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Reichert WM, editor. Indwelling Neural Implants: Strategies for Contending with the In Vivo Environment. Boca Raton (FL): CRC Press/Taylor & Francis; 2008.

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Indwelling Neural Implants: Strategies for Contending with the In Vivo Environment.

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Chapter 7Soft, Fuzzy, and Bioactive Conducting Polymers for Improving the Chronic Performance of Neural Prosthetic Devices

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7.1. INTRODUCTION

Microfabricated electrodes for stimulating and recording signals from individual neurons have facilitated direct electrical connections with living tissue. While these devices have worked reasonably well in acute applications, chronically implanted electrodes have had more limited success [1,2]. To improve the long-term integration of these devices, coatings have been developed to accommodate the differences in mechanical properties, bioactivity, and mechanisms of charge transport between the engineered electronic device and living cells [3–10]. Conducting polymers can be directly deposited onto electrode surfaces with precisely controlled morphologies. The coatings lower the impedance of the electrodes and provide a mechanical buffer between the hard device and the soft tissue. These coatings can be tailored to incorporate and deliver pharmacological agents such as anti-inflammatory drugs and neurotrophic factors. In vivo studies to date have shown that these coatings improve the long-term recording performance of cortical electrodes [11].

In this review we first discuss the development of neural prosthetic devices, including the history of their development, issues associated with the electrode–tissue interface, inflammation and neural loss in the tissue near the electrode surface, the mechanical property differences between the probe and the tissue, the geometry of the probe, and materials used to modify the electrode surface. We then discuss the design of materials for the electrode–tissue interface to help these probes function more effectively over the long term. These materials are intended to improve device performance by creating a mechanically compliant (soft), high-surface-area (fuzzy), low-impedance electrode–tissue interface that can have controlled biological functionality. We conclude by describing the results of work to date that have focused on the design, synthesis, and characterization of electrode interface materials, with particular attention to the use of conducting polymers that have been shown to significantly improve the electrical properties at these interfaces.

7.2. OVERVIEW OF NEURAL PROSTHETIC DEVICES

Over the past 50 years, many different types of neural prosthetic devices have been used to record and stimulate neural signals in the central nervous system (CNS) and peripheral nervous system (PNS) [12,13]. Implanted microelectrodes of various designs, including microwires [14,15] and micromachined electrodes (Utah electrodes and Michigan electrodes) are currently being used for the treatment of deafness (cochlear implants) [16] and Parkinson’s disease (deep brain stimulation) [17,18].

The elastic modulus of silicon is near 100 GPa, whereas that of brain is on the order of 100 kPa, akin to Jell-O [19,20]. This corresponds to a stiffness variation of approximately six orders of magnitude. The mismatch of stiffness at the tissue–device interface has the potential to create large interfacial strains during the lifetime of chronic implants. To avoid sharp interfaces between materials, it is necessary to provide a gradient of mechanical properties (Figure 7.1a). This interfacial mechanical mismatch has been addressed by applying deposition of electrospun nanofibers of polymer on the microelectrode arrays [3], and genetically engineered protein polymer materials have been applied to the devices to mediate the mechanical differences between the device and brain tissue with structural stability of natural silk with responsive properties (Figure 7.1b) [21]. Flexible polyimide-based microelectrode arrays are being fabricated that minimize the stress resulting from micromotion of stiff probes [22]. Recently, the Kipke group developed a cortical probe design that has the electrodes held off to the side of the main probe body [23] to float the electrode on a thin supporting member. The thicker, main part of the probe would still be stiff enough to facilitate insertion into tissue. Histological studies have shown a significant decrease in the amount of inflammation seen around the smaller lateral shank [23]. Although this design has not yet been implemented on functional probes, these results support the idea that electrodes with reduced stiffness increase the integration of the electrode into the tissue.

FIGURE 7.1. (a) Schematic of a modulus gradient in a polymer film that could help mediate the large differences in mechanical properties between a stiff prosthetic device and soft tissue [3].

FIGURE 7.1

(a) Schematic of a modulus gradient in a polymer film that could help mediate the large differences in mechanical properties between a stiff prosthetic device and soft tissue [3]. (b) Scanning electron microscope images of nanofibrous coatings of an SLPF (more...)

To understand the effects of the device design and the insertion method on cellular sheath development, a study of chronically implanted, single-shanked, chisel-tipped microelectrodes with trapezoid and square cross sections as well as blade-type, single-shanked microelectrodes from the Center for Neural Communication Technology (CNCT) (University of Michigan) was performed. These studies demonstrated that long-term reactivity is essentially independent of electrode size, shape, surface texture, and insertion method [24,25]. Clinical applications of neuroprosthetic devices could, however, benefit from miniaturization of these devices since the volume of surrounding tissue associated with early reactive response is associated with the amount of tissue damage during insertion.

During the insertion of electrodes into the brain, rupturing of blood vessels and damage to surrounding tissue is inevitable. For example, a typical Michigan probe measuring 15 μm × 60 μm × 2 mm will displace 0.0018 mm3 of cortical tissue, resulting in displacement of about 50 neurons and 400,000 synapses, assuming that the human cortex contains approximately 30,000 neurons and 0.24 billion synapses in a cubic millimeter [26]. The mechanical trauma of electrode insertion initiates the wound healing process and subsequent cellular encapsulation around the implant, leading to decreased signals and eventual failure of communication between neurons and electrodes. It is hypothesized that the primary role of the glial scar is to separate the damaged tissue from the rest of the body to maintain the blood–brain barrier and prevent lymphocyte infiltration [27]. Szarowski et al. showed that this response includes an early reactive component observed immediately upon device insertion and a sustained reactive component that develops with time and is maintained as long as the implant is present [24].

The sustained reactive response, a major source of failure in chronically implanted electrodes, is associated with the characteristics of the surface materials of the implants. Sapphire [28] and alumina (Al2O3) [29] for the substrate of micro-machined electrodes and polyesterimide-coated gold wire [30] have been used to improve performance of the electrodes. Electrostatic layer-by-layer (LbL) self-assembly techniques have been used for the surface modification of neural electrodes [31]. They used alternating polyelectrolytes, either polyethyleneimine or chitosan, and proteins, either laminin or gelatin, to fabricate multilayer films on silicon wafers. Huber et al. suggested other techniques such as adsorption, covalent coupling, and electrochemical polymerization that could be used to coat laminin-derived peptides on glassy carbon surfaces [32]. Silk-like polymers containing fibronectin fragments (SLPF) and nonapeptide (CDPGYIGSR) were immobilized into polypyrrole (PPy)-conducting polymers to enhance electric transportation and adhesion of neurons [4]. Microcontact printing, a new technique of chemically and molecularly patterning surfaces on a submicrometer scale, was also used to modify silicon substrates with polylysine [33]. Other studies have been performed with coatings containing living cells designed to better integrate the devices with living tissue. For example, Martin and Tresco implanted electrodes coated with olfactory ensheathing cells derived from the olfactory bulbs of adult Fischer 344 rats into the cerebral cortex for a period of 2 weeks. It was found that the reactivity surrounding cellular-coated electrodes was significantly reduced compared to the controls [34].

7.3. RECORDING SITE–NEURON INTERFACE

Signal transmission along neurons is the result of ionic currents generated by movement of ions via specific ion channels in the cellular membrane. When a neuron generates an action potential, the current flows within the cell and leaks though regions of membrane. The flow of current across the membrane generates a complex potential field around the neuron. The magnitude and orientation of this field depend on the size, geometry, and location of the cell and the time course of depolarization [35]. The time course of this potential difference is what gives rise to the traditional extracellular action potential. The action potentials are transient waveforms with a typical duration of 1 msec.

Extracellular recording methods have been used to obtain data about the properties of CNS structures, including the mapping of field potentials from a single neuron to the excitability of CNS dendrites and studying behaviorally related discharge patterns of CNS neurons [36]. In addition, extracellular recording makes it possible to monitor the activity of multiple neurons in the vicinity of the electrode. Although conduction at the electrode–electrolyte interface is not yet fully understood, it involves a capacitive mechanism (charging and discharging of the electrode double layer) and a Faradaic mechanism (chemical oxidation and reduction). In general, the diffusion and recombination of ionic species from the liquid is a nonlinear charge-transfer process [37,38]. Figure 7.2 shows Michigan electrodes chronically implanted into the auditory cortex of a guinea pig. Five to ten seconds of continuous neural recordings are collected using a National Instruments data acquisition card coupled to a Plexon multichannel acquisition processor running RASPUTIN. The auditory cortex recordings are driven by 200-ms noise bursts presented 2 per second using a digital to analog converter (Tucker-Davis Technologies). The round cap on the guinea pig’s head covers and protects the chronic electrodes. The electrode connector is placed under the cap. The rod-shaped screw next to the cap of the electrodes functions as a ground when the neural spikes are recorded (Figure 7.2).

FIGURE 7.2. Michigan electrodes chronically implanted into auditory cortex of guinea pig and extracellular recording of neural signals.

FIGURE 7.2

Michigan electrodes chronically implanted into auditory cortex of guinea pig and extracellular recording of neural signals.

It is known that the amplitude of the extracellular spike is much smaller than the corresponding intracellular spike, because the magnitude of the extracellular current is smaller to begin with and diminishes rapidly as a function of distance from the cell [39]. To record extracellular potentials with the greatest signal and spatial selectivity, electrodes are required to have recording sites with dimensions as small as 20 μm, and it is highly recommended that they be placed as close as possible to the firing neuron. Henze et al. claims that at distances >50 μm from the soma, individual spikes could be recognized with extracellular recordings (the reliability of unit separation for these signals is decreased significantly). At distances >140 μm from the soma, extracellular spikes could not be distinguished from the background noise level [40].

7.4. BIOMATERIAL COATINGS ON IMPLANTED ELECTRODES

In considering the design of materials for the interface between living tissue and an engineered electronic device, it is necessary to consider the dramatic differences in structure and properties between these two systems. Living tissue is wet, whereas biomedical devices are usually solids. Tissue is usually quite soft, whereas devices are often hard. Tissue conducts electron charge by ionic transport, whereas devices conduct charge with electrons and holes. Any material that functions at the interface between the biotic tissue and abiotic device will therefore need to take these large variations of behavior into account. Conducting polymers are particularly attractive for this purpose because they have mechanical properties between those of the metal electrode and the tissue and are able to facilitate charge transport with various cationic and anionic species. They can also be complexed with a variety of biologically active counter-ions. The ultimate limits of conducting polymers for use in biosensors were discussed by Goepel, who described the concept of a “molecular wire” that could facilitate signal transduction from biological tissue to an electrically conducting metal substrate [41].

Our research group has been developing materials and processes for improving the long-term performance of electronic biomedical devices, with particular interest in the microfabricated, implantable cortical probes that have been developed at the University of Michigan. We began by focusing on genetically engineered protein polymers that combine amino acid sequences from structural proteins such as silk and elastin with binding sequences from matrix proteins such as fibronectin and laminin. It was found that continuous films of these polymers were not as useful as filamentous structures that could be created by a novel process called electrospinning. Electrospinning creates high-surface-area structures that promote cellular ingrowth (Figure 7.3). This open microstructure also made it possible to maintain carrier transport to the device [42] and provided a mechanically compliant surface for interfacing with the soft tissue of the brain [3]. Electrospun filaments can also be loaded with biologically active molecules for controlled release into tissue [10,43–45].

FIGURE 7.3. SEM images of electrospun filaments of protein polymers with tailored morphologies.

FIGURE 7.3

SEM images of electrospun filaments of protein polymers with tailored morphologies. The diameters of the filaments created by electrospinning can be controlled by changing the solution concentration, electric field, and distance from the substrate. The (more...)

However, one important limitation of the electrospinning process is that the polymer coats the entire probe with fibers. Indeed, any type of nonconducting polymer coating will not be useful for these microfabricated electronic devices, since the electrodes need to remain extremely sensitive for single-cell recording. For improving the electrical transport properties of the probe, and for controlling the composition of the material in the vicinity of the recording site, we have since developed schemes that have allowed them to exert more direct control on the environment closer to the active sites on the probe. Specifically, we have investigated conducting polymers that can be electrochemically deposited directly on the electrodes [4].

A key requirement for a polymer to become intrinsically conductive is the molecular orbital overlap to allow formation of a delocalized molecular wave function. Conducting polymers have a delocalized, extended φ-bonded system of electrons resulting from the conjugation of alternating single and double bonds along the molecular backbone. Applications of conducting polymers include analytical devices and biosensors [46–51]. They have afforded new surface modification strategies for functionalization of conventional metal electrodes for creating pharmacological and toxicological biochemical sensors. Conducting polymers have also been utilized for light-emitting diodes [52], photovoltaic solar cells [53], lightweight batteries [54], antistatic coatings, and electrochromic devices [55].

Among the currently available inherently conducting polymers (Figure 7.4a), PPy (Figure 7.4b) has been extensively investigated because it is highly conductive, easily oxidized, and electropolymerizable from water. The deposited PPy film is reasonably stable and adherent to the electrode [56,57]. The biocompatibility of PPy has been investigated in vitro and in vivo [5,58,59]. PPy has been used to modify bioelectronic devices [4,60–62]P and to create biosensors by immobilization of biological elements [63] including enzymes [64,65], antibodies [66,67], and DNA [67].

FIGURE 7.4. Structures of conducting polymers (a) and schematics of the oxidative electrochemical polymerization of polypyrrole (b) and poly(3,4-ethylenedioxythiophene) (c).

FIGURE 7.4

Structures of conducting polymers (a) and schematics of the oxidative electrochemical polymerization of polypyrrole (b) and poly(3,4-ethylenedioxythiophene) (c). The polymers have a net positive charge in their electrically active state and are typically (more...)

Electrochemical polymerization is the preferred technique for the synthesis of conducting polymers for coatings on biomedical devices, since it is reproducibly controlled, provides the highest conductivities, and deposits the polymer only on specified areas. As shown in the reaction scheme (Figure 7.4b,c), the pyrrole and ethylenedioxythiophene (EDOT) monomer are electrochemically polymerized into the polymer PPy and poly(3,4-ethylenedioxythiophene) (PEDOT). The oxidation of monomers by the application of current to the monomer solution forms cationic radicals that eventually lead to polymerization. Electrochemical polymerization of conducting polymers is performed by application of constant current (galvanostatic), constant potential (potentiostatic), or potential scanning or sweeping methods. The total electrical charge passing through the electrode drives this stoichiometric reaction, leading to coatings of precisely defined composition, thickness, and micro-structure. The polymerization reaction is typically performed in a three-electrode cell including a counter electrode (anode), a reference electrode, and a working electrode (cathode). Metals such as gold, titanium, platinum, and chromium are often used for the counter electrode [68].

Cui et al. established that electrochemical polymerization can be used to deposit coatings of electrically conducting polymers such as PPy directly onto metal neural electrode sites [4] (Figure 7.5). Cui et al. also found that the impedance magnitude can be tailored by changing the applied current and that the lowest impedance could be obtained with a PPy thickness of 10 to 12 μm, depending on the dopant. This optimum thickness is apparently related to the fact that the impedance is profoundly influenced by charge carrier transport across the electrode–tissue interface. Thinner films are flat, and the effective surface area at the interface between the electrode sites and neural tissue is relatively small. During growth, the surface roughens, increasing the effective surface area of the interface [4] (Figure 7.6).

FIGURE 7.5. Optical micrographs of electropolymerization of PPy on a neural prosthetic device seen from the above (a) and from the side (b).

FIGURE 7.5

Optical micrographs of electropolymerization of PPy on a neural prosthetic device seen from the above (a) and from the side (b). The thickness of the conducting polymer film can be precisely tailored by controlling the solution concentration, current (more...)

FIGURE 7.6. SEM images of PPy–SLPF-coated neural electrodes.

FIGURE 7.6

SEM images of PPy–SLPF-coated neural electrodes. From (a to d), the deposition time was increased at constant current, so the total amount of charge that passed through during the deposition increased. (a) Bare gold, (b) 1 mC, (c) 4 mC, (d) 10 (more...)

Neural electrodes are often used for chronic applications that demand the chemical and electrical stability of the implanted materials. Yamato et al. found that PPy has limited electrical response under cyclic voltammetry because of its poorly defined chemical structure [69]. Heywang and Jonas reported the synthesis of a variety of poly(alkylenedioxythiophenes), including PEDOT [70]. PEDOT exhibits improved conductivity and thermal stability because the dioxylethylene bridging groups across the 3 and 4 positions of the EDOT monomer block the possibility of α-β′ coupling. Yamato et al. reported that PEDOT–polystyrenesulfonate (PSS) was more chemically stable than PPy–PSS [69]. After polarization for 16 h at 400 mV and pH 7.5, only 5% of the original electrochemical activity of PPy–PSS remained, whereas PEDOT–PSS retained 89% of its original activity [69]. Along with PPy, the PEDOT family of polymers has also been intensively investigated for neural prosthetic applications [71–74]. Xiao et al. also investigated PEDOT derivatives such as poly(hydroxymethylated-3,4-ethylenedioxylthiophene) (PEDOT-MeOH) and PEDOT doped with its derivative sulfonatoalkoxyethlyenedioxythiophene (SEDOT) to overcome the low water solubility of EDOT that results in limited immobilization of biologic agents into PEDOT [72–74].

It has been reported that the effective surface area of the neural electrode site is crucial in determining its electrical properties and in providing a gradient of mechanical properties that promotes better integration with cells [5,71]. Yang et al. explored a number of methods to create features of well-defined size and high surface area of conducting polymers using templating techniques. A variety of nanostructured conducting polymer morphologies have been fabricated as coatings including the nanomushroom, nanohair, and nanopore structures [6,7,75,76] (Figure 7.7). Highly ordered microporous PEDOT-conducting polymer coatings fabricated using polystyrene latex sphere templates had the lowest impedance value (~10 kΩ), with 300-nm scale-polystyrene spheres [6]. These spheres had the smallest diameters that still retained relatively high order in the film, creating an open-cell foam that facilitates charge transport. The oxidation potential for microporous PEDOT–LiClO44B was −0.7 V versus a saturated calomel electrode (SCE), which is significantly lower than that of microporous PPy–LiClO4 (−0.4 V versus SCE) and PEDOT–PSS (−0.4 V versus SCE) and PPy–PSS (0.1 V versus SCE) [71]. These lower oxidation potentials are critical in preventing degradation from biological reducing agents in living tissue [76]. Yang et al. also developed PEDOT films that can be electrochemically deposited through self-assembled nonionic surfactant films on surfaces of neural electrodes [77], following methods reported by Hulvat and Stupp [78].

FIGURE 7.7. SEM images of a variety of nanostructured conducting polymer morphologies fabricated using templating techniques including nanomushroom (a); nanohair (b); and nanopore (c) structures.

FIGURE 7.7

SEM images of a variety of nanostructured conducting polymer morphologies fabricated using templating techniques including nanomushroom (a); nanohair (b); and nanopore (c) structures.

The performance of polymer-coated electronic devices involves the transfer of charge through a variety of interfaces and through materials with different species that are the mobile charge carriers. In metal electrodes charge transport involves the motion of electrons, whereas in conducting polymers the charge is predominantly transported by local positive charges that move along the molecular backbone (holes or polarons) (Figure 7.8). In water-laden tissue, charge transport involves the motion of positively charged cations or negatively charged anions (Figure 7.8). Transfer reactions involving exchange of charge between these various carriers occur at the metal–polymer and polymer–tissue interfaces [38]. It is reasonable to expect that the detailed composition and microstructure of these various interfaces will be critical for the performance of electronic biomedical devices implanted in living, ionically conducting tissue. In particular, it is anticipated that increasing the effective surface area will allow for many more opportunities for such charge transfer to occur, significantly lowering the overall impedance of the device.

FIGURE 7.8. Schematic diagram showing the mechanisms of charge transport near the electrode–tissue interface under different potential bias.

FIGURE 7.8

Schematic diagram showing the mechanisms of charge transport near the electrode–tissue interface under different potential bias. With negative bias, there is electron transport through the polysilicon connectors and metal electrode. There is a (more...)

Electrochemical impedance spectroscopy measurements have shown that the charge transport characteristics of the polymer coatings are extremely sensitive to the surface morphology. The rough, fuzzy structure of the electrochemically deposited conducting polymer leads to a significant (~2 orders of magnitude) decrease in the sample impedance. Quantitative comparisons of the impedance response as a function of temporal frequency have shown correlations with the surface topology of the coatings as measured by atomic force microscopy (AFM) [79]. It has also been shown that the microstructure and electrical properties of the coatings correlate with their mechanical behavior, as measured by nanoindentation [80]. The fuzzy, low-impedance coatings also have the lowest effective modulus, consistent with a more open physical architecture.

PEDOT-coated microelectrodes have recently been studied in vivo. The results showed high-quality spike recordings at 6 weeks postimplant from PEDOT-coated electrodes. The signal-to-noise ratios for the PEDOT-coated sites were higher than the uncoated iridium electrode sites, and there was a 20% increase in the average number of units recorded per site [11].

Polymer processing methods have proven to be relatively easy to scale up and have been used on many types of probe geometries. In addition to chronic probes being actively designed and fabricated by the CNCT at Michigan, multishank neural probes (Utah electrode arrays [UEAs]) intended for implantation into feline dorsal root ganglia (DRG) have also been coated with conducting polymers [81,82]. The conducting polymers have also been investigated for applications such as deep brain stimulation (DBS), cochlear implants, and pacemakers. In the future it may also be possible to adapt similar approaches to other devices for which it is important to maintain electrical contact with living tissue such as electroencephalogram skin sensors or glucose sensors.

7.4.1. Incorporation of Biologic Species into Conducting Polymers

A biosensor is a device that has a biological sensing element capable of producing a signal that is a function of the concentration of a specific chemical or set of chemicals. Specifically, the monitoring of metabolites such as glucose, urea, cholesterol, and lactate is of central importance in clinical diagnostics. Conducting polymers have attracted much interest as suitable immobilization matrices for bio-sensors because the electrochemical incorporation of bio-species into the conducting polymers permits the localization of these molecules on the electrodes of any size or geometry. A number of techniques such as physical adsorption [83], entrapment [84], cross-linking [85], and covalent bonding [86] have been used to immobilize biological molecules in conducting polymers.

Biomaterials need to be designed to physically support tissue growth and to elicit desired receptor-specific responses from specific cell types. One way to achieve these requirements is to incorporate biological molecules into synthetic materials. It has been shown that PPy is able to support the in vitro growth and differentiation of multiple cell types including neurons [58] and endothelial cells [87,88]. By choosing appropriate biologically active dopants, the properties of conducting polymers can be tailored for specific cell and tissue interactions.

For example, Collier et al. showed that hyaluronic acid, a hydrated glycosamino-glycan found in almost all extracellular tissues in the body, can be polymerized with PPy [89]. They assessed in vitro cellular response with PC 12 (pheochromocytoma) cells and in vivo response of the subcutaneous implantation of these materials in rats [89]. They suggested that the conductivity loss of a hyaluronic–PPy composite can be circumvented by using a bilayer approach. Garner et al. studied the incorporation of heparin, a potent anticoagulant known to promote endothelial cell growth, into PPy and assessed the resulting materials as substrates for endothelial cell growth. They suggested that when polymers were grown under conditions of equal total charge but at different current densities, those grown at higher current densities incorporated more heparin because the charge neutralization of PPy by heparin molecules that were already incorporated into PPy required some rearrangement of conformation that was kinetically limiting at higher rates of polymerization (i.e., higher current density) [87].

The incorporation of biomolecules into conducting polymers has been used for surface modification of implanted neural microelectrodes. Cui et al. synthesized PPy containing biomolecules including a silk-like protein with fibronectin fragments (SLPF) and a nonapeptide CDPGYIGSR onto microelectrode sites [4]. Incorporation of the nonapeptide CDPGYIGSR into PPy facilitated the growth of neuroblastoma cells. In addition, the higher surface area of the electrode sites facilitated charge transport, which is crucial for effective neural communication [4]. The stability of PPy–CDPGYIGSR coatings was tested in deionized water soaking experiments, where it was found that the peptides entrapped in PPy did not diffuse away within 7 weeks. More intensive in vivo assessment was subsequently performed on these materials, and it was found that significantly more neurofilament-positive staining was present on the peptide-coated electrodes than controls, indicating that the coatings had established strong connections with neurons. After 1 week in deionized water, 83% of the coated electrodes showed positive immunostaining for neurofilaments, while only 10% of the uncoated electrodes had this staining. At 2 weeks, 67% of the coated and 5% of the uncoated electrodes showed neurofilament staining [5].

Kim et al. incorporated nerve growth factor (NGF) as a counter-ion in the electrochemical deposition of PPy and PEDOT and evaluated the ability of NGF-incorporated PPy to elicit specific biological interactions with the neurons [90]. Impedance measurements at the biologically relevant frequency of 1 kHz revealed that the minimum impedance of the NGF-modified PPy film, 15 k., was lower than the minimum impedance of the peptide-modified PPy film (100 k.). The PC-12 cells attached to the conductive PPy and PEDOT had extended neurites, indicating that the NGF in the polymer film remained biologically active (Figure 7.9). Thus, the incorporation of NGF can modify the biological interactions of the electrode without compromising the conductive properties of the polymeric film [90].

FIGURE 7.9. PC12 cells seeded on control PEDOT (A) as a control, on PEDOT–collagen (B), and PEDOT–NGF (C).

FIGURE 7.9

PC12 cells seeded on control PEDOT (A) as a control, on PEDOT–collagen (B), and PEDOT–NGF (C). Cells were incubated with each polymer substrate for 48 hours prior to imaging. Cells with neurites indicate that the NGF incorporated in the (more...)

Richardson et al. incorporated the neurotrophin NT-3 into PPy films and showed increased neurite outgrowth from auditory neuron explants in vitro [168]. Electrical stimulation was also shown to have a beneficial impact on neural regeneration. These films are being investigated as coatings on cochlear implants.

7.4.2. Hydrogels

Hydrogels have been widely used in biomaterials applications because they can be highly swollen with water and have mechanical properties that are similar to those of living tissue. Alginate is a well-known polysaccharide obtained from brown algae that is widely used for drug delivery and tissue engineering. Alginate often serves as a delivery vehicle of cells for tissue engineering applications because of its bio-compatibility, low toxicity, and simple gelation with divalent cations such as CaP2+, MgP2+, and SrP2+. Alginates are a family of natural copolymers of β-D-mannuronic acid (M) and α-L-guluronic acid (G) [91]. A potential limitation of alginate gels in tissue engineering is the inherent resistance to protein adsorption and cellular attachment because of its hydrophilic character and lack of specific integrin-binding sequences such as the RGD tripeptide from fibronectin or the IKVAV and YIGSR sequences from laminin [92]. This nonfouling property may be useful for limiting the extent of glial scar formation on the neural electrodes after implantation.

Alginate hydrogels cross-linked with calcium sulfate (CaSO4) have been used as cell delivery vehicles because of their slow process of gelation. Alginate beads are commonly produced by fast gelation with calcium chloride (CaCl2) since it is more easily controlled than CaSO4-induced gelation. However, this rapid gelation often results in a nonuniform structure [93]. To overcome the gradients of cross-link density and mechanical properties due to CaCl2, CaCO3 has been used to form more structurally uniform gels for use as scaffolding materials [94]. In the presence of other ions such as Na+, ionically cross-linked alginate gels may lose mechanical strength or may swell via a process involving lost divalent ions into the surrounding medium and subsequent dissolution. The extent to which an alginate gel exhibits swelling depends on the ratio of Na+ to Ca2+ and the composition of the alginate itself [95].

To improve integration with living tissue, hydrogels have been used to deliver and release growth factors and bioactive agents. Hydrogels containing biotrophic molecules such as vascular endothelial growth factor (VEGF) [96], basic fibroblast growth factor (bFGF) [97], epidermal growth factor (EGF) [98], and bone morphogenetic protein (BMP) [99] have been developed.

Kim et al. developed hydrogel coatings to improve the functionality of chronically implanted neural recording electrodes. An alginate hydrogel was used as the coating material on microelectrode arrays for better integration and mechanical buffering between the electrodes and CNS tissue (Figure 7.10). Kim et al. have confirmed that alginate hydrogels can be coated onto the surfaces of neural probes by a dipping process. The coating thickness can be varied from less than a micron to over 100 μm. During tissue insertion, problems can occur if the hydrogel coatings are too soft. In this case the coating can be sheared off the probe, remaining at the surface of the cortex. However, it has been found that it is possible to dehydrate the hydrogel and then allow the material to reabsorb water after insertion. The hydrogels have been shown to readily reswell after insertion into agar tissue phantoms (Figure 7.10) [9].

FIGURE 7.10. (a) Hydrogel coating deposited onto a microfabricated neural probe.

FIGURE 7.10

(a) Hydrogel coating deposited onto a microfabricated neural probe. (b) Multilayered hydrogel coating prepared by sequential dipping. Scale bar is 50 μm. (Adapted from Kim, D. H. et al., Biomaterials 27(15), 3031–3037, 2006. With permission.) (more...)

Acute studies of in vivo recordings from hydrogel-coated probes implanted in guinea pig brain were recently conducted by Kim et al. in collaboration with David J. Anderson and James Wiler at the Kresge Hearing Research Institute. Kim et al. examined the quality of recording as a function of the thickness using 200-msec noise bursts and found a significant reduction in the number of high-quality units recorded even for relatively thin (5 to 10 μm) films (Figure 7.11). Evidently, even these relatively thin hydrogels can push the neurons away from the electrode site. These results clearly demonstrated the importance of the local proximity of cells to the electrode surface and confirmed the need to establish intimate connections with the neurons to maintain signal quality [8]. Kim et al. also evaluated the use of PEDOT conducting polymer to maintain the functionality of hydrogel-coated microelectrodes. PEDOT deposited on the electrode site restored the lost functionality of the electrodes caused by hydrogel coatings as shown by the number of clearly detectable units and the signal-to-noise ratio (Figure 7.12).

FIGURE 7.11. Decrease in fraction of clearly detectable units in guinea pig cortex for microfabricated neural probes coated with hydrogels of various thickness.

FIGURE 7.11

Decrease in fraction of clearly detectable units in guinea pig cortex for microfabricated neural probes coated with hydrogels of various thickness.

FIGURE 7.12. Average signal-to-noise ratios with various coatings including no coating, hydrogel (HG) coating, PEDOT deposition on the electrode sites, and PEDOT deposition on the electrode sites under the HG coatings (one-way ANOVA).

FIGURE 7.12

Average signal-to-noise ratios with various coatings including no coating, hydrogel (HG) coating, PEDOT deposition on the electrode sites, and PEDOT deposition on the electrode sites under the HG coatings (one-way ANOVA).

In designing materials that can facilitate charge transport through low-density structures, Kim et al. also found that conducting polymers can be grown in the hydrogels, resulting in open, extended networks that grow out from the electrode site into the hydrogel matrix. The conducting polymer precipitates onto the low-density hydrogel network as a scaffold. The result is an extremely interconnected, nanoporous, high-surface-area film (Figure 7.13). The impedances of these porous conducting polymer films are around three orders of magnitude less than the initial impedance of the metal electrode (Figure 7.14). Kim et al. also found that these films can be readily dehydrated and reswollen without apparent influence on the microstructure or electronic transport properties [9]. Furthermore, the conducting polymers could still be readily grown through the hydrogel after disrupting the microstructure by freeze-drying. Impedance measurements at the biologically important frequency of 1 kHz showed that the minimum impedance of this polymer- modified hydrogel was 7 k. (Figure 7.14). This was much lower than the minimum impedance of PPy-coated electrodes (~100 k.) [9].

FIGURE 7.13. Optical microscope images of (a) film-like structure of conducting polymer PPy deposited on the electrode sites without a hydrogel scaffold.

FIGURE 7.13

Optical microscope images of (a) film-like structure of conducting polymer PPy deposited on the electrode sites without a hydrogel scaffold. (b) Schematic of the film-like conducting polymer PPy. (c) Conducting polymer PPy grown through a hydrogel scaffold. (more...)

FIGURE 7.14. Impedance of PPy conducting polymers with different morphologies including bare gold, PPy, PPy in lyophilized hydrogel (LHG), and PPy in hydrogel (HG).

FIGURE 7.14

Impedance of PPy conducting polymers with different morphologies including bare gold, PPy, PPy in lyophilized hydrogel (LHG), and PPy in hydrogel (HG).

Anti-inflammatory steroids such as dexamethasone (DEX) are of interest to reduce the tissue reaction. Kim et al. investigated the release of DEX from alginate hydrogel matrices and compared it with that from free nanoparticles (NPs) and NPs immobilized in the hydrogel matrix [100]. DEX-loaded poly(lactic-co-glycolic acid) (PLGA) NPs with typical particle size ranging from 400 to 600 nm were prepared using a solvent evaporation technique. The in vitro release of DEX from NPs entrapped in the hydrogel showed that 90% of the drug was released over 2 weeks (Figure 7.15). The impedance of NP-loaded hydrogel coatings on microfabricated neural probes was measured and showed negligible increases over 3 weeks (Figure 7.16). In vivo impedance of chronically implanted electrodes loaded with DEX was maintained at a relatively constant level, while that of control electrodes increased by three times about 2 weeks after implantation until it stabilized at approximately 3 M. [100] (Figure 7.16).

FIGURE 7.15. Amount of dexamethasone released from hydrogel without being incorporated into nanoparticles, from free nanoparticles, and from nanoparticles encapsulated into a hydrogel.

FIGURE 7.15

Amount of dexamethasone released from hydrogel without being incorporated into nanoparticles, from free nanoparticles, and from nanoparticles encapsulated into a hydrogel. (Adapted from Kim, D. H. et al., Biomaterials 27(15), 3031–3037, 2006. (more...)

FIGURE 7.16. Impedance as a function of time for a dexamethasone-loaded, hydrogel-coated neural probe implanted into guinea pig cortex, compared with an uncoated control.

FIGURE 7.16

Impedance as a function of time for a dexamethasone-loaded, hydrogel-coated neural probe implanted into guinea pig cortex, compared with an uncoated control.

Tresco’s group at the University of Utah has provided quantitative information about the nature and extent of cellular reactivity that occurs in vivo after the hydrogel-coated probes have been implanted into rat cortex for various periods of time (1 week, 2 weeks, and 12 weeks). They have confirmed the formation of a most proximal layer of microglia directly on the probe surface, followed by a layer of reactive astrocytes. There was also a reduction in the number of neurons in the corresponding area close to the probe. The microglia remained most active when the probes were mechanically tethered to the skull, supporting the hypothesis that mechanical strains from micromotion of the probe are involved in exacerbating the chronic inflammation [101]. Their most recent data also confirmed a substantial decrease in the amount of inflammation around hydrogel-coated probes.

7.4.3. Nanotubes

There is currently considerable interest in the development of carbon-based nanotubes for a variety of applications. Several groups have investigated the use of carbon nanotubes for the surfaces of neural prosthetic devices. It has been found that polymer-modified carbon nanotubes can promote the differentiation and proliferation of NG108-15 and PC12 neuroblastoma cells [102,103]. Although there is some ability to functionalize the surfaces of these carbon nanotubes, the interactions of these stiff, rigid rods with living cells has not yet been determined. Another limitation is that most current methods of preparing nanotubes lead to a distribution in diameters and thus variations in physical properties (such as metallic or semiconducting). Methods to separate carbon nanotube mixtures based on their diameter should make it possible to better control their properties in the future [104]. Hybrid materials can also be produced. For example, it has recently been reported that nanocomposite coatings can be prepared by the electrochemical polymerization of conducting polymers such as PEDOT around carbon nanotubes [105].

Abidian et al. successfully established methods for the preparation of drug-loaded conducting polymer nanotubes [10]. The fabrication process involved the electrospinning of a biodegradable polymer (PLGA) into which a drug (dexa-methasone) had been incorporated followed by electrochemical deposition of the conducting polymer around the drug-loaded electrospun biodegradable polymers (Figure 7.17). The diameters of the electrospun nanofibers ranged from 40 to 500 nm, with the majority between 100 and 200 nm. The wall thickness of the PEDOT nanotubes varied from 50 to 100 nm, and the nanotube diameter ranged from 100 to 600 nm (Figure 7.17a,b). By controlling the polymerization time, tubular structures with thin walls (shorter deposition time) or thick walls (longer deposition time) could be reproducibly prepared. The initial impedance of the bare gold sites before surface modification was 800 ± 20 k. for acute probes (1250 μm2). This value of impedance was decreased to a minimum of 8 ± 2 k. by formation of conducting polymer nanotubes on the electrode sites.

FIGURE 7.17. (a) SEM image of PEDOT nanotubes prepared by electrochemical polymerization around PLGA nanofibers.

FIGURE 7.17

(a) SEM image of PEDOT nanotubes prepared by electrochemical polymerization around PLGA nanofibers. (b) Close-up view of the end of a single PEDOT nanotube after removal of the nanofiber template. (c) Schematic of the actuation of a nanotube under an (more...)

Abidian [10] also found that individual drugs and bioactive molecules could be precisely released at desired points in time by using electrical stimulation of nanotubes (Figure 7.17c,d). When a conducting polymer is exposed to an external electrical potential, ions will diffuse in or out of the material to balance the local electrostatic charge [106]. The extent of expansion or contraction depends on the number, size, and mobility of ions exchanged [107]. Electrochemical actuators using conducting polymers based on this principle have been developed by several investigators [108–110].

In collaboration with Joseph Corey, it has also been successfully demonstrated that aligned conducting polymer nanotubes can directionally guide the neurite out-growth of DRG explants and PC12 cells (Abidian et al, unpublished).

7.4.4. Hybrid Live Cell-Conducting Polymer Coatings For Neural Electrodes

The pursuit of a long-term, therapeutic brain–device interface continues to motivate advancements in the design and development of implantable neural prosthetic devices. However, unpredictable device performance associated with limited bio-compatibility and poor tissue integration remains a barrier to successful testing and implementation [14,24,25,111–113]. To function properly once implanted, neural prosthetic devices rely on their ability to establish and maintain direct, functional communication with neurons in the surrounding tissue. This highlights the central importance of an intimate electrode–tissue interface and presents a challenge to bridging the biotic–abiotic interface by joining electronically and ionically conductive systems [115,116]. Intimate integration of electrodes with surrounding tissue will facilitate charge transfer between electrodes and target cells as well as increase biocompatibility.

Integration at the device–tissue interface can be increased through the use of bioactive or biomimetic materials that can physically and biochemically interact with surrounding tissue [117–119]. However, for use in implanted bioelectric devices these materials should also maintain or improve the electronic–ionic communication between the device electrodes and the tissue. Therefore, in recent years studies have focused on developing strategies to increase tissue integration, electrical sensitivity, and charge transfer capacity at the device–tissue interface through the use of inherently conductive polymers and conducting polymer–protein composites as low-electrical-impedance, bioactive coatings for microelectrodes on biomedical devices [59,62,74,89,103,120,121].

Numerous studies indicate that neural electrode functionality can be increased by modifying the surface of the electrode sites with low-impedance conductive polymer coatings with nanoscale roughness or porosity [4,6,7,11,79] and through the incorporation of cell adhesion peptides [4], proteins [31,90,122], or anti-inflammatory drugs [100,123]. Together these studies suggest that the most benefit could be gained by multifunctional modifications of the electrode surface that have increased electrical activity, bioactivity, mechanical softness, and topological features on a similar scale to that of cells in tissues and cell surface and extracellular matrix structures. These ideas also evoke the possibility that the biofunctionality and bio-compatibility of the electrode could be further increased by incorporating living cells or cellular components into an electrode coating to exploit unique cellular physiology and signal transduction capabilities.

Recent studies have found that living neural cells can be incorporated directly into a matrix of the conducting polymer, PEDOT, while still maintaining cell viability and signal transduction capabilities. This has resulted in the generation of functional hybrid PEDOT–neural cell electrode coatings as well as a method of using neural cells to “template” PEDOT films to create highly biomimetic conductive substrates with cell-shaped features that are also cell attracting. Electrical characterization of the conducting polymer matrix containing live neural cells suggested a relationship between the electrode and neural cells that is distinct from a more typical configuration used for electrically interfacing neurons in which neural cells are cultured on or near metal electrodes. Intimate interactions between the conducting polymer and the neuronal membrane were revealed as PEDOT covered delicate filopodia and neurites. This unique cell-conducting polymer–electrode interface may be an ideal candidate material for the development of a new generation of bio-sensors and “smart” bioelectrodes. The ability to polymerize PEDOT in the presence of living cells has been confirmed in vitro around living cells [165] as well as in vivo through living tissue [166].

In earlier studies described by the Wallace group, PPy was used to generate a novel biosensor composed of PPy doped with erythrocytes for detection of blood Rh-factor via the Rhesus factor antigens on the cell surface [124]. The erythrocyte-containing PPy bound three times as much antibody as unmodified PPy as detected by ELISA and resistometry. This study indicates that cell-conducting polymer matrices can perform sensitive and specific biosensing. However, Campbell et al. [124] found that the presence of erythrocytes within the PPy matrix did not alter the electrical properties of the PPy. This is likely because unlike neural cells, erythrocytes are nonadherent and nonelectrically active cells. The incorporation of electrically responsive, electrode-adherent cells into a conducting polymer matrix provides for an additional opportunity to utilize both the biochemical and electrochemical qualities of the incorporated cells for sensing purposes.

In preparation for experiments involving the polymerization of PEDOT in the presence of living cells, it was first determined whether PEDOT and its monomer EDOT were toxic to cells. Previous studies have shown that cells can be cultured for days to weeks on PEDOT and PPy with little or no toxicity [58,77]. However, the effect of the EDOT monomer on cell viability was not known. It was found that neural cells (SH-SY5Y neuroblastoma-derived cell line) could be exposed to as much as 0.01 M EDOT and 0.02 M PSS (polyanionic dopant in monomer solution) for as long as 72 hours while maintaining 80% cell viability (Figure 7.18). In our studies, cells were typically exposed to EDOT for less than 10 minutes, so the cytotoxicity was negligible.

FIGURE 7.18. Working concentrations of EDOT monomer are not cytotoxic.

FIGURE 7.18

Working concentrations of EDOT monomer are not cytotoxic. MTT cytotoxicity assay for exposure of SH-SY5Y neural cells to increasing concentrations of EDOT in monomer solution (all with 0.02 M PSS) for 0 to 72 hours. (Adapted from Richardson-Burns, S. (more...)

PEDOT was electrochemically polymerized directly in the presence of live neural cells cultured on electrodes (Au, Au/Pd, or ITO) using 0.5 to 1 uA/mm2 galvano-static current from a monomer solution containing 0.01 M EDOT and 0.02 M PSS in phosphate buffered saline (PBS) (Figure 7.19a,b). This resulted in the formation of PEDOT on the electrode, surrounding and embedding the cells (Figure 7.19c,d,e). The morphology and topology of the PEDOT polymerized around the neural cells was assessed using optical microscopy and scanning electron microscopy (SEM). After deposition, PEDOT appeared as a dark, opaque substance around the cells. The cells themselves and their nuclei remained intact throughout and following polymerization (Figures 7.19c and 7.19e, respectively). Interestingly, PEDOT deposition was prohibited in areas where cells were evidently strongly adhered to the substrate (Figure 7.19c). Using SEM, it was found that the PEDOT on the electrode and around the cells displays the fuzzy, nodular surface topology that is typical of PEDOT (Figure 7.20A). The polymer also appeared to wrap around the exterior of the cells and their extensions (Figure 7.20B,C).

FIGURE 7.19. The conducting polymer PEDOT can be electrochemically polymerized in the presence of living cells.

FIGURE 7.19

The conducting polymer PEDOT can be electrochemically polymerized in the presence of living cells. (a) Diagram representing the electrochemical deposition cell and the neural cell monolayer cultured on the surface of the metal electrode prior to polymerization. (more...)

FIGURE 7.20. SEM images of PEDOT polymerized around neural cells.

FIGURE 7.20

SEM images of PEDOT polymerized around neural cells. (A) PEDOT polymerized in the presence of mouse primary dissociated cortical cultures (MCC). PEDOT (rough, nodular texture) covers the electrode surface as well as some cellular processes. (B) Higher-magnification (more...)

Neural cells partially embedded in PEDOT maintained their viability for almost 1 week, suggesting that the PEDOT matrix was not a significant barrier to cell nutrient transport. However PEDOT-surrounded neurons eventually began to die by apoptosis that can be triggered as long as 24 to 72 hours after the initial insult [125,126]. One of the more dramatic findings in cells surrounded by PEDOT was substantial disruption of the cytoskeleton, specifically a loss of F-actin stress fibers that can be detected as early as 2 hours after polymerization and is complete by 24 hours after polymerization (Figure 7.21A–H). These fibers are physically and biochemically associated with integrins and other protein complexes at focal adhesions that are the primary mediators of cell surface interactions with the extracellular matrix and neighboring cells [127]. This observation provides insight about the molecular-level interactions at the plasma membrane–PEDOT interface and suggests that the presence of the polymer so near the membrane may disrupt integrin signaling and focal adhesion maintenance [128,129]. Furthermore, loss of F-actin stress fibers is indirect evidence of the activation of biochemical signaling pathways involving focal adhesion kinase (FAK), jun-N terminal kinase (JNK), and src (tyrosine kinase oncogene), each of which has been implicated in apoptosis [130–132]. This may explain why cells embedded in PEDOT undergo apoptosis. It also gives us molecular targets for future studies to attempt to interrupt focal adhesion disruption, block development of cytoskeletal abnormalities, and rescue cells from death. In addition, studies of neuronal apoptosis have indicated that alterations in actin cytoskeletal morphology can be associated with oxidative stress in neurons [133]. The PEDOT electropolymerization process may involve production of free radicals at or near the surface of the neurons on the electrode, so future studies will also explore whether oxidative stress plays a role in disruption of the actin cytoskeleton in neurons embedded in PEDOT.

FIGURE 7.21. The F-actin cytoskeleton is disrupted in cells embedded in the PEDOT matrix.

FIGURE 7.21

The F-actin cytoskeleton is disrupted in cells embedded in the PEDOT matrix. (A) Bright-field image of SY5Y cells cultured on electrode (negative control; no PEDOT, not exposed to current). (B) SY5Y cells cultured on electrode (negative control; no PEDOT, (more...)

PEDOT was polymerized in the presence of live neural cells to generate conductive polymer substrates with biomimetic topology consisting of cell-shaped holes and imprints on the same scale as cell surface features. Following polymerization of PEDOT around the neurons, the cells and cell material were removed from the PEDOT matrix using enzymatic and mechanical disruption. This resulted in a neural cell-templated, fuzzy PEDOT material with a combination of nanometer and micrometer scale features (Figure 7.22A,B,C). The neural cell-templated polymer topography included neuron-shaped holes (Figure 7.22A) and tunnels, crevasses, and caves (Figure 7.22B,C) resulting from conductive polymer molded around cell bodies and extended neurites. Through use of this method, evidence was found of intimate contact at the interface between the PEDOT matrix and plasma membrane of the cells as exemplified in Figure 7.22, in which the PEDOT (dark substance) revealed nanometer-scale tendrils at the leading edge of a neurite (Figure 7.21D,E,F).

Figure 7.22. Neuron-templated PEDOT films are cell-attracting, conductive substrates.

Figure 7.22

Neuron-templated PEDOT films are cell-attracting, conductive substrates. (A) Optical image of neural cell-templated PEDOT film on electrode shows numerous cellshaped holes and neurite-templated channels left behind following removal of cells from the (more...)

It was hypothesized that the biomimetic surface of the cell-templated PEDOT would be attractive to cells because of its nanometer scale fuzziness and the unique cell-shaped holes and imprints. Therefore it was tested whether new cells seeded on top of the cell-templated PEDOT would show evidence of repopulating the cellshaped holes or of increased adhesion to the cell-templated surface. It was found that SY5Y cells cultured on the neuron-templated PEDOT substrate showed a preference for adhering to the cell-templated zones over the regions of untemplated PEDOT (Figure 7.22A,B,C). A subset of cells did seem to repopulate the cell-shaped holes of the film (Figure 7.22A,B,C), however these cells did not settle down into the exact position of the original cells used for templating. These findings suggest that when implanted in tissue, this cell-templated polymer surface may encourage cells in the host tissue to adhere near or within the cell-shaped holes and send processes into the tunnels and crevasses. This would provide for very intimate contact between cells and the conductive polymer, making possible continuous electrical contact between the electrode and the tissue. The use of different methods for removing cells results in variation in the amount of cell material that remains associated with the PEDOT matrix. This provides an opportunity for spatially localized biochemical control of interactions between target cells and the electrode at the cellular-and subcellular-length scale. When coupled with the mechanical control provided by the cytomimetic topology, tailoring of the biochemistry of the cell-templated surface could make possible precise manipulation and tracking of neurite guidance, growth, and signal transduction.

7.4.5. Surface Chemical Characterization by X-Ray Photoelectron Spectroscopy (XPS)

Recently, an XPS study comparing Baytron P, a form of commercially available PEDOT–PSS, and electrochemically deposited PEDOT–PSS was completed. The following sections gives a background into previous XPS work completed on PEDOT with counter-ions, followed by studies on PEDOT degradation mechanisms.

7.4.5.1. PEDOT and Counter-Ions

The majority of XPS characterization has been completed on the commercially available PEDOT–PSS, Baytron P, because of its use in organic electronic devices [134]. The characteristic regions normally used for PEDOT analysis are the carbon (C 1s), oxygen (O 1s), and sulfur (S 2p) regions. These initial studies focused on the effect of different dopants on the PEDOT binding energy in an effort to deduce how the counter-ion was binding with PEDOT.

Initial XPS characterization was carried out on chemically polymerized PEDOT via iron(III) tris-p-toluene sulfonate. PEDOT was found to have peaks at 289.8 ± 0.2 eV in the C 1s region, 538.4 ± 0.2 eV in the O 1s region, and 168.2 eV in the S 2p region [135]. The PEDOT was then doped with the large polymeric anion PSS or the small anion tosylate (p-methyl benzyl sulfonate) (TsO). The normal peak positions for poly (sodium 4-styrenesulphonate), PSSNa+, are 285.0 eV (aliphatic carbon), 284.68 eV (aromatic carbon), 285.16 eV (C-S), 531.72 eV (O 1s in SO3−), 168.23 eV (S 2p3/2 in SO3−), and 1071.76 eV (Na+) [136].

Xing et al. found that the use of anionic counter-ions resulted in the oxidation of PEDOT, thus broadening the C 1s peaks. The C 1s for PEDOT–PSS was found at 289.4 eV, while the peak for the PEDOT–TsO was at 288.8eV [135]. The addition of PSS resulted in a second peak at 536.4 ± 0.2 eV and a slight shift in the PEDOT peak (538.4 ± 0.2 eV) in the O 1s region, while the use of TsO resulted in a second peak at 535.8 eV along with the PEDOT peak at 538.2 eV in the O 1s range. Both PSS and TsO exhibited two peaks at 172 eV and 168.2 eV in the S 2p range [135]. Additional tosylate work has also been performed by Kim et al. [137].

Greczynski et al. continued to study the effect of counter-ions on PEDOT by studying the dopants poly(4-styrenesulfonic acid) (PSSH) and PSSNa+ [138,139]. Commercial PEDOT–PSS was found to have S 2p peaks at 164.5 and 165.6 eV (spin-orbit coupling splitting of the PEDOT sulfur) and a peak at 169 eV (PSS). Peak deconvolution analysis showed the S 2p3/2 of PSSH to be at 168.8 eV and the S 2p3/2 of PSSNa+ at 168.4eV [138,139]. Greczynski et al. found that the S 2p spectrum is complicated by the presence of PEDOT. This manifests itself in an asymmetric tail at higher binding energies, a general shift to higher binding energies, and broad binding energy distributions, resulting from a positive charge delocalized over multiple and adjacent rings. Analysis of the O 1s peak found that the peak was composed of multiple peaks at 532.4 eV (oxygen double bonded to sulfur in PSSH), 533.5 eV (hydroxyl-oxygen atoms), 531.9 eV (PSSNa+), and 533.7 eV (oxygen atoms in the dioxyethylene bridge of PEDOT) [138,139].

Zotti et al. studied the doping structure relationship of electrochemically polymerized PEDOT with p-toluenesulfonic acid (TosH), sodium toluenesulfonate (TosNa), PSSH, and PSSNa+ [140]. This study, which also compared the chemically polymerized PEDOT–PSS to the electrochemically polymerized PEDOT–PSS, found that the electrochemically polymerized PEDOT/PSS contained more PEDOT relative to PSS than the chemically polymerized version through comparison of the PSS to thiophene ring (PEDOT) ratios. Evidently, electrochemically polymerized PEDOT–PSS was more compact, allowing for a smaller distance between chains, resulting in a reduction of the charge hopping distance. It is also probable that a single PSS polyanion connects multiple PEDOT chains, ultimately leading to higher conductivity [140].

An S 2p region comparison of the counter-ions between the two electrochemically polymerized PEDOT–PSSH and PEDOT–TosH samples yielded peaks at 167.8 eV (SO3−H+) and 166.8 eV (SO3−PEDOT+) for the PEDOT–PSSH and at 166.8eV (SO3−PEDOT+) for the PEDOT–TosH. From the integrated peak intensities, the relative quantities of the counter-ions were deduced. The PSS counter-ion was found to be present in a greater amount than the TosH counter-ion because only half of the PSS can be used to neutralize the PEDOT charge, whereas the amount of TosH utilized is only that which is needed to neutralize the PEDOT [140,141]. The greater amount of PSS results in a decrease in conductivity due to the increase in the average charge hopping distance [140].

Based on the PSS to thiophene ring ratio, PEDOT–PSSH had a smaller amount of sulfonate in the film than the PEDOT–PSSNa+ samples, and a greater quantity of PEDOT within the PEDOT–PSSH samples was found than was present in the PEDOT–PSSNa+. PSSH was therefore found to dope the PEDOT better than the PSSNa+ [140].

XPS has also been utilized to deduce the chemical binding energies of PEDOT after surface treatments, such as an acid or heat treatment, and under different solvent conditions. The Greczynski et al. study described the effects of hydrochloric acid (HCl) treatment and thermal treatments on the commercial PEDOT–PSS [138,139]. The effects of using different solvent solutions containing sorbitol, N-methylpyrrolidone (NMP), and isopropanol when heated were studied by Jönsson et al. [167].

7.4.5.2. PEDOT Degradation

PEDOT degradation studies using XPS have also been conducted. These studies focus on the atmospheric [141], UV-light [141,142], and electron bombardment degradation mechanisms [141,143].

Atmospheric effects on PEDOT and PSS, specifically the appearance of nitrogen, were studied by Crispin et al. [141]. The appearance of nitrogen within PEDOT–PSS films is thought to be a result of atmospheric ammonia molecules (NH3) reacting with water and the sulfonic acid group of PSS to form a hydroxide, which further reacts to form ammonium salts. The formation of ammonium salts induces desulfonation and thus aging of the PSS, which has been known to occur with exposure to light and heat [141].

Marciniak et al. [142] explored ultraviolet (UV) light degradation on EDOT, EDOT–SO2, PEDOT–PF6, and PEDOT–C14H29. After UV exposure, the presence of SO2 peaks indicated that photo-oxidation had occurred, resulting in shorter conjugation lengths that ultimately reduced conductivity. Chain scission was also found to occur. Marciniak et al. suggest that the degradation pathway of PEDOT is similar to the pathway in polythiophenes, in which energy transfer to the oxygen follows the π–π* transition, and then the oxygen finally reacts with the conjugated chain [142].

Electron bombardment studies were carried out on films of commercial PEDOT–PSS, PSSH, and EDOT to study the degradation effects on the polymer’s structure [141,143]. The result of this study showed that the energy used (3 eV [current density: 1 μA/cm2] for 67 h), which is less than typical energies used in organic light-emitting devices, was enough to trigger degradation of PEDOT. The degradation of the PEDOT thus involves not only the deterioration of the charge transport mechanism resulting in lower conductivities, but also the formation of free oxygen and sulfur atoms that in turn could cause breakdowns in other layers of the device because of their highly reactive nature [143].

In addition to deducing information on various counter-ion bonding, surface treatments, and degradation pathways, XPS has also been used to verify the presence of additives within PEDOT or PEDOT derivatives, such as PEDOT–PSS with poly(ethylene glycol) [144], adenosine triphosphate (ATP) [145], gold NPs [146], and PEDOT-coated latex spheres [147]. Other studies have focused on PEDOT binding with substrate materials, such as aluminum [148] and indium tin oxide [52,149].

7.4.5.3. Current Work

Figure 7.24 shows a schematic of the structure of an electrochemically polymerized PEDOT–PSS film, as well as a spun-cast film from a chemically polymerized PEDOT–PSS suspension. The PEDOT chains are relatively rigid, whereas the PSS is more flexible. The anionic PSS chains are expected to aggregate around the PEDOT. In the electrochemically polymerized film, the reaction proceeds by charge transport from the metal electrode out toward the surrounding solution. Presumably, the closest chains connect directly to the metal, with bonds that have a certain degree of charge transfer depending on the differences in electronegativity between the metal substrate and the PEDOT moieties. As the film thickens and densifies, occasional branching points allow for a rough surface texture as indicated. However since charge must be transferred from the metal substrate to the surrounding solution for the conducting polymer film to electrochemically deposit, efficient electrical pathways need to maintained.

Figure 7.23. XPS spectra of (a) carbon (C 1s), (b) oxygen (O 1s), and (c) sulfur core levels (S s2p) for electrochemically polymerized PEDOT–PSS (solid) and Baytron P (dashed).

Figure 7.23

XPS spectra of (a) carbon (C 1s), (b) oxygen (O 1s), and (c) sulfur core levels (S s2p) for electrochemically polymerized PEDOT–PSS (solid) and Baytron P (dashed). The changes in relative peak intensity indicate that there is more PSS relative (more...)

For the PEDOT/PSS film prepared by chemical oxidation, the polymer chains are first formed in solution and are surrounded by PSS to help keep the molecules suspended. The film is formed by evaporation onto the substrate. In this case, close interactions between chains are less likely to form, and there may be a preferred orientation of the chain backbone in the plane of the film. There is also substantially more PSS in the film, as confirmed by XPS studies (Figure 7.23). Because of these differences in structure, electrochemically polymerized films typically show much better electrical properties than solution-deposited films of PEDOT–PSS and are therefore generally preferred for biomedical device applications.

Figure 7.24. Schematic diagram comparing the microstructure of electrochemically polymerized PEDOT (top) with solution-cast films of commercially chemically polymerized PEDOT–PSS suspension (Baytron P).

Figure 7.24

Schematic diagram comparing the microstructure of electrochemically polymerized PEDOT (top) with solution-cast films of commercially chemically polymerized PEDOT–PSS suspension (Baytron P). Since the electrochemically polymerized PEDOT requires (more...)

7.4.6. Biological Conjugated Polymers: Melanins

Although PEDOT is not known to be found in its native state in living systems, examples exist of conjugated polymers with quite similar chemistries that are common in nature. Important examples are the melanins, which function as the dark, light-absorbing molecules that color hair and skin. Melanin derivatives are also found in certain tissues such as the eyes and ears and in the substania nigra of the human brain. It is known that reduced levels of melanin production in the brain correlate with certain disease states. For example, the loss of neuromelanin in the brain is associated with Parkinson’s disease [150]. Although natural and synthetic melanins have been studied relatively extensively in the literature, their detailed biological function and potential use in biomedical devices is still not well established [151–153].

Eumelanin and pheomelanin, the two main forms of melanin, are both composed of aromatic, fused bicyclic repeating units. Eumelanin contains both 5,6-dihydroxyindole (DHI) and 5,6-dihydroxyindole-2-carboxylic acid (DHICA) repeating units. As shown in Figure 7.25, these monomers have two oxygen atoms directly attached to a conjugated molecular backbone. This chemistry is also found in EDOT. It is therefore reasonable to anticipate that melanin may have electrical activity similar to that of PEDOT. Also, since eumelanin is an all natural, totally biologically derived polymer, it may have advantages for interfacing with living tissue. However, it may also prove to be less chemically stable over the long term.

Figure 7.25. Chemical structures of melanin and PEDOT.

Figure 7.25

Chemical structures of melanin and PEDOT. Note that both molecules share a conjugated molecular backbone, and they have oxygen atoms pendent to the backbone that in PEDOT are known to donate electrical charge and improve the electrical properties and (more...)

Zielinski and Pande reported the first electrochemical synthesis of eumelanin in 1990 [154]. Horak and Weeks synthesized melanin from DHI in phosphate buffer in 1993 [155]. Since then eumelanin has been synthesized using 3,4-dihydroxy-L-phenylalanine (L-dopa) (Figure 7.26) [151–153] and DHI monomers [156,157]. L-dopa is the natural precursor for eumelanin but does not contain an indde unit. Cyclization of the molecule occurs during polymerization to form both types of eumelanin repeating units shown in Figure 7.25. Compared to L-dopa, the structure of DHI has more of a resemblance to the repeating unit structure of eumelanin, but DHI is unstable in air and requires an inert atmosphere during all synthetic steps. Although DHICA is also a repeating unit in natural eumelanin, the direct electrochemical polymerization of DHICA has not yet been reported.

Figure 7.26. Optical micrograph of electrochemically polymerized thin film of synthetic melanin.

Figure 7.26

Optical micrograph of electrochemically polymerized thin film of synthetic melanin.

Studies of the electrical and optical properties of eumelanin imply that it is a semiconducting polymer [158,159]. Free-standing films of electropolymerized dopa-melanin were shown to have a conductivity of 1.4 × 10−6 S/cm [153], while dopa-melanin polymerized oxidatively has a conductivity of 6.4 × 10−8 S/cm [160]. Many synthetic eumelanins have also demonstrated photoconductivity [153,158]. There are still interesting open questions about the chemical structures that are formed during eumelanin polymerization. It appears that eumelanin consists of oligomers that contain many chemically distinct species. These species could include DHI, DHICA, and the oxidized forms of these molecules. Also, it seems that the monomer units bond to each other through many different types of atoms and that there is a general lack of organization within the oligomers [161,162]. Since the type of microstructure that forms should affect the electrical properties of the eumelanin film, controlling the structure to create long, completely conjugated polymer chains might result in a more electrically useful material.

Although it has not yet been studied in much detail, there have been conflicting studies concerning the effects of synthetic eumelanin on living cells. Ostergren et al. reported the ability to load PC12 cells with synthetic dopa-melanin without affecting the viability of the cells [163]. However, Li et al. showed that dopa-melaninloaded SK-N-SH cells demonstrate increased cellular stress and that death occurs via apoptotic mechanisms [164]. The increased cell death is believed to be at least partially due to the formation of hydroxyl radicals. If this is the case, it may be possible to modify the hydroxyl groups on eumelanin to eliminate the formation of free radicals.

7.5. CONCLUSIONS

The electroactive, biomimetic conducting polymer and hybrid conductive polymer–live cell electrode coatings described here represent novel strategies for addressing shortcomings at the electrode–tissue interface. This technology should help facilitate the establishment of long-term, bidirectional communication between host cells and implanted microelectrode-based biomedical devices that is critical for realization of functional body–machine interfaces. These studies further elucidate the factors that are important in the design and development of novel electroactive biomaterials intended for direct, functional contact with living electrically active tissues including the CNS, PNS (peripheral nervous system), sensory organs, and heart.

ACKNOWLEDGMENTS

We gratefully acknowledge the financial support of our research from the National Institutes of Health National Institute of Neurological Disorders and Stroke (NIH-NINDS-N01-NS-1-2338), the National Science Foundation (DMR-0518079), the University of Michigan College of Engineering Gap funding award, and the U.S. Army Multidisciplinary Research Initiative (MURI) program on Bio-Integrating Structural and Neural Prosthetic Materials. We have benefited from productive collaborations with Joseph Corey, Eva Feldman, Yoash Raphael, Bryan Pfingst, Wayne Aldridge, Frank Pelosi, Daryl Kipke, Paul Cederna, and Steve Goldstein at the University of Michigan. We have also worked in collaboration with Patrick Tresco at Utah, Greg Kovacs at Stanford, Jim Weiland at USC, Richard Normann at Utah, and Richard Stein at the University of Alberta. A variety of undergraduate research assistants provided laboratory assistance, including Wynn Koehler (MSE), Matt Meier (BME), Matthew Lapsley (MSE), Tani Kahlon (MSE), Mark Ferrall (LS&A), Michelle Leach (BME), Clair Harris (LS&A), Eric Tannebaum (LS&A), Deepa Rengaraj (MSE), Amber Brannan (Rose-Hulman Institute of Technology), Catherine Burk (BME), Kyle Roebuck (BME), Jingga Morry (BME), Brian Foster (EECS), Elizabeth Flak (MSE), Kate Gallup (MSE), Sejal Tailor (MSE), Beneque Cousin (MSE), Daniel Margul (BME), Olivia Kao (BME), and Grace Hu (MSE). Many of these undergraduate students were supported by the University of Michigan Undergraduate Research Opportunity Program (UROP). We also recognize contributions from Jayne Choi, Peter Keshtkar, and Max Betzig from Greenhills High School in Ann Arbor, who participated in a summer research outreach program coordinated by Rachel Goldman of the MSE Department. We also recognize the efforts of Rickard Axelsson, who visited our laboratory from Olle Inganas’ group in Sweden. Several invention disclosures related to this research activity have been filed with the University of Michigan Office of Technology Transfer, and patents are pending at the U.S. Patent and Trademark Office. Various aspects of this work are under active consideration for potential commercialization. David C. Martin, Sarah Richardson-Burns, and Jeffrey L. Hendricks are cofounders of Biotectix LLC, a recently formed start-up company that has obtained exclusive options to license aspects of this work from the University of Michigan.

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