochemistry of a robust neural interface. Electrochem. Soc.
Interface 26, 49–51. https://doi.org/10.1149/2.F05173if.
Tan, D.W., Schiefer, M.A., Keith, M.W., Anderson, J.R., Tyler, D.J., 2015. Stability andselectivity of a chronic, multi-contact cuﬀ electrode for sensory stimulation in humanamputees. J. Neural Eng. 12, 026002. https://doi.org/10.1088/1741-2560/12/2/
026002.
Tarler, H., Grill, W.M., Mortimer, J.T., 1995. Comparison between monopolar and tripolar conﬁgurations in chronically implanted nerve cuﬀ electrodes. In: Proceedingsof 17th International Conference of the Engineering in Medicine and Biology Society.
Presented at the 17th International Conference of the Engineering in Medicine andBiology Society. IEEE, Montreal, Que., Canada. pp. 1093–1094. https://doi.org/10.
1109/IEMBS.1995.579515.
Tarler, M.D., Mortimer, J.T., 2004. Selective and independent activation of four motor
fascicles using a four contact nerve-cuﬀ electrode. IEEE Trans. Neural Syst. Rehabil.
Eng. 12, 251–257. https://doi.org/10.1109/TNSRE.2004.828415.
Tarler, M.D., Mortimer, J.T., 2003. Comparison of joint torque evoked with monopolar
and tripolar-cuﬀ electrodes. IEEE Trans. Neural Syst. Rehabil. Eng. 11, 227–235.
https://doi.org/10.1109/TNSRE.2003.816867.
Taylor, J., Donaldson, N., Winter, J., 2004. Multiple-electrode nerve cuﬀs for low-velocity
and velocity-selective neural recording. Med. Biol. Eng. Comput. 42, 634–643.
https://doi.org/10.1007/BF02347545.
Taylor, J., Metcalfe, B., Clarke, C., Chew, D., Nielsen, T., Donaldson, N., 2015. A summaryof current and new methods in velocity selective recording (VSR) of electroneurogram (ENG). In: 2015 IEEE Computer Society Annual Symposium on VLSI.
Presented at the 2015 IEEE Computer Society Annual Symposium on VLSI (ISVLSI).
IEEE, Montpellier, France. pp. 221–226. https://doi.org/10.1109/ISVLSI.2015.34.
Thomas, P.K., 1963. The connective tissue of peripheral nerve: an electron microscope
study. J. Anat. 97, 35–44.
Thota, A.K., Kuntaegowdanahalli, S., Starosciak, A.K., Abbas, J.J., Orbay, J., Horch, K.W.,
Jung, R., 2015. A system and method to interface with multiple groups of axons inseveral fascicles of peripheral nerves. J. Neurosci. Methods 244, 78–84. https://doi.
org/10.1016/j.jneumeth.2014.07.020.
Tian, P., Yi, W., Chen, C., Hu, J., Qi, J., Zhang, B., Cheng, M.M.-C., 2018. Flexible 3D
carbon nanotubes cuﬀ electrodes as a peripheral nerve interface. Biomed.
Microdevices 20. https://doi.org/10.1007/s10544-018-0268-6.
Tos, P., Ronchi, G., Papalia, I., Sallen, V., Legagneux, J., Geuna, S., Giacobini‐Robecchi,
M.G., 2009. Chapter 4 Methods and protocols in peripheral nerve regeneration experimental research: part I—experimental models. International Review ofon the Utah Slant Electrode Array. J. Neural Eng. 8, 045004. https://doi.org/10.
1088/1741-2560/8/4/045004.
Sharp, A.A., Fromherz, S., 2011. Optogenetic regulation of leg movement in midstage
chick embryos through peripheral nerve stimulation. J. Neurophysiol. 106,2776–2782. https://doi.org/10.1152/jn.00712.2011.
Silva, A.P.D., Jordão, C.E.R., Fazan, V.P.S., 2007. Peripheral nerve morphometry: comparison between manual and semi-automated methods in the analysis of a small
nerve. J. Neurosci. Methods 159, 153–157. https://doi.org/10.1016/j.jneumeth.
2006.06.012.
Silveira, C., Brunton, E., Spendiﬀ, S., Nazarpour, K., 2018. Inﬂuence of nerve cuﬀ channel
count and implantation site on the separability of aﬀerent ENG. J. Neural Eng. 15,046004. https://doi.org/10.1088/1741-2552/aabca0.
Sohal, H.S., Vassilevski, K., Jackson, A., Baker, S.N., O’Neill, A., 2016. Design and microfabrication considerations for reliable ﬂexible intracortical implants. Front. Mech.
Eng. China 2. https://doi.org/10.3389/fmech.2016.00005.
Somann, J.P., Albors, G.O., Neihouser, K.V., Lu, K.-H., Liu, Z., Ward, M.P., Durkes, A.,
Robinson, J.P., Powley, T.L., Irazoqui, P.P., 2018. Chronic cuﬃng of cervical vagus
nerve inhibits eﬀerent ﬁber integrity in rat model. J. Neural Eng. 15, 036018. https://doi.org/10.1088/1741-2552/aaa039.
Song, K.-I., Park, S.E., Lee, S., Kim, H., Lee, S.H., Youn, I., 2018. Compact optical nerve
cuﬀ electrode for simultaneous neural activity monitoring and optogenetic stimulation of peripheral nerves. Sci. Rep. 8. https://doi.org/10.1038/s41598-018-33695-2.
Sparks, S., Evans, D., Byars, D., 2014. A low cost, high ﬁdelity nerve block model. Crit.
Ultrasound J. 6, 12. https://doi.org/10.1186/s13089-014-0012-2.
Spearman, B.S., Desai, V.H., Mobini, S., McDermott, M.D., Graham, J.B., Otto, K.J., Judy,
J.W., Schmidt, C.E., 2018. Tissue-engineered peripheral nerve interfaces. Adv. Funct.
Mater. 28, 1701713. https://doi.org/10.1002/adfm.201701713.
Srinivasan, A., Guo, L., Bellamkonda, R.V., 2011. Regenerative microchannel electrodearray for peripheral nerve interfacing. In: 2011 5th International IEEE/EMBS
Conference on Neural Engineering. Presented at the 5th International IEEE/EMBS
Conference on Neural Engineering (NER 2011). IEEE, Cancun. pp. 253–256. https://doi.org/10.1109/NER.2011.5910535.
Srinivasan, A., Tahilramani, M., Bentley, J.T., Gore, R.K., Millard, D.C., Mukhatyar, V.J.,
Joseph, A., Haque, A.S., Stanley, G.B., English, A.W., Bellamkonda, R.V., 2015.
Microchannel-based regenerative scaﬀold for chronic peripheral nerve interfacing in
amputees. Biomaterials 41, 151–165. https://doi.org/10.1016/j.biomaterials.2014.
11.035.
Srinivasan, A., Tipton, J., Tahilramani, M., Kharbouch, A., Gaupp, E., Song, C.,
Venkataraman, P., Falcone, J., Lacour, S.P., Stanley, G.B., English, A.W.,
Bellamkonda, R.V., 2016. A regenerative microchannel device for recording multiple
single-unit action potentials in awake, ambulatory animals. Eur. J. Neurosci. 43,
474–485. https://doi.org/10.1111/ejn.13080.
Sta, M., Cappaert, N.L.M., Ramekers, D., Baas, F., Wadman, W.J., 2014. The functionaland morphological characteristics of sciatic nerve degeneration and regeneration
after crush injury in rats. J. Neurosci. Methods 222, 189–198. https://doi.org/10.
1016/j.jneumeth.2013.11.012.
Standards and Guidances for Neurological Devices, 2018. US Food Drug Adm. [WWWDocument] URL https://www.fda.gov/MedicalDevices/
ProductsandMedicalProcedures/NeurologicalDevices/ucm528789.htm (Accessed
2.2.19). .
Stein, R.B., Charles, D., Davis, L., Jhamandas, J., Mannard, A., Nichols, T.R., 1975.
Principles underlying new methods for chronic neural recording. Can. J. Neurol. Sci.
J. Can. Sci. Neurol. 2, 235–244. https://doi.org/10.1017/S0317167100020333.
Stein, R.B., Charles, D., Gordon, T., Hoﬀer, J.-A., Jhamandas, J., 1978. Impedance
properties of metal electrodes for chronic recording from mammalian nerves. IEEE
Trans. Biomed. Eng. BME-25, 532–537. https://doi.org/10.1109/TBME.1978.
326287.
Stein, R.B., Nichols, T.R., Jhamandas, J., Davis, L., Charles, D., 1977. Stable long-term
recordings from cat peripheral nerves. Brain Res. 128, 21–38. https://doi.org/10.
1016/0006-8993(77)90233-5.
Stein, R.B., Oğuztöreli, M.N., 1978. The radial decline of nerve impulses in a restricted
cylindrical extracellular space. Biol. Cybern. 28, 159–165. https://doi.org/10.1007/
BF00337137.
Stein, R.B., Weber, D.J., Aoyagi, Y., Prochazka, A., Wagenaar, J.B.M., Shoham, S.,
Normann, R.A., 2004. Coding of position by simultaneously recorded sensory neurones in the cat dorsal root ganglion: coding of dorsal root ganglion neurones. J.
Physiol. 560, 883–896. https://doi.org/10.1113/jphysiol.2004.068668.
Stieglitz, T., Beutel, H., Meyer, J.-U., 1997. A ﬂexible, light-weight multichannel sieve
electrode with integrated cables for interfacing regenerating peripheral nerves. Sens.
Actuators Phys. 60, 240–243. https://doi.org/10.1016/S0924-4247(97)01494-5.
Stieglitz, T., Ruf, H.H., Gross, M., Schuettler, M., Meyer, J.-U., 2002. A biohybrid system
to interface peripheral nerves after traumatic lesions: design of a high channel sieve
electrode. Biosens. Bioelectron. 17, 685–696. https://doi.org/10.1016/S09565663(02)00019-2.
Stieglitz, T., Schuetter, M., Koch, K.P., 2005. Implantable biomedical microsystems forneural prostheses. IEEE Eng. Med. Biol. Mag. 24, 58–65. https://doi.org/10.1109/
MEMB.2005.1511501.
Stieglitz, T., Schuettler, M., Rubehn, B., Boretius, T., Badia, J., Navarro, X., 2011.
Evaluation of polyimide as substrate material for electrodes to interface the peripheral nervous system. In: 2011 5th International IEEE/EMBS Conference on NeuralEngineering. Presented at the 5th International IEEE/EMBS Conference on NeuralEngineering (NER 2011). IEEE, Cancun. pp. 529–533. https://doi.org/10.1109/NER.
2011.5910602.
Stieglitz, T., Schuettler, M., Schneider, A., Valderrama, E., Navarro, X., 2003. Noninvasive
measurement of torque development in the rat foot: measurement setup and resultsfrom stimulation of the sciatic nerve with polyimide-based cuﬀ electrodes. IEEE

34

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

(25 electrodes/mm 2) penetrating microelectrode array for recording and stimulating
sub-millimeter neuroanatomical structures. J. Neural Eng. 10, 045003. https://doi.
org/10.1088/1741-2560/10/4/045003.
Waters, R.L., McNeal, D., Perry, J., 1975. Experimental correction of footdrop by electrical stimulation of the peroneal nerve. J. Bone Joint Surg. Am. 57, 1047–1054.
Weber, D.J., Stein, R.B., Everaert, D.G., Prochazka, A., 2007. Limb-state feedback fromensembles of simultaneously recorded dorsal root ganglion neurons. J. Neural Eng. 4,
S168–S180. https://doi.org/10.1088/1741-2560/4/3/S04.
Weinberg, S.L., 1993. Helical Nerve Electrode. US5251634. .
Wellman, S.M., Eles, J.R., Ludwig, K.A., Seymour, J.P., Michelson, N.J., McFadden, W.E.,
Vazquez, A.L., Kozai, T.D.Y., 2018. A materials roadmap to functional neural interface design. Adv. Funct. Mater. 28, 1701269. https://doi.org/10.1002/adfm.
201701269.
Wells, M., Goldstein, L., 2010. The polony phantom: a cost-eﬀective aid for teaching
emergency ultrasound procedures. Int. J. Emerg. Med. 3, 115. https://doi.org/10.
1007/s12245-009-0156-1.
Weremfo, A., Carter, P., Hibbert, D.B., Zhao, C., 2015. Investigating the interfacial
properties of electrochemically roughened platinum electrodes for neural stimulation. Langmuir 31, 2593–2599. https://doi.org/10.1021/la504876n.
White, R.L., Gross, T.J., 1974. An evaluation of the resistance to electrolysis of metals foruse in biostimulation microprobes. IEEE Trans. Biomed. Eng. BME-21, 487–490.
https://doi.org/10.1109/TBME.1974.324339.
Wieringa, P.A., Wiertz, R.W.F., de Weerd, E., Rutten, W.L.C., 2010a. Bifurcating microchannels as a scaﬀold to induce separation of regenerating neurites. J. Neural Eng. 7,016001. https://doi.org/10.1088/1741-2560/7/1/016001.
Wieringa, P.A., Wiertz, R.W.F., de Weerd, E.L., Rutten, W.L.C., 2010b. In vitro veriﬁcation of a 3-D regenerative neural interface design: examination of neurite growth andelectrical properties within a bifurcating microchannel structure. Proc. IEEE 98,
389–397. https://doi.org/10.1109/JPROC.2009.2038950.
Wodlinger, B., Durand, D., 2010. Peripheral nerve signal recording and processing for
artiﬁcial limb control. 2010 Annual International Conference of the IEEE Engineeringin Medicine and Biology. Presented at the 2010 Annual International Conference of
the IEEE Engineering in Medicine and Biology 6206–6209. https://doi.org/10.1109/
IEMBS.2010.5627735.
Wodlinger, B., Durand, D.M., 2009. Localization and recovery of peripheral neuralsources with beamforming algorithms. IEEE Trans. Neural Syst. Rehabil. Eng. 17,
461–468. https://doi.org/10.1109/TNSRE.2009.2034072.
Wodlinger, B., Dweiri, Y., Durand, D.M., 2015. Biochips. Implantable Biomedical
Microsystems. Elsevier, pp. 203–214. https://doi.org/10.1016/B978-0-323-26208-8.
00009-1.
Woloszko, J., Cahalan, P.T., Hendriks, M., 1998. Biocompatible Medical Lead.
US5741319. .
Wright, J., Maceﬁeld, V.G., van Schaik, A., Tapson, J.C., 2016. A review of control
strategies in closed-loop neuroprosthetic systems. Front. Neurosci. 10. https://doi.
org/10.3389/fnins.2016.00312.
Wurth, S., Capogrosso, M., Raspopovic, S., Gandar, J., Federici, G., Kinany, N., Cutrone,A., Piersigilli, A., Pavlova, N., Guiet, R., Taverni, G., Rigosa, J., Shkorbatova, P.,
Navarro, X., Barraud, Q., Courtine, G., Micera, S., 2017. Long-term usability and biointegration of polyimide-based intra-neural stimulating electrodes. Biomaterials 122,
114–129. https://doi.org/10.1016/j.biomaterials.2017.01.014.
Xiang, Z., Yen, S.-C., Sheshadri, S., Wang, J., Lee, S., Liu, Y.-H., Liao, L.-D., Thakor, N.V.,
Lee, C., 2016. Progress of ﬂexible electronics in neural interfacing - a self-adaptive
non-invasive neural ribbon electrode for small nerves recording. Adv. Mater. 28,
4472–4479. https://doi.org/10.1002/adma.201503423.
Yaghouby, F., Shafer, B., Asgari, S., Vasudevan, S., 2018. An experimental model forassessing long-term safety and eﬃcacy of vagus nerve stimulation, in: neuroscience
meeting planner. In: Presented at the Society for Neuroscience. San Diego, CA. p.
589.21.
Yim, S., Oh, Y.-E., Choi, W., Park, H., Jeong, J., Ihn, Y., Hwang, D., Oh, S.-R., Kim, J., Kim,
K., 2018. Preliminary results of a handheld nerve electrode insertion device. In: 2018
IEEE International Conference on Robotics and Automation (ICRA). Presented at the2018 IEEE International Conference on Robotics and Automation (ICRA). IEEE,
Brisbane, QLD. pp. 1504–1510. https://doi.org/10.1109/ICRA.2018.8460882.
Yizhar, O., Fenno, L.E., Davidson, T.J., Mogri, M., Deisseroth, K., 2011. Optogenetics inneural systems. Neuron 71, 9–34. https://doi.org/10.1016/j.neuron.2011.06.004.
Yoshida, K., Farina, D., Akay, M., Jensen, W., 2010. Multichannel intraneural and intramuscular techniques for multiunit recording and use in active prostheses. Proc.
IEEE 98, 432–449. https://doi.org/10.1109/JPROC.2009.2038613.
Yoshida, K., Hennings, K., Kammer, S., 2006. Acute performance of the thin-ﬁlm longitudinal intra-fascicular electrode. In: The First IEEE/RAS-EMBS International
Conference on Biomedical Robotics and Biomechatronics, 2006. BioRob 2006.
Presented at the The First IEEE/RAS-EMBS International Conference on Biomedical
Robotics and Biomechatronics, 2006. BioRob 2006. IEEE, Pisa, Italy. pp. 296–300.
https://doi.org/10.1109/BIOROB.2006.1639102.
Yoshida, K., Pellinen, D., Pivin, D., Rousche, P., Kipke, D., 2000. Development of the thinﬁlm longitudinal intra-fascicular electrode. In: Proceedings of the 5th Annual
Conference of the International Functional Electrical Stimulation Society. Presented
at the International Functional Electrical Stimulation Society. Denmark. pp. 4.
Yu, H., Xiong, W., Zhang, H., Wang, W., Li, Z., 2014. A parylene self-locking cuﬀ electrodefor peripheral nerve stimulation and recording. J. Microelectromech. Syst. 23,
1025–1035. https://doi.org/10.1109/JMEMS.2014.2333733.
Yuste, R. (Ed.), 2011. Imaging: a Laboratory Manual. Cold Spring Harbor Laboratory
Press, Cold Spring Harbor, N.Y.
Zabara, J., 1985. Peripheral control of hypersynchronous discharge in epilepsy.
Electroencephalogr. Clin. Neurophysiol. 61, S162. https://doi.org/10.1016/00134694(85)90626-1.

Neurobiology. Elsevier, pp. 47–79. https://doi.org/10.1016/S0074-7742(09)87004-9.
Towne, C., Montgomery, K.L., Iyer, S.M., Deisseroth, K., Delp, S.L., 2013. Optogenetic
control of targeted peripheral axons in freely moving animals. PLoS One 8, e72691.
https://doi.org/10.1371/journal.pone.0072691.
Triangle BioSystems International (2019) [WWW Document], n.d. URL https://www.
trianglebiosystems.com/index.html (Accessed 3.29.19).
Tyler, D.J., 2018. Electrodes for the neural interface. Neuromodulation. Elsevier, pp.
239–274. https://doi.org/10.1016/B978-0-12-805353-9.00020-6.
Tyler, D.J., Durand, D.M., 2003. Chronic response of the rat sciatic nerve to the ﬂat interface nerve electrode. Ann. Biomed. Eng. 31, 633–642. https://doi.org/10.1114/1.
1569263.
Tyler, D.J., Durand, D.M., 2002. Functionally selective peripheral nerve stimulation witha ﬂat interface nerve electrode. IEEE Trans. Neural Syst. Rehabil. Eng. 10, 294–303.
https://doi.org/10.1109/TNSRE.2002.806840.
Tyler, D.J., Durand, D.M., 1997a. Alteration of neural geometry for selective nerve stimulation. In: Proceedings of the 19th Annual International Conference of the IEEE
Engineering in Medicine and Biology Society. “Magniﬁcent Milestones and Emerging
Opportunities in Medical Engineering” (Cat. No.97CH36136). Presented at the 19th
Annual International Conference of the IEEE Engineering in Medicine and Biology
Society. “Magniﬁcent Milestones and Emerging Opportunities in Medical
Engineering”. IEEE, Chicago, IL, USA. pp. 2002–2003. https://doi.org/10.1109/
IEMBS.1997.758736.
Tyler, D.J., Durand, D.M., 1997b. A slowly penetrating interfascicular nerve electrode forselective activation of peripheral nerves. IEEE Trans. Rehabil. Eng. 5, 51–61. https://doi.org/10.1109/86.559349.
Tyler, D.J., Peterson, E.J., Brill, N., White, K., 2011. Increased selectivity of clinicalperipheral nerve interfaces. In: 2011 5th International IEEE/EMBS Conference on
Neural Engineering. Presented at the 5th International IEEE/EMBS Conference on
Neural Engineering (NER 2011). IEEE, Cancun. pp. 257–260. https://doi.org/10.
1109/NER.2011.5910536.
Valle, G., Mazzoni, A., Iberite, F., D’Anna, E., Strauss, I., Granata, G., Controzzi, M.,
Clemente, F., Rognini, G., Cipriani, C., Stieglitz, T., Petrini, F.M., Rossini, P.M.,
Micera, S., 2018a. Biomimetic intraneural sensory feedback enhances sensation
naturalness, tactile sensitivity, and manual dexterity in a bidirectional prosthesis.
Neuron 100https://doi.org/10.1016/j.neuron.2018.08.033. 37-45.e7.
Valle, G., Petrini, F.M., Strauss, I., Iberite, F., D’Anna, E., Granata, G., Controzzi, M.,
Cipriani, C., Stieglitz, T., Rossini, P.M., Mazzoni, A., Raspopovic, S., Micera, S.,
2018b. Comparison of linear frequency and amplitude modulation for intraneuralsensory feedback in bidirectional hand prostheses. Sci. Rep. 8. https://doi.org/10.
1038/s41598-018-34910-w.
Varejão, A.S.P., Meek, M.F., Ferreira, A.J.A., Patrı́cio, J.A.B., Cabrita, A.M.S., 2001.
Functional evaluation of peripheral nerve regeneration in the rat: walking track
analysis. J. Neurosci. Methods 108, 1–9. https://doi.org/10.1016/S0165-0270(01)00378-8.
Vasudevan, S., Kumsa, D., Takmakov, P., Welle, C.G., Hammer, D.X., 2016. Real time
imaging of peripheral nerve vasculature using optical coherence angiography. In:
Madsen, S.J., Yang, V.X.D., Jansen, E.D., Luo, Q., Mohanty, S.K., Thakor, N.V. (Eds.),
Presented at the SPIE BiOS. San Francisco, California, United States. pp. 96901D.
https://doi.org/10.1117/12.2213160.
Vasudevan, S., Patel, K., Welle, C., 2017. Rodent model for assessing the long term safety
and performance of peripheral nerve recording electrodes. J. Neural Eng. 14, 016008.
https://doi.org/10.1088/1741-2552/14/1/016008.
Vasudevan, S., Vo, J., Shafer, B., Nam, A.S., Vakoc, B.J., Hammer, D.X., 2019. Toward
optical coherence tomography angiography-based biomarkers to assess the safety of
peripheral nerve electrostimulation. J. Neural Eng. https://doi.org/10.1088/17412552/ab1405.
Veltink, P.H., van Alsté, J.A., Boom, H.B.K., 1989. Multielectrode intrafascicular andextraneural stimulation. Med. Biol. Eng. Comput. 27, 19–24. https://doi.org/10.
1007/BF02442165.
Veraart, C., Grill, W.M., Mortimer, J.T., 1993. Selective control of muscle activation witha multipolar nerve cuﬀ electrode. IEEE Trans. Biomed. Eng. 40, 640–653. https://doi.
org/10.1109/10.237694.
Veraart, C., Raftopoulos, C., Mortimer, J.T., Delbeke, J., Pins, D., Michaux, G., Vanlierde,A., Parrini, S., Wanet-Defalque, M.-C., 1998. Visual sensations produced by optic
nerve stimulation using an implanted self-sizing spiral cuﬀ electrode. Brain Res. 813,
181–186. https://doi.org/10.1016/S0006-8993(98)00977-9.
Vijayaraghavan, S., Huq, R., Hausman, M.R., 2014. Methods of peripheral nerve tissue
preparation for second harmonic generation imaging of collagen ﬁbers. Methods 66,
246–255. https://doi.org/10.1016/j.ymeth.2013.08.012.
Vince, V., Brelen, M.E., Delbeke, J., Colin, I.M., 2005. Anti-TNF-α reduces the inﬂammatory reaction associated with cuﬀ electrode implantation around the sciatic
nerve. J. Neuroimmunol. 165, 121–128. https://doi.org/10.1016/j.jneuroim.2005.
04.019.
VNS Therapy System Epilepsy Physician’s Manual (US), 2018.
VNS Therapy, 2019. LivaNova. [WWW Document]URL. https://us.livanova.cyberonics.
com/healthcare-professionals/.
Vuckovic, A., Tosato, M., Struijk, J.J., 2008. A comparative study of three techniques fordiameter selective ﬁber activation in the vagal nerve: anodal block, depolarizing
prepulses and slowly rising pulses. J. Neural Eng. 5, 275–286. https://doi.org/10.
1088/1741-2560/5/3/002.
Wadhwa, R., Lagenaur, C.F., Cui, X.T., 2006. Electrochemically controlled release of
dexamethasone from conducting polymer polypyrrole coated electrode. J. Control.
Release 110, 531–541. https://doi.org/10.1016/j.jconrel.2005.10.027.
Wark, H.A.C., Sharma, R., Mathews, K.S., Fernandez, E., Yoo, J., Christensen, B., Tresco,P., Rieth, L., Solzbacher, F., Normann, R.A., Tathireddy, P., 2013. A new high-density

35

Journal of Neuroscience Methods 332 (2020) 108523

C.E. Larson and E. Meng

doi.org/10.1109/TNSRE.2010.2091429.
Zhong, Y., Bellamkonda, R.V., 2007. Dexamethasone-coated neural probes elicit attenuated inﬂammatory response and neuronal loss compared to uncoated neuralprobes. Brain Res. 1148, 15–27. https://doi.org/10.1016/j.brainres.2007.02.024.
Zitnik, R.J., 2011. Treatment of chronic inﬂammatory diseases with implantable medical
devices. Ann. Rheum. Dis. 70, i67–i70. https://doi.org/10.1136/ard.2010.138677.

Zanos, T.P., 2019. Recording and decoding of vagal neural signals related to changes inphysiological parameters and biomarkers of disease. Cold Spring Harb. Perspect.
Med., a034157. https://doi.org/10.1101/cshperspect.a034157.
Zariﬀa, J., Nagai, M.K., Schuettler, M., Stieglitz, T., Daskalakis, Z.J., Popovic, M.R., 2011.
Use of an experimentally derived leadﬁeld in the peripheral nerve pathway discrimination problem. IEEE Trans. Neural Syst. Rehabil. Eng. 19, 147–156. https://36