Journal of Medical and Biological Engineering, 31(3): 151-160
151
Review: Bioengineering Approaches for Guided Peripheral
Nerve Regeneration
Pei-Hwa Wang1
I-Ling Tseng1
Shan-hui Hsu2,3,4,*
1
Department of Animal Science and Technology, National Taiwan University, Taipei 106, Taiwan, ROC
Institute of Polymer Science and Engineering, National Taiwan University, Taipei 106, Taiwan, ROC
3
Research Center for Developmental Biology and Regenerative Medicine, National Taiwan University, Taipei 106, Taiwan, ROC
4
Institute of Biomedical Engineering, National Chung Hsing University, Taichung 402, Taiwan, ROC
2
Received 5 Apr 2011; Accepted 27 May 2011; doi: 10.5405/jmbe.902
Abstract
Peripheral nerve injury is a serious health concern for society, affecting 2.8% of trauma patients, many of whom
acquire long-term disability, and the related socioeconomic costs are relatively expensive. Current techniques in
peripheral nerve repair include the use of autografts and nerve conduits to bridge the nerve gap. Bioengineers have
developed a variety of nerve conduits to improve nerve regeneration, as well as employed functionalized bioactive
additives to the nerve conduits. Within such a discipline, adequate evaluation methods are extremely important for
correctly assessing nerve repair and regeneration. There are histological, electrophysiological or functional analyses.
This article reviews the research efforts that integrate bioengineering approaches.
Keywords: Peripheral nerve injury, Nerve conduit, Electrophysiological measurement, Functional recovery
1. Introduction
The nervous system comprises two main parts: the central
nervous system (CNS) and the peripheral nervous system
(PNS), which also includes the autonomic nervous system and
the somatic nervous system. Compared to the CNS, the
microenvironment surrounding an injury site in the PNS is
often more permissive to axonal regeneration. Peripheral nerve
injury (PNI) is a serious health concern for society, affecting
2.8% of trauma patients, many of whom acquire long-term
disability, and the related socioeconomic costs are relatively
expensive [1]. Peripheral nerve injury results in partial or
complete interruption of normal physiology of the nerve. The
research efforts in peripheral nerve repair have significantly
increased during the recent years. Concerning in vivo studies in
peripheral nerve repair, a PubMed analysis of a random sample
of 1500 research papers on nerve regeneration showed that
more than 90% of them chose the rat as the animal model. The
main reason is the relatively larger nerve size compared with
mice and more standardized functional tests [2]. Various large
animal models have been employed for nerve regeneration,
such as rabbits, sheep, pigs and primates. These models do not
have standardized functional tests, and investigators have to
establish such tests in the future. On the other hand, the nerve
regeneration rate in these animals is slower than that in rats,
* Corresponding author: Shan-hui Hsu
Tel: +886-2-33665313; Fax: +886-2-33665237
E-mail: [email protected]
due to the larger nerve diameter. As clinically translational
studies progress, large animal models have to be employed.
Their anatomy of nerves may be more similar to those of
humans, and their function needs to be investigated thoroughly.
A more appropriate animal model should be developed in the
future in order to study nerve regeneration across the long-gap
defect of a large diameter nerve. This review attempted to
cover various bioengineering aspects in peripheral nerve
reconstruction.
2. Current techniques and concepts in peripheral
nerve repair
The recovery of peripheral nerve lesions often requires a
graft to bridge the gap when the gap is large than 5 mm in
length. Although nerve autograft is the first-choice strategy
(gold standard) in reconstructions, lack of a graft material
remains a concern. Currently, implantation of the
bioengineered nerve conduits has been developed in the field
of regenerative medicine, which can be used clinically as an
alternative to nerve autograft.
2.1 Autologous nerve grafts (autografts)
Up to the present, autologous nerve grafts as the gold
standard in peripheral nerve repair have served as ideal nerve
conduits because they provide a permissive and stimulating
scaffold including Schwann cell basal laminae, neurotrophic
factors, and adhesion molecules [3]. Autologous nerve grafts
often give satisfactory results, as reported in many studies
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regarding peripheral nerve regeneration [4]. Donor nerves can
be taken from the less functional nerves such as sural nerves,
superficial cutaneous nerves or lateral and medial antebrachii
cutaneous nerves [4].
However, there are still many disadvantages of using
autologous nerve grafts. For example, lack of graft materials,
donor site loss-of-function, formation of potential painful
neuroma, and differences in structure and size between donor
and recipient grafts are the drawbacks of autologous nerve
grafts [5]. Furthermore, the length of available autologous
nerve grafts is limited, thus this is a problem in situations
where extensive reconstruction is required. For example, a
suitable nerve graft may not be always available in diabetic
patients. As a result, in the last decade increasing efforts have
been made to understand nerve regeneration and find
alternatives to the autologous nerve grafts. Studies on artificial
nerve conduits for nerve regeneration have extended and
focused on the applications of various conduit materials and
bioactive additives in order to avoid sacrifice of a donor nerve.
2.2 Nerve conduits
The design of artificial nerve conduits has advanced in the
past decades. Their availability and ease of fabrication are
better than those of autologous nerve grafts. However, nerve
regeneration efficacy and functional recovery are often inferior
to those of autografts. A wide variety of materials has been
evaluated in nerve repair. They can be roughly divided into two
categories: native materials and synthetic materials. The
designs of lumen fillers have also been demonstrated as
indispensable to enable nerve regeneration across large defect
gaps. For instance, neurotrophic factors, cellular components,
and extracellular molecules are all additives that can fill in the
nerve conduits.
2.2.1 Nerve conduit materials for peripheral nerve repair
Many biological materials have been utilized extensively
for nerve guidance conduit, such as vein [6,7], muscle [8], small
intestine submucosa [9] and amnion membrane [10,11]. All of
these materials lead to great results, but the disadvantage is still
the limited source of these tissues. As a result, a large number of
purified extracellular matrix (ECM) or glycosaminoglycan are
considered as another choice to fabricate nerve conduits. The
major components in ECM are collagen, laminin, and
fibronectin. They can regulate neural activity and regeneration.
The FDA-approved NeuraGen tube fabricated from cross-linked
bovine collagen I is commercially available. The use of these
tubes in a monkey median nerve transection model was first
described by Archibald et al. [12,13], where they showed
comparable performance to nerve autografts, and ultimately
superiority to direct suturing of nerve stumps. Clinical studies
using NeuraGen tubes for repair of nerve gaps up to 18 mm in
human hands also showed promising results [14].
In addition to the biological materials, the artificial
synthetic nerve conduit is another alternative for peripheral
nerve defect repair. Due to their specific properties, such as
tailored degradation time by altering molecular weights or
compositions, synthetic polymers are better manipulated than
natural materials. Synthetic nerve conduits can be divided to
biodegradable and non-biodegradable ones. Silicone and
expanded
polytetrafluoroethylene
(PTFE)
are
nonbiodegradable. Such non-biodegradable grafts often result in
chronic tissue response or nerve compression [15]. Neither
silicone nor PTFE showed good recovery over defects
exceeding 4 cm [16]. The drawbacks of non-biodegradable
materials have led to the development of biodegradable nerve
conduits.
Polyesters,
including
poly(glycolic acid)
(PGA),
poly(lactic acid) (PLA), polycaprolactone (PCL) and their
copolymers, are widely used in biomedical applications, some
of which have been FDA-approved, because of their
biocompatibility and safety in vivo. They are commonly
synthesized in vitro by ring-opening polymerization, and
degraded in vivo by hydrolysis of the ester linkages. The most
appealing property of polyesters is their specifically tailored
properties, such as degradation rate and mechanical strength
through copolymerization, which make their potential
drawback in releasing acidic degradation by-products
negligible. For example, the degradation activity and rigidity of
the poly(lactide-co-glycolide) (PLGA) copolymer are highly
dependent on its molecular weight and crystallinity, which are
easily controlled by altering the ratio of glycolide to lactide
monomers during synthesis [17]. However, among the wide
variety of well-manipulated polyesters, only PGA and
poly(lactide-co-epsilon-caprolactone) (PLCL), for now, have
been developed into nerve conduits and marketed as Neurotube
and Neurolac, respectively, which are FDA-approved and
commercially available. Continuing efforts are still invested in
order to develop the next generation of polyesters-based nerve
conduits.
2.2.2 Surface modifications in nerve conduits
In addition to the properties of nerve conduit materials that
change the mechanical tolerance, biocompatibility and
absorbability, the internal surface modifications of nerve
conduits are also important. Surface modifications can adjust
the scaffold permeability and guidance structure. The nerve
conduits should be semipermeable to provide adequate
diffusion of oxygen and metabolites between the conduit lumen
and external environment. The guidance structures mimicking
the natural fibrous structure of the ECM, like nanofibrous
scaffolds and microgrooves, are suitable for cell attachment and
proliferation [18,19].
Lu and colleagues [20] fabricated controllable asymmetric
PCL membranes, and cells proliferated well due to the
directional transport characteristics. PLA nerve conduits with
designed micropores and surface microgrooves showed
enhanced ability in the repair of 10-mm sciatic nerve
transection defects in rats [21]. Christopherson and colleagues
[22] indicated that, as the fiber diameter decreased, higher
degrees of proliferation and cell spreading and lower degree of
cell aggregation were observed. PLGA microgrooved films
exerted a guidance effect on both early-stage neurite outgrowth
and neurite elongation [23]. In summary, surface modification
in internal lumen surface is important, and it can direct the
axonal outgrowth and enhance the nerve regeneration process.
Bioengineering in Peripheral Nerve Regeneration
2.2.3 Functionalized bioactive nerve conduit luminal additives
for axon regeneration
Only the empty synthetic nerve conduits often do not
achieve the goal of promoting nerve regeneration across long
lesion gaps. As a result, many studies have incorporated
functionalized bioactive additives in the lumen of the conduits.
As mentioned, growth/neurotrophic factors, cells, nucleic acids
and ECM molecules such as collagen, laminin and fibronectin
are often involved.
(a) Neurotrophic factors
Neurotrophic factors play important roles in stimulating
nerve regeneration when used in nerve repair. They can control
the survival, migration, proliferation and differentiation of
various neural cell types [24]. In particular, nerve growth factor
(NGF), neurotrophin-3 (NT-3), glial cell line-derived
neurotrophic factor (GDNF) and fibroblast growth factors
(acidic FGF and basic FGF) are the most common choices.
Lee and colleagues [25] used heparin to stabilize NGF and
to control its diffusion from a fibrin matrix. The nerve silicone
conduit, which was loaded in the walls of the conduit with 20
or 50 ng/ml of NGF, produced similar outcomes in terms of
number of regenerating nerve fibers and fiber diameter as
compared with an autograft in a 13-mm rat sciatic nerve injury
model. As a result, both 20- and 50-ng/mL NGF delivery
systems could promote a similar level of nerve regeneration as
the autograft control. On the other hand, the empty conduits
and conduits comprising the delivery system alone without
NGF were significantly less effective. Kokai et al. [26] used the
GDNF-embedded nerve conduits to bridge a 1.5-cm defect in
the male Lewis rat for a 16-week period. GDNF is a candidate
trophic factor for motoneurons during axonal regeneration. It
belongs to a subfamily within the transforming growth factor
superfamily. Kokai et al.’s results showed that the measured
gastrocnemius twitch force in animals treated with GDNF was
significantly higher than that in negative controls and was not
significantly different from that of the isograft-positive control
group.
Chitosan
nerve
conduits
filled
with
a
heparin-incorporated fibrin-fibronectin matrix serving as
delivery systems for basic fibroblast growth factor (bFGF)
were used for repairing 10-mm sciatic nerve defects in adult
rats. Three months post-operation, the conduction velocity and
the muscle restoration rate in animals of the experimental group
were significantly higher than those of the PBS control group
[27].
These results clearly indicate the possibilities of including
neurotrophic factors into nerve conduits and its effectiveness in
assisting nerve regeneration and partial functional recovery in
the PNS.
(b) Cellular components
Cell transplantation in a bioartificial conduit might be a
suitable strategy to create a favorable environment for nerve
regeneration. Transplanted cells may secrete neurotrophic
factors to enhance regeneration. Furthermore, cellular
components may also support and guide the regeneration of
injured nerves [28].
Schwann cells play an important role in supporting axonal
regrowth and migration following PNI. They can release
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bioactive factors and promote blood vessel formation [29].
When the peripheral nerve is injured, Schwann cells in the
injured region will produce various ECM molecules such as
laminin and collagen. Moreover, after Wallerian degeneration,
Schwann cells form bands of Büngner that serve as a guiding
rail for regrowing axons [30]. Cheng and Chen [31] seeded
Schwann cells on PLGA fiber and membrane, then wrapped
them as conduits to bridge a 20-mm critical gap of rabbit sciatic
nerve. The conduit promoted nerve regeneration, and the
number of axons and the area of nerve fibers were comparable
to the autograft group after 3 months. To make nerve
regeneration in defects of larger than 3 cm in rabbit peroneal
nerves, luminal fillers such as Schwann cells were shown to
promote axonal outgrowth even for gaps over 6 cm [32,33].
However, for the treatment of acute injuries, the use of
autologous cultured Schwann cells may be impractical, because
of the technical difficulties and the time required for harvesting
and expanding this slow-growing type of cell. Mosahebi et al.
[34] indicated that transplanting a sufficient number of
Schwann cells into a conduit may need ten weeks to culture
these cells.
Mesenchymal stem cells (MSCs) are multipotent stem
cells derived from various sources such as bone marrow. They
have been shown to transdifferentiate into neural cells [35].
Human umbilical cord-derived MSCs (UC-MSCs) that are
available from cell banks can be induced to differentiate into
various cell types, thereby making them practical potential
sources for cell-based therapies. Matsuse et al. [36] reported
that cells with Schwann cell properties as well as the ability to
support axonal regeneration and myelin reconstruction could be
successfully induced from UC-MSCs to promote functional
recovery after peripheral nerve injury. There are also reports
that adipose-derived stem cells may provide an effective cell
population, without the limitations of the donor-site morbidity
associated with isolation of Schwann cells, and could be a
clinically translatable route towards new methods to enhance
peripheral nerve repair [37,38]. Moreover, neural stem cells
may serve as seed cells of peripheral nerve tissue engineering
and be used in artificial nerve to repair nerve defects [39,40].
In summary, using transplanted cells in a bioartificial
conduit might be a practical strategy for improving peripheral
nerve regeneration. It has become evident that molecular and
cellular elements in addition to the conduit are needed in order
to mimic a nerve microenvironment and to promote axonal
regrowth.
(c) Extracellular components
ECM molecules like collagen, laminin and fibronectin
have been utilized as additives in nerve conduit lumen for
decades. They can regulate cell viability and myelination of
peripheral nerves [41]. Laminin can increase Schwann cell
attachment and induce neurite outgrowth in vitro [42].
Favorable functional recovery was also observed in defects of
up to 8 cm in canine peroneal nerves if PGA–collagen nerve
conduits were filled with laminin-coated collagen fibers or
laminin-impregnated collagen sponge prior to implantation
[43]. Such favorable nerve regeneration may reflect the typical
condition faced in human peripheral nerve injuries and indicate
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Table 1. Advantages and disadvantages of different methods used to evaluate the peripheral nerve regeneration.
Evaluation methods
Histology
Advantages
It is the most common method to evaluate peripheral
nerve regeneration.
Electron microscopy is a useful tool for investigating the
early stages of nerve damage and regeneration at a
subcellular level.
Magnetic resonance neurography
It is a non-invasive method to evaluate the nerve
The high cost restrains the willingness of the researchers.
regeneration. A longitudinal investigation on one animal
can be conducted over the long term.
Electrophysiological evaluation
methods
It can provide quantitative measurement of nerve activity
in the normal and experimental groups.
Serial electrophysiological studies yield important
information about the longitudinal nerve regeneration
and reinnervation.
Significant variations are usually observed in small
laboratory animals and on different days in the same
animal, which are influenced by the errors in measuring
the exact length of the nerve unless dissected after animal
sacrifice. Results can be disturbed by variations in body
temperature of the animal.
Kinematic gait analysis
It can accurately assess normal and abnormal rat gait.
The gait analysis data do not always correspond to the
true functional recovery.
There is no standard instrument that can be applied to
larger animals.
the importance of luminal fillers in enhancing PNS
regeneration. Fibronectin is a large ECM glycoprotein, which
promotes cell adhesion and migration [44]. Whitworth and
colleagues [45] demonstrated the ability of fibronectin to
support a significantly faster rate of axonal growth than
freeze-thawed muscle grafts within the first 10 days
post-surgery. Furthermore, from day 15 to day 60, fibronectin
conduits resulted in comparable amounts of regenerating axons
and Schwann cells in the conduits as compared to nerve grafts.
To sum up, ECM components may be another important factor
in peripheral nerve repair.
3. Evaluation
methods
in
peripheral
regeneration experimental research
nerve
In peripheral nerve regeneration study, adequate
evaluation methods are extremely important to assess the
peripheral nerve regeneration. Vleggeert-Lankamp [46] scored
the methods for grafting of the rat sciatic nerve with a conduit.
The evaluation of nerve fiber count, nerve fiber density, nerve
histological success ratio, compound muscle action potential,
muscle weight, and muscle tetanic force were demonstrated to
have resolving power. Most research studies have used three
most frequently methods for evaluating nerve recovery, i.e.
morphological analysis, gait analysis and electrophysiological
evaluation. The common evaluation methods are listed in
Table 1, together with their advantages and disadvantages.
Each is described in the following sections.
3.1 Morphological techniques
3.1.1 Histology evaluation method
Nerve histology has been the most common method of
evaluating peripheral nerve regeneration. Quantitative
estimation of nerve fiber morphology is a key tool in nerve
regeneration research [47,48]. The most important parameters
that can be used for the assessment of nerve fibers are number
of fibers, diameter of fibers, density of fibers, the ratio of the
axon diameter to the myelinated fiber diameter, and the ratio of
the total myelinated fiber area to the total nerve tissue area.
Disadvantages
Quantitative estimation of nerve fiber morphology might
have bias in different parameters.
In most studies, morpho-functional correlation is poor,
and thus the relationship between the morphology and
functional studies should be more carefully interpreted.
Although different types of fixatives can be used for
nerve, 4% paraformaldehyde in phosphate-buffered
saline (PBS) is mostly used. To obtain good histological
quality, perfusion is not required, and specimen immersion in
the fixative solution is enough to fix the nerve sample [48].
Nerves are then embedded by paraffin or cryo-embedding and
transversely sectioned. Subsequently, tissues are postfixed in
an osmium tetroxide buffer waiting for dehydration. Tissues
are then stained with a toluidine blue or phenylenediamine
solution for light microscopy or with uranyl acetate and
aqueous lead citrate for electron microscopy [47]. Previously,
nerve fibers were counted by hand, but now, digital images are
collected and imported into imaging software for evaluation.
In most cases, not all nerve fibers are counted, just those in
sample areas, after which the total number is computed from
the sample area and the total area of the nerve cross-section.
For example, Kiyotani and colleagues [49] recorded the
morphometric features by 35-mm reversal fim. These films
were scanned and transformed into image files by a computer.
Using software (Adobe Photoshop), these files were then
processed for analysis.
The diameter of fibers is always smaller in regenerated
compared with untreated nerves, and the diameter size will
increase over time. Perfusion-fixed nerve fibers tend to be
somewhat smaller in diameter (maximum 12.5 µm in the rat
sciatic nerve and its branches) [50] in comparison with
immersion-fixed nerve fibers (maximum 15.5 µm in the rat
sciatic nerve and its branches) [51]. Vleggeert-Lankamp [46]
suggested that variations in the diameter of regenerated nerves
were too small to use the diameter size as a useful tool to
compare with the nerve grafts, but it is an excellent tool to
demonstrate differences in untreated and synthetic nerve
grafted rats.
The nerve fiber density is calculated as the number of
nerve fibers per square millimeter or another square measure.
In general, the density in autografted nerves is lower compared
with that in a synthetic nerve conduit. Evans et al. [52]
described that the density in regenerated nerves decreased with
time after surgery.
Bioengineering in Peripheral Nerve Regeneration
The ratio of the axon diameter to the myelinated fiber
diameter is called G-ratio. It was considered to be a useful
parameter to assess the maturity of nerve fibers.
Vleggeert-Lankamp [46] showed that the G-ratio in untreated
nerves was smaller than that in the regenerated nerve fibers,
indicating that regenerated nerves had a relatively thinner
myelin sheath. Raimondo et al. [48] compared the myelinated
nerve fibers in resin-embedded and paraffin-embedded normal
rat radial nerves, and the G-ratios were 0.72 ± 0.06 and
0.71 ± 0.05 (mean ± SD), respectively. den Dunnen and
colleagues [53] demonstrated that G-ratios obtained 2 years
after surgery were equal to values in untreated nerves. This
would imply that the relation between axon and myelin sheath
would decrease on maturation. Several reports have different
opinions with respect to whether the G-ratio is larger in
untreated or in regenerated nerves [54,55,56].
The N-ratio is calculated as the total myelinated fiber area
divided by the total nerve tissue area. The N-ratio represents
the total mass of the regenerating nerve in the tube, and is a
parameter indicative of both the number of sprouting events
and the degree of maturation of the regenerating nerve [57]. A
low N-ratio was indicative of a relatively large amount of
fibrous tissue [58]. Researchers reported that the N-ratio in
normal nerves was significantly higher [54,55,56]. However,
the N-ratio increased with time after surgery [56]. Regarding a
comparison between the G-ratio and N-ratio, VleggeertLankamp [46] indicated that N-ratio was more recommended
to use as a promising parameter to evaluate the morphological
repair in the future.
3.1.2 Magnetic resonance (MR) neurography
Currently, MR neurography can diagnose neuroma in
continuity, detect nerve regeneration in cases of traumatic injury,
and demonstrate abnormal signal in diseased peripheral nerves.
MR neurography is used to perform non-invasive and long-term
tracking for peripheral nerve injury. The images can correlate
with functional evaluation data. Howe and colleagues [59]
developed a new method using T2 or diffusion-weighted MR
imaging protocols to detect the nerve in the rabbit forelimb. As
the technology advanced, high-resolution coils and appropriate
image processing made the T2-based protocols [60,61] better in
many peripheral nerve pathological conditions, such as brachial
plexus injury, carpal tunnel syndrome, ulnar nerve compression,
lower-extremity nerve compression, cervical radiculopathy and
acute axonotmetic nerve injury.
Using the rat sciatic nerve crush model, Cudlip et al. [62]
demonstrated that results from the walking track analysis of
the sciatic nerve functional deficit correlated well with MR
neurography. Other studies also proved the degree of nerve
damage and nerve regeneration process could be evaluated by
signal intensities on T2-weighted images and the time course
of T2 values and ratios [63,64]. MR neurography findings, the
electrophysiological examination and histology data, in
denervation and reinnervation are linked together [65].
The sensitivity of MR image can predict the peripheral
nerve injury and diagnose the nerve regeneration conditions, as
shown in Figure 1. In the clinical study, it supplies a possibility
that patients do not need to undergo invasive surgery and MR
155
image can quickly detect the degree of axonal damage as early
as 24 hours after the injury. This application makes the
peripheral nerve regeneration studies predict the process of
nerve recovery more accurately. However, there was only one
study using MRI to track the nerve regeneration in a conduit
[66].
Figure 1. One example showing the sagittal view of the MR image
taken at 3 months for rabbit sciatic nerve bridged by a
polylactide nerve conduit. The long white arrows indicate the
location of regenerated nerve (dark line) inside the conduit.
The implanted conduit was 27 mm in length.
3.2 Electrophysiological evaluation
Electrophysiological measurements are commonly used
in determining peripheral nerve disorders in animal models.
These tests provide a variety of quantitative measures,
including
sensory
and
motor
nerve
conduction,
electromyography, spinal reflex tests and motor and sensory
evoked potentials (SEPs). Not all electrophysiology
experiments are invasive. Some low invasive methods can
avoid sacrificing animals and allow serial evaluation of nerve
reinnervation.
When nerve is stimulated and muscles are recorded at a
distance by electrodes, the compound muscle action potential
(CMAP) is achieved. The targets frequently used in sciatic
nerve model are the gastrocnemius, tibial or plantar muscles.
CMAP can detect the latency and the amplitude of the signal.
The latency and amplitude are often expressed as the ratio of
the experimental side to the contralateral side. Many studies
indicated that normal animals had shorter latency and higher
amplitude of CMAP [67,68]. Most studies showed significant
differences between the normal and regenerated nerves.
Another common parameter is the mean nerve conduction
velocity (NCV). To measure the NCV, the nerve has to be
electrically stimulated at one point and the electrical activity
has to be measured at a distinct point. Bipolar hooked platinum
stimulating and recording electrodes are regularly used for this
purpose and are placed before and after the area of interest (the
nerve graft). The NCV is calculated as the ratio of the
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conducting distance and the latency time to the peak of the
maximal action potential. The NCV is strongly dependent on
temperature [69]. The lower the temperature is, the lesser the
NCV is. In the in vivo experiments, a heating lamp was often
used to keep the body temperature at approximately 37°C.
When the axon diameter and myelination are changed, NCV
and the latency of the action potentials are altered, and cannot
return to the normal values for a long period of time after
successful regeneration [70]. The significant variations in
NCV were frequently observed between small laboratory
animals and on different evaluation time in the same animal,
which are influenced by errors in measurement of nerve length
(unless done under dissection) and to variations in body
temperature [71]. As a result, NCV is considered less useful
than the amplitude for evaluating nerve regeneration.
Electromyography (EMG) records the electrical activity
of muscles using implanted intramuscular electrodes in
animals which are awake and freely moving [72,73]. An
evaluation of the extent of myo-electrical activation in large
regions of muscles and co-ordination between flexor and
extensor muscles can be detected through EMG signals. After
sciatic nerve injury and repair, EMG patterns in the operated
hind limb are abnormal, coactivation of the gastrocnemius and
tibialis anterior muscle is present, and the burst activity is
poorly adjusted to the phases of the step cycle. Such irregular
patterns are attributed to the nonselective reinnervation of
muscles by regenerated motor axons [74]. In some cases, EMG
patterns in the grafted rat are highly abnormal, but the
qualitative intensity of the muscular response is not
significantly different between regenerated and normal nerves
[75,76]. Scholle et al. [73] reported that simultaneous
video-recording of animal movement allows a temporal
correlation of kinematic and EMG parameters. Implantation of
electrodes in the rat hindlimb muscles allowed serial
recordings and analyses for the temporal course of
reinnervation [77], as well as the activation pattern during
locomotion [74]. With EMG signals, it is hard to obtain the
graded voluntary muscle contraction in targeted muscles, and
when muscles are highly activated, investigators can easily
underestimate the activity of the muscle [78]. To stabilize the
use of EMG is still a great challenge which EMG should be
further carefully confirmed.
Sensory evoked potentials (SEPs) can be obtained by a
rather complex method, which allows evaluation of the whole
neural pathway, from brain to peripheral nerve. The electrical
stimulation site of SEPs in experimental animals with sciatic
nerve injury is directly applied to the sciatic nerve distal to the
graft by using a needle electrode. Using screw electrodes
placed on the left cerebral sensorimotor cortex, SEPs can be
recorded. It is evaluated based on the amplitude and latency.
SEP recording requires applying multiple stimuli and
averaging ten to hundreds of responses. Because of the
difficulty of the technique and more variations due to
anesthesia levels and central synaptic efficacy in different
animals, SEPs are less suitable to determine axonal
regeneration than nerve conduction tests. However, the method
can be worthwhile for examination of short nerves [79],
proximal lesions of the peripheral nerve, sensory or motor
central disturbances, and changes in central somatotopic maps
[80]. Especially, it is the most persuasive evaluation for the
injury and diseases of the spinal cord [81,82].
Electrophysiological evaluation involves different methods
and provides a quantitative measure of nerve activity in the
normal and pathological states. Clinical electrophysiological
examination is applied to obtain information regarding
localization, pathophysiology, and injury severity level. Serial
electrophysiological investigations yield important messages
about the time-course of regeneration and reinnervation, and are
able to reveal temporal differences with the repaired time.
3.3 Kinematic gait analysis
Functional recovery is one of the primary goals of
peripheral nerve regeneration study. Although a large number of
methods have been used to evaluate the nerve functions, their
validity is still questioned. In the last decade, the most often
used sciatic nerve model is the rat. Rat gait analysis has been
developed for thirty years, but there are still some limitations to
analysis. As a result, several new sensitive quantitative methods
are designed by using computerized gait analysis.
The pioneering works of de Medinaceli’s laboratory
develop a quantitative method to evaluate the sciatic nerve
function in rats, which is known as the sciatic functional index
(SFI). It is a method to measure the footprints from the
walking rats in a walkway [83]. Since then, SFI has been used
extensively in studies of peripheral nerve injury and repair,
because the method is inexpensive, convenient and has been
regarded as very accurate and reliable [84,85]. Three different
footprint parameters are measured, including distance from the
heel to the third toe, print length (PL); distance from the first
to the fifth toe, toe spread (TS); and distance from the second
to the fourth toe, intermediary toe spread (ITS). All three
parameters are combined by the formula derived by
researchers [86], SFI is calculated as follows: All three
parameters are taken from the experimental (E) and normal
(N). An index of zero reflects normal function, and -100
theoretically represents complete loss of function.
SFI  38.3(
EPL  NPL
ETS  NTS
EIT  NIT
)  109 .5(
)  13.3(
)  8.8
NPL
NTS
NIT
Despite SFI analysis being used by most researchers,
there are many problems that have limited its experimental use.
Before experiments, the animals have to be allowed to practice
walking on the track, so that they can be more familiar with
the track. When recording an animal’s footprints, some prints
are not measurable owing to the smearing of the print, the
development of flexion contractures, autotomy, dragging of the
tail across the print, or contamination with other footprints.
Because of the difficulty in SFI measurement, Bervar [87]
used a new alternative video analysis of static footprint
analysis, so called static sciatic index (SSI). This method is
technically easier to perform than the walking footprint video
and can accurately quantify the degree of functional loss after
sciatic nerve injury in the rat.
Bioengineering in Peripheral Nerve Regeneration
Another functional index is toe out angle (TOA), defined
as the degree of angle between the direction of progression and
a reference line in the sole of the foot (from the calcaneus to
the tip of the third digit) [88]. Varejão and colleagues [88]
found a good correlation between SFI and TOA measurements
in terms of predicting functional recovery.
As the technology has fast progressed, the computerized
analyzer has been developed. Bozkurt and colleagues [89]
demonstrated that CatWalk could simultaneously measure
dynamic as well as static gait parameters, for instance,
intensity of the hind paws, individual paw parameters,
dynamic paw parameters, coordination-related parameters. It
may be used as a complementary approach to other functional
testing paradigms.
The sciatic nerve model of rat has been used most
commonly so far, and the functional evaluation system in rats
is more complete than in other animals. Hsu and colleagues
[66] studied the rabbit sciatic nerve model. The motion angle
of the rabbit was evaluated and the electrophysiological
measurements and histological analysis had positive
correlation with this method. As for the walkway, there are
such devices for large animals, too. For example, the
Animal Walkway™ System can automatically calculate the
gait parameters, analyze the relationship between multiple foot
strikes and save time on analysis, but it is not for used the
sciatic nerve model. This system is usually utilized to study
laminitis, osteoarthritis, and hip dysplasia in cats, dogs, horses,
cows and sheep. For future nerve regeneration studies, the
functional evaluation methods for large animals have to do
more research and development.
Acknowledgment
The authors were sponsored by the National Science
Council through a medical device development grant (NSC
99-2321-B-002-043).
References
[1]
[2]
[3]
[4]
[5]
[6]
[7]
[8]
[9]
4. Future perspectives on peripheral nerve repair
Ideally, the interventions for peripheral nerve injury
should be initiated early and take into account the level of
injury. Although the physician cannot master the biological
changes inside a damaged nerve, the therapy should start as
soon as possible to keep more neurons alive, encourage axons
to cross longer gaps, maximize the accuracy of target
reinnervation and manage the neuropathic pain.
Despite the extensive amount of work on developing and
searching among a variety of materials as the candidate for
nerve conduits, no single material has been shown to have the
potential to overtake the nerve autograft in terms of nerve
regenerative potential and function. On the other hand, past
studies have highlighted various design modifications that
would contribute to improving the performance of artificial
nerve conduits. Processing techniques could be utilized to
develop internal conduit microstructures that came close to
mimicking the natural fibrous structure of the ECM [90].
However, the continued lack of success rate of nerve
regeneration over long gaps strongly suggests that merely
providing physical guidance through a conduit is innately
insufficient, and additional trophic supports (neurotrophic
factors, cellular components and ECM molecules) in the form
of biochemical or biophysical signals is required.
157
[10]
[11]
[12]
[13]
[14]
[15]
[16]
[17]
[18]
J. Noble, C. A. Munro, V. S. Prasad and R. Midha, “Analysis of
upper and lower extremity peripheral nerve injuries in a
population of patients with multiple injuries,” J. Trauma, 45:
116-122, 1998.
P. Tos, G. Ronchi, I. Papalia, V. Sallen, J. Legagneux, S. Geuna
and M. G. Giacobini-Robecchi, “Methods and protocols in
peripheral nerve regeneration experimental research: Part I –
Experimental models,” Int. Rev. Neurobiol., 87: 47-79, 2009.
K. G. Almgren, “Revascularization of free peripheral nerve
grafts. An experimental study in the rabbit,” Acta. Orthop. Scand.
Supp., 154: 1-104, 1975.
E. O. Johnson and P. N. Soucacos, “Nerve repair: experimental
and clinical evaluation of biodegradable artificial nerve guides,”
Injury, 39: S30-S36, 2008.
S. Mackinnon and A. L. Dellon, “Clinical nerve reconstruction
with a bioabsorbable polyglycolic acid tube,” Plast. Reconstr.
Surg., 85: 419-424, 1990.
R. L. Walton, R. E. Brown, W. E. Matory Jr., G. L. Borah and J.
L. Dolph, “Autogenous vein graft repair of digital nerve defects
in the finger: a retrospective clinical study,” Plast. Surg., 89:
944-949, 1989.
G. Risitano, G. Cavallaro, T. Merrino, S. Coppolono and F.
Ruggeri, “Clinical results and thoughts on sensory nerve repair
by autologous vein graft in emergency hand reconstruction,”
Chir. Main., 21: 194-197, 2002.
B. Battiston, P. Tos, T. R. Cushway and S. Geuna, “Nerve repair
by means of vein filled with muscle grafts: I. Clinical results,”
Microsurgery, 20: 32-36, 2000.
S. F. Badylak, R. Record, K. Lindberg, J. Hodde and K. Park,
“Small intestinal submucosa: a substrate for in vitro cell
growth,” J. Biomater. Sci. Polym. Ed., 9: 863-878, 1998.
M. F. Meek, J. H. Coert and J. P. Nicolai, “Amnion tube for
nerve regeneration,” Plast. Reconstr. Surg., 107: 622-623,
2001a.
J. Mohammad, J. Shenaq, E. Rabinovsky and S. Shenaq,
“Modulation
of
peripheral
nerve
regeneration:
a
tissue-engineering approach. The role of amnion tube nerve
conduit across a 1-centimeter nerve gap,” Plast. Reconstr. Surg.,
105: 660-666, 2000.
S. J. Archibald, C. Krarup, J. Shefner, S. T. Li and R. D.
Madison, “A collagen-based nerve guide conduit for peripheral
nerve repair: an electrophysiological study of nerve regeneration
in rodents and nonhuman primates,” J. Comp. Neurol., 306:
685-696, 1991.
S. J. Archibald, J. Shefner, C. Krarup and R. D. Madison,
“Monkey median nerve repaired by nerve graft or collagen nerve
guide tube,” J. Neurosci., 15: 4109-4123, 1995.
W. W. Ashley Jr., T. Weatherly and T. S. Park, “Collagen nerve
guides for surgical repair of brachial plexus birth injury,” J.
Neurosurg., 105: 452-456, 2006.
S. Panseri, C. Cunha, J. Lowery, U. Del Carro, F. Taraballi, S.
Amadio, A. Vescovi and F. Gelain, “Electrospun micro- and
nanofiber tubes for functional nervous regeneration in sciatic
nerve transections,” BMC Biotechnol., 8: 39, 2008.
S. Stanec and Z. Stanec, “Reconstruction of upper-extremity
peripheral-nerve injuries with ePTFE conduits,” J. Reconstr.
Microsurg., 14: 227-232, 1998.
X. S. Wu, “Synthesis and properties of biodegradable
lactic/glycolic acid polymers,” in: D. L. Wise (Ed.),
Encyclopedic Handbook of Biomaterials and Bioengineering,
Part A: Materials, New York: Marcel Dekker, 1015-1054, 1995.
T. J. Sill and H. A. von Recum, “Electrospinning: applications in
drug delivery and tissue engineering,” Biomaterials, 29:
158
J. Med. Biol. Eng., Vol. 31 No. 3 2011
1989-2006, 2008.
[19] S. h. Hsu and H. C. Ni. “Fabrication of the microgrooved/
microporous polylactide substrates as peripheral nerve conduits
and in vivo evaluation,” Tissue Eng. Part A., 15: 1381-1390,
2009.
[20] C. M. Lu, C. J. Chang, C. M. Tang, H. C. Lin, S. C. Hsieh, B. S.
Liu and W. C. Huang, “Effects of asymmetric polycaprolactone
discs on co-culture nerve conduit model,” J. Med. Biol. Eng., 29:
76-82, 2009.
[21] H. C. Ni, Z. Y. Lin, S. h. Hsu and I. M. Chiu, “The use of air
plasma in surface modification of peripheral nerve conduits,”
Acta Biomater., 6: 2066-2076, 2010.
[22] G. T. Christopherson, H. Song and H. Q. Mao, “The influence of
fiber diameter of electrospun substrates on neural stem cell
differentiation and proliferation,” Biomaterials, 30: 556-564,
2009.
[23] L. Yao, S. Wang, W. Cui, R. Sherlock, C. O’Connell, G.
Damodaran, A. Gorman, A. Windebank and A. Pandit, “Effect of
functionalized micropatterned PLGA on guided neurite growth,”
Acta Biomater., 5: 580-588, 2009.
[24] E. J. Huang and L. F. Reichardt, “Neurotrophins: roles in
neuronal development and function,” Annu. Rev. Neurosci., 24:
677-736, 2001.
[25] A. C. Lee, V. M. Yu, J. B. Lowe, M. J. Brenner, D. A. Hunter, S.
E. Mackinnon and S. E. Sakiyama-Elbert, “Controlled release of
nerve growth factor enhances sciatic nerve regeneration,” Exp.
Neurol. ,184: 295-303, 2003.
[26] L. E. Kokai, D. Bourbeau, D. Weber, J. McAtee and K. G. Marra,
“Sustained growth factor delivery promotes axonal regeneration
in long gap peripheral nerve repair,” Tissue Eng. Part A., In
Press., 2011.
[27] H. Han, Q. Ao, G. Chen, S. Wang and H. Zuo, “A novel basic
fibroblast growth factor delivery system fabricated with
heparin-incorporated fibrin-fibronectin matrices for repairing rat
sciatic nerve disruptions,” Biotechnol. Lett., 32: 585-591, 2010.
[28] M. Tohill and G. Terenghi, “Stem-cell plasticity and therapy for
injuries of the peripheral nervous system,” Biotechnol. Appl.
Biochem., 40: 17-24, 2004.
[29] B. Schlosshauer, E. Muller, B. Schroder, H. Planck and H. W.
Muller, “Rat Schwann cells in bioresorbable nerve guides to
promote and accelerate axonal regeneration,” Brain Res., 963:
321-326, 2003.
[30] R. P. Bunge, “The role of the Schwann cell in trophic support
and regeneration,” Proc. Argenteuil Symposium on Brain Repair,
S19-S21, 1993.
[31] B. Cheng and Z. R. Chen, “Fabricating autologous tissue to
engineer artificial nerve,” Microsurgery, 22: 133-137, 2002.
[32] B. Strauch, D. M. Rodriguez, J. Diaz, H. L. Yu, G. Kaplan and D.
E. Weinstein, “Autologous Schwann cells drive regeneration
through a 6-cm autogenous venous nerve conduit,” J. Reconstr.
Microsurg., 17: 589-595, 2001.
[33] F. Zhang, B. Blain, J. Beck, J. Zhang, Z. R. Chen, Z. W. Chen
and W. C. Lineaweaver, “Autogenous venous graft with
one-stage prepared Schwann cells as a conduit for repair of long
segmental nerve defects,” J. Reconstr. Microsurg., 18: 295-300,
2002.
[34] A. Mosahebi, M. Simon, M. Wiberg, R. Martin and G. Terenghi,
“Retroviral labeling of Schwann cells: in vitro characterization
and in vivo transplantation to improve peripheral nerve
regeneration,” Glia, 34: 8-17, 2001.
[35] E. K. F. Yim, S. W. Pang and K. W. Leong, “Synthetic
nanostructures inducing differentiation of human mesenchymal
stem cells into neuronal lineage,” Exp. Cell Res., 313:
1820-1829, 2007.
[36] D. Matsuse, M. Kitada, M. Kohama, K. Nishikawa, H.
Makinoshima, S. Wakao, Y. Fujiyoshi, T. Heike, T. Nakahata, H.
Akutsu, A. Umezawa, H. Harigae, J. Kira and M. Dezawa,
“Human umbilical cord-derived mesenchymal stromal cells
differentiate into functional Schwann cells that sustain
peripheral nerve regeneration,” J. Neuropathol. Exp. Neurol., 69:
973-985, 2010.
[37] L. Y. Santiago, J. Clavijo-Alvarez, C. Bravfield, J. P. Rubin and
K. G. Marra. “Delivery of adipose-derived precursor cells for
peripheral nerve repair,” Cell Transplant, 18: 145-158, 2009.
[38] P. G. di Summa, P. J. Kingham, W. Raffoul, M. Wiberg, G.
[39]
[40]
[41]
[42]
[43]
[44]
[45]
[46]
[47]
[48]
[49]
[50]
[51]
[52]
[53]
[54]
[55]
[56]
Terenghi and D. F. Kalbermatten. “Adipose-derived stem cells
enhance peripheral nerve regeneration,” J. Plast. Reconstr.
Aesthet. Surg. (JPRAS), 63: 1544-1552, 2010.
B. F. Guo and M. M. Dong, “Application of neural stem cells in
tissue- engineered artificial nerve,” Otolaryngol. Head Neck
Surg., 140: 159-164, 2009.
K. Y. Fu, L. G. Dai, I. M. Chiu, J. R. Chen and S. h. Hsu,
“Sciatic nerve regeneration by microporous nerve conduits
seeded with glial cell line-derived neurotrophic factor or
brain-derived neurotrophic factor gene transfected neural stem
cells,” Artif. Organs, 35: 363-372, 2011.
J. L. Podratz, E. Rodriguez and A. J. Windebank, “Role of the
extracellular matrix in myelination of peripheral nerve,” Glia, 35:
35-40, 2001.
C. Deister, S. Aljabari and C. E. Schmidt, “Effects of collagen 1,
fibronectin, laminin and hyaluronic acid concentration in
multi-component gels on neurite extension,” J. Biomater. Sci.
-Polym. Ed., 18: 983-997, 2007.
K. Matsumoto, K. Ohnishi, T. Kiyotani, T. Sekine, H. Ueda, T.
Nakamura, K. Endo and Y. Shimizu, “Peripheral nerve
regeneration across an 80-mm gap bridged by a polyglycolic
acid (PGA)-collagen tube filled with laminin-coated collagen
fibers: a histological and electrophysiological evaluation of
regenerated nerves,” Brain Res., 868: 315-328, 2000.
K. M. Yamada, “Cell-surface interactions with extracellular
materials,” Annu. Rev. Biochem., 52: 761-799, 1983.
I. H. Whitworth, R. A. Brown, C. Dore, C. J. Green and G.
Terenghi, “Orientated mats of fibronectin as a conduit material
for use in peripheral nerve repair,” J. Hand Surg. Br., 20:
429-436, 1995.
C. L. Vleggeert-Lankamp, “The role of evaluation methods in
the assessment of peripheral nerve regeneration through
synthetic conduits: a systematic review,” J. Neurosurg., 107:
1168-1189, 2007.
D. A. Hunter, A. Moradzadeh, E. L. Whitlock, M. J. Brenner, T.
M. Myckatyn, C. H. Wei, T. H. Tung, and S. E. Mackinnon,
“Binary imaging analysis for comprehensive quantitative
histomorphometry of peripheral nerve,” J. Neurosci. Methods.,
166: 116-124, 2007.
S. Raimondo, M. Fornaro, F. Di Scipio, G. Ronhi, M. G.
Giacobini-Robecchi and S. Geuna, “Methods and protocols
in peripheral nerve regeneration experimental research: Part II –
morphological techniques,” Int. Rev. Neurobiol., 87: 81-103,
2009.
T. Kiyotani, M. Teramachi, Y. Takimoto, T. Nakamura, Y.
Shimizu and K. Endo, “Nerve regeneration across a 25-mm gap
bridged by a polyglycolic acid-collagen tube: a histological and
electrophysiological evaluation of regenerated nerves,” Brain
Res., 740: 66-74, 1996.
H. Schmalbruch, “Fiber composition of the rat sciatic nerve,”
Anat. Rec., 215: 71-81, 1986.
J. C. Mira, “Quantitative studies of the regeneration of rat
myelinated nerve fibres: variations in the number and size of
regenerating fibres after repeated localized freezings,” J. Anat.,
129: 77-93, 1979.
G. R. Evans, K. Brandt, A. D. Niederbichler, P. Chauvin, S.
Herrman, M. Bogle. L. Otta, B. Wang and C. W. Jr. Patrick,
“Clinical long-term in vivo evaluation of poly(L-lactic-acid)
porous conduits for peripheral nerve regeneration,” J. Biomater.
Sci. Polym. Ed., 11: 869-878, 2000.
W. F. den Dunnen, B. Van der Lei, J. M. Schakenraad, E. H.
Blaauw, I. Stokroos, A. J. Pennings and P. H. Robinson,
“Long-term evaluation of nerve regeneration in a biodegradable
nerve guide,” Microsurgery, 14: 508-515, 1993.
W. F. den Dunnen, I. Stokroos, E. H. Blaauw, A. Holwerda, A. J.
Pennings, P. H. Robinson and J. M. Schakenraad,
“Light-microscopic and electron-microscopic evaluation of
short-term nerve regeneration using a biodegradable
poly(DL-lactide-epsilon-caprolacton) nerve guide.,” J. Biomed.
Mater. Res., 31: 105-115, 1996a.
G. D. Sterne, R. A. Brown, C. J. Green and G. Terenghi,
“Neurotrophin-3 delivered locally via fibronectin mats enhances
peripheral nerve regeneration,” Eur. J. Neurosci., 9: 1388-1396,
1997.
S. Wang, A. C. Wan, X. Xu, S. Gao, H. Q. Mao, K. W. Leong
Bioengineering in Peripheral Nerve Regeneration
[57]
[58]
[59]
[60]
[61]
[62]
[63]
[64]
[65]
[66]
[67]
[68]
[69]
[70]
[71]
[72]
[73]
and H. Yu, “A new nerve guide conduit material composed of a
biodegradable
poly(phosphoester),”
Biomaterials,
22:
1157-1169, 2001.
S. Itoh, K. Takakuda, S. Ichinose, M. Kikuchi and K.
Schinomiya, “A study of induction of nerve regeneration using
bioabsorbable tubes,” J. Reconstr. Microsurg., 17: 115-123,
2001.
W. F. den Dunnen, B. Van der Lei, J. M. Schakenraad, I.
Stokroos, E. Blaauw, H. Bartels, A. J. Pennings and P. H.
Robinson, “Poly(DL-lactide-epsilon- caprolactone) nerve guides
perform better than autologous nerve grafts,” Microsurgery, 17:
348-357, 1996b.
F. A. Howe, A. G. Filler, B. A. Bell and J. R. Griffiths,
“Magnetic resonance neurography, Magn. Reson. Med.,
28: 328-338, 1992.
A. G. Filler, F. A. Howe, C. E. Hayes, M. Kliot, H. R. Winn, B.
A. Bell, J. R. Griffiths and J. S. Tsuruda, “Magnetic resonance
neurography,” Lancet., 341: 659-661, 1993.
F. A. Howe, D. E. Saunders, A. G. Filler, M. A. McLean, C.
Heron, M. M. Brown and J. R. Griffiths, “Magnetic resonance
neurography of the median nerve,” Br. J. Radiol., 67:1169-1172,
1994.
S. A. Cudlip, F. A. Howe, J. R. Griffiths and B. A. Bell,
“Magnetic resonance neurography of peripheral nerve following
experimental crush injury, and correlation with functional
deficit,” J. Neurosurg., 96: 755-775, 2002.
E. Yamabe, T. Nakamura, K. Oshio, Y. Kikuchi, H. Ikegami and
Y. Toyama, “Peripheral nerve injury: Diagnosis with MR
imaging of denervated skeletal muscle-experimental study in
rats,” Radiology, 247: 409-417, 2008.
J. Shen, C. P. Zhou, X. M. Zhonh, R. M. Guo, J. F. Griffith, L. N.
Cheng, X. H. Duan and B. L. Liang, “MR neurography: T1 and
T2 measurements in acute peripheral nerve traction injury in
rabbits,” Radiology, 254: 729-738. 2010.
M. Bendszus, C. Wessig, L. Solymosi, K. Reiners and M.
Koltzenburg, “MRI of peripheral nerve degeneration and
regeneration: correlation with electrophysiology and histology,”
Exp. Neurol., 188: 171-177, 2004.
S. h. Hsu, S. H. Chan, C. M. Chiang, C. C. C. Chen and C. F.
Jiang, “Peripheral nerve regeneration using a microporous
polylactic acid asymmetric conduit in a rabbit long-gap sciatic
nerve transection model,” Biomaterials, 32: 3764-3775, 2011.
S. Itoh, K. Takakuda, S. Kawabata, Y. Aso, K. Kasai, H. Itoh and
K. Shinomiya, “Evaluation of cross-linking procedures of
collagen tubes used in peripheral nerve repair,” Biomaterials, 23:
227-238, 2002.
S. Wang, Q. Cai, J. Hou, J. Bei, T. Zhanh, J. Yang and Y. Wan,
“Acceleration effect of basic fibroblast growth factor on the
regeneration of peripheral nerve through a 15-mm gap,” J.
Biomed. Mater. Res. A., 66: 522-531, 2003.
A. S. Paintal, “Conduction in mammalian nerve fibers,” in: J. E.
Desmedt (Ed.), New Developments in Electromyography and
Clinical Neurophysiology, Basel: Karger, 2: 19-41, 1973.
K. Fugleholm, H. Schmalbruch and C. Krarup, “Post
reinnervation maturation of myelinated nerve fibers in the cat
tibial nerve: chronic electrophysiological and morphometric
studies,” J. Peripher. Nerv. Syst., 5: 82-95, 2000.
X. Navarro and E. Udine, “Methods and protocols in peripheral
nerve regeneration experimental research: Part III.
Electrophysiological evaluation,” Int. Rev. Neurobiol., 87:
105-126, 2009.
F. Biedermann, N. P. Schumann, M. S. Fischer and H. C. Scholle,
“Surface EMG recordings by using a miniaturized matrix
electrode: a new technique for small animals,” J. Neurosci.
Methods, 97: 69-75, 2000.
H. C. Scholle, F. Biedermann, D. Arnold, H. A. Jinnah, R.
Grassme and N. P. Schumann, “A surface EMG multi-electrode
technique for characterizing muscle activation patterns in mice
during treadmill locomotion,” J. Neurosci. Methods, 146:
159
174-182, 2005.
[74] A. Gramsbergen, J. I. Jkema-Paassen and M. F. Meek, “Sciatic
nerve transection in the adult rat: abnormal EMG patterns during
locomotion by aberrant innervation of hindleg muscles,” Exp.
Neurol., 161: 183-193, 2000.
[75] M. F. Meek, J. F. van der Werff, J. P. Nicolai and A.
Gramsbergen, “Biodegradable p(DLLA-epsilon-CL) nerve
guides versus autologous nerve grafts: electromyographic and
video analysis,” Muscle Nerve, 24: 753-759, 2001b.
[76] L. J. Chamberlain, I. V. Yannas, H. P. Hsu, G. R. Strichartz and
M. Spector, “Near-terminus axonal structure and function
following rat sciatic nerve regeneration through a collagen-GAG
matrix in a ten-millimeter gap,” J. Neurosci. Res., 60: 666-677,
2000.
[77] A. W. English, Y. Chen, J. S. Carp, J. R. Wolpaw and X. Y. Chen,
“Recovery of electromyographic activity after transaction and
surgical repair of the rat sciatic nerve,” J. Neurophysiol., 97:
1127-1134, 2007.
[78] S. J. Day and M. Hulliger, “Experimental simulation of cat
electromyogram: evidence for algebraic summation of
motor-unit action-potential trains,” J. Neurophysiol., 97:
1127-1134, 2001.
[79] H. Wang, E. J. Sorenson, R. J. Spinner and A. J. Windebank,
“Electrophysiologic findings and grip strength after nerve
injuries in the rat forelimb,” Muscle Nerve, 38: 1257-1265,
2008.
[80] S. Barbay, E. K. Peden, G. Falchook and R. J. Nudo, “An index
of topographic normality in rat somatosensory cortex:
application to a sciatic nerve crush model,” J. Neurophysiol., 88:
1339-1351, 2002.
[81] R. Nashmi, H. Imamura, C. H. Tator and M. G. Fehlings, “Serial
recording of somatosensory and myoelectric motor evoked
potentials: role in assessing functional recovery after graded
spinal cord injury in the rat,” J. Neurotrauma, 14: 154-159,
1997.
[82] G. García-Alías, E. Verdú, J. Forẻs, R. López-Vales and X.
Navarro, “Functional and electrophysiological characterization
of photochemical graded spinal cord injury in the rat,” J.
Neurotrauma, 20: 501-510, 2003.
[83] L. de Medinaceli, W. J. Freed and R. J. Wyatt, “An index of the
functional condition of rat sciatic nerve based on measurements
made from walking tracks,” Exp. Neurol., 77: 634, 1982.
[84] C. M. Nichols, T. M. Myckatyn, S. R. Rickman, K. Fox, T.
Hadlock and S. E. Mackinnon, “Choosing the correct functional
assay: a comprehensive assessment of functional test in the rat,”
Behav. Brain. Res., 163: 143-158, 2005.
[85] A. S. P. Varejão, M. F. Meek, A. J. A. Ferreira, J. A. B. Patrício
and A. M. Cabrita, “Functional evaluation of peripheral nerve
regeneration in the rat: walking track analysis,” J. Neurosci.
Methods, 108: 1-9, 2001.
[86] J. R. Bain, S. E. Mackinnon and D. A. Hunter, “Functional
evaluation of complete sciatic, peroneal, and posterior tibial
nerve lesions in the rat,” Plast. Reconstr. Surg., 83: 129-136,
1989.
[87] M. Bervar, “Video analysis of standing: an alternative footprint
analysis to assess functional loss following injury to the rat
sciatic nerve,” J. Neurosci. Meth., 102: 109-116, 2000.
[88] A. S. P. Varejão, A. M. Cabrita, S. Geuna, P. Melo-Pinto, V. M.
Filipe, A. Gramsbergen and M. F. Meek, “Toe out angle: a
functional index for the evaluation of sciatic nerve recovery in
the rat model,” Exp. Neurol., 183: 695-699, 2003.
[89] A. Bozkurt, R. Deumens, J. Scheffel, D. M. O’Dey, J. Weis, E. A.
Joosten, T. Führmann, G. A. Brook and N. Pallua, “CatWalk gait
analysis in assessment of functional recovery after sciatic nerve
injury,” J. Neurosci. Meth., 173: 91-98, 2008.
[90] S. Agarwal, J. H. Wendorff and A. Greiner, “Use of
electrospinning technique for biomedical applications,” Polymer,
49: 5603-5621, 2008.
160
J. Med. Biol. Eng., Vol. 31 No. 3 2011