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Designing Tyrosine-Derived Polycarbonate Polymers
for Biodegradable Regenerative Type Neural
Interface Capable of Neural Recording
Dan Lewitus, R. Jacob Vogelstein, Gehua Zhen, Young-Seok Choi, Member, IEEE, Joachim Kohn,
Stuart Harshbarger, and Xiaofeng Jia
Abstract—Next-generation neuroprosthetic limbs will require
a reliable long-term neural interface to residual nerves in the
peripheral nervous system (PNS). To this end, we have developed novel biocompatible materials and a fabrication technique
to create high site-count microelectrodes for stimulating and
recording from regenerated peripheral nerves. Our electrodes are
based on a biodegradable tyrosine-derived polycarbonate polymer
system with suitable degradation and erosion properties and a
fabrication technique for deployment of the polymer in a porous,
degradable, regenerative, multiluminal, multielectrode conduit.
The in vitro properties of the polymer and the electrode were tuned
to retain mechanical strength for over 24 days and to completely
degrade and erode within 220 days. The fabrication technique
resulted in a multiluminal conduit with at least 10 functioning
electrodes maintaining recording site impedance in the single-digit
kOhm range. Additionally, in vivo results showed that neural
signals could be recorded from these devices starting at four weeks
postimplantation and that signal strength increased over time.
We conclude that our biodegradable regenerative-type neural
interface is a good candidate for chronic high fidelity recording
electrodes for integration with regenerated peripheral nerves.
Index Terms—Biodegradable, electrode, interface, neuroprosthetics, peripheral nerve, regenerative.
I. INTRODUCTION
pproximately 1.8 million people were estimated to suffer
limb loss (excluding finger and toe amputees) in 2010 [1]
and more than 185 000 new amputations would be performed in
A
Manuscript received June 22, 2010; revised September 15, 2010 and
November 18, 2010; accepted November 30, 2010. Date of publication
December 10, 2010; date of current version April 08, 2011. This work was supported by the Johns Hopkins University Applied Physics Laboratory under the
Defense Advanced Research Projects Agency 2009 Revolutionizing Prosthetics
program under contract N66001-06-C-8005. The work of X. Jia was also supported in part by the National Institute of Health under Grant RO1 HL071568
and in part by the American Heart Association under Grant 09SDG1110140.
The views, opinions, and/or findings contained in this article/presentation are
those of the author/presenter and should not be interpreted as representing
the official views or policies, either expressed or implied, of the Defense Advanced Research Projects Agency or the Department of Defense. Distribution
Statement “A” (Approved for Public Release, Distribution Unlimited).
D. Lewitus and J. Kohn are with the New Jersey Center for Biomaterials,
Rutgers, The State University of New Jersey, Piscataway, NJ 08854 USA.
R. G. Vogelstein and S. Harshbarger are with the Johns Hopkins University
Applied Physics Laboratory, Laurel, MD 20723 USA.
G. Zhen is with Department of Orthopaedic Surgery, Johns Hopkins University School of Medicine, Baltimore, MD 21205 USA.
Y.-S. Choi was with the Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205 USA. He is now
with the Neural Interface Research Team, Electronics and Telecommunications
Research Institute, Daejeon 305700, Korea .
X. Jia is with Department of Biomedical Engineering, Physical Medicine and
Rehabilitation, Johns Hopkins University School of Medicine, Baltimore, MD
21205 USA (e-mail: xjia1@jhmi.edu).
Color versions of one or more of the figures in this paper are available online
at http://ieeexplore.ieee.org.
Digital Object Identifier 10.1109/TNSRE.2010.2098047
the United States [2]. While many of these patients will receive
prostheses, the rate of prosthetic rejection remains unsatisfactory at 22%–50%, due in part to lack of functional gain [3]. The
two main factors limiting the functionality of prosthetic upper
limbs are the lack of a high-bandwidth information channel conveying the user’s movement intent and the lack of a high-resolution sensory feedback pathway from the artificial limb. To address these needs, research groups have begun to investigate the
feasibility of peripheral nerve interfaces for feedback and control of upper extremity prostheses [3]–[6].
Peripheral nervous system (PNS) interfaces for neural
prosthetics provide a reasonable compromise between highly
invasive cortical implants [7], [8] and noninvasive control
schemes such as those relying on electromyography (EMG)
[9], [10]: they achieve higher signal selectivity than EMG but
do not provide access to the high-level movement parameters
observable in motor cortex. Another option being explored is
a procedure known as targeted reinnervation, where residual
nerve fibers are meticulously separated and attached to muscle
tissue to allow higher fidelity EMG electrode recording [11].
Because the residual peripheral nerves serve limited purpose
after upper-limb amputation, a potentially attractive option is
to implant multichannel electrodes in the residual components
of the peripheral nerves to address the input–output bandwidth
limitations described above. The most invasive form of PNS
interfaces are regenerative electrodes [12], [13], which require
severed nerves to grow and regenerate through an array of electrodes. The benefit of this approach is that it enables direct and
stable contact between individual recording sites and growing
fascicles, and consequently provides some fidelity advantages
over other devices relying on EMG signal. Our group and
others have previously demonstrated recording from residual
nerves with intraneural devices in human amputees with the
use of longitudinal intrafascicular interfaces (LIFEs) [3]–[6]
and microelectrode arrays (MEAs) [14], [15].
The most popular design in regenerative electrodes is
the sieve electrode of various forms [16], [17]. Recently,
long-channel arrays with regenerating nerves growing along
individual channels with embedded recording sites have been
described. Such channels are primarily based on silicon fabrication techniques (photolithography with material deposition)
[18], [19], allowing a larger density of recording sites and a
high signal to noise ratio [20]. These devices retain their compartmental structure indefinitely, possibly leading to long-term
failure due to compressive forces on the maturing nerve, and the
fact that individual fascicles and axons will most likely remain
individualized and isolated from nearby tissue and fluid. Moreover, the constant presence of rigid materials has been known to
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LEWITUS et al.: DESIGNING TYROSINE-DERIVED POLYCARBONATE POLYMERS FOR BIODEGRADABLE REGENERATIVE TYPE NEURAL INTERFACE
205
Fig. 1. Structure of tyrosine-derived terpolymers terpolymers. In our study, the R pendant chain is ethyl.
cause long-term deterioration in maturing regenerating nerves
[21], [22]. The use of softer materials reduces the forces acting
on the nerves and resultant tissue damage [23], yet does not
provide sufficient structural support for the regenerating tissue
to retain its conformation [17], [24]. To address some of these
issues, Cho et al. [25] have lithographically fabricated electrodes and embedded them in a polysaccharide conduit with an
outer shell made of polyurethane, a nondegradable, nonporous
material.
Our aim is to develop a polymeric system that will allow
us to design a biodegradable regenerative-type neural interface (bRTNi) that combines the advantages of many of the
approaches described above. Our bRTNi is a multiluminal
conduit with embedded electrodes. Taking inspiration from
rolled film and multiluminal permeable biodegradable conduit
[26]–[29] together with the efficiency of LIFEs electrodes
[3]–[5], [12], [30]–[33], we have created a regenerative neural
interface. The neural interface acts as a multiluminal nerve
regeneration conduit with embedded electrodes for recording.
Once the polymer degrades, the healthy regenerated nerve
should have embedded electrodes remaining within, allowing
chronic recording. Our polymer system is based on the most
recent generation of the tyrosine-derived polycarbonates. This
family of polymers developed over the last 25 years by us
and other groups [34]. They have been extensively shown to
be biocompatible and biodegradable [35]. Furthermore, acidic
environments are not formed upon breakdown into metabolites
[36], reducing the chances of causing additional inflammation
within injured tissue. The synthesis of copolymers, and more
recently terpolymers with the addition of polyethylene glycol
(PEG) has shown to modify various polymer physical properties
while remaining biocompatible [36]–[38]. We hypothesized
that modification of the polymer backbone chemistry would
allow us to tune the polymer properties to be compatible with
both nerve tissue regeneration rate (1 mm/day) [39], [40] and
mechanical properties. The new polymer, developed along
with a novel design and fabrication technique, will create a
biodegradable scaffold for potential chronic neural recordings.
The functionality of the bRTNi design will be evaluated both
in vitro and in vivo.
II. METHODS
A. Polymers and Preparation of Polymer Films
The polymers based on the amino acid L-tyrosine were
synthesized via solution polycondensation reactions according
to previously published procedures [38], [41]. Polymers
were repeatedly precipitated in a suitable nonsolvent and
vacuum dried to constant weight. All polymers consist of the
basic structure poly (DTR-co-XX%-DT-co-YY%-PEGMW
carbonate)s, shown in Fig. 1, with varying amount of all
three monomers. To simplify the identification of tyrosine-derived polycarbonates, the following notation in the
form of RXXYY(ZK) will be used to specify different poly
(DTR-co-XX%-DT-co-YY%-PEGMW carbonate)s. As an
example, poly (DTE-co-25%-DT-co-2%-PEG2K carbonate)
will have a notation of E2502 (2K), where E stands for ethyl.
For in vitro evaluation of polymer degradation, mass loss, and
mechanical properties, porous solvent cast polymer films were
prepared by the particulate leaching technique. Polymer powder
with the addition of 20% by weight of glucose (G7021, Sigma
Aldrich, St. Louis, MO) was dissolved at 10% w/v in tetrahydrofuran (THF, VWR Scientific Products, Radnor, PA) and shaken
for 4 h until the polymer was completely dissolved. Solutions
were cast into PTFE dishes and left to evaporate overnight under
nitrogen in a chemical hood, followed by seven days of drying in
vacuum at 40 . Films were cut into 5 20 mm samples and
placed in deionized water for three days at room temperature
to allow glucose particulate leaching. Resulting films measured
thick.
200
B. Polymer Degradation, Mass Loss In Vitro, Mechanical
Testing and Morphology Changes
5 40 mm film strips were cut from cast films and placed
in 20 mL scintillation vials (Fisher Scientific, Pittsburg, PA)
containing prewarmed phosphate buffer saline (PBS, pH 7.4,
Sigma Aldrich, St. Louis, MO) and incubated at 37 . At prewere visually indetermined time points, samples
spected for integrity, taken out of incubation and used for mechanical testing, then lyophilized for 96 h to remove all water.
1.5 mL of N,Ndimethylformamide (DMF) containing 0.1% trifluoroacetic acid (TFA) was then added to the lyophilized samfilples and shaken for 6 h, followed by filtration (0.45
ters, Whatman, NJ) and analysis of molecular weight using gel
permeation chromatography (GPC) using previously described
procedures [38], [41]. Molecular weights of degraded samples
were compared to that of pristine polymer film.
To evaluate the mass loss from the polymer films, aliquots
from degradation media were treated with 1 N NaOH to
hydrolyze the terpolymer fragments to DT and PEG, then
neutralized by acidification with 12 N HCl followed by high
pressure liquid chromatography (HPLC). HPLC analysis gave
the amount of DT in the supernatant. The HPLC system consisted of a Waters Alliance 2695 Module, Waters 2487 Dual
Absorbance Ultraviolet (UV) Detector and Empower Pro
Software (Waters Corporation, Milford, MA). The column was
particle
a Perkin-Elmer Pecosphere C18 (33 4.6 mm; 3
size; 0258-1064, Waltham, MA). The mobile phase was a
mixture of water and methanol containing 0.1% TFA with a
gradient of 80:20–40:60 in 6 min. The concentration of DT
in the hydrolyzed aliquot was determined from a DT standard
calibration curve. The amount of DT that had leached into the
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Fig. 2. Fabrication steps of rolled conduit. (1) PDMS mold. (2) placing of wire
electrodes. (3) Casting of polymer solution. (4) Formed film with embedded
wires. (5) Rolled film, proximal nerve end. (6) Rolled film, distal nerve end. (7)
View of constructed conduit; left side attaches to the proximal end of the nerve,
right to the distal.
buffer was a good approximation of the extent of erosion of the
polymer.
Cast films were tested using a servohydraulic biaxial mechanical testing system coupled with a computerized data acquisition system (MTS, Eden Prairie, MN). Using a 100 Newton
load cell, a tensile test was performed uniaxially at a rate of
10 mm/min, five samples per condition in prewarmed media
at 37 . Degradation samples were removed from incubation
media, were measured in the thickness and width dimensions,
and placed in the sample holder for testing. Ultimate tensile
strength and strain are reported. Thickness measurements were
recorded to determine whether swelling occurred during the incubation time in the media.
The effect of particulate leaching and incubation on the morphology of film samples was visualized using scanning electron
microscopy (SEM, Amray 1830, Bedford, MA). Incubated film
was gently washed with deionized water and lyophilized to remove all water then mounted and sputter-coated with gold palladium (SCD 004 Sputter Coating Unit, Balzars, Liechtenstein)
prior to imaging.
C. bRTNi Fabrication and Impedance In Vitro
in diameter (776000,
Teflon coated Pt/Ir microwires 100
A-M systems, Sequim, WA.) were used as electrodes. Wires
were cut to a total length of 150 mm. To provide a recording
site, 1–2 mm of wire was exposed at a distance of 1–3 mm from
the end of the wire. This allowed for the recording site to be at
the inner part of the conduit rather than its face to avoid false
recording from nerves that did not regenerate. Indication of regeneration was assumed once recording occurred, since it required growth of tissue into the conduit. Fabrication steps of
the bRTNi are as shown in Fig. 2. The description is as follows:
1) a polydimethylsiloxane (Sylgard 184, Fisher Scientific, Pittsburgh, PA) negative mold was fabricated using a custom-made
grooves with
aluminum-machined positive mold with 500
100
depth and 100
width; the width of the PDMS mold
was trimmed to 5 mm; 2) the distal end of the wires were tightly
inserted into the mold groves and held by the PDMS structure; not all grooves were filled with wires for these preliminary
studies, leaving blank polymer channels that allowed for maintenance of the open architecture of the conduit and generally improving regeneration through a multi-lumen structure [28]; 3) a
polymer solution containing porogen was dispensed using a syringe with a 27 g needle on top of the mold (0.3 mL), covering
the wires and filling the empty grooves, then dried overnight
under a chemical hood; 4) the formed film was adhered to the
wires and gently removed from the mold using forceps; 5) the
film was rolled to reach 2.5–3.0 mm in diameter, matching the
diameter of the sciatic nerve of a rabbit [42]. Excess material
was cut and the edge of the film was attached to the main conduit through solvent welding with THF. Illustration of rolled
film with embedded wires is seen in Fig. 2 (steps 5 and 6).
To allow additional attachment points and support for the severed nerve ends to the edge of the conduit, two polymer “flaps”
were attached via solvent welding to the sides of the conduit.
Once all solvent-based procedures were finished, the structure
for seven days to remove exwas placed under vacuum at 40
cess THF. The wires protruding from the proximal side of the
conduit were bound together above the structure in a manner
that allowed spacing between them, reducing (to the extent feasible) the obstruction of the distal end of the conduit by the
wires. Wires were placed through a flexible biomedical silicone tubing (807900, A-M systems, Sequim, WA) to bundle and
protect them together. This allowed placing the connector subcutaneously for periodical recordings. Connection of the proximal end of the wires was made by soldering 5 mm of exposed
wires directly to a 16 channel chronic connector with a protective cap (ZCA-OMN16, Tucker Davis Technologies, Alachua,
FL). To avoid infiltration of foreign elements and fluids to the
leading wires and tubes, medical grade silicone adhesive was
used for sealing (MED ADH 4100 RTV, Bluestar Silicones, East
Brunswick, NJ). Removal of glucose from the conduits and generation of porosity was done by sinking tubes in deionized water
for 72 h followed by drying under vacuum for one day, then
sterilization in ethylene oxide followed by 10–14 days under
vacuum at room temperature.
In vitro impedance was measured using a commercial
impedance meter designed for microelectrodes (Electrode
Impedance Tester, Model 2900, AM-systems, Sequim, WA).
Using 2 model bRTNis, each with five embedded electrodes,
the distal ends of electrodes were soldered to a pin dip connector (Mill-Max, Oyster Bay, NY) for ease of connection
to a measuring device. Measurements were performed by
immersing probes in 50 mL of PBS at room temperature using
Ag/AgCl reference electrode (BASi, West Lafayette, IN) placed
50 mm from the tested electrode. Impedance was measured
with an applied 1000 Hz, 20 nA sine wave immediately after
fabrication, particulate leaching, and one week of incubation in
PBS at 37 .
D. In Vivo Feasibility Study: Implantation,
Electrophysiological Recording and Signal Processing
We have evaluated the implantation procedure and signal
recording potential of fabricated devices using an experimental
protocol approved by the Johns Hopkins Animal Care and Use
Committee (ACUC) and the US Army Medical Research and
LEWITUS et al.: DESIGNING TYROSINE-DERIVED POLYCARBONATE POLYMERS FOR BIODEGRADABLE REGENERATIVE TYPE NEURAL INTERFACE
Material Command Animal Care and Use Review Office. All
procedures were compliant with NIH guidelines.
Three New Zealand rabbits, weighing an average of 4.1 kg
(3.8–4.4 kg), were anesthetized by administering ketamine
(10 mg/kg) and xylazine (5 mg/kg) intravenously, followed
by 1.0%–2.0% isoflurane/oxygen applied using a facemask,
adjusted to maintain the anesthetized state of the animal
throughout the procedure. Strict sterile surgical rules were
followed during all experiments. Rabbits were shaved and
prepared in the usual fashion. An incision was made along
the posterior of the left hind leg at 5 cm and the sciatic nerve
was exposed. A tunnel was formed under the skin to allow the
covered electronic connector to be placed subcutaneously. The
sciatic nerve was cut 1 cm proximal to the bifurcation point of
the sciatic nerve in the popliteal fossa. The epineurium of both
nerve stumps were sutured first to the device flaps using 9-0
suture under microscope, four times on each side, then directly
to the face of the nerve guide conduit. The muscular and skin
incisions were closed by layers. The animals had free access
to food and water before and after the experiments, and were
subjected to a 12 h day/night cycle in a quiet environment.
At predetermined periods, the rabbit was reanesthetized
with ketamine and xylazine, shaved and prepared in the usual
fashion. At the site of the distal connector touched by the
surgeon’s finger, a new 0.8 cm incision was made along the
skin in the left hind leg. The connector was carefully exposed
and interfaced with a Tucker Davis Technologies ZIF16 headstage connected via a preamplifier. The neuron spike signal
in the sciatic nerve from the electrode was recorded continuously for 30 min with 24.14 kHz sampling frequency (RX 5,
Tucker-Davis Technologies, Alachua, FL), downsampled to
12 kHz, and fourth-order Butterworth bandpass filtered with
forward-backward, zero-phase lag with cutoff frequencies of
700–2000 Hz. A wavelet-based denoising method [43] was
used to increase the signal-to-noise ratio. The skin incision was
closed by layers. The firing rates of the action potentials were
analyzed offline and quantified with MATLAB 7.0.
E. Statistical Methods
Statistical analysis was performed using a computerized statistical package (Statistics Program for the Social Sciences version 16.0, Chicago, IL). Group parametric values were reported
as
and repeated measure of general linear analysis was performed for parametric data with Student’s t-test for
continuous variables.
III. RESULTS
A. Polymer Molecular Weight, Mass Loss, Mechanical
Properties and Morphology Changes
All polymers used had starting molecular weight averages of
over 150 kD. Fig. 3(A) illustrates the loss in molecular weight
(MW) during the first four weeks for series of polymers initially evaluated for use for the bRTNi fabrication. It is clear that
all polymers experienced a sharp and immediate loss in MW
within the first week, followed by a slower pace of degradation.
Polymers which lost more than 50% of their original MW after
one week of incubation were omitted from further evaluation.
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Fig. 3. Fractional molecular weight loss versus time. (A) Four week data collected all polymers. (B) 32 weeks of data collected for promising polymers
E1001(2K) and E1002(2K). R values for logarithmic regression are 0.90 and
0.99, respectively. (C) Fraction mass loss and linear regression for E1001 (2K)
and E1002 (2K). R values are 0.85 and 0.47, respectively.
The two promising polymers, E1001(2K) and E1002(2K), were
continually evaluated for up to 230 days as shown in Fig. 3(B).
By the last time point, the E1002(2K) had completely eroded
in the media (see mass loss data) and its molecular weight was
reduced by over 90%. The E1001(2K) lost over 85% of its original molecular weight, but did not erode in a significant manner.
These findings are confirmed by the mass loss data presented in
Fig. 3(C). Clearly the E1001(2K) has minimally eroded. The
films were still intact (though fragile to touch). In contrast, the
E1002(2K) had completely eroded, and linear regression of the
E1002 (2K) data reveals total erosion time of 210 days.
Ultimate stress and strain of incubated films are shown in
Fig. 4. The mechanical properties test required measurement
of dimensions and mounting onto tensile testing clamps, requiring sample mechanical integrity. The zero time point is
post glucose leaching. With the faster degrading polymers
(those with 15% DT or more), the films collapsed during the
second collection. For that reason, only the data for the slower
degrading polymers is presented, together with a single example of E1502(2K). The E1001(2K), even though was slower
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IEEE TRANSACTIONS ON NEURAL SYSTEMS AND REHABILITATION ENGINEERING, VOL. 19, NO. 2, APRIL 2011
Fig. 5. (A) Dimensional retention (lack of swelling) of E1002(2K) films. Incubated sample thickness was measured against the undegraded samples. (B)
SEM images of film samples after particulate leaching, formation of pores is
visible (B) after fabrication (C), (D), (E) 10, 21, and 28 days incubation, respectively) connections between pores are formed, materializing as fissures. Scale
bar 100 .
m
Fig. 4. Over three weeks of ultimate strain (top) and stress (below) results for
E1002(2K) compared to E1001(2K) and E1502(2K).
to degrade according to GPC data, was too weak to handle after
17 days. The E1002(2K) kept its integrity for an additional
week. The surprising difference in mechanical properties retention could be attributed to the elevated amount of PEG in
the E1002(2K). Even through the addition of PEG accelerates
the degradation rate of the polymer; it also acts as a plasticizer,
increasing the compliance of the polymer and allowing additional time of use. The ultimate stress was usually followed by
a reduction in strength until complete failure. While visually
inspecting the film during testing, a small tear was initially
observed accompanying the reduction in strength, followed by
tear propagation and strength reduction till complete failure.
Dimensional retention is presented as a measure of the effect
of incubation time on the morphology of degraded films in
Fig. 5(A). This data was collected from measurements performed prior to the mechanical testing. Data depicts that the
fabrication of the films (prepared at different batches) was
consistent, and swelling did not occur (no statistical difference
was detected). In the SEM scans in Fig. 5, visual change in
porosity is observed after particulate leaching [Fig. 5(B)]. With
additional incubation time, connecting fissures between pores
are observed, possibly leading to loss of mechanical strength
[Fig. 5(C) and (D)].
B. bRTNi Fabrication and Impedance In Vitro
Images of fabricated probes are shown in Fig. 6. In Fig. 6(A),
the proximal part of the bRTNi is shown. A structure of 12 channels is visible. 10 channels are formed by active electrode wires
Fig. 6. (A) Cross section view to the proximal part of regenerative electrode
tube, arrows mark electrode location, circles mark blank polymer channels. (B)
Profile of bRTNi showing lead wires encased in a protective tube. (C) Uncapped
ZIF-16 connector. (D) bRTNi with added flaps positioned after implantation.
Inserts show distal and proximal end of nerves in respect to the device. (E) Representative specimen of harvested bRTNi after eight weeks.
(marked with arrows). Two are blank polymer (marked with circles). In the background, the lead tube containing the wires is
visible. In Fig. 6(B), the same bRNTi is shown from its profile.
Fig. 6(C) shows the ZIF connector. Fig. 6(D) illustrates the final
structure of the design placed implanted within a rabbit sciatic
nerve. Fig. 6(E) illustrates an explanted device after eight weeks.
bRNTi is still intact, and nerve shows signs of integration in the
device.
Recorded impedance values are presented in Table I. In 2 of
the 10 electrodes, the initial impedance was three orders of magnitude higher than in the average of the rest. Once particulate
leaching was initiated, all impedance values dropped to a similar range. It is possible that during the fabrication, a thin layer
of polymer solution surrounded some of the electrodes. As the
LEWITUS et al.: DESIGNING TYROSINE-DERIVED POLYCARBONATE POLYMERS FOR BIODEGRADABLE REGENERATIVE TYPE NEURAL INTERFACE
209
Fig. 7. (A) Representative 10 s plot of single channel four weeks postimplantation. (B) Representative 10 s plot of single channel eight weeks postimplantation. (C) Representative plots of multichannels of peripheral nerve spiking activities eight weeks postimplantation. (D) Firing rates of multichannels eight weeks
postimplantation.
TABLE I
bRTNi IMPEDANCE CHANGE AFTER PARTICULATE LEACHING
At two weeks after implantation, no obvious neural spike
signal was detected. Signals started to appear at four weeks
postimplantation [Fig. 7(A)], and became stronger at eight
weeks postimplantation [Fig. 7(B)–(D)]. The average firing
rate of the multichannel recorded signals was higher at eight
weeks postimplantation (21.7 2.2 spikes/s) than four weeks
.
postimplantation (12.9 1.9 spikes/s)
IV. DISCUSSION
polymer is a dielectric material, it may have caused the formation of an insulating layer around the recording site. Once the
polymer is exposed to the aqueous environment, this thin layer
is removed (via degradation/erosion), allowing the exposure of
the recording site and reduction in impedance values.
C. In Vivo Electrophysiological Feasibility Study
Representative neuronal spikes are shown in Fig. 7. Neural
, and
spike signals with signal-to-noise ratios
peak-to-peak amplitudes from 40–120
were successfully
recorded for at least eight weeks following chronic implantation.
Our study demonstrates that a biodegradable terpolymer is
able to be tuned to degrade at a rate that is slow enough to possibly match neural regeneration, yet fast enough to allow nerve
freedom once regeneration is complete. Our polymer system is
based on novel variation of the tyrosine-derived polycarbonate
terpolymers. These are known to be biocompatible [36], [44],
[45] and their tunable properties are primarily influenced by
polymer composition [36], [38], [46], [47]. Unlike conventional
biodegradable polymers, they do not form acidic environments
as they degrade, preventing localized toxicity [36], [48]. The
addition of PEG blocks modulated water absorption of the
polymer due to their intrinsic hydrophilic properties [47],
which in turn modulated degradation rate and mechanical
properties (enhanced flexibility). The addition of DT monomer
increases the initial rigidity of the polymer, yet also significantly increases the degradation rate of the polymer. The DTE
block allows for stability and strength. Fine tuning of the three
monomer compositions is required to achieve optimization
of degradation rate and structural properties for a specific
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applications. A wide variety of polymers and designs have been
applied for degradable conduits. From simple hollow tubes
[33] to multiluminal or grooved tubes [49], and fiber-filled
tubes [50], [51], where the lumens and fibers act as guiding
objects for axonal regrowth. Interestingly, the degradation rates
of the polymers have not been reported. In one particular study,
where the degradation rate of collagen tubes was associated
with the quality of critical gap regeneration, a 2–3 week half
life for implants was found to be ideal. Our chosen polymer,
E1002(2K), was found in vitro to lose 50% of its molecular
weight within two weeks, maintain its mechanical integrity
for over three weeks, and completely erode within 25 weeks.
Feng et al. suggest preferred degradation of the polymer within
12 weeks [31]. Consistent with other studies [52], [53], our
degradation/erosion evaluations were performed in vitro. In the
case of mass loss, in vivo reduction in polymer mass may be
accelerated due to greater mass transport and the presence of
solubilizing factors [54].
Among the many technologies explored for direct neuronal
interfacing, the regenerative device could have great benefits
due to its exploitation of the regenerative property of amputated
peripheral nerves. This can occur even without the presence of a
distal stump and specifically by the assistance of muscular reinnervation [30], [55], [56]. The use of longitudinal regenerative
channels with recording ability has been of high interest and
could result in high fidelity recording; a few designs of this nature have been suggested [19], [25], yet with none of them is
the conduit is fully degradable. We have used Pt/Ir based wires
for their availability, simplicity of use, and proven record as
functional intrafascicular electrodes [32]. Our fabrication technique allows for future adoption of use of polyLIFE [57] as the
recording moiety (replacing the Pt/Ir wires in the conduit). Replacing the current Pt/Ir wires with polyLIFE could provide: 1)
an increase in the mechanical match between nerves and electrodes remaining after conduit resorption, 2) an increase in the
number of electrodes available for recording in the conduit due
to their smaller size, and 3) minimized obstruction of the distal
end of the conduit by the lead wires to the connector. This will
result in a fully degradable conduit with remaining free electrodes, and could be made possible through minor modifications
such as change in groove depth and spacing in the PDMS mold
design.
An ideal regenerative scaffold should obtain mechanical
properties that are at the same order of magnitude of those
of the inherent tissue, allowing healthy regeneration through
initial protection of the growing organ. Avoidance of stress
shielding at the latter stages of regeneration is also required.
In environments where external forces may be applied on the
implantation site, failure of the device itself (an “adhesive”
failure) is preferred over the failure of the regenerating tissue (a
“cohesive” failure). This minimizes the potential of inflicting
additional damage on the regenerating tissue. The strength of a
regenerative device should reduce with time in accordance to
the tissue becoming healthier and stronger. These were guidelines used in the selection criteria for the polymer. Healthy
peripheral nerves under tension exhibit an initial toe region for
the first 5%–15% of strain [58] and a 12 MPa [59] ultimate
strength. We identified that the E1002(2K) maintained strength
that is just under that of a nerve for over three weeks (12 to 4
MPa in 24 days). The E1002(2K) strain fits within the range
of the toe region of a nerve (12% to 4%), thus avoiding a
mechanical properties mismatch. Reports that severed nerves
retain over 50% of their recovered tensile strength within 1–4
weeks of repair (i.e., approximately 6 MPa) [60] allow us to
assume that our polymer is suitable for use as a regenerative
construct for injured nerves.
Measurements of sample dimensions prior to mechanical
testing revealed little swelling of the E1001(2K) and E1002(2K)
during the incubation period. The lack of swelling indicates
dimensional stability that is essential for nerve guide conduits,
as polymer swelling could lead to steric obstruction of nerve
growth and even nerve compression [61].
We have been able to use simple a polymer fabrication technique to demonstrate the potential in fabricating a high electrode
count biodegradable conduit. In vitro impedance data revealed
recording potential that is maintained over time. Action potentials from severed sciatic nerves were successfully recorded in
real time. We have combined best-of-breed concepts from existing regeneration-type sieve electrodes, nerve guidance channels and biodegradable polymers. In this work, we did not address the long term biocompatibility effect of the implantation
on the physiological health of the tissue. However, this study
was intended as a proof of design with a limited in vivo characterization. Our main goal was to develop a biodegradable regenerative-type neural interface and successfully demonstrate
the recording potential of our design, which were both achieved.
Future work will allow our design to further serve its multiple
function potential by: 1) providing a microstructured mechanical scaffold for regenerating fibers, 2) promoting neurite ingrowth through the application of electric fields, 3) applying
chemical mediators to allow for unrestricted axonal growth, and
4) positioning active electrode sites within closer proximity of
nerve fibers. In addition, successful development of this technology will have implications for nerve repair in clinical settings, as variations of the proposed devices could be used in
place of autologous nerve grafts. Future long-term and larger
in vivo studies, including quantitative biocompatibility research
are needed to verify the long-term in vivo effect.
V. CONCLUSION
This study has shown that our biodegradable polymer created via our unique fabrication technique obtains the mechanical and degradation retention properties required from nerve regeneration apparatus. An eight-week successful recording session has demonstrated that our biodegradable regenerative type
neural interface is a good candidate for chronic high fidelity
recording electrodes for integration in the regeneration of peripheral nerves. This study also provides additional new control sources for the future development of new nerve signal controlled prosthetics.
ACKNOWLEDGMENT
The authors would like to thank Dr. R. Rojas for his help in
the polymers synthesis within this study and M. Fifer for his
editorial support.
LEWITUS et al.: DESIGNING TYROSINE-DERIVED POLYCARBONATE POLYMERS FOR BIODEGRADABLE REGENERATIVE TYPE NEURAL INTERFACE
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Dan Lewitus received the B.S. degree in plastics engineering from the Shenkear College of Engineering
and Design, Israel, in 2000, and the M.S. degree in
plastics engineering from the University of Massachusetts, Lowell, in 2004. He is currently working
toward the Ph.D. degree in biomedical engineering
at the New Jersey Center for Biomaterials, Rutgers
University, New Brunswick, NJ.
His research involves developing biodegradable
polymer systems for neural prosthetics both in the
central and peripheral nervous systems. In between
his degrees and prior to his Ph.D. studies, he worked both in Israel and Germany
on developing of performance enhancing additives for the polymer processing
industry.
R. Jacob Vogelstein received the Sc.B. degree in
neuroengineering from Brown University, Providence, RI, and the Ph.D. degree in biomedical
engineering from the Johns Hopkins University
School of Medicine, Baltimore, MD.
He is the Assistant Program Manager for Applied
Neuroscience at the Johns Hopkins University
(JHU) Applied Physics Laboratory and an Assistant
Research Professor in Electrical Engineering at
the JHU’s Whiting School. He has worked on
neuroscience technology for over a decade, focusing
primarily on neuromorphic systems and closed-loop brain–machine interfaces.
His research has been featured in a number of prominent scientific and engineering journals including the IEEE TRANSACTIONS ON NEURAL SYSTEMS AND
REHABILITATION ENGINEERING, the IEEE Transactions on Biomedical Circuits
and Systems, and the IEEE TRANSACTIONS ON NEURAL NETWORKS.
Gehua Zhen received the M.D. degree in clinical medicine from Shanghai Medical University,
Shanghai, China, in 1997. She completed her
surgery residency and fellowship and was an
attending surgeon in the Shanghai 6th People’s
Hospital, Shanghai. She was a postdoctoral fellow in
Johns Hopkins University School of Medicine from
2005 to 2010.
She has been a Faculty in the Department of Orthopedic Surgery, Johns Hopkins University School
of Medicine, Baltimore, MD, since 2010.
Young-Seok Choi (M’07) received the B.S. degree in electronic and electrical engineering from
Hanyang University, Seoul, Korea, in 2000, and the
Ph.D. degree in electrical and computer engineering
from Pohang University of Science and Technology
(POSTECH), Pohang, Korea, in 2007. From 2007
to 2010, he was a Postdoctoral Fellow in the Department of Biomedical Engineering, Johns Hopkins
University School of Medicine, Baltimore, MD.
Since July 2010, he is with the Neural Interface
Research Team, Electronics and Telecommunications Research Institute, Daejeon, Korea, where he is currently a Senior
Researcher. His current research interests include neural signal processing,
brain–computer interface, neural interface, and neural engineering.
Joachim Kohn is a Board of Governors Professor of Chemistry at Rutgers University. He has served as Director of the New Jersey Center for Biomaterials
since its establishment in 1997. He is a Fellow of the American Institute for
Medical and Biological Engineering (AIMBE) and the Chair of the International
College of Fellows of Biomaterials Science and Engineering. He is the principal
investigator of several leading federally-funded R&D programs: NIH-funded
postdoctoral training program in Tissue Engineering, NIH funded National Resource for Polymeric Biomaterials (RESBIO), the DoD-funded Center for Military Biomaterials Research (CeMBR) and the DoD-funded Armed Forces Institute for Regenerative Medicine (AFIRM). His research interests focus on the
development of new biomaterials. He pioneered the use of combinatorial and
computational methods for the optimization of biomaterials for specific medical
applications. He is mostly known for his seminal work on "pseudo-poly(amino
acid)s". He has published over 200 scientific manuscripts and reviews, and holds
45 patents.
Stuart Harshbarger, photograph and biography not available at the time of
publication.
Xiaofeng Jia received the M. D. degree in clinical
medicine from Zhejiang Medical University, Zhejiang, China, in 1994, the M.S. degree in surgery
from Shanghai Medical University, Shanghai, China,
in 1997, and the Ph.D. degree in surgery (orthopedics) from Fudan University, Shanghai, China, in
2003.
He has been a Faculty in the Department of
Biomedical Engineering, Physical Medicine and
Rehabilitation, Johns Hopkins University School
of Medicine, Baltimore, MD, since 2007. He completed his surgery residency in the Huashan Hospital and Orthopedic Surgery
fellowship in the Shanghai 6th People’s Hospital, Shanghai, China. Later,
he was an Attending Orthopedic Surgeon in Shanghai 6th People’s Hospital
and Zhongshan Hospital, Shanghai. His current research interests include
novel application of neuro-electrophysiology for detection and restoration of
peripheral nerve and spinal cord injury, therapeutic hypothermia of brain and
spinal cord after global ischemia injury.
Dr. Jia is a member of the American Academy of Orthopaedic Surgeons,
American Association for Hand Surgery, Society of Critical Care Medicine,
American Heart Association. He is a recipient of the 2008 Annual Research
Awards from American Association for Hand Surgery, the 2009 Top 20 Scientific Exhibits and the 2008 Finalist of Best Overall Poster Award of American
Academy of Orthopaedic Surgeons, the 2007 Cardiopulmonary, Perioperative
and Critical Care Junior Investigator Travel Award from American Heart Association, the 2008 Finalist of Research Citation Award from Society of Critical
Care Medicine.