Conducting polymers in biomedical engineering

ARTICLE IN PRESS

Prog. Polym. Sci. 32 (2007) 876–921 www.elsevier.com/locate/ppolysci

Conducting polymers in biomedical engineering Nathalie K. Guimarda, Natalia Gomezb, Christine E. Schmidtb,c, a

Chemistry Department, University of Texas at Austin, Austin, TX, USA Chemical Engineering Department, University of Texas at Austin, Austin, TX, USA c Biomedical Engineering Department, University of Texas at Austin, Austin, TX, USA b

Received 17 April 2007; received in revised form 23 May 2007; accepted 24 May 2007 Available online 11 June 2007

Abstract Conducting polymers (CPs) were first produced in the mid-1970s as a novel generation of organic materials that have both electrical and optical properties similar to those of metals and inorganic semiconductors, but which also exhibit the attractive properties associated with conventional polymers, such as ease of synthesis and flexibility in processing. The fact that several tissues are responsive to electrical fields and stimuli has made CPs attractive for a number of biological and medical applications. This review provides information on desirable CP properties specific to biomedical applications and how CPs have been optimized to generate these properties. The manuscript first introduces different types of CPs, their unique properties and their synthesis. Then specific information is provided on their modification for use in applications such as biosensors, tissue engineering, and neural probes. Although there remain many unanswered questions, particularly regarding the mechanisms by which electrical conduction through CPs affects cells, there is already compelling evidence to demonstrate the significant impact that CPs are starting to make in the biomedical field. r 2007 Elsevier Ltd. All rights reserved. Keywords: Electroactive biomaterial; Neural probes; Biosensors; Tissue engineering; Polypyrrole; Polythiophene

Contents 1. 2. 3. 4.

The discovery of conducting polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 877 Synthesis of conducting polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 877 Conductivity and doping of conducting polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 879 Use and modification of conducting polymers for biomedical applications . . . . . . . . . . . . . . . . . . . . . . . . . 882 4.1. General modification strategies for conducting polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 883 4.2. Biosensor applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 885 4.3. Tissue engineering applications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 893 4.3.1. Polypyrrole . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 893

Abbreviations: CP, conducting polymer; CV, cyclic voltammetry; DRG, dorsal root ganglion; FN, fibronectin; HA, hyaluronic acid; NGF, nerve growth factor; PANI, polyaniline; PC12, rat pheochromocytoma cell line; PEDOT, poly(3, 4-ethylenedioxythiophene); PLA, poly(lactic acid); PLGA, poly(lactic-co-glycolic acid); PPy, polypyrrole; PSS, poly(styrene sulfonate); PT, polythiophene; PVA, poly(vinyl alcohol); RGD, arginine–glycine–aspartic acid (peptide sequence); TCPS, tissue culture polystyrene Corresponding author. Biomedical Engineering Department, University of Texas at Austin, Austin, TX, USA. E-mail address: [email protected] (C.E. Schmidt). 0079-6700/$ - see front matter r 2007 Elsevier Ltd. All rights reserved. doi:10.1016/j.progpolymsci.2007.05.012

ARTICLE IN PRESS N.K. Guimard et al. / Prog. Polym. Sci. 32 (2007) 876–921

5.

6.

877

4.3.2. Polyaniline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 900 4.3.3. Polythiophene and novel conducting polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 903 4.4. Neural probe applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 903 4.4.1. Polypyrrole . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 904 4.4.2. Poly(3,4-ethylenedioxythiophene) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 906 4.5. Other applications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 908 Challenges and future directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 910 5.1. Electrical properties. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 911 5.2. Biological and physical properties. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 913 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 914 Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 914 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 914

1. The discovery of conducting polymers The electrically conducting polymer (CP) polypyrrole (PPy) dates back to the 1960s, but little was understood about the polymer at this time and the discovery was essentially lost [1]. It was only in 1977, when Alan MacDiarmid, Hideki Shirakawa, and Alan Heeger reported a 10 million-fold increase in the conductivity of polyacetylene doped with iodine (Fig. 1A), that the first inherently conductive polymer was recognized [2,3]. Although polyacetylene, a non-cyclic polyene, is still one of the most studied polymers in this field, it has significant limitations, such as difficulty with processing and high instability in air. Unlike polyacetylene, polyphenylenes, which are cyclic polyenes, are known to be thermally stable as a result of their aromaticity [4]. Consequently, the development of such aromatic CPs

for different applications has received much attention. Polyheterocycles, such as PPy, polythiophene (PT), polyaniline (PANI), and poly(3,4-ethylenedioxythiophene) (PEDOT) (Fig. 1B), developed in the 1980s, have since emerged as another class of aromatic CPs that exhibit good stabilities, conductivities, and ease of synthesis [5,6]. Refer to Table 1, which lists a broad range of different CPs and their conductivities. 2. Synthesis of conducting polymers CPs can be synthesized either chemically or electrochemically, with each having advantages and disadvantages as summarized in Table 2 [7]. Different methods of chemical synthesis include either condensation polymerization (i.e., step growth polymerization) or addition polymerization

O

O n Polyacetylene (PA)

N H

n

S

Polypyrrole (PPy)

n

Polythiophene (PT)

S

Poly(3,4-ethylene dioxythiophene) (PEDOT)

H N

n

H N

N

N

n

Polyaniline (PANI) Fig. 1. Chemical structures of common conductive polymers: (A) polyacetylene, the first documented conducting polymer and (B) most commonly explored conducting polymers for biomedical applications: polypyrrole, polythiophene, poly(3,4-ethylenedioxythiophene), and polyaniline.


878 Table 1 Conductivity of common CPsa Conducting polymer

Polyacetylene (PA) Polyparaphenylene (PPP) Polyparaphenylene sulfide (PPS) Polyparavinylene (PPv) Polypyrrole (PPy) Polythiophene (PT) Polyisothionaphthene (PITN) Polyaniline (PANI) a

Maximum conductivity (S/cm)b

Type of doping

200–1000 500 3–300 1–1000 40–200 10–100 1–50 5

n,p n,p p p p p p n,p

Reference Electrode (e.g., Ag/AgCl, saturated calomel electrode)

Working Electrode (e.g., ITO or metal) Counter Electrode (e.g., platinum mesh)

Reproduced from [18]. S ¼ Siemens.

Electrolyte Solution (monomer and dopant)

Fig. 2. Three electrode setup for electrochemical synthesis: reference electrode, working electrode (where polymerization occurs), and counter electrode all submersed in a monomer and electrolyte solution.

b

Table 2 Comparison of chemical and electrochemical CP polymerization Polymerization approach Chemical polymerization

Advantages

Larger-scale

Electrochemical polymerization

production possible Post-covalent modification of bulk CP possible More options to modify CP backbone covalently

Thin film

synthesis possible Ease of synthesis Entrapment of molecules in CP Doping is simultaneous

Disadvantages

Cannot make thin films

Synthesis more complicated

Difficult to

remove film from electrode surface Post-covalent modification of bulk CP is difficult

(i.e., chain growth polymerization). Condensation polymerization proceeds via the loss of small molecules, such as hydrochloric acid or water. Radical, cation, and anion polymerizations are all examples of addition polymerization, which are distinguished by the respective radical, cation, or anion intermediate state of the live or reactive end of the polymer chain during synthesis. Chemical synthesis not only provides many different possible routes to synthesize a variety of CPs, but also permits the scale-up of these materials, which is currently not possible with electrochemical synthesis.

Electrochemical synthesis is a common alternative for making CPs, particularly because this synthetic procedure is relatively straightforward [7,8]. Electrochemical preparation of CPs dates back to 1968 when ‘‘pyrrole black’’ was formed as a precipitate on a platinum electrode by exposing an aqueous solution of pyrrole and sulfuric acid to an oxidative potential [9]. Today, electrochemical polymerization is performed using a three-electrode configuration (working, counter, and reference electrodes) in a solution of the monomer, appropriate solvent, and electrolyte (dopant) (Fig. 2). Current is passed through the solution and electrodeposition occurs at the positively charged working electrode or anode. Monomers at the working electrode surface undergo oxidation to form radical cations that react with other monomers or radical cations, forming insoluble polymer chains on the electrode surface (Fig. 3). A number of important variables must be considered, including deposition time and temperature, solvent system (water content), electrolyte, electrode system, and deposition charge. Each of these parameters has an effect on film morphology (thickness and topography), mechanics, and conductivity, which are properties that directly impact the utility of the material for biomedical applications. For example, a non-protic, non-nucleophilic solvent generates stronger and more conductive CPs because protic solvents, like nucleophilic solvents, can generate side reactions with the growing CP chain, limiting and disrupting chain growth. The most significant difference between electrochemical and chemical methods of CP synthesis is that very thin CP films on the order of 20 nm can be produced using the electrochemical technique, whereas powders or very thick films are typically


-e

879

X = NH, S, O

X

X

X

H

H

X X

X

-2H+

X

H

X

X

H

X

-e X

X

X

X

X

n

X

X

-2H+

X

H

X

X

n

H n

X

Fig. 3. Mechanism for heterocycle polymerization via electrochemical synthesis. X ¼ NH, S, or O. This pathway is initiated by the oxidation of a monomer at the working electrode to give a cation species, which can react with a neutral monomer species or radical cation oligomeric species to generate the polymer. Adapted from [10].

produced with chemical polymerization. All CPs can be synthesized chemically, but electrochemical synthesis is limited to those systems in which the monomer can be oxidized in the presence of a potential to form reactive radical ion intermediates for polymerization. The standard CPs (i.e., PPy, PT, PANI, PEDOT) can be polymerized both chemically and electrochemically; however, several novel CPs with modified monomers are only amenable to chemical polymerization. 3. Conductivity and doping of conducting polymers CPs are unique in that they are organic chains of alternating double- and single-bonded sp2 hybridized atoms, which endow the polymer with metallike semiconductive properties. The series of alternating single and double bonds, which is generated by electron cloud overlap of p-orbitals to form p molecular orbitals, is referred to as a conjugated system. Studies have demonstrated that planar conformation of the alternating double-bond system, which maximizes sideways overlap between the p molecular orbitals, is critical for conductivity. This p-bonded system is further described in terms of electronic wave functions that are delocalized over the entire chain. This delocalization allows

charge mobility along the polymer backbone and between adjacent chains, but delocalization is limited by both disorder and Coulombic interactions between electrons and holes. Prior to doping, these systems are insulative (1010 S/cm; S ¼ 1/O); however, the electrical conductivity of polyheterocyclic films, for instance, can be augmented up to 12 orders of magnitude (102 S/cm) depending on the polymer system and the type and extent of doping, as described later [10]. Conductivity is a measure of electrical conduction and thus a measure of the ability of a material to pass a current. Generally, materials with conductivities less than 108 S/cm are considered insulators, materials with conductivities between 108 and 103 S/cm are considered semiconductors, and materials with conductivities greater than 103 S/cm are considered conductors. Conductivity (s) is the inverse of resistivity (r) and therefore has the units of inverse ohms (O1), also known as Siemens (S), per unit of distance, generally centimeters (i.e., S/cm). Resistivity is determined from a material’s resistance (R), which is often measured using the 4-point probe technique [11]. The 4-point-probe technique involves applying a constant current (I) across two electrodes at the surface of a material and then measuring the change

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in potential (V), where V ¼ IR, between another pair of electrodes. The following equations then apply: r ¼ ðR AÞ=l

(1)

rs ¼ ðR wÞ=l

(2)

where r is the bulk or volume resistivity, rs is the surface resistivity, R is the resistance of a uniform specimen (also called sheet or surface resistance, Rs), A is the cross-sectional area, l is the length between electrodes measuring current, and w the sample width. Bulk resistivity takes into account film thickness whereas surface resistivity does not. For a film having the same width and length (i.e., a square), surface resistivity is equal to surface resistance and is usually reported in units of O/square, simply in reference to the square surface dimensions. Resistance in O/square (l ¼ w) can be converted to bulk resistivity by multiplying by sample thickness. Eqs. (1) and (2) are limited to square or rectangular films and apply only to thin films (thickness 5 spacing between electrodes). For arbitrary-shaped films, the van der Pauw equation can be used to more accurately calculate bulk resistivity from measured resistance [11]. Cyclic voltammetry (CV) is a technique often used in conjunction with the 4-point probe method to characterize a material’s redox properties. The setup for generating a cyclic voltammogram is similar to the three-electrode configuration for electrochemical synthesis of CPs, where a CP sample is deposited on the working electrode. CV cannot be used to quantitatively measure conductivity, as with the 4-point probe technique, but can be used to assess whether a material is in fact conductive; this technique can also provide information on stability of electroactivity as well as oxidation/reduction potential and reversibility. CV is also more sensitive and thus can be used to gauge electroactivity of less conductive materials. Doping is the process of oxidizing (p-doping) or reducing (n-doping) a neutral polymer and providing a counter anion or cation (i.e., dopant), respectively. Upon doping, a CP system with a net charge of zero is produced due to the close association of the counter ions with the charged CP backbone. This process introduces charge carriers, in the form of charged polarons (i.e., radical ions) or bipolarons (i.e., dications or dianions), into the polymer (Fig. 4). The attraction of electrons in one repeat-unit to the nuclei in

*

X

X

X

X

*

X n

electron acceptor

*

X

X

X Polaron

X

*

X n

electron acceptor

*

X

X

X Bipolaron

X

*

X n

Fig. 4. Introduction of polaron and bipolaron lattice deformation upon oxidation (p-type doping) in heterocyclic polymers. X ¼ S, N, or O. A polaron or radical cation is introduced into the conjugated backbone after the loss of an electron. When oxidation of the same segment of the conjugated backbone occurs the unpaired electron of the polaron is lost and a dication (i.e., bipolaron) is formed.

neighboring units yields charge mobility along the chains and between chains, often referred to as ‘‘electron hopping’’. The ordered movement of these charge carriers along the conjugated CP backbone produces electrical conductivity. The smaller the band gap (i.e., distance between conducting band and valence band) energy for a CP the more conductive it is considered to be. Fig. 5 illustrates the change in band structure of PPy and PT subsequent to doping. There are many factors that influence this band gap and thus conductivity, including dopant, oxidation level/doping percentage, and synthesis method and temperature, therefore there are often discrepancies among results presented by different research groups [10,12–14]. The reader is referred to Heeger [15,16] and Bredas [17] for detailed explanations of conductivity mechanisms. Doping can be performed chemically or electrochemically and is dependent on oxidation potential. The oxidation potential of oligomers decreases as the number of monomers in the chain increases; therefore during electrochemical synthesis doping occurs as polymerization proceeds because the oxidation potential for doping the CP polymer is


881

Fig. 5. The valence-effective Hamiltonian band structure evolution of PPy (top) and PT (bottom) upon doping. PPy (top): (A) undoped and (B) intermediate doping level. Formation of non-interacting bipolarons at 0.45 eV above the valence band (VB) and 0.9 eV below the conductive band (CB). (C) 33% doping level (experimentally obtained with electrochemical doping). Formation of bipolaron bands with width of 0.25 eV. (D) 100% doping level per monomer. Merging of bipolaron bands with VB and CB. Note the decrease in band gap from 4 to 1.4 eV. Adapted from [17]. PT (bottom): (A) undoped. (B) 0.1% doping level with polaron states in the gap. (C) Few (1–20%) percent doping level, with the formation of non-interacting bipolarons at 0.61 eV above VB and 0.71 eV below CB. (D) 30% doping level, where the bipolaron states overlap and form two bands. (E) Hypothetical 100% doping level, with quasi-metallic behavior. Adapted from [10].

lower than that required for polymerizing the CP. Conductivity can be augmented by increasing the doping percentage and varying the dopant (Table 3). The chemical nature of the dopant not only affects electroactivity, but also affects surface and bulk structural properties. In addition, small and large dopants can both modulate electrical conductivities and surface structural properties, but larger dopants, such as hyaluronic acid (HA), can change polymer density and more dramatically

affect characteristics such as surface topography and physical handling properties [18]. When CPs are doped, the reactive species used to dope the CP oxidizes or reduces the CP and leaves behind an anion or cation, respectively. The reactive species is sometimes, but not always, the same as the dopant counter ion left behind. For instance, in the case of doping with I2, the dopant is I 3 or I5 and when FeCl3 is used as a reactive species, Cl or FeCl 4 is the dopant, whereas doping with BF4



Table 3 Doping levels and conductivities of a variety of CPsa Polymer

Dopant (X)

Structure

Conductivity (S/cm)

Polypyrrole (PPy)

CF3SO 3

(C4H3N)X0.3

ClO 4

(C4H3N)X0.3

150 (film) 10 (pp) 100 (film)

SO3CF 3 None BF 4 or PF6

(C4H2S)X0.3 (C4H2S)X0.01 (C4H2S)X0.06

10–20 (pp) 1.5 107 (pp) 0.02 (pp)

Poly(3-methylthiophene)

SO3CF 3 SO3CF 3 None PF6

(C5H4S)X0.3 (C5H4S)X0.5 (C5H4S)X0.005 (C5H4S)X0.12

30–50 (pp) 100 (pp) 107 (pp) 1 (pp)

Poly(3,4-dimethylthiophene) Polyfuran Polyazulene

SO3CF 3 SO3CF 3 ClO4

(C6H6S)X0.3 (C4H2O)X0.3 (C10H6)X0.25

10–50 (pp) 20–50 (pp) 102–101 (pp)

Polythiophene (PT)

Film: electrochemically polymerized films; pp: pressed pellet from chemically synthesized powder. a Adapted from [9].

generates BF 4 counter anions. CPs can be doped with a variety of molecules, such as small salt ions, peptides, or polymers, including polysaccharides and proteins. Chloride anions are commonly used to dope CPs because of their biological compatibility, generating, for example, oxidized PPy doped with chloride (i.e., PPyCl). To incorporate molecules that are not capable of redox chemistry, such as most biological dopants, it is necessary to synthesize/dope CPs electrochemically. The biological molecule to be used as a dopant must be charged and present in solution with the CP monomer as electrochemical polymerization occurs. Alternatively, the biological molecule can simply be entrapped in the presence of another dopant during synthesis. It should be noted that large molecule dopants (e.g., poly(styrene sulfonate) (PSS), high-molecularweight HA) are to some extent physically trapped and thus more stably integrated in the CP. These large biomolecules are not as readily leached out or exchanged under normal conditions or under application of an electrical potential. Small dopants, however, can more easily diffuse out of the CP and can be exchanged with other ions within the surrounding environment. Thus, CP properties (e.g., volume) can be regulated via the dynamic doping and de-doping process which occurs during oxidation and reduction of the CP. The reader is referred to the two-volume Handbook of Conducting Polymers [15], which provides a comprehensive review of the fundamental char-

acteristics of CPs, including their synthesis and electroactive properties. 4. Use and modification of conducting polymers for biomedical applications CPs have electrical and optical properties similar to those of metals and inorganic semiconductors, but also exhibit the attractive properties associated with conventional polymers, such as ease of synthesis and processing [16]. This unique combination of properties has given these polymers a wide range of applications in the microelectronics industry, including battery technology, photovoltaic devices, light emitting diodes, and electrochromic displays (reviewed in [19]), and more recently in the biological field. Research on CPs for biomedical applications expanded greatly with the discovery in the 1980s that these materials were compatible with many biological molecules such as those used in biosensors. By the mid-1990s CPs were also shown, via electrical stimulation, to modulate cellular activities, including cell adhesion, migration, DNA synthesis, and protein secretion [20–23]. Specifically, many of these studies involved nerve, bone, muscle, and cardiac cells, which respond to electrical impulses. Most CPs present a number of important advantages for biomedical applications, including biocompatibility, ability to entrap and controllably release biological molecules (i.e., reversible doping), ability to transfer charge from a biochemical reaction, and the potential to easily alter the


electrical, chemical, physical, and other properties of the CPs to better suit the nature of the specific application. These unique characteristics are useful in many biomedical applications, such as biosensors, tissue-engineering scaffolds, neural probes, drug-delivery devices, and bio-actuators (Table 4). Other electroactive materials, in addition to CPs, have been used in biological applications. For example, electrets (materials with a quasi-permanent surface charge provided by trapped charge carriers) and piezoelectric materials (materials which generate transient electrical charges under mechanical deformation) have been used in tissue engineering as nerve conduits [24,25]. Metals and semiconductors such as gold, iridium, and silicon have been used in neural probes and sensors [26,27]. However, CPs exhibit many advantages over these materials. In comparison, CPs are inexpensive, easy to synthesize, and versatile because their properties can be readily modulated by the wide range of molecules that can be entrapped or used as dopants. In addition, CPs permit control over the level and duration of electrical stimulation for tissue engineering applications, a limitation of electrets. CPs can also be tailored to create substrates with high surface area, a key aspect to decreasing impedance in neural probes. In particular, CPs can have as much as 50 times the surface area of bare iridium electrodes [28]. Furthermore, CPs can be precisely deposited on metal electrodes for biosensors, and can be intimately interfaced with biomolecules for a more effective transduction mechanism [29]. 4.1. General modification strategies for conducting polymers As already mentioned, CPs offer many advantages over other materials, with electrical activity being the most attractive property. However, there is always the desire to further optimize a material when targeting a specific application. The two common properties desired for all biomedical applications, aside from conductivity, are biocompatibility and redox stability, but beyond these needs, CP modification tends to be specific for the application. In particular, most research has focused on biological and physical modification of CPs. For biosensors, it is important to tune the hydrophobicity, conductivity, and reactive functionalities for modification of CPs to successfully incorporate biomolecules and to improve detection

883

of binding events. For tissue engineering, CP properties including biomolecule functionalization, surface roughness, hydrophobicity, three-dimensional geometry, redox stability, and degradability are critical. Neural probe applications require materials with high surface area, hydrophobicity, and cell specificity to improve and maintain good signal-to-noise ratio for detection of neuron signals. A popular strategy for optimizing the biological properties of a CP is the incorporation of bioactive molecules. This can be achieved through a number of techniques, including physical adsorption, entrapment, doping, and covalent attachment of desired biomolecules (Fig. 6). Physical adsorption is straightforward, but the adsorbed molecule can dissociate and render the material ‘‘inactive’’. Another non-covalent method is entrapment, which can be achieved by having the desired molecule present in the monomer/electrolyte solution during synthesis. The process of doping CPs, necessary to induce conductivity, can also be exploited to modify CPs non-covalently and to introduce new properties for a desired application. The range of possible dopants is vast as long as the selected dopant is charged. Alternatively, covalent methods can be used to more permanently functionalize CPs. The monomer can be synthesized with desired functional groups and then polymerized. Post-polymerization covalent modification is also possible, but is more challenging for insoluble polymers. It is important to note that the steric effects of any incorporated functional group may disrupt the planarity of the conjugated system, which could in turn decrease conductivity. Functionalization of CPs with different biomolecules has allowed biomedical engineers to modify CPs with biological sensing elements, and to turn on and off different signaling pathways to create CPs that enhance adhesion and proliferation of a variety of cell types and improve their biocompatibility. It is just as important to consider physical and electrical properties of CPs as their chemical composition. For example, many CPs are crystalline and have limited porosity. To overcome the drawbacks associated with crystalline materials there have been efforts to coat some CPs with or covalently tether CPs to a more malleable material, such as a polyester (e.g., poly(lactic-co-glycolic acid) (PLGA)). In general, manipulation of CP properties (e.g., roughness/topography, porosity, hydrophobicity, mechanical strength, malleability, degradability, redox stability, conductivity) can be achieved



Table 4 Conducting polymers in biological applications Application

Description of application

Tissue engineering

Biocompatible, biodegradable scaffolds containing stimuli to enhance tissue regeneration

Biocompatibility Good

Implantable electrodes for recording or stimulating neurons, primarily in the brain

Biocompatibility Good

Neural probes

Advantages of conducting polymers

Devices containing biomolecules as sensing elements, integrated with electrical transducers

Bio-actuators

Devices to create mechanical force that could be used as ‘‘artificial muscle’’type actuators

biodegradable Not highly porous Hydrophobicity

Decreased electrical contact at interface

PPy and derivatives

[18,22,23,106,129,130,131,133–140, 146,147,151–154,159-162] PANI [171–175] PT and derivatives [176] Novel CPs [177]

PPy [28,132,178–181] PEDOT [183–186]

synthesis on metal electrodes Increased surface area (decreased impedance)

Ability to entrap

Devices for storage and controlled release of drugs

conductivity

Drug Delivery

Not

Polymers currently explored [Ref]

Good stability Electrochemical

Biosensors

conductivity Possible modification to include chemical cues

Limitations of conducting polymers

biomolecules in films Possible surface modification Efficient electric charge transfer from bioreactions Electrochemical synthesis on metal electrodes

Ability to entrap

biomolecules Controlled release with reduction

Biocompatibility Good conductivity

Hydrophobicity

Hydrophobicity

dopant uptake/ release (control volume) Lightweight Work at body temp. and with body fluids

via previously mentioned chemical means, such as the incorporation of molecules as dopants or via entrapment, or through covalent insertion of

can denature entrapped proteins Rapid release

Short-term redox

Can control

See Table 5

can denature entrapped proteins Diffusion barriers for entrapped enzymes

stability Delamination of CP films Response limited by ion mobility

PPy [190] PEDOT [189]

PPy [193–195,197,198] PANI [196,198,199] Polymer-carbon nanotube composites [196,198]

functional groups into the CP backbone that may, for example, increase conductivity or permit degradation. Micropatterning and modification of


885

Fig. 6. Examples of modification strategies used with conducting polymers. CPs have been modified chemically and physically using a number of approaches as a means to change conductivity, bioactivity, and physical topography/geometry. A few common modification approaches are depicted, including non-covalent chemical methods (molecule entrapment and adsorption), covalent chemical conjugation, and micropatterning using standard lithography techniques.

synthesis parameters, such as using surfactants and regulating temperature and deposition charge for electrochemical synthesis of CPs, are strategies that have also been used to successfully manipulate several physical properties. CPs have thus far proven straightforward to modify, like other polymers. However, it is difficult to compare modified CPs from different research groups because of the large number of techniques available for modifying CPs, which result in little consistency from one investigation to the next. Although a wide range of CP variants has been explored for a number of applications, as described in more detail below, considerable research remains to be performed.

Analyte

Sensing element Transducer (CP)

Biochemical Signal Electronic signal

Electronics Fig. 7. Schematic of a biosensor. A biological sensing element (e.g., enzyme, antibody) detects a specific analyte producing a biochemical signal that is transferred to the transducer (e.g., conducting polymer), which ultimately produces a digital electronic signal that is proportional to the amount of analyte present.

4.2. Biosensor applications The first biosensing device was created by integrating an enzyme into an electrode [30], and since that time, much progress has been made in monitoring and diagnosing metabolites (e.g., glucose, hormones, neurotransmitters, antibodies, antigens) for clinical purposes. A biosensor is composed of a sensing element (i.e., biomolecule) and a transducer [29]. The sensing element interacts with the analyte of interest producing a chemical signal that is transmitted to the transducer, which ultimately transforms the input into an electrical signal (Fig. 7). CPs are extensively used as transducers that integrate the signals produced by biological sensing

elements such as enzymes (Table 5). Depending on how the chemical signal is sensed and transmitted, biosensors can be divided into several categories: amperometric (measures current), potentiometric (measures potential), conductometric (measures change in conductivity), optical (measures light absorbance or emission), calorimetric (measures change in enthalpy), and piezoelectric (measures mechanical stress) (reviewed in [29]). The most common types of transducers are amperometric and potentiometric. An amperometric biosensor measures the current produced when a specific product is oxidized or reduced (e.g., redox reaction of a substrate in an enzyme) at a


886

Table 5 Examples of biosensors using conducting polymersa Analyte (sensing element)

Types of sensor

Glucose (glucose oxidase)

Amperometric Potentiometric

PPy [20,21,49,51–54,56,105] PANI [68,216,217] PT [68,87]

Cholesterol (cholesterol oxidase/esterase)

Amperometric

PPy [63,218,219] PANI [104]

L-lactate

Amperometric

PPy [220] PANI [221,222] PT [88]

Urea (urease)

Amperometric Potentiometric Conductometric

PPy [40,41,223]

DNA (DNA hybridization)

Amperometric Gravimetric

PPy [50,64–67,78,79,81–84] PT [90,91]

a

(lactate oxidase/dehydrogenase)

Polymers explored [Ref.]

Modified from [28].

Substrate

Enzyme

Product

Conducting Polymer (reduced)

Conducting Polymer (oxidized) electrons

Measures current

Fig. 8. Schematic of electron transfer in an amperometric biosensor. An enzyme catalyzes a redox reaction of a specific analyte, which results in the reduction of the conducting polymer (transducer) and the measurement of a current.

constant applied potential [29] (Fig. 8). The CP mediates the electron transfer (e.g., via hydrogen peroxide) between an enzyme, such as an oxidase or dehydrogenase, and the final electrode; however, the exact mechanism is not completely understood [29]. Redox mediators such as ferrocene, viologen, Prussian Blue, or their derivatives are used to improve electron transfer from the biochemical reaction to the CP and therefore improve sensor sensitivity and selectivity. These redox mediators can be entrapped, incorporated as dopants, or chemically conjugated to the monomer [31–39]. Potentiometric biosensors use ion-selective electrodes as physical transducers. For example, detection of urea by ureases is performed via the production

of NH3, which interacts with PPy to produce an electrical signal. This signal could be a product of a change in pH and the subsequent ion mobility in the polymer matrix triggered by an equilibration of the dopants with the free ions in solution [40,41]. A key aspect in biosensor applications is the integration of the electrical component (i.e., CP) with the biological recognition components. The immobilization of bioactive macromolecules in or on electrically conductive polymers has been extensively explored in an effort to provide intimate contact between these two elements [29,42–46]. This section focuses on describing the different available techniques for the immobilization of biologically active molecules on CPs. For this immobilization, it is critical to maintain the activity of the molecules, increase stability, and ensure accessibility of the analyte to perform biological events such as hybridization of complementary oligonucleotides, antigen–antibody binding, or enzyme-catalyzed reactions. Table 6 summarizes the main categories of immobilization techniques of biological sensing elements on CPs. Two main classes are distinguished: non-covalent and covalent modifications. Non-covalent modifications include adsorption, physical entrapment, and affinity binding. Covalent immobilization includes all techniques that create a covalent bond between the conducting substrate


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Table 6 Immobilization techniques of biomolecules on conducting polymers for biosensing devices Immobilization technique Non-covalent techniques Adsorption

Principles of immobilization

Electrostatic forces, hydrogen bonding, Van der Waal’s forces, etc.

Advantages

Simple

Limitations

Biomolecule loss (desorption) over time

Limited control over immobilization

Random orientation on surface Entrapment

Molecule incorporation during electropolymerization

Simple Good proximity between elements

Potential loss of biomolecule activity

Steric and diffusion constraints Requires high biomolecule concentration

Affinity binding

High affinity interactions such as avidin-biotin

Control over molecule

Covalent techniques Chemical conjugation

Surface chemical reaction between functional groups

orientation High accessibility of analytes Minimal loss of biomolecule activity

Tighter control over

immobilization High accessibility of analytes Minimal biomolecule loss over time Control over biomolecule orientation

and the biomolecule via functional moieties. Recent reviews provide multiple examples of these methods [47,48], in addition to the ones described below. Physical adsorption is the simplest method of immobilization and one of the first approaches used for biosensors. As an example, glucose oxidase has been adsorbed onto PPy for an amperometric sensor and was shown to detect glucose over a wide range of concentrations (2.5–30 mM) using dimethylferrocene as an electron transfer mediator [49]. DNA biosensors have also been created using adsorption techniques. To achieve this, DNA has been indirectly adsorbed to PPy surfaces via mercapto-oligonucleotide probe immobilization onto Au–Ag nanocomposites adsorbed onto PPy [50]. Although adsorption is simple, controlling the concentration of the immobilized compound is difficult and immobilization is not stable because of the weak non-covalent forces involved, which decrease the lifetime of the biosensor [48]. Another drawback is that compound adsorption occurs as a

Requires pre-immobilization of one of the affinity molecules (e.g., biotin)

Complex Conditions are not always appropriate for biomolecules

Potential loss of biomolecule activity

monolayer, which limits the quantity of sensing element. An alternative to adsorption is physical entrapment of the desired biomolecule during electropolymerization, which is one of the most extensively used techniques. During this process monomer, dopant, and biomolecules are mixed in a single solution used for electrochemical polymerization. This process is usually performed under mild conditions (i.e., neutral pH, aqueous, low oxidation potentials) without chemical reactions that could alter the activity of proteins, and only requires a single step for both polymerization and molecule immobilization. For this reason CPs, such as PPy, are frequently used to entrap biomolecules [46]. Many early applications successfully entrapped glucose oxidase (GOx) in PPy films [20,21,51,52]. Improvements continue to be made on glucose biosensors, which also serve as models for other biosensors. Recent advancements include the GOxinitiated polymerization of pyrrole to form sensitive



PPy-coated GOx nanoparticles that act as glucose biosensors [53], and a PPy-coated ultramicroelectrode glucose sensor fabricated using photolithography with nA/(mmol L) sensitivity [54]. Wang and coworkers [55] discovered that the sensitivity of biosensors utilizing entrapped enzymes is enhanced with increasing microscopic roughness of the electrode’s surface; therefore these authors have targeted high surface area PPy-coated platinum electrodes. Other enzymes such as horseradish peroxidase, phosphorylases, polyphenol oxidase lactate dehydrogenase, and deaminases have also been entrapped in CP films for biosensors [56–61]. For example, a PANI-derivative polymer was mixed with polylysine to render it water-insoluble for aqueous-based biosensors and then subsequently modified with horseradish peroxidase (via entrapment) [58]. PEDOT has also been used to entrap enzymes. For instance, the ability of PEDOT (synthesized by chemical vapor-phase polymerization) to shrink to 5% of its original thickness once washed with ethanol has been exploited to entrap horseradish peroxidase [62]. Detection of cholesterol has been performed in amperometric biosensors by entrapment of cholesterol oxidase/esterase in derivatives of PPy and polynaphthalene [63]. Entrapment methods have also been used for immobilization of antibodies [64] and DNA [64–67]. Most entrapment methods involve codeposition during electrochemical synthesis. An alternative to this is sol–gel encapsulation, which has been performed using silica and PT or silica and PANI to entrap glucose oxidase for glucose sensing [68]. Entrapment of living cells within CPs has also been reported in an effort to create novel biosensors. Detection of dopamine was achieved by entrapment of cells extracted from banana pulp in PPy films in which the enzyme polyphenol oxidase catalyzes the conversion of dopamine to quinine with a corresponding consumption of oxygen [69]. This study was based on the finding that the browning reaction of the banana tissue was produced by the conversion of dopamine to quinine by polyphenol oxidase [70]. In a different study, whole intact human erythrocytes containing viable antigens were entrapped within a poly(vinyl sulfonate)- and heparin sulfate-doped PPy electroactive matrix from which signals were induced directly using resistometry techniques. These composites of cells and conductive matrices served as proof-ofprinciple for potential biosensors for blood group

determination [71,72], which could eventually be used by the medical profession to transfuse human blood and blood products to patients. Although entrapment is a popular immobilization technique, it has some important limitations. For example, the hydrophobic nature of the polymer compromises the quaternary structure of proteins, decreasing their biological activity. To overcome this limitation, new alternatives have focused on creating more hydrophilic polymers using modified monomers, such as pyrrole rings with long hydrophilic chains [73,74]. Also, entrapment methods require a high concentration of the biomolecule (0.2–3.5 mg/mL), which is not always available and increases the cost of the process. Finally, the entrapment procedure diminishes the accessibility of analytes to the sensing element and thus affects affinity complex formation (e.g., antibody–antigen, hybridization of nucleotides). As a result, other immobilization techniques, such as affinity binding and covalent modification, have been explored to overcome these limitations. Affinity binding methods are based on immobilizing molecules on the surface of CPs via strong non-covalent interactions. As a conventional approach in this category, the use of the avidin–biotin complex presents advantages over other immobilization techniques because of the extremely specific and high-affinity interactions between biotin and the glycoprotein avidin (Ka ¼ 1015 mol–1 L) [75]. This technique allows control over orientation of the immobilized molecules by adjusting the location of the binding elements, which increases the activity and accessibility of the biological sensing elements. In contrast with conventional grafting, this approach can also be applied to prepare assemblies containing multilayers of biological molecules. The use of an avidin-immobilized biotin association was achieved by copolymerizing pyrrole monomers with biotinylated hydrophilic pyrrole monomers with different poly(ethylene glycol) (PEG) lengths as spacer arms (Fig. 9) [76]. The amount of anchored avidin on the hydrophilic PPy was modulated by the immobilized biotin density and by the length of the PEG spacer arm linking the biotin moiety to the PPy backbone. Recently, the biotin–avidin affinity immobilization technique was utilized in combination with scanning electrochemical microscopy technology to deposit and functionalize biotinylated PPy to create biologically active microspot biosensors [77]. The design of complex biological architectures was further demonstrated in


889

O HN

NH

S N-(CH2)3-(O-CH2-CH2)2-O-(CH2)3-NH-CO-(CH2)4 1 O HN

NH

S N-(CH2)3-(O-CH2-CH2)2-O-(CH2)3-NH-CO-(CH2)5-NH-CO-(CH2)4 2 Fig. 9. Hydrophilic pyrrole-biotin monomers with different lengths of the spacer arms: 1 (13 atoms) and 2 (20 atoms) [76]. Reproduced from Torres-Rodriguez et al. by permission of Elsevier Science Ltd.

1

-1 polypyrrole-biotin/avidin

polypyrrole-biotin

4

2 Cpbio -3

3 Cpbio Detection - Hybridization

Biosensor

polypyrrole-biotin/avidin/ODN-probe Cpbio ODN-target Cpbio

polypyrrole-biotin/avidin/ODN-probe Cpbio

Fig. 10. Schematic of the oligonucleotide (ODN) sensor, ODN sensing and sensor regeneration, where Cpbio and Cp*bio represent the biotinylated DNA probe and complementary biotinylated DNA probe, respectively [78]. Reproduced from Dupont-Filliard et al. by permission of Elsevier Science Ltd.

the fabrication of PPy chips (Fig. 10) [78] and PPy nanowires [79], in which the successive immobilization of avidin and biotinylated oligonucleotides

allowed the detection of hybridization events. Thiophenes have also been biotinylated and incorporated into CPs used for biosensors, which rely on a



transduction mechanism induced by the binding of avidin to produce a change in polymer electroactivity. In particular, avidin has been detected by drastic changes in the cyclic voltammograms of the polymer, which is attributed to the bound protein blocking ion motion [80]. There are other affinity binding molecular complexes, similar to the avidin–biotin complex, that have been explored for the immobilization of DNA to CP surfaces. Cosnier et al. [81] developed a unique intercalator (i.e., a molecule that inserts itself into double-stranded DNA)-based immobilization technique to amperometrically detect singlestranded DNA. Target DNA strands are detected when they form DNA duplexes with labeled DNA probes and are subsequently immobilized by the affinity binding of the intercalator, which is covalently bound to PPy. Not only can these PPy surfaces be used for the detection of any singlestranded DNA, but detection limits are very low (1 pg/mL). Alternately, a label-free DNA biosensor detects hybridization based on the electrostatic changes in the ion-exchange kinetics at a PPycoated microelectrode surface. Pyrroles functionalized with a phosphonic acid group are grafted to the PPy surface so that DNA probes can be immobilized via electrostatic complexation of the DNA phosphate group with phosphonic acid groups via magnesium cations. In addition to having low detection limits, these sensors offer label-free detection and can be regenerated for repeated use [82]. An attractive alternative to affinity binding for biomolecule attachment involves the introduction of appropriate functional groups into CP backbones or the surface modification of the polymers, followed by covalent bonding (i.e., grafting) of bioactive macromolecules to the surfaces. In comparison to adsorption, entrapment, and affinity binding immobilization, this approach is typically more robust and stable to external environmental factors, allows high loading, and increases biosensor lifetime; however, it is usually more complex and sometimes requires reaction conditions not suitable for biomolecules. Compared to entrapment methods, surface chemical conjugation increases the accessibility of the analytes and enhances the formation of affinity interactions. Conducting copolymers containing covalently substituted monomers have been fabricated as a means to facilitate the immobilization of biomolecules. One strategy for this type of immobilization is

to functionalize CP monomers prior to polymerization. For example, copolymers of PPy and oligonucleotides bearing a pyrrole group have been reported for DNA sensors [83,84]. Other copolymers have been formed with monomers functionalized with amine or carboxylic acid groups, which subsequently react with functional groups on the biomolecules to be immobilized. As an example, immunosensors for detecting Listeria monocytogenes were created by covalently binding the Listeria monoclonal antibody to a copolymer of carboxylic acid-functionalized PPy and regular PPy [85,86]. Carboxylic acid-functionalized PT has also been developed and used for glucose oxidase and lactate oxidase immobilization [87,88]. A novel use for carboxylic acid substituted PT is P-aminophenyl-a-D-mannopyranose immobilization, which binds selectively to several viruses. Therefore this novel glycoPT can be used to detect viruses [89]. Protection of electron withdrawing functional groups, (e.g., carboxylic acid), on monomers is not uncommon because they hinder polymerization. For example, the protection of carboxylic acid functionalized thiophene with substituted benzyl groups facilitated the electrochemical synthesis of a PT derivative, which later could be used for DNA immobilization to detect DNA [90]. DNA immobilization on PT has also been achieved via covalent linkage to sulfonyl chloride functionalized PT, synthesized through the co-polymerization of methylthiophene and sulfonamide (protected sulfonyl chloride) functionalized thiophene [91]. Other procedures incorporate activated esters such as N-hydroxysuccinimide by functionalizing thiophene or pyrrole monomers [81,92–94] (Fig. 11). This functional group reacts with amines in proteins and other biomolecules under mild conditions. An interesting extension of this type of functionalization was reported for the formation of PPy nanoparticles for visual diagnostic assays [95]. In this investigation, pyrrole and N-succinimidyl ester pyrrole were polymerized in the presence of silica nanoparticles, which were subsequently used for immobilization of human serum albumin. A more recent approach for chemical grafting is the use of photo-activated chemistries. For example, electropolymerization of pyrrole–benzophenone polymers has been reported. The benzophenone group can be activated with UV irradiation after polymerization for reaction with groups present in biomolecules (Fig. 12) [96,97]. Another conjugation method is chemical grafting after polymerization of unmodified CPs. One common


891

Fig. 11. Immobilization of DNA on PPy derivatives with N-hydroxysuccinimide groups. Polypyrrole copolymers functionalized with ferrocenyl groups and an activated ester reacted with oligonucleotides with a terminal amino group. Upon addition of complementary oligonucleotide sequences and hybridization, a large modification of redox activity of ferrocenyl groups occurs, which is determined by amperometric methods [93]. Reproduced from Korri-Youssouri et al. by permission of Elsevier Science Ltd.

BM O

C

hv

C OH

BM C O

C O

O

O

N

N

POLYMER n

n

ELECTRODE

Fig. 12. Immobilization of biomolecules (BM) on polypyrrole– benzophenone substrates. The biomolecule is grafted to the polymer upon UV irradiation [96]. Reproduced from Cosnier and Senillou et al. by permission of Royal Society of Chemistry.

method for post-CP modification is the use of glutaraldehyde (often in conjunction with bovine serum albumin). Glutaraldehyde crosslinking techniques have been used to immobilize enzymes, such as glucose oxidase, glutamate oxidase, lactate oxidase, xanthine oxidase, choline oxidase, alcohol oxidase, uricase, trypsin, and acetylcholinesterase on PPy [34,36–38,98–103]. Cholesterol esterase, cholesterol oxidase, and peroxidase have all been immobilized on PANI using glutaraldehyde crosslinking as well [104]. Another good example of postpolymerization covalent modification is the immobilization of glucose oxidase and viologen on PPy via acrylic acid grafting [105]. Regular PPy substrates were immersed in acrylic acid solutions and irradiated with UV light. This reaction facilitated

the insertion of acrylic acid residues on the surface of PPy, which introduced carboxylic acid functional groups used for the subsequent immobilization of glucose oxidase (Fig. 13). CPs modified in this fashion retained relatively high conductivity (only 10–50% decrease). These surface-modified polymers typically have higher conductivities compared to most bulk-modified polymers. For example, polymers with substituted monomers exhibit a significant decrease in conductivity (3–5 orders of magnitude) [106]. Regardless of the immobilization technique employed there have been efforts to miniaturize biosensors in the last few years. A couple of examples of microelectrode biosensors employing affinity or entrapment immobilization techniques have been previously discussed. Often these microelectrode biosensors are made with surface micromachining techniques, but a novel technique developed by Liu et al. [107] involves consecutive platinization and pyrrole polymerization on a silicone substrate. Biomolecules can be immobilized in a number of ways; however, covalent immobilization was easier and produced better results. The advantages of this biosensor fabricated with bulk micromachining technology include its compatibility with complementary metal oxide semiconductor (CMOS) technology, smaller sensitive surface area, lower detection limit, and larger sensitivity per unit area. In addition to the methods described above, there are other novel modifications of CPs for sensing applications. A recent study described the use of caffeine-imprinted PPy for a piezoelectric caffeine sensor [108]. Molecular imprinting techniques were used by electropolymerizing PPy in the presence of caffeine and removing the template caffeine


892 PPy

COOH

UV + AAc

A

COOH

C

N

R1

N

R

(WSC)

O 40min, 25°C

COOH

1 h, 4 °C

B

O COOH

C

O H N

E

H2N

E

C

COOH

C

O

O COOH

24 h, 4 °C, pH = 7.4

O

C

N

HN

R

R1

COOH

Fig. 13. Post-polymerization covalent immobilization of enzymes, E, on PPy. (A) Graft copolymerization of acrylic acid (AAc) on PPy film surface. (B) Preactivation with water-soluble carbodiimide (WSC). (C) Glucose oxidase immobilization on polypyrrole matrix [105]. Reproduced from Cen et al. by permission of Elsevier Science Ltd.

molecules at the end of the polymerization via washing. This allowed the formation of unique three-dimensional sites used for the selective recognition of caffeine, a process applicable to the food and pharmaceutical industries. Following the same concept of imprinting, polythiophene polymers modified with zwitterionic, peptide-like side chains have been used in sensors by imprinting biomolecules via electrostatic and hydrogen binding. These sensors have been tested for both DNA and protein detection [109]. Another unique biosensor technique involves coating gold electrodes, upon which aptamers have been covalently immobilized, with ferrocene-modified PT. In this instance, the bioactive molecule (the aptamer) is not immobilized directly to the CP; however, the CP surrounds the bioactive molecule allowing the CP to act as an electrical transducer [110]. A similar strategy was used for the detection of double-stranded DNA [111]. In other studies, PPy has been polymerized in the interstitial spaces of a hydrogel containing immobilized enzymes. Because of the highly hydrophilic environment provided by the hydrogel, the performance and characteristics of the biosensor were greatly improved [112]. More specifically, PPy hydrogels, such as PPy–alginate [113] and PPy–polyacrylamide [114], have recently been explored to optimize GOx entrapment, retention, and function.

PPy–alginate gels have also been reported for the entrapment of algal Chlorella vulgaris cells to detect algal alkaline phosphatase activity. The alginate component of the matrix serves to create a more cell-compatible environment, while pyrrole increased the overall stability of the biosensor [115]. PPy–methacrylate/tetraethylene glycol-based hydrogels have also been explored for potential application in vivo as subcutaneous implantable biochips to monitor biomolecules, such as glucose and lactate [102]. Recent advances in carbon nanotubes (CNTs) include the incorporation of CNTs into a number of CP-based biosensors. For example, preliminary studies have been performed exploring the properties of both PPy/CNT and PANI/CNT devices as pH sensors [116]. One application used DNA-doped PPy in conjunction with CNTs for label-free detection of DNA. In particular, the unique properties of the PPy–CNTs allowed the detection of hybridization reactions with complementary DNA sequences via a decrease in impedance [117]. Alternatively, similar DNA sensors have been created from a composite of PPy and CNTs functionalized with carboxylic groups to covalently immobilize DNA onto CNTs [118,119]. CNTs have also been incorporated into biosensors as nanotube arrays onto which enzymes, such as GOx, can be immobilized along with a CP [120] and as a PPy


dopant [121,122]. In general, the presence of CNTs tends to increase the overall sensitivity and selectivity of biosensors. 4.3. Tissue engineering applications The development of biosensors has created a foundation for using CPs in many biological environments. In fact it was a logical next step to explore CPs as biomaterials for different cell types and functions because previous studies had shown that cells such as fibroblasts, neurons, and osteoblasts responded to electrical fields created by electrets [24,123] or between electrodes in vitro and in vivo [124–126]. The general CP properties desired for tissue engineering applications include conductivity, reversible oxidation, redox stability, biocompatibility, hydrophobicity (40–701 water contact angle promotes cell adhesion), three-dimensional geometry, and surface topography. However, there are always exceptions, as described below. 4.3.1. Polypyrrole PPy was one of the first CPs studied for its effect on mammalian cells [22]. To date, PPy has been reported to support cell adhesion and growth of a number of different cell types, including endothelial cells [22,127,128], rat pheochromocytoma (PC12) cells [71,129], neurons and support cells (i.e., glia, fibroblasts) associated with dorsal root ganglia (DRG) [130,131], primary neurons [132,133], keratinocytes [134], and mesenchymal stem cells [135]. As previously stated, PPy in its simplest unmodified form can be synthesized to have a small molecule anion (e.g., Cl) as a dopant, which generally is not considered to contribute additional biological properties to PPy. Although such versions of PPy have been studied, most research has focused on either the non-covalent or covalent modification of PPy to optimize interactions with specific cell types and other material properties that are important for PPy’s use in vivo. The reader is referred to an excellent review on PPy and its interactions with biological tissues [136]. Several studies have demonstrated cell and tissue compatibility of PPy in vitro and in vivo. For example, one of the earlier studies showed that PPy doped with p-toluene sulfonate (PPyTS) is cytocompatible with mouse fibroblasts and neuroblastoma cells [137]. In vivo, there is minimal tissue response to implanted PPy [130,137] and PPy can support the regrowth of regenerating axons, for

893

example [138–130]. Because the advantage of using CPs in tissue-engineering applications is the ability to subject cells to an electrical field, studies have also addressed cell compatibility when a current or voltage is applied to PPy [129,137]. Although there is some evidence of cytotoxicity after long time exposure to current (e.g., 96 h exposure to 1 mA), shorter periods of exposure to current have not been found to have negative effects on cells in culture [137]. In addition to the biocompatibility of PPy, a number of studies have shown that electrical stimulation using PPyTS can modulate cellular response. Most of these early studies simply relied on the passive adsorption of biomolecules from serum or the coating of purified protein solutions to the surface of PPy. In one of the first studies performed to assess the suitability of CPs for sustaining cell growth and controlling cell function, aortic endothelial cells were cultured on fibronectin (FN)-coated PPy films and exposed to oxidizing potentials [22]. Oxidized PPy resulted in cell spreading, whereas reduction of PPy to its neutral state caused cell rounding (Fig. 14) and a concomitant drop (98%) in DNA synthesis. Despite the change in cellular morphology, cell viability (490%) and adhesion were very good on both oxidized and neutral PPy. Other studies have shown that electrical stimulation of PPy in its oxidized form can also be used to modulate cell function [129]. PC12 cells seeded on electrochemically synthesized PSS-doped PPy (PPyPSS) films having a resistivity of 1 kO were found to exhibit a 91% increase in median neurite length when a positive potential of 100 mV was passed through the PPy for 2 h (Fig. 15). These studies demonstrate that cell growth and function can be drastically enhanced at the interface of PPy undergoing electrical stimulation. Although very little is known about the effects of an electric potential and/or an electrical field on cell activity, it is speculated that reduction of PPy and electrical conduction through PPy can both affect cells in several ways. For example, the process of neutralization of PPy, in which a reducting potential is applied, causes the expulsion of negative ions in the case of small dopants or the uptake of positive ions from the medium in the case of large, entrapped dopants. For PPyPSS, an uptake of positive ions such as Na+ from the medium is speculated to affect several processes, including protein adsorption and the cell cycle [22]. Along



Fig. 14. Photomicrograph of endothelial cells cultured on fibronectin-coated PPyTS for 4 h: (A) PPy in native oxidized state and (B) PPyTs reduced by application of 0.5 V for 4 h ( 700) [22]. Reproduced from Wong et al. by permission of the National Academy of Sciences.

these lines, the effects of electrical stimulation of PPyPSS on FN adsorption and thus cell shape and growth were further elucidated [140]. Stimulation of these PPy films during protein incubation increased FN adsorption to these surfaces from both purified FN and heterogenous serum-containing solutions. The authors also observed PC12 cell adhesion and demonstrated that with the aid of electrical stimulation FN adsorption to PPy films could be increased, producing a surface that subsequently enhanced neurite extension. An alternative to adsorbing bioactive molecules to PPy is the entrapment of such molecules during electrochemical synthesis either as the dopant or in addition to the dopant. Recently this technique was used to entrap large molecules, poly(vinyl alcohol) (PVA), within PPyClO4 (tetraethylammonium perchlorate was the oxidant) [141]. The authors were able to generate a substrate with high surface area

Fig. 15. PC12 cell differentiation on PPyPSS in the (A) absence and (B) presence of electrical stimulation. (A) Cells grown for 48 h but not subjected to electrical stimulation are shown for comparison. (B) PC12 cells were grown on PPy for 24 h in the presence of nerve growth factor, then exposed to electrical stimulation (100 mV) across the polymer film. Images were acquired 24 h after stimulation. Scale bar ¼ 100 mm [129]. Reproduced from Schmidt et al. by permission of the National Academy of Sciences.

that promoted enhanced PC12 cell adhesion and proliferation compared to TCPS controls. However, as commonly seen with the entrapment of large biomolecules, conductivity of PPyClO4 with PVA entrapped was decreased 10-fold compared to PPy doped with toluenesulfonic acid controls. Despite this decrease its conductivity was still significant (7 S/cm). There have also been many studies that have entrapped biomolecules, such as adenosine 50 -triphosphate (ATP) [142–144] and nerve growth factor (NGF) [71] in PPy and other CPs for both drug delivery and tissue engineering applications. In particular, co-entrapment of NGF and dextran sulfate, followed by the controlled release of NGF


upon PPy reduction, was shown to stimulate PC12 cell neurite extension [71]. Very recently a study found that neurite outgrowth from auditory neurons could be specifically targeted upon electrical stimulation of PPy entrapping neurotrophin-3 [145]. The dopant introduces new properties into the material; thus, PPy can be optimized to promote the growth of different cell types or to induce specific aspects of wound healing simply by varying the dopant. For example, HA-doped PPy has been synthesized and explored for tissue-engineering applications because of HA’s inherent role in wound healing and angiogenesis [18]. Conductivity of electrochemically polymerized PPyHA films was reported to be greatly reduced (3.08 103 S/cm) compared to the conductivity of PPyPSS films (9.25 S/cm). To improve conductivity, PPyHA bilayer films were fabricated and shown to have conductivities comparable to PPyPSS (8.02 S/cm). These films had PSS as the dopant in the bottom layer (to retain conductivity) and HA as the dopant only in a thin layer on the top (to provide biological activity). In vivo studies showed that the PPyHA bilayer films were biocompatible and promoted vascularization as a result of the presence of HA (Fig. 16) [18]. Other studies have also explored modification of PPy via different dopants, including doping with dermatan sulfate for increasing keratinocyte viability [134], doping with heparin for increasing endothelial cell proliferation [127,128], and doping with laminin-derived peptides to control neuron and astrocyte adhesion [132]. It is important to note that both surface topography and conductivity are often significantly altered when biomolecules are exclusively used as dopants [18,134]. Although large biomolecule dopants are known to significantly decrease conductivity and have more of an effect on topography compared to small dopants, the exact effects of particular dopants are not very predictable (Fig. 17) [134]. Dopants can also be used as intermediate ‘‘tethers’’ to allow further modification of PPy with biomolecules. A good example of this technique is doping with poly(glutamic acid) (PGlu) to provide a handle (i.e., carboxylic acid pendant group) for future functionalization [131]. The carboxylic acid groups on PGlu could be covalently linked using carbodiimide chemistry to any amino group, such as those found in polylysine and laminin (Fig. 18). DRG neurons were observed to preferentially adhere to and extend neurites on the PPy surfaces modified with polylysine and/or laminin using this

895

Fig. 16. In vivo tissue response to (A) PPyPSS films and (B) PPyHA bilayer films. Films were implanted in subcutaneous pouches in rats. Tissues surrounding the material were harvested after 2 weeks, fixed, imbedded and stained with hematoxylin and eosin. The heavy black lines in the images are the films. Blood vessels are denoted by arrows. Scale bar is 100 mm [18]. Reproduced from Collier et al. by permission of John Wiley & Sons, Inc.

technique. This functionalization technique localizes the bioactive molecules at the surface, while in theory maintaining the bulk properties of PPyPGlu; however, it is possible that the dopant (PGlu) itself reduces bulk conductivity compared to large polymeric anions containing aromatic rings or small anions. This novel technique is advantageous in that a single material can be used to easily incorporate a range of different biomolecules, including those that are not charged. Besides using dopants to modify PPy properties, there are other emerging non-covalent approaches to further modify PPy for biomedical applications. For instance, one approach to modifying PPy noncovalently was achieved using a peptide, selected


896

0.4

I / mA

0.2 0.0 -0.2 -0.4 -0.6 0.4

I / mA

0.2 0.0 -0.2 -0.4 -0.6 -1.0

-0.8 -0.6 -0.4 -0.2

0.0

0.2

E / V vs Ag/AgCl

0.4

-1.0

-0.8

-0.6

-0.4

-0.2

0.0

0.2

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E / V vs Ag/AgCl

Fig. 17. Topography of thick PPy films as assessed using scanning electron microscopy (SEM) (top) and corresponding cyclic voltammograms (CVs), indicating electrical activity of the materials (bottom). (A) PPyCl. (B) PPy doped with poly(vinyl sulfate). (C) PPy doped with dermatan sulfate. (D) PPy doped with collagen (inset: thin film of collagen-doped PPy). A narrow CV spectrum correlates to decreased electroactivity. Scale bars are 100 mm [134]. Reproduced from Ateh et al. by permission of Mary Ann Liebert, Inc.

from phage display libraries, that specifically binds to PPyCl [146]. This study demonstrated that the peptide could be used to modify the PPyCl surface with the arginine–glycine–aspartic acid (RGD) peptide and promote PC12 cell adhesion in serum-free media, whereas no cell adhesion was seen on unmodified surfaces. An advantage to this surface modification technique is that, in theory, it should not

modify bulk properties such as conductivity and like the PGlu approach described above, could be used to modify PPy with a range of different biomolecules that do not need to be charged. In addition to non-covalent approaches to modifying PPy as discussed above, modification via covalent bonds has also been extensively explored. Multiple techniques have been reported,


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Fig. 18. (A) Schematic of polypyrrole doped with poly(glutamic acid) (abbreviated as pPy[pGlu] in the figure) followed by the covalent attachment of polylysine (pLys) and/or laminin (Lmn). (B) Phase contrast and fluorescence images of DRGs adhered to the surface of pPy[pGlu]-X, where X ¼ 1. pLys, 2. Lmn, and 3. pLys-Lmn. Cell nuclei were labeled with DAPI (blue florescence). All images were taken at 10 magnification and the 200 mm scale bar applies to all images shown [131]. Reproduced from Song et al. by permission of Elsevier Science Ltd.



some of which are described here. Modification of the b-positions on PPy was achieved by creation of strong sulfide bonds with cysteines [147,148]. These cysteines served as ‘‘handles’’ to attach bioactive molecules via peptide bonds or peptides containing a terminal cysteine (S) group that could be directly bound to PPy. In particular, osteoblast adhesion was enhanced up to 230% on RGDS peptidemodified PPy surfaces. Covalent modification of pyrrole at the N-position is another common method of functionalizing PPy, although this modification does decrease the conductivity of PPy more than b substitution [7]. The most prevalent technique for N-functionalization consists of hydrolyzing 1-(2-cyanoethyl) pyrrole to 1-(2-carboxyethyl) pyrrole [106,149,150]. The carboxylic acid functional groups can be activated for future protein immobilization with hydroxysuccinimide (NHS) in the presence of a water-soluble carbodiimide before [150] or after pyrrole polymerization [106]. For example, the cell adhesive RGD peptide sequence was covalently conjugated to the surface of NHSfunctionalized PPyCl [106,151]. Although, as expected, the conductivity of these films decreased from 2.81 102 to 4.65 102 S/m, human umbilical vein endothelial cell (HUVEC) adhesion and proliferation were improved. Other covalent techniques create covalent bonds only at the PPy surface, rather than modifying PPy in bulk. In one of these studies, NGF was surfaceimmobilized on PPyPSS (thickness ¼ 200 nm), using an intermediate photo-crosslinker consisting of polyallylamine conjugated to an arylazido functional group [152]. On exposure to UV light, NGF was fixed to the substrate via photoactivation and non-specific reaction of the arylazido moieties. Similar levels of neurite extension were observed for PC12 cells and DRG neurons in the presence of immobilized NGF compared to positive controls with soluble NGF. Additionally, electrical stimulation experiments were conducted with the modified polymer and revealed a 50% increase in neurite outgrowth in PC12 cells compared to experiments without electrical stimulation. This novel modification of PPyPSS provides both electrical and biological stimulation by presenting tethered growth factors and producing only a small decrease in the material’s conductivity (modified PPy ¼ 9.372 S/ cm; unmodified PPy ¼ 14.572 S/cm), which represents a major advantage when compared to other covalent techniques that produce a decrease in conductivity by orders of magnitude.

In another investigation, HA was attached to PPyTS via grafting of acrylate groups and silanization with amine-containing silanes [153,154]. HA was subsequently immobilized using carbodiimide chemistry for the reaction of amine groups from the grafting procedure with the carboxylic groups from HA. These studies are similar to the reported PPyHA films using HA as a dopant [18], described earlier, but in this approach the authors use a covalent attachment method at the surface of PPy to avoid bulk incorporation of HA and to better preserve conductivity [153,154]. As predicted, this modification approach resulted in good conductivity (only a 4–38% decrease, depending on the degree of functionalization), as well as unaffected mechanical properties of the polymer and increased hydrophilicity of the surface. An increased hydrophilicity enhances interactions between the CP and extracellular matrix components. The authors found that HA-modified PPy supported PC12 cell adhesion and when this material was sulfated, platelet adhesion was reduced in vivo. In general, covalent surface modifications increase the stability of the biomolecules and have a minor impact on the CP conductivity (compared to dopant and entrapment methods). A great number of studies have focused on optimizing PPy–cell interactions by the incorporation of biomolecules, but it is necessary also to consider other material properties such as redox stability, mechanical properties, degradability, and surface topography. In particular, the development of a PPy analog that would decrease the redox potential of PPy and render it stable and more conductive under the oxidative conditions of biological systems was investigated [155]. PPy not only has a relatively high oxidation potential and so is susceptible to polymer breakdown due to overoxidation, but there is also a significant amount of a–b0 coupling, which induces structure disorder that disrupts electroactivity and is the primary site of polymer breakdown. PPy derivatives, poly(3,4ethylenedioxypyrrole) (PEDOP) and poly(3,4-propylenedioxypyrrole) (PProDOP), were synthesized to address this latter issue and were found to have a good range of conductivities (50 S/cm) and good oxidative stability in the presence of dithiothrietol and glutathione. In addition to fine tuning the electrical properties of PPy, it is important to consider its mechanical properties. As noted earlier, unmodified PPy, like most other CPs, is crystalline and brittle, which does


not make it an ideal candidate for tissue scaffold materials. Several research groups have explored in vitro and in vivo properties of PPy-coated polyester fabrics. PPy coating of substrates has been achieved non-covalently by both chemical [156] and electrochemical [157] deposition; however, these materials are susceptible to delamination. Alternatively, covalent modification of phosphorous trichlorideactivated polyester fabric with N-modified PPy generates PPy-coated polyester that is much less susceptible to delamination [158]. As expected, PPycoated polyester fabrics are only slightly conductive (surface resistivity of 104–105 O/square); however, these materials are flexible and exhibit good in vitro and in vivo biocompatibility [159–161]. Degradability and surface topography are also important parameters for interfacing CPs with cells. The development of a bioresorbable form of PPy was obtained by electrochemically synthesizing small, slightly water-soluble PPy chains (1700– 3200 Da), which would enable the gradual erosion and renal clearance of the polymer to occur in vivo [162]. The monomer unit of this modified PPy was functionalized at the b position with either a methyl ester or a carboxylic acid side chain. The methyl ester-modified polymer was soluble in organic solvents and eroded much more slowly (6% mass loss over 80 days) compared to the carboxylic acidmodified form, which was insoluble in organic

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solvents and slightly soluble in water with a degradation rate of about 27% over 80 days. Although these novel polymers are not inherently degradable (i.e., the backbones of the polymers are not actually cleaved), the ability of the polymers to erode achieves the same goal. Along similar lines, other studies have introduced the property of biodegradation by creating blends of PPy with degradable polymers such as polylactide (PLA). For example, PPy nanoparticle–PLA composites were created and shown to be both degradable and conductive (resistivity ranging from 2 107–15 O/ square) [23]. Both parameters could be modulated by varying the amount of PPy in the blend, which additionally impacted topography (Fig. 19). The authors found that fibroblasts attached and grew on the PPy nanoparticle–PLA composites with an increased viability when stimulated at currents of 10–50 mA. This material offers the advantage of permitting electrical stimulation at biologically significant currents, maintaining electroconductivity for long times (15% of conductivity retained after 1000 h), as well as being both degradable and biocompatible in vivo [23,163,164]. Several studies have exclusively explored the role of surface topography for CPs. For example, admicellar polymerization has been used to precisely control PPy film thickness and to modulate surface topography (Fig. 20), thereby influencing mesenchymal

Fig. 19. SEM images of the surface of PPy nanoparticle-PLA composite membranes with various PPy content: (A) 5%, (B) 7%, (C) 9%, (D) 17% [23]. Reproduced from Shi et al. by permission of Elsevier Science Ltd.



stem cell adhesion and differentiation to osteoblasts [135]. In another approach, cellulose fibers were used as a substrate to generate nanoscale topography within PPy [165]. The morphologically complex nanofeatures of the cellulose were preserved and transferred to PPy coatings by means of polymerization-induced adsorption. Microfabrication techniques have also been used for creating PPy topographical patterns that provide contact guidance cues to primary neurons [133]. In particular, patterning of 1 and 2 mm wide PPyPSS microchannels was obtained using electropolymerization and electron beam lithography. To demonstrate the effect of these surface patterns on cells, embryonic hippocampal neurons were cultured on patterned PPy and found to polarize (i.e., define an axon) faster on this modified material, having a two-fold increase in the number of cells with axons compared to cells cultured on unmodified PPyPSS (Fig. 21). There have been other micro- or nano-patterned CP surfaces created for potential tissue engineering applications, including three dimensional honeycomb, porous CP films [166,167] and microsized circles and microfluidic mimics of the vascular

network [168]. In a slightly different approach, electrochemical coating of PPy and PPy derivatives onto metallic surfaces was used as a technique to impart metallic surfaces with topography and functionality in the hope of generating more biocompatible and biofunctional interfaces on metallic medical implants such as stents [169]. 4.3.2. Polyaniline The exploration of PANI for tissue engineering applications has progressed more slowly than the development of PPy for similar applications; however, recently there has been more evidence for the ability of PANI and PANI variants to support cell growth. The cell compatibility of only a few different cell lines has been explored, including cardiac myoblasts and PC12 cells. It has been suggested that perhaps the compatibility of PANI is specific to particular cells [170]. A handful of strategies have been explored for the development of PANI with good biocompatibility, conductivity, and mechanical properties. These strategies most often involve noncovalent and covalent techniques, as is the case for strategies used for PPy modification.

Fig. 20. Representative atomic force microscopy images of PPy thin films made by admicellar polymerization (AP) and chemical polymerization without surfactant (no SDS) on polystyrene dishes. (A) 20 103 M pyrrole (AP). (B) 35 103 M pyrrole (AP). (C) 50 103 M pyrrole (AP). (D) 5 103 M pyrrole (no SDS). The vertical scale was adjusted to 1000 nm. The thickness ranged from 35 to 154 nm for 20 103 M (AP), 48–466 nm for 35 103 M (AP), 85–697 nm for 50 103 M (AP), and 46–199 nm for chemical polymerization without surfactant (no SDS) [135]. Reproduced from Castano et al. by permission of Wiley-VCH Verlag GmbH & Co. KGaA.


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Fig. 21. Patterning of PPy to create microchannels for contact guidance of neurons. Phase-contrast (left) and fluorescence (right) photomicrographs of hippocampal neurons on PPy. (A,B) Cells cultured on 2 mm wide and 200 nm deep PPy microchannels. (C,D) Cells cultured on unmodified PPy. The green labeling (Alexa 488) corresponds to Tau-1 (axonal marker) immunostaining. Cells polarized (i.e., established a single axon) more readily on microchannels than on unmodified PPy. Scale bar ¼ 20 mm. Images are at the same magnification [133]. Reproduced from Gomez et al. by permission of John Wiley & Sons, Inc.

The demonstration of PANI’s biocompatibility in vivo has sparked interest in its use for tissue engineering applications. Studies have assessed the in vivo response to implants of different oxidation states of PANI (emeraldine, nigraniline, and leucoemeraldine) [171]. In general, there was an absence of significant inflammation at the implant sites. There were, however, thin layers of fibrous tissue encapsulating these unmodified PANI implants and immune response cells (i.e., mast cells) were observed. There were no signs of abnormality of muscle and adipose tissues in the vicinity of the implants. Although this study lacked controls (e.g., comparison with acceptable implant materials such as PLGA) and detailed quantitative assessment of compatibility, the study nonetheless had promising

results and encouraged further exploration of the application of PANI in biological systems. Cell biocompatibility of doped PANI (with HCl) has been observed in vitro [170]. However, an initial lag in cell (cardiac myoblast) growth rate was found in the presence of doped PANI; it was proposed that this correlated to acid leaching from doped films and a subsequent decrease in conductivity. Although a few studies have suggested the biocompatibility of unmodified PANI, as described above, there have also been conflicting reports that demonstrated poor cell adhesion and growth on this unmodified CP [172]. For this reason, other methods have been sought to modify PANI to render it biocompatible while maintaining the desirable electrical properties of the material.



Fig. 22. SEM image of PANI films. (A) PANI immersed in N-methylpyrrolidinone (NMP) for 300 s followed by dipping in water. (B) PANI immersed in 0.03 g/mL Pluronic polymer/NMP solution followed by dipping in water [173]. Reproduced from Li et al. by permission of Elsevier Science Ltd.

Fig. 23. (A) SEM image of PANI-gelatin blend fibers with ratio 45:55. Original magnification is 5000 . (B) Morphology of H9c2 myoblast cells at 20 h post-seeding on 45:55 PANI-gelatin blend fiber. Staining is for nuclei-bisbenzimide and actin cytoskeletonphalloidin; fibers autofluoresce. Original magnification is 400 . (C) SEM images of H9c2 cells cultured on 45:55 PANI-gelatin blend fibers [175]. Reproduced from Li et al. by permission of Elsevier Science Ltd.

For example, hydrophilicity of PANI has been modified by a two-liquid adsorptive entrapment of poly(ethylene oxide)–poly(propylene oxide)–poly (ethylene oxide) (PEO–PPO–PEO) triblock copolymer. This technique results in the entrapment of PPO within PANI, while the PEO ends remain exposed at the surface (Fig. 22). The exposure of the hydrophilic PEO segments consequently decreases subsequent protein adsorption. Because this technique targets the PANI surface, bulk conductivity is maintained (0.6 S/cm) [173]. Similarly, PANI– chitosan nanocomposites have been explored to increase biocompatibility and enhance surface characteristics [174]. PANI–gelatin crosslinked composites have been demonstrated to have good biocompatibility in vivo. In addition, gelatin has desirable mechanical properties that allow it to be electrically spun into fibers to generate three-dimensional scaffolds (Fig. 23) [175]. Varying the ratio of PANI to gelatin allowed for the

optimization of conductivity and fiber diameter, where cells were found to prefer smaller diameter fibers. The optimization of PANI biocompatibility has been targeted by covalent modification as well. There are a number of methods to modify a CP covalently and thus functionalize the surface; however, the primary concern is to avoid disrupting the conductivity of the CP. Graft copolymerization of emeraldine PANI with acrylic acid was used to immobilize collagen, which was found to reduce inflammation in vivo [171]. Direct modification of the aniline monomer to create a functional group (methyl chloride) that can be linked to any amino acid has also been investigated [172]. In this approach the conjugation of PANI was not interrupted by the addition of a copolymer and therefore, the conductivity was better maintained (4–10 S/cm versus about 10 S/cm for unmodified PANI). Functionalization of PANI with tryptophan


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composed of heterocycles, which were tethered together by degradable ester linkers (Fig. 24a) [177]. The goal was to create a polymer that would be conductive, thus permitting in vivo electrical activation of cells, and also be degradable, eliminating the need to remove these scaffolds surgically. The overlap of conjugated regions would enable inter and intra ‘‘electron hopping’’ to give rise to conductivity and the polymer would degrade in the presence of esterases in vivo. The goal was realized; however, doping of the polymer to render it conductive (104 S/cm), was again only achieved with iodine. Despite this drawback, the undoped version of this polymer was degradable (Fig. 25) and also biocompatible in vivo and in vitro [177]. These results were considered promising and currently research is focused on the synthesis and characterization of a modified polymer that will be degradable, more conductive and doped by biocompatible chloride anions. A novel polymer, poly(dialcoholdimethylquaterthiophene-co-adipic acid) (Fig. 24b), has been synthesized and is currently being characterized by Schmidt and coworkers.

was demonstrated with this technique, and resulted in good PC12 cell attachment, proliferation, and response to NGF. 4.3.3. Polythiophene and novel conducting polymers PPy and PANI remain the most extensively studied CPs for tissue engineering purposes to date, but a few other CPs have been explored, such as polythiophene (PT) and novel CPs. In one study, a small library of polymers consisting of a PT backbone with oligosiloxane grafted to the b position of the thiophene rings was generated [176]. The authors created different PT homopolymers by varying the chain length of the oligosiloxane, but they also created copolymers with oligosiloxane-grafted thiophene and 3-methylthiophene. Conductivity of the PT homopolymers decreased in general as the length of the oligosiloxane increased (maximum conductivity of 1.7 106 S/cm), but the corresponding copolymers were more conductive (maximum conductivity of 5 105 S/cm). Unfortunately, these polymers were doped with iodine, which is toxic to cells. In vitro biocompatibility studies with human ovarian cancer cells (HeLa) were only performed on undoped polymers. Perhaps the biggest limitation of CPs for in vivo applications is their inherent inability to degrade. Thus far two examples of modified PPy have been mentioned, which attempt to overcome this drawback: slightly water-soluble oligopyrrole variants and PPy nanoparticle-PLA composites [23,162]. Although these PPy variants do not have degradable backbones, they have been designed to be small enough, such that upon the surface erosion of the material, the CP can be renally cleared. Rivers et al. reported an alternative approach in which the CP backbone was designed to degrade. This CP, biodegradable electrically conducting polymer (BECP), was designed to have conducting regions

4.4. Neural probe applications Many of the advance made in CPs for tissue engineering applications, particularly with respect to neurons, are relevant to the development of optimized neural electrodes. Especially important to this area of research is the need to intimately interface electrodes with neural tissue and to relay signals efficiently between the cells and the electrode, thus integrating the device seamlessly with the native neuronal signaling network. CPs are attractive candidates for interfacing electrodes with neurons because they can achieve high surface area, helping to promote effective ion exchange between recording sites and the surrounding tissue. The goal

O H O

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Fig. 24. Chemical structures of (A) biocompatible electrically conductive polymer (BECP) and (B) poly(5,5000 -dialcohol-3,3000 -dimethylquaterthiophene-co-adipic acid) (PAMQAA).


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Wavelength (nm) Fig. 25. Degradation studies of BECP films (3.0 cm 2.2 cm 2.0 mm) were performed in PBS with and without cholesterol esterase for 2 weeks. The supernatants of the solutions were evaluated by measurement of their UV–vis absorption. The aromatic rings found in the polymer and its degraded species absorb at 340 nm in PBS and the esterase absorbs at 280 nm. The supernatant of the polymer+esterase shows a strong absorption peak at 340 nm compared to the supernatant of the polymer without the esterase. This demonstrates that BECP is degradable by enzymes that are known to be secreted during in vivo wound healing events. Note: The y-axis is reported in absorbance units (a.u.) [177]. Reproduced from Rivers et al. by permission of Wiley-VCH Verlag GmbH & Co. KGaA.

is to increase surface area of the recording site, while maintaining a sufficiently small geometric area to isolate the action potential from a single neuron. A larger surface area results in an increase in capacitance, which corresponds to a decrease in impedance, thus improving signal-to-noise ratio. Ideally, a neural probe would maximize neural signals recorded, minimize noise, maintain high capacitance, and remain conductive over the long term. Although PPy is commonly explored for coating neural probes, more recent studies have focused on the polythiophene derivative PEDOT (poly(3,4-ethylenedioxythiophene)) because of its stable oxidative state and higher conductivity. 4.4.1. Polypyrrole Similar to research progress in biosensor and tissue engineering applications, most of the initial studies exploring the application of CPs to neural probes focused on PPy. Most CPs explored for neural probe applications have been modified to improve neural signal detection. In one study, the

authors designed a neural probe, which could also be used as a neural scaffold, from PPy doped with PSS or sodium dodecylbenzenesulfonate (NaDBS) [178]. This research group was interested in investigating the effectiveness of an implant specially fabricated from the electrochemical deposition of PPy onto a patterned gold template. The dopant and the temperature at which electrochemical deposition proceeded were varied. PPy doped with NaDBS was more conductive than PPyPSS, which could increase signal transportation from the cells to the electrode. With regard to temperature, PPy films synthesized using either dopant but at a lower temperature (4 1C) were rougher than the same films made at 25 1C. This is a significant observation because rougher topography correlates to increased surface area, which would increase signal conduction by increasing the interface with neurons. In vitro, neural networks consisting of glia, axons, dendrites, and synapses were observed for all PPy films (Fig. 26). In vivo, the implants with rougher surfaces had increased implant integration and decreased gliosis compared to controls [178]. PPy augmented with biological moieties may offer advantages for neural probe applications. Several studies have focused on doping PPy with peptides derived from extracellular matrix proteins including laminin-derived peptides, such as p31 (CDPGYIGSR) [132,179], p20 (RNIAEIIKDI) [132] and YIGSR-containing sequences (DCDPGYIGSR) [180], in addition to FN peptide variants such as the silk-like polymer having FN fragments (SLPF) [179], which is a synthetic copolymer consisting of amino acid sequence blocks of silk fibroin and the cell attachment domain of FN (RGD). Careful selection of the bioactive molecule to be incorporated at the electrode surface is essential to enhance neuron adhesion and growth and thereby increase the signal received from neurons, while simultaneously minimizing astrocyte adhesion, which would interfere with neuron signaling. In one example, a silicon-based, 4-pronged neural probe was micropatterned with a layer of gold. PPy doped with either SLPF or laminin-derived nonapeptide p31 was deposited on the gold recording sites [179]. Not only did the entrapment of these peptide sequences enhance cell attachment, growth, and migration, but at a particular intermediate deposition time, the surface area and the conductivity of PPy were maximized. An increase in surface area (Fig. 27) aided integration of the probe into the


Fig. 26. PPy surfaces. The surface texture of PPy can be controlled through electrochemical polymerization temperature. (A) Smooth morphology when PPyNaDBS is polymerized at 24 1C. (B) Rough morphology when PPyNaDBS is polymerized at 4 1C. Scale bars are 200 mm. (C) Fluorescently labeled explanted cortical neurons growing and forming networks on a PPyNaDBS surface after 21 days. Green: neurons, red: glia, blue: nuclei. Scale bar is 50 mm [178]. Reproduced from George et al. by permission of Elsevier Science Ltd.

neural tissue and, along with the increase in conductivity, this improved signal transportation. Finally, the authors demonstrated that glial cells

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preferentially attached to PPySLPF (compared to gold), and neuroblastoma cells adhered significantly better to PPyCDPGYIGSR as compared to PPyCH3COO, further suggesting the enhancement of electrode-neuron interactions in the presence of PPy doped with bioactive peptides [179]. This work was extended to monitor in vivo neural recordings from DCDPGYIGSR-modified PPy probes. Even though conductivity decreased as a result of the entrapped molecules within PPy, impedance values were still low enough to detect neural activity one week after neural probe implantation. Unfortunately, the PPyDCDPGYIGSR coating was not sufficient to prevent gliosis after two weeks of implantation and further work is being performed to minimize astrocyte and fibroblast encapsulation of neural probes [180]. Further efforts to optimize cell attachment to PPy neural probes have been made. A study comparing PPy doped with two different laminin sequences or both of these laminin sequences together was performed to optimize neural interfacing [132]. The p31 laminin fragment is known to promote cell binding, whereas the p20 laminin fragment specifically promotes neurite initiation and outgrowth. As expected, neuron density was greatest on laminin-doped controls, with the combination of p31 and p20 dopants giving rise to the second highest neuron density compared to PPy doped with p31 or p20 individually. Neurites extended equally well on PPy doped with p20 and PPy doped with both p31 and p20. Although these laminin-modified PPy surfaces exhibited lower astrocyte cell adhesion compared to gold surfaces, there was a larger number of astrocytes versus neurons adhered to the laminin-modified PPy (Fig. 28). Consequently, the optimization of this system is still in progress with more recent work having been directed toward gaining finer control over neuronal signal detection [Stauffer and Cui, personal communication, 2006]. Just as positive cues can be entrapped in PPy electrodes, these investigators have found that detectable neuronal waveforms can be inhibited with the entrapment and release of an antagonist of alpha-amino-3-hydroxy-5-methyl-4-isoxazoleproprionic (AMPA)/kainite receptors from PPy electrodes (Fig. 29). In addition to modification of the CP with peptides, deposition of the CP within a hydrogel network is another attractive strategy to better integrate the CP with target cells. Hydrogels are attractive because they are biocompatible, porous,



Fig. 27. SEM images of PPySLPF-coated electrode sites. From (A) to (D) the deposition time increased, corresponding to a total charge passed of (A) 0 mC, (B) 1 mC, (C) 4 mC, and (D) 10 mC. The area of the uncoated electrode is 1250 mm2 [179]. Reproduced from Cui et al. by permission of John Wiley & Sons, Inc.

and can be tailored to possess the mechanical properties of the surrounding tissue (e.g., brain tissue), which would create a better electrode–cell interface. In a recent study, PPyPSS was electrochemically grown in alginate gels coated on electrodes [181]. PPy was found to grow vertically from the gold patterned surface up through the gel, demonstrating controlled PPy deposition (to one specific recording site). This would permit the modification of each site with different bioactive agents. Increasing deposition time of PPy decreased the impedance of PPy–alginate blends. At the biologically relevant frequency of 1 kHz this impedance value (7 kO) after 40 min of deposition was found to be even less than that of unmodified PPy. Moreover, PPy deposited within a hydrogel served to increase the surface area of the PPy film. Optimizing freeze-drying conditions allowed for control over hydrogel porosity; larger pores are important for growth of cells to improve signal transfer and vascularization of the hydrogel. Overall, these studies demonstrated that PPy–alginate-coated electrode recording sites could transport charge as efficiently as uncoated gold probes [181].

4.4.2. Poly(3,4-ethylenedioxythiophene) PPy is often the first CP explored for new applications because it is so well understood; however, as previously mentioned, PPy is susceptible to irreversible oxidation [155]. In one study, PPyPSS was found to retain only 5% of its original charge after polarization for 16 h [182]. Maintaining a high capacitance over long periods is necessary to record chronic neural activity. PEDOT has recently been explored as an alternative to PPy because it is much more stable to oxidation and more conductive than PPy. Unlike PPy, PEDOT was found to retain 89% of its conductivity under similar conditions [182]. Two studies have characterized the electrochemical deposition of PEDOT and PEDOT– MeOH on neural probes [183,184]. Both materials resulted in improved electrochemical stability and increased surface area compared to controls, which lowered impedance. Both forms of PEDOT were successfully doped with the laminin-derived DCDPGYIGSR peptide and rat glial cells preferentially grew on PEDOT–DCDPGYIGSR. PEDOT–MeOH has not been tested in vivo, but the advantage of using this CP over regular PEDOT is


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Fig. 28. Fluorescence (left) and corresponding SEM (right) images of neurons on different PPy surfaces. (A,B) PPy doped with p31 and p20. (C,D) PPy doped with p31. (E,F) PPyPSS [132]. Reproduced from Stauffer and Cui by permission of Elsevier Science Ltd.

Fig. 29. A polypyrrole based electrically controlled release system applied in a cultured neuronal network. (A) the arrows indicate electrodes that have been modified with a PPy film loaded with neurochemical CNQX (6-cyano-7-nitroquinoxaline-2,3-dione), a specific antagonist of AMPA/kainate receptors. CNQX is released directly from the electrodes and affects cells in the nearby region. (B) Neuronal waveforms recorded near a PPy/CNQX-modified electrode can be seen before release and (C) is not seen after release. (Unpublished work by William Stauffer and X. Tracy Cui, Bioengineering Department, University of Pittsburgh.)



that its monomer is more soluble in water, which would permit polymer synthesis in the aqueous systems that are necessary for incorporation of many biomolecules [183,184]. As for PPy-coated electrodes, PEDOT-coated electrodes require maximum surface area to minimize impedance. Surfactants have commonly been used in other applications to precisely control material surface structure; thus, the use of surfactants was recently explored in the synthesis of CP films on neural probe sites [185]. The authors found that the surfactant-templated PEDOT surface had a minimum impedance of 35 kO, which was less than that of controls (PEDOT and PPy). Additionally, CV analysis demonstrated that these templated films had increased charge capacity and were more electrochemically stable than controls. These improved properties are likely a result of the increased surface area (as confirmed by SEM imaging) generated by templating PEDOT. Although poly (oxythylene)10-oleyl ether surfactant is considered toxic to cells, careful selection of the surfactant concentration and proper washing permitted survival of SH-SY5Y human neuroblastoma cells on templated PEDOT [185]. In a subsequent study, the efficacy of surfactanttemplated PEDOT electrodes for recording chronic neural activity was compared to that of uncoated iridium sites over six weeks [186]. Parameters such as impedance, root mean square noise, signal amplitude, and signal-to-noise ratio were recorded. As expected, the PEDOT-coated electrode sites had lowered impedance, and thus reduced noise and decreased signal loss, compared to control iridium sites. Unfortunately, the advantages of reduced noise and decreased signal loss were mitigated by encapsulation of the electrode, and thus signal-tonoise ratios were similar for both PEDOT and control electrode sites. Future studies are focused on depositing agents to minimize the immune response, reduce encapsulation, and induce nerve growth toward the electrode [186]. 4.5. Other applications In addition to biosensors, tissue engineering, and neural probes, there are other important investigations of CPs for biological applications. These include employment of CPs as drug-delivery mediators, actuators, and antioxidants. This section briefly describes the most representative advances in these areas. As explained below, many of the drug

delivery and actuator approaches are based on the reduction of CPs, which triggers expulsion of ions (i.e., de-doping) and an associated change in volume [187]. With respect to drug delivery applications, electrical stimulation of CPs has been used to release a number of therapeutic proteins and drugs including, for example, NGF [71], dexamethasone [188,189], and heparin [190]. An early investigation demonstrated entrapment and electrically stimulated release of bovine serum albumin and NGF from PPy doped with polyelectrolytes (e.g., dextran sulfate) [71]. The use of polyelectrolytes induced high water content in the CP, allowing an easy release of the entrapped protein. For this release, PPy was reduced with negative potential, producing a rapid expulsion of anions in less than one minute. The retained activity of the released NGF was corroborated with a neurite extension assay with PC12 cells. Although protein entrapment in CPs often affects the protein’s folding and activity because of the hydrophobicity of the polymer (as discussed in the biosensor section), the use of polyelectrolytes and high water content in this study might have overcome this limitation. Another recent study investigated the release of NGF from PPy by using biotin as a co-dopant during the electrical polymerization [191]. In this investigation, NGF was biotinylated and immobilized to streptavidin entrapped within PPy films doped with both biotin and dodecylbenzenesulfonate. The authors showed that NGF was exclusively released when PPy was electrically stimulated. In particular, a 3 V pulse for 150 s resulted in a release of 22 ng/cm2 of NGF from the surface of the polymer. The activity of the released protein was confirmed by observing PC12 cell neurite extension in the presence of the released NGF. This approach provides great flexibility in the number and types of drugs that could be delivered using the conventional biotin–streptavidin strategy. The release of heparin from hydrogels immobilized onto PPy films can also be triggered by electrical stimulation [190]. PVA hydrogels were covalently immobilized onto PPy films via grafting of aldehyde groups to PPy and chemical reaction of these with hydroxyl groups from the hydrogel (Fig. 30). An accelerated release of heparin from the hydrogel was reported when PPy was electrically stimulated. The authors discussed various mechanisms that could influence these results, including changes in pH as a result of electrolysis,


909

O OH UV induced graft copolymerization O

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Fig. 30. Controlled release of heparin from poly(vinyl alcohol) (PVA) hydrogels immobilized on PPy. (A) Post-polymerization of PPy to incorporate aldehyde groups. (B) Covalent immobilization of PVA hydrogels containing heparin on PPy substrates. Controlled release of heparin was obtained by electrical stimulation of PPy [190]. Reproduced from Li et al. by permission of John Wiley & Sons, Inc.

electrophoresis, changes in temperature, and the polyanionic nature of heparin. A recent publication demonstrated the use of CP nanotubes for drug release, in which PEDOT was polymerized on top of electrospun PLGA fibers (100 nm diameter) loaded with dexamethasone (Fig. 31) [189]. PLGA was subsequently removed to produce PEDOT nanotubes encapsulating small molecules. The PEDOT nanotubes released the drug in a controlled fashion upon electrical stimulation, probably as a consequence of expansion/reduction of polymer cavities produced by the expulsion of anions. The change in volume of CPs upon electrical stimulation has also been exploited for the development of actuators to create both drug delivery devices [192] and ‘‘artificial muscle’’-type (i.e., electrochemical–mechanical) devices. For example, the actuating properties of CPs have been harnessed to release drugs from reservoirs covered by thin PPy bi-layer flaps upon application of a small potential to the PPy [192]. In artificial muscle applications, two layers of a CP are placed in a triple layer arrangement separated by a non-conductive material, as illustrated in Fig. 32 [193,194]. When current

is applied across the two CP films, one of the films is oxidized and the other is reduced. The oxidized film experiences an inflow of dopant ions and an associated expansion, whereas the reduced film expels ions and shrinks [193,195]. The combined effect is translated into a mechanical force that bends the polymer, which has been compared to the mechanisms in natural muscles. PPy, PANI, PPy– PANI composites, and composites of these polymers with carbon nanotubes (CNTs) (e.g., PANI–CNT, PANI–CNT–PPy) have all been explored for their ability to function as actuators. Of these materials, PPy–PANI produced the highest workper-cycle [196–199]. Also, the addition of CNTs to PANI fibers was found to increase the electromechanical actuation because the CNTs improved the mechanical, electronic, and electrochemical properties of PANI fibers [200]. These types of devices have potential as actuators for many biomedical applications, such as steerable catheters for minimally invasive surgery [201], micropumps and valves for labs-on-a-chip [202,203], blood vessel connectors (Fig. 33), and microvalves for urinary incontinence (reviewed by Smela [204]).



Fig. 31. PEDOT nanotubes for drug controlled release. (A) Electrospun PLGA fibers are loaded with drug (dexamethasone). (B) Degradation of PLGA fibers release the drug. (C) PEDOT is electropolymerized around PLGA fibers loaded with drug. (D) After degradation of PLGA fibers, PEDOT nanotubes are loaded with drug. (E) PEDOT nanotubes do not release drug in a neutral electrical condition. (F) PEDOT nanotubes release the drug upon external electrical stimulation with a positive voltage, which produces contraction of the nanotubes and the subsequent expulsion of the drug. (G) SEM image. Electropolymerized PEDOT nanotubes on the electrode site of an acute neural microelectrode after removing the PLGA core fibers. (H) SEM image. Higher magnification of a single PEDOT nanotube on a neural microelectrode array [189]. Reproduced from Abidian et al. by permission of Wiley-VCH Verlag GmbH Co. KGaA.

Finally, the ability of CPs to scavenge harmful free radicals from the environment has recently been explored as an additional benefit of these materials in biological applications [205]. In particular, polymers such as PANI and PPy in their neutral form were found to react effectively with 2–4 free radicals per monomer unit. The effectiveness of CPs to scavenge free radicals results from their ability to be easily oxidized and therefore does not necessitate electrical stimulation. The authors of this investigation highlighted the important role of this special property of CPs in biomedical applications where tissues experience high oxidative stress. 5. Challenges and future directions In roughly 30 years, dozens of CPs have been discovered and characterized. Thus far only a small

number of these (excluding variants of unmodified CPs) have been found to be well suited for biomedical applications. Targeted attributes include biocompatibility, conductivity, and stability in biological systems. A number of CPs have yet to be explored for biocompatibility. Perhaps there is no need to extend the library of CPs available for biomedical applications if CPs, such as PPy and PEDOT, can be modified to generate the ideal CP for specific biomedical applications. But what is an ‘‘ideal’’ CP, given the target application? Some of the most significant unanswered questions for applying CPs to biomedicine are: What is the range of conductivity necessary for tissue engineering? How can a specific cell be targeted for adhesion and/or regeneration? How can neuronal signaling be enhanced? How can a CP be made to degrade and still maintain electroactivity? How can the redox activity


PPy

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Fig. 32. Schematic of artificial muscle devices. (A) When current is applied, the left PPy film acts as anode and swells by the entry of the hydrated counter ions. Simultaneously, the right film acts as cathode and contracts by the expulsion of the counter ions. These volume changes and the constant length of the non-conducting film promotes the movement of the triple layer device. (B) By changing the direction of current the movement takes place in the opposite direction [194]. Reproduced from Otero et al. by permission of Elsevier Science Ltd.

and conductivity be preserved in the long term? How can entrapment of hydrophilic molecules be maximized? A more detailed discussion of these challenges and efforts to overcome them will follow. 5.1. Electrical properties Two of the more fundamental goals in this field have been to optimize conductivity and maximize electrical stability, which would improve biosensing and neural probe measurements, as well as enhance tissue regeneration [155]. The issue of CP conductivity is an especially challenging one because it is not clear what conductivities should be targeted given particular conditions. There is, however, evidence of an upper limit on useful CP conductivities in order to avoid protein denaturation and cell

death, as well as a lower limit at which little or no useful conductance effects are seen. These boundaries are likely dependent on cell type and applied current or potential. Although these boundaries generally remain undefined, it would be of great benefit to the biomedical community to explore/ define them for particular applications involving CPs. It can be stated, however, that CP conductivities ranging from 104 to 9 S/cm have been found to enhance cell growth to varying extents (see tissue engineering section), albeit parameters in each instance are rarely held constant. When considering the effects of CP conductivity it must be recalled that the current or potential applied to the CP can be varied and also contributes to the field effects that cells and proteins experience in the presence of a CP. Therefore, it is not surprising that there seems



Fig. 33. PPy blood vessel connector insertion surgery. (A,B) A PPy–Au bilayer device in the reduced state is curled into a roll that is inserted half-way into one of the ends of the severed blood vessel. (C,D) The other half of the device is inserted inside the other end of the vessel and the bilayer expands a few minutes after PPy is oxidized. The PPy tube holds the two parts of the blood vessel together during healing, which replaces sutures. The connector is non-thrombogenic and very thin to avoid restricting the space inside the blood vessel [204]. Reproduced from Smela by permission of Wiley-VCH Verlag GmbH Co. KGaA.

to be an optimal current or potential range to maintain cell viability. For instance, Shi and coworkers reported that fibroblast viability is maximized at currents between 10 and 50 mA for a PPy–PLA composite (3% PPy) with resistivity of 1 103 O/square [23]. Optimization of a CP’s conductivity for a particular application is an issue that begs attention on many levels. With regard to biosensors, it is essential to increase the charge transfer (i.e., conductivity) and

electrical wiring between enzymes and the CP. Common redox mediators such as ferrocene are typically used as counter ions during polymerization, but these compounds suffer from poor stability. To overcome this limitation, the functionalization of pyrrole monomers with ferrocene and viologen groups has been studied, but some of these monomers are difficult to electropolymerize, and as a result copolymerization with regular pyrrole is sometimes necessary. In addition to charge transfer mechanisms, other transduction mechanisms, such as conductimetric, impedimetric, or optical mechanisms, have been investigated. For example, impedance spectroscopy has been used to monitor the change in capacitance of PPy films on biological events such as antigen–antibody binding [206]. As another example, PPy films doped with CNTs functionalized with oligonucleotides have been successfully implemented for DNA biosensors using direct impedance measurements [117]. Optical biosensors measure the amount of light absorbed or emitted as a result of a biochemical reaction, and these devices can be directly coupled to fiber optics. As an illustration of this approach, biotinylated PPy was polymerized on an indium tin oxide (ITO)-coated optical fiber and used for binding avidin and biotinylated peroxidase; detection was performed via chemiluminescence [207]. In another approach, a PT variant is used as an optical transducer. DNA hybridization can be detected via fluorescence measurements as a result of the inherent physical interaction of this PT variant with double-stranded DNA versus singlestranded DNA, which brings about a change in fluorescence. This detection system results in very low detection limits (1021 M) from large sample volumes [208,209]. The development of methods to improve the transduction mechanism for biosensor applications continues to be an active area of research. Maximum conductivity is also critical for tissue engineering and neural probe applications. For instance, several studies have attempted to maintain conductivity while attempting to preferentially target a specific cell type through the incorporation of bioactive peptides. For instance, laminin peptides have been incorporated into PPy to target neural cells and not astrocytes, which are associated with undesired glial scar formation [132,179,180]. Unfortunately, these materials are not selective enough for neurons and over time the neural signal decreases. Such decreases in conductivity continue


to be a challenge in neural probe applications as well as in tissue engineering applications as a result of non-specific cell encapsulation, the entrapment of large molecules, and the oxidizing biological environment, which all interfere with conductivity. Recently, Martin and coworkers proposed a novel method of bypassing the insulating scar tissue that forms at device–tissue interfaces [210]. This method consists of in situ polymerization of ethylenedioxythiophene around living neural tissue to form an electrically conducting PEDOT network which branches from the electrode out into the extracellular space where it is in intimate contact with cells. This technique is still in the early stages of development, but it offers a biocompatible method for integrating electrodes into a target tissue such that conductivity is maintained. The electrical properties of CPs make them exceptionally promising materials for use in bioactuator applications. CP-based actuators can be electrically controlled, require low voltage (less than 1 V), can be continuously switched between maximum and minimum strains, and can work with liquid electrolytes, such as body fluids; however, there is still a need to increase ion mobility in the polymers to achieve faster responses.

5.2. Biological and physical properties The hydrophobicity of CPs is a major limitation for successfully entrapping proteins and maintaining their bioactivity, which is particularly critical for biosensors and drug delivery applications. Different approaches have been studied to overcome this drawback, including the use of monomers that contain both redox centers and hydrophilic chains [31,73,74], blends with polyelectrolytes [71], and composite devices containing hydrogels and CPs [112,211,212]. Surface properties are critical for tissue engineering and neural probe applications, in which the goal is to promote the attachment of specific subsets of cells. However, it is still not clear what the desirable properties, such as slight hydrophobicity, surface roughness, and conductivity, are for optimal growth of certain cells. Some studies suggest that desired properties are specific to cell type, but additional studies need to be performed, holding more parameters constant, to determine a set of guidelines for different cell types. As the modification of CPs continues to diversify their properties,

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more cell types will be analyzed for their interaction with CPs. As noted previously, surface area is of great importance for many biomedical applications. Increasing the surface area of a biosensor or neural probe, for instance, will increase the signal detected and therefore lower the detection limit. As alluded to in the application section, CPs already serve to increase the surface area of inorganic conducting surfaces; however, there have been great strides to increase the surface area even further. Currently, the most common strategy is to electrospin CPs to create nanofibers, and some applications involving such surfaces have been mentioned. Nonetheless, it has been and continues to be a challenge to create nanofibers from most (unmodified) CPs due to their insolubility. In an effort to overcome this issue Chronakis et al. used poly(ethylene oxide) as a carrier to successfully electrospin PPy [213]. Recently, a novel approach was developed, which uses V2O5 to initiate PPy or PEDOT nanofiber formation directly from pyrrole and 3,4-ethylenedioxythiophene monomers, respectively [214,215]. Although CPs generally have fairly good mechanical properties, ideally CPs used for tissue engineering and neural probes would be less brittle, more malleable, and degradable. Several studies attempted to address the former two challenges by coating PPy on malleable polyester fabrics; however, despite improved mechanical properties, conductivity dropped significantly and some delamination occurred over time [158,161]. Other methods for improving the mechanical properties of CPs include creating composites or blends and doping with large molecules that possess the desired mechanical properties [175]. These processes unfortunately result in interference with ‘‘electron hopping’’ within the CP due to the presence of insulating molecules. Similar issues arise for the few attempts that have been made to synthesize degradable CPs [162,177]. Therefore, improving mechanical properties of CPs while maintaining conductivity still remains an important goal. For bioactuator applications, advantages of CPs include their strength, their ability to function at room or physiological temperatures, and that they can be microfabricated and are lightweight [204]. However, there are important hurdles in this field as well, beginning with the necessity of a thorough understanding and modeling of the actuation mechanism, which would allow a high degree of control by modifying the CP properties. A key



technical challenge is the delamination of the CP from the electrode surface because of continuous cycling. 6. Conclusion Conducting polymers (CPs), unlike many other materials, have uses in a diverse array of applications ranging from photovoltaic devices to nerve regeneration. The unique property that ties all these applications together is the conductivity of the CP. CPs are organic in nature, making them more likely to be biocompatible; further, the presence of a conjugated backbone within the polymer endows it with the ability to conduct electrons, like metals/ semiconductors, and unlike any other polymer. In addition to these highly desirable properties, the ease of preparation and modification of CPs have made them a popular choice for many applications. This is especially true in biomedicine, where many applications benefit from the presence of conductive materials, whether for biosensing or for control over cell proliferation and differentiation. Despite the vast amount of research already conducted on CPs, for biomedical applications, the field is still growing and many questions remain to be answered.

[3]

[4]

[5]

[6]

[7]

[8] [9]

[10]

[11]

[12]

Acknowledgments

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We would like to recognize Joo-Woon Lee for initial discussions and ideas regarding the content of the article. We would like to acknowledge Davidson Day Ateh, Tracy Cui, Paul Matthew George, Diane Hoffman-Kim, Robert Langer, Peter Lelkes, Dave Martin, Tayhas Palmore, Eli Ruckenstein, Vassilios Sikavitsas, Elizabeth Smela, Gordon Wallace, Joyce Wong, and Ze Zhang for contributions of figures. We would also like to thank and acknowledge Maeve Cooney for assistance with figure copyrights, schematics, and editing. Finally, we would like to thank the many members of the Schmidt lab who provided valuable feedback and input on this project.

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References [1] Street GB. Polypyrrole: from powders to plastics. In: Skotheim TA, editor. Handbook of conducting polymers, vol. I. New York: Marcel Dekker; 1986. p. 265–91. [2] Shirakawa H, Louis EJ, MacDiarmid AG, Chiang CK, Heeger AJ. Synthesis of electrically conducting organic

[15]

[16]

[17]

[18]

[19]

[20]

polymers: halogen derivatives of polyacetylene, (CH)x. J Chem Soc Chem Commun 1977:578–80. Heeger AJ. Semiconducting and metallic polymers: the fourth generation of polymeric materials (Nobel Lecture). Angew Chem Int Ed 2001;40:2591–611. Feast WJ. Synthesis of conducting polymers. In: Skotheim TA, editor. Handbook of conducting polymers, vol. I. New York: Marcel Dekker; 1986. p. 1–43. Kundu K, Giri D. Evolution of the electronic structure of cyclic polythiophene upon bipolaron doping. Am Inst Phys 1996;105:11075–80. Hong SY, Marnick DS. Understanding the conformational stability and electronic structures of modified polymers based on polythiophene. Macromolecules 1992:4652–7. Diaz AF, Bargon J. Electrochemical synthesis of conducting polymers. In: Skotheim TA, editor. Handbook of conducting polymers, vol. I. New York: Marcel Dekker; 1986. p. 81–115. Diaz AF, Kanazawa KK. Electrochemical polymerization of pyrrole. J Chem Soc Chem Commun 1979:635. Dall’Olio A, Dascola G, Varacco V, Bocchi V. Electron paramagnetic resonance and conductivity of an electrolytic oxypyrrole [(pyrrole polymer)] black. C R Acad Sci Ser C 1968;267:433–5. Tourillon G. Polythiophene and its derivatives. In: Skotheim TA, editor. Handbook of conducting polymers, vol. I. New York: Marcel Dekker; 1986. p. 293–350. Pauw JLvd. A method of measuring specific resistivity and Hall effect of discs of arbitrary shape. Phillips Res Rep 1958;13:1–9. Kaneto K, Yoshino K, Inuishi Y. Electrical and optical properties of polythiophene prepared by electrochemical polymerization. Solid State Commun 1983;46:389–91. Chung T-C, Kaufman JH, Heeger AJ, Wudl F. Charge storage in doped poly(thiophene): optical and electrochemical studies. Phys Rev B 1984;30:702–10. Wallace G, Kane-Maguire L. Conducting polymers. In: Wnek GE, Bowlin GL, editors. Encyclopedia of biomaterials and biomedical engineering. New York: Marcel Dekker, Inc.; 2004. p. 374–83. Heeger AJ. Polyacetylene: new concepts and new phenomena. In: Skotheim TA, editor. Handbook of conducting polymers, vol. II. New York: Marcel Dekker; 1986. p. 729–56. Heeger AJ. Semiconducting and metallic polymers: the fourth generation of polymeric materials. Synth Met 2002;125:23–42. Bredas JL. Electronic structure of highly conducting polymers. In: Skotheim TA, editor. Handbook of conducting polymers, vol. II. New York: Marcel Dekker; 1986. p. 859–913. Collier JH, Camp JP, Hudson TW, Schmidt CE. Synthesis and characterization of polypyrrole–hyaluronic acid composite biomaterials for tissue engineering applications. J Biomed Mater Res 2000;50:574–84. Gurunathan K, Murugan AV, Marimuthu R, Mulik UP, Amalnerkar DP. Electrochemically synthesized conducting polymeric materials for applications towards technology in electronics, optoelectronics and energy storage devices. Mater Chem Phys 1999;61:173–91. Foulds NC, Lowe CR. Enzyme entrapment in electrically conducting polymers. J Chem Soc Faraday Trans 1986;82: 1259–64.

ARTICLE IN PRESS N.K. Guimard et al. / Prog. Polym. Sci. 32 (2007) 876–921 [21] Umana M, Waller J. Protein modified electrodes: the glucose/oxidase/polypyrrole system. Anal Chem 1986;58: 2979–83. [22] Wong JY, Langer R, Ingber DE. Electrically conducting polymers can noninvasively control the shape and growth of mammalian cells. Proc Natl Acad Sci USA 1994;91:3201–4. [23] Shi G, Rouabhia M, Wang Z, Dao L H, Zhang Z. A novel electrically conductive and biodegradable composite made of polypyrrole nanoparticles and polylactide. Biomaterials 2004;25:2477–88. [24] Aebischer P, Valentini RF, Dario P, Domenici C, Galleti PM. Piezoelectric guidance channels enhance regeneration in the mouse sciatic nerve after axotomy. Brain Res 1987;436:165–8. [25] Valentini RF, Sabatini AM, Dario O, Aebischer P. Polymer electrec guidance chanels enhance peripheral nerve regeneration in mice. Brain Res 1989;480:300–4. [26] Turner JN, Shain W, Szarowski DH, Andersen M, Martins S, Isaacsin M, et al. Cerebral astrocyte response to micromachined silicon implants. Exp Neurol 1999;156: 33–49. [27] Weidland JD, Anderson DJ. Chronic neural stimulation with thin-film, iridium oxide electrode. IEEE Trans BioMed Eng 2000;47:911–8. [28] Cui X, Hetke JF, Wiler JA, Anderson DJ, Martin DC. Electrochemical deposition and characterization of conducting polymer polypyrrole/PSS on multichannel neural probes. Sensors Actuators A 2001;93:8–18. [29] Gerard M, Chaubey A, Malhotra BD. Application of conducting polymers to biosensors. Biosenors Bioelectron 2002;17:345–59. [30] Clark LC, Lyons C. Electrode systems for continuous monitoring in cardiovascular surgery. Ann NY Acad Sci 1962;102:29–45. [31] Cosnier S, Dawod M, Gorgy K, Da Silva S. Synthesis and electrochemical characterization of a new electropolymerizable hydrophilic viologen designed for enzyme wiring. Microchim Acta 2003;143:139–45. [32] Chen J, Too CO, Wallace GG, Swiegers GF, Skelton BW, White AH. Redox-active conducting polymers incorporating ferrocenes: preparation, characterization and biosensing properties of ferrocenylpropyl and -butyl polypyrroles. Electrochim Acta 2002;47:4227–38. [33] Chen J, Too CO, Wallace GG, Swiegers GF. Redox-active conducting polymers incorporating ferrocenes, 2: preparation and characterisation of polypyrroles containing propyl- and butyl-tethered [1.1]ferrocenophane. Electrochim Acta 2004;49:691–702. [34] Cete S, Yasar A, Arslan F. An amperometric biosensor for uric acid determination prepared from uricase immobilized in polypyrrole film. Artif Cells Blood Substitutes Biotechnol 2006;34:367–80. [35] Arslan F, Yasar A, Kilic E. Preparation of Pt/polypyrrole–ferrocene hydrogen peroxide sensitive electrode for the use as a biosensor. Russian J Electrochem 2006;42: 137–40. [36] Arslan F, Yasar A, Kilic E. An amperometric biosensor for xanthine determination prepared from xanthine oxidase immobilized in polypyrrole film. Aritif Cell Blood Substitutes Immobilization Biotechnol 2006;34: 11–126.

915

[37] Alves WA, Fiorito PA, Cordoba de Torresi SI, Torresi RM. Design of molecular wires based on supramolecular structures for application in glucose biosensors. Biosensors Bioelectron 2006;22:298–305. [38] Fiorito PA, Brett CMA, Cordoba de Torresi SI. Polypyrrole/copper hexacyanoferrate hybrid as redox mediator for glucose biosensors. Talanta 2006;69:403–8. [39] Li J-P, Gu H-N. A selective cholesterol biosensor based on composite film modified electrode for amperometric detection. J Chinese Chem Soc (Taipei, Taiwan) 2006;53: 575–82. [40] Pandey PC, Mishra AP. Conducting polymer-coated enzyme microsensor for urea. Analyst 1988;113:329–31. [41] Gambhir A, Gerard M, Mulchandani A, Malhotra BD. Co-immobilization of urease and glutamate dehydrogenase in electrochemically prepared polypyrrole–polyvinyl sulphonate films. Appl Biochem Biotechnol 2001;96:249–57. [42] Palmisano F, Zambonin PG, Centonze D. Amperometric biosensors based on electrosynthesised polymeric films. Fresenius J Anal Chem 2000;366:586–601. [43] Habermuller L, Mosbach M, Schuhmann W. Electrontransfer mechanisms in amperometric biosensors. Fresenius J Anal Chem 2000;366:560–8. [44] Bakker E, Telting-Diaz M. Electrochemical sensors. Anal Chem 2002;74:2781–800. [45] Schuhmann W. Amperometric enzyme biosensors based on optimised electron-transfer pathways and non-manual immobilisation procedures. Rev Mol Biotechnol 2002;82:425–41. [46] Cosnier S. Biosensors based in electropolymerized films: new trends. Anal Bioanal Chem 2003;377:507–20. [47] Malhotra BD, Chaubey A, Singh SP. Prospects of conducting polymers in biosensors. Anal Chim Acta 2006;578:59–74. [48] Ahuja T, Mir IA, Kumar D, Rajesh. Biomolecular immobilization on conducting polymers for biosensing applications. Biomaterials 2007;28:791–805. [49] Tamiya E, Karube I, Hattori S, Sizuki M, Yokoyama K. Micro glucose sensors using electron mediators immobilized on a polypyrrole-modified electrode. Sensors Actuators 1989;18:297–307. [50] Fu Y, Yuan R, Chai Y, Zhou L, Zhang Y. coupling of a reagentless electrochemical DNA biosensor with conducting polymer film and nanocomposite as matrices for the detecion of the HIV DNA sequences. Anal Lett 2006;39:467–82. [51] Trojanowicz M, Matuszewski W, Podsiada M. Enzyme entrapped polypyrrole modified electrode for flow-injection determination of glucose. Biosensors Bioelectron 1990;5:149–56. [52] Fortier G, Brassard E, Belanger D. Optimization of a polypyrrole glucose oxidase sensor. Biosensors Bioelectron 1990;5:473–90. [53] Ramanavicius A, Kausaite A, Ramanaviciene A. Polypyrrole-coated glucose oxidase nanoparticles for biosensor design. Sensors Actuators B 2005;B111–B112:532–9. [54] Zhu M, Jiang Z, Jing W. Fabrication of polypyrrole–glucose oxidase biosensor based on multilayered interdigitated ultramicroelectrode array with contained trenches. Sensors Actuators B 2005;B110(2):382–9. [55] Wang J, Myung NV, Yun M, Monbouquette HG. Glucose oxidase entrapped in polypyrrole on high-surface area Pt


[56]

[57]

[58]

[59]

[60]

[61]

[62]

[63]

[64]

[65]

[66]

[67]

[68]

[69]

[70] [71]

N.K. Guimard et al. / Prog. Polym. Sci. 32 (2007) 876–921 electrodes: a model platform for sensitive electroenzymatic biosensors. J Electroanal Chem 2005;575:139–46. Serradilla RS, Lopez RB, Mora DN, Mark HB, Kauffmann JM. Hydrogen peroxide sensitive amperometric biosensor based on horseradish peroxidase entrapped in a polypyrrole electrode. Bionsensors Bioelectron 2002;17: 921–8. Llaudet E, Botting NP, Crayston JA, Dale N. A threeenzyme microelectrode sensor for detecting purine release from central nervous system. Bionsensors Bioelectron 2003;18:43–52. Ngamna O, Morrin A, Moulton SE, Killard AJ, Smyth MR, Wallace GG. An HRP based biosensor using sulphonated polyaniline. Synth Met 2005;153:185–8. Abu-Rabeah K, Polyak B, Ionescu RE, Cosnier S, Marks RS. Synthesis and characterization of a pyrrole-alginate conjugate and its application in a biosensor. Biomacromolecules 2005;6:3313–8. Cristea C, Mousty C, Cosnier S, Popescu IC. Organic phase PPO biosensor based on hydrophilic films of electropolymerized polypyrrole. Electrochim Acta 2005;50: 3713–8. Zhu L, Lu Y. Electrochemically embedded enzyme in polypyrrole film and its application in biosensor. Gongneng Cailiao 2005;36:619–21. Chen J, Winther-Jensen B, Lynam C, Ngamna O, Moulton S, Zhang W, et al. A simple means to immobilize enzyme into conducting polymers via entrapment. Electrochem Solid State Lett 2006;9(7):H68–70. Vidal JC, Garcia-Ruiz E, Espuelas J, Aramendia T, Castillo JR. Comparison of biosensors based in entrapment of cholesterol oxidase and cholesterol esterase in electropolymerized films of polypyrrole and diaminonaphtalene derivative for amperometric determination of cholesterol. Anal Bioanal Chem 2003;377:273–80. Li CM, Sun CQ, Song S, Choong VE, Maracas G, Zhang XJ. Impedance labelless detection-based polypyrrole DNA biosensor. Front Biosci 2005;10:180–6. Rodriguez MI, Alocilja EC. Embedded DNA–pyrrole biosensor for rapid detection of Esherichia coli. IEEE Sens J 2005;5:733–6. Chen Y, Elling, Lee Y-l, Chong S-C. A fast sensitive and label free electrochemical DNA sensor. J Phys: Conf Ser 2006;34:204–9. Jiang X, Lin X. Overoxidizing polypyrrole film directed DNA immobilization for construction of electrochemical micro-biosensors and simultaneous determination of serotonin and dopamine. Anal Chim Acta 2005;537: 145–51. Yamagishi FG, Stanford Jr TB, VanAst CI. Biosensors from conducting polymer transducers and sol–gel encapsulated bioindicator molecules. Proc Electrochem Soc 2001; 2001–18:213–23. Deshpande MV, Hall EA. An electrochemically grown polymer as an immobilization matrix for whole cells: applications in an amperometric dopamine sensor. Biosensors Bioelectron 1990;5:431–48. Sidwell JS, Rechnitz GA. Bananatrode, an electrochemical biosensor for dopamine. Biotechnol Lett 1985;7:419–22. Hodgson AJ, John MJ, Campbell T, Georgevich A, Woodhouse S, Aoki T, et al. Integration of biocomponents with synthetic structures—use of conducting polymer

[72]

[73]

[74]

[75] [76]

[77]

[78]

[79]

[80]

[81]

[82]

[83]

[84]

[85]

[86]

polyelectrolyte composites. Proc SPIE Int Soc Opt Eng 1996;2716:164–76. Campbell TE, Hodgson AJ, Wallace GG. Incorporation of erythrocytes into polypyrrole to form the basis of a biosensor to screen for rhesus(D) blood groups and rhesus(D) antibodies. Electroanalysis 1999;11:215–22. Mousty C, Galland B, Cosnier S. Electrogeneration of a hydrophilic cross-linked polypyrrole film for enzyme electrode fabrication: application to the Amperometric detection of glucose. Electroanalysis 2001;13:186–90. Fabiano S, Tran-Minh C, Piro B, Dang LA, Pham MC, Vittori O. Poly 3,4-ethylenedioxythiophene as an entrapment support for amperometric enzyme sensor. Mater Sci Eng C 2002;21:61–7. Wilchek M, Bayer EA. The avidin–biotin complex in bioanalytical applications. Anal Biochem 1988;171:1–32. Torres-Rodriguez LM, Billon M, Roget A, Bidan G. Electrosynthesis of a biotinylated polypyrrole film and study of the avidin recognition by QCM. J Electroanal Chem 2002;523:70–8. Evans SAG, Brakha K, Billon M, Mailley P, Denuault G. Scanning electrochemical microscopy (SECM): localized glucose oxidase immobilization via the direct electrochemical microspotting of polypyrrole–biotin films. Electrochem Commun 2005;7:135–40. Dupont-Filliard A, Billon M, Livache T, Guillerez S. Biotin/avidin system for the generation of fully renewable DNA sensor based on biotinylated polypyrrole film. Anal Chim Acta 2004;515:271–7. Ramanathan K, Bangar MA, Yun M, Chen W, Myung NV. Bioaffinity sensing using biologically functionalized conducting-polymer nanowire. J Am Chem Soc 2005;127:496–7. Mouffouk F, Brown SJ, Demetriou AM, Higgins SJ, Nichols RJ, Rajapakse RM, et al. Electrosynthesis and characterization of biotin-funtionalized poly(terthiophene) copolymers and their response to avidin. J Mater Chem 2005;15:1186–96. Cosnier S, Ionescu RE, Herrmann S, Bouffier L, Demeunynck M, Marks RS. Electroenzymatic polypyrrole-intercalator sensor for the determination of West Nile virus cDNA. Anal Chem 2006;78:7054–7. Riccardi CdS, Yamanaka H, Josowicz M, Kowalik J, Mizaikoff G, Kranz C. Label-free DNA detection based on modified conducting polypyrrole films at microelectrodes. Anal Chem 2006;78:1139–45. Livache T, Roget A, Dejean E, Barthet C, Bidan G, Teoule R. Preparation of a DNA matrix via an electrochemically directed copolymerization of pyrrole and oligonucleotides bearing a pyrrole group. Nucleic Acids Res 1994;22: 2915–21. Lasalle N, Roget A, Livache T, Mailley P, Veil E. Electropolymerisable pyrrole-oligonucleotide: synthesis and analysis of ODN hybridisation by fluorescence and QCM. Talanta 2001;55:993–1004. Minett AI, Barasci JN, Wallace GG. Immobilisation of anti-Listeria in a polypyrrole film. React Funct Polym 2002;53:217–27. Minett AI, Barasci JN, Wallace GG. Coupling conducting polymers and mediated electrochemical responses for the detection of Listeria. Anal Chim Acta 2003; 475:37–45.

ARTICLE IN PRESS N.K. Guimard et al. / Prog. Polym. Sci. 32 (2007) 876–921 [87] Shimomura M, Kojima N, Oshima K, Yamauchi T, Miyauchi S. Covalent immobilization of glucose oxidase on film prepared by electrochemical copolymerization of thiophene-3-acetic acid and 3-methylthiophene for glucose sensing. Polym J (Tokyo, Japan) 2001;33:629–31. [88] Welzel HP, Kossmehl G, Engelmann G, Neumann B, Wollenberer U, Scheller F, et al. Reactive groups on polymer covered electrodes, Part 4—lactate oxidase-biosensor based on electrodes modified by polythiophene. Macromol Chem Phys 1996;197:3355–63. [89] Jeffries-El M, Zaiger K, McCullough RD. Design and synthesis of novel glycopolythiophenes. PMSE Preprints 2002;86:166. [90] Kang SK, Kim J-H, An J, Lee EK, Cha J, Lim G, et al. Synthesis of polythiophene derivatives and their applications for electrochemical DNA sensor. Polym J (Tokyo, Japan) 2004;36:937–42. [91] Gautier C, Cougnon C, Pilard J-F, Casse N. Label-free detection of DNA hybridization based on EIS investigation of conducting properties of functional polythiophene matrix. J Electroanal Chem 2006;587:276–83. [92] Hiller M, Kranz C, Hubber J, Bauerle P, Schuhmann W. Amperometric biosensors produced by immobilization of redox enzymes at polythiophene-modified electrode surfaces. Adv Mater 1996;8:219–22. [93] Korri-Youssouri H, Makrouf B. Electrochemical biosensing of DNA hybridization by ferrocenyl groups functionalized polypyrrole. Anal Chim Acta 2002;469:85–92. [94] Bera-Aberem M, Ho H-A, Leclerc M. Functional polythiophene as optical chemo- and biosensors. Tetrahedron 2004;60:11169–73. [95] Azioune A, Slimane AvB, Hamou LA, Pleuvy A, Chehimi MM, Perruchot C, et al. Synthesis and characterization of active ester-functionalized polypyrrole–silica nanoparticles: application to the covalent attachment of proteins. Langmuir 2004;20:3350–6. [96] Cosnier S, Senillou A. An electrogenerated poly(pyrrolebenzophenone) film for the photografting of proteins. Chem Commun 2003:414–5. [97] Konry T, Novoa A, Shemer-Avni Y, Hanuka N, Cosnier S, Lepellec A, et al. Optical fiber immunosensor based on a poly(pyrrole-benzophenone) film for the detection of antibodies to viral antigen. Anal Chem 2005;77: 1771–9. [98] Biloivan OA, Verevka SV, Dzyadevych SV, JaffrezicRenault N, Zine N, Bausells J, et al. Protein detection based on microelectrodes with the PPy[3,3-co (1,2-C2B9H11)]2 solid contact and immobilized proteinases: preliminary investigations. Mater Sci Eng C 2006; 26:574–7. [99] Hamdi N, Wang J, Walker E, Maidment NT. An electroenzymatic L-glutamate microbiosensor selective against dopamine. J Electroanal Chem 2006;591:33–40. [100] Guerrieri A, Lattanzio V, Palmisano F, Zambonin PG. Electrosynthesized poly(pyrrole/poly(2-naphthol) bilayer membrane as an effective anti-interference layer for simultaneous. Biosensors Bioelectron 2006;21:1710–8. [101] Carelli D, Centonze D, De Giglio A, Quinto M, Zambonin PG. An interference-free first generation alcohol biosensor base don a gold electrode modified by an overoxidized nonconducting polypyrrole film. Anal Chim Acta 2006;565: 27–35.

917

[102] Guiseppi-Elie A, Brahim S, Slaughter G, Ward KR. Design of a subcutaneous implantable biochip for monitoring of glucose and lactate. IEEE Sensors J 2005;5:345–55. [103] Gade VK, Shirale DJ, Gaikwad PD, Savale PA, Kakde KP, Kharat HJ, et al. Immobilization of GOD on electrochemically synthesized Ppy–PVS composite film by cross-linking via glutaraldehyde for determination of glucose. React Funct Polym 2006;66:1420–6. [104] Singh S, Solanki PR, Pandey MK, Malhotra BD. Cholesterol biosensor based on cholesterol esterase, cholesterol oxidase and peroxidase immobilized onto conducting polyaniline films. Sensors Actuators B 2006;B115: 534–41. [105] Cen L, Neoh KG, Kang ET. Surface functionalization of polypyrrole film with glucose oxidase and viologen. Biosensors Bioelectron 2003;18:363–74. [106] Lee JW, Serna F, Nickels J, Schmidt CE. Carboxylic acid-functionalized conductive polypyrrole as a bioactive platform for cell adhesion. Biomacromolecules 2006;7: 1692–5. [107] Liu J, Bian C, Han J, Chen S, Xia S. A silicone-based bulk micromachined amperometric microelectrode biosensor with consecutive platinization and polymerization of pyrrole. Sensors Actuators B 2005;B106:591–601. [108] Ebarvia BS, Cabanilla S, Sevilla III F. Biomimetic properties and surface studies of a piezoelectric caffeine sensor based on electrosynthesized polypyrrole. Talanta 2005;66: 145–52. [109] Bjork P, Persson NK, Peter K, Nilson R, Asberg P, Inganas O. Dynamics of complex formation between biological and luminescent conjugated polyelectrolytes—a surface plasmon resonance study. Biosensors Bioelectron 2005;20:1764–71. [110] Le Floch F, Ho H-A, Leclerc M. Label-free electrochemical detection of protein based on ferrocene-bearing cationic polythiophene and aptamer. Anal Chem 2006;78:4727–31. [111] Le Floch F, Ho H-A, Harding-epage P, Bedard M, NeaguPlesu R Leclerc M. Ferrocene-functionalized cationic PT for the label-free electrochemical detection of DNA. Adv Mater 2005;17:1251–4. [112] Parthasarathy RV, Martin CR. Synthesis of polymeric microcapsule arrays and their use for enzyme immobilization. Nature 1994;369:298–301. [113] Ionescu RE, Abu-Rabeah K, Cosnier S, Marks R S. Improved enzyme retention from an electropolymerized polypyrrole–alginate matrix in the development of biosensors. Electrochem Commun 2005;7:1277–82. [114] Retama JR, Mecerreyes D, Lopez-Ruiz B, Lopez-Cabarcos E. Synthesis and characterization of semiconducting polypyrrole/polyacrylamide microparticles with GOx for biosensor applications. Colloids Surf A 2005;270–271:239–44. [115] Ionescu RE, Abu-Rabeah K, Cosnier S, Durrieu C, Chovelon J-M, Marks RS. Amperometric algal Chlorella vulgaris cell biosensors based on alginate and polypyrrole–alginate gels. Electroanalysis 2006;18:1041–6. [116] Ferrer-Anglada N, Kaempgen M, Roth S. Transparent and flexible carbon nanotube/polypyrrole and carbon nanotube/polyaniline pH sensors. Phys Stat Sol B 2006; 243:3519–23. [117] Cai H, Xu Y, He PG, Fang YZ. Indicator Free DNA Hybridization detection by impedance measurement based on the DNA-doped conducting polymer film formed on the


[118]

[119]

[120]

[121]

[122]

[123]

[124]

[125]

[126]

[127]

[128]

[129]

[130]

[131]

[132]

[133]

N.K. Guimard et al. / Prog. Polym. Sci. 32 (2007) 876–921 carbon nanotube modified electrode. Electroanalysis 2003; 15:1864–70. Cheng G, Zhao J, Tu Y, He P, Fang Y. A sensitive DNA electrochemical biosensor based on magnetite with a glassy carbon electrode modified by mulit-walled nanotubes in polypyrrole. Anal Chim Acta 2005;533:11–6. Xu Y, Ye X, Yang L, He P, Fang Y. Impedance DNA biosensor using electropolymerized polypyrrole/mulitwalled carbon nanotubes modified electrode. Electroanalysis 2006;18:1471–8. Qu L, He P, Li L, Gao M, Wallace G, Dai L. Aligned/ micropatterned carbon nanotube arrays: surface functionalization and electrochemical sensing. Proc SPIE Int Soc Opt Eng 2005:5732 (Quantum Sensing and Nanophotonic Devices II, 84–92). Wang J, Musameh M. Carbon-nanotubes doped polypyrrole glucose biosensor. Anal Chim Acta 2005;539: 209–13. Tsai Y-C, Li S-C, Liao S-W. Electrodeposition of polypyrrole-multiwalled carbon nanotube-glucose oxidase nanobiocomposite film for the detection of glucose. Biosensors Bioelectron 2006;22:495–500. Fine G, Valentini RF, Bellamkonda R, Aebischer P. Improved nerve regeneration through piezoelectric vinylidenefluoride–trifluoroethylene copolymer guidance channels. Biomaterials 1991;12:775–80. Jaffe LF, Poo MM. Neurites grow faster towards the cathode than the anode in a steady field. J Exp Zool 1979;209:115–28. Giaever I, Keese CR. Monitoring fibroblast behavior in tissue culture with an applied electric field. Proc Natl Acad Sci USA 1984;81:3761–4. Kerns JM, Pavkovic IM, Fakhouri AJ, Wickersham KL, Freeman JA. An experimental implant for applying a DC electrical field to peripheral nerve. J Neurosci Methods 1987;19:217–23. Garner B, Georgevich A, Hodgson AJ, Liu L, Wallace GG. Polypyrrole–heparin composites as stimulus-responsive substrates for endothelial cell growth. J Biomed Mater Res 1999;44:121–9. Garner B, Hodgson AJ, Wallace GG, Underwood PA. Human endothelial cell attachment to and growth on polypyrrole–heparin is vitronectin dependent. J Mater Sci: Mater Med 1999;10:19–27. Schmidt CE, Shastri VR, Vacanti JP, Langer R. Stimulation of neurite outgrowth using an electrically conducting polymer. Proc Natl Acad Sci USA 1997;94:8948–53. Wang X, Gu X, Yuan C, Chen S, Zhang P, Zhang T, et al. Evaluation of biocompatibility of polypyrrole in vitro and in vivo. J Biomed Mater Res 2004;68A:411–22. Song H-K, Toste B, Ahmann K, Hoffman-Kim D, Palmore GT R. Micropatterns of positive guidance cues anchored to polypyrrole doped with polyglutamic acid: a new platform for characterzing neurite extension in complex environments. Biomaterials 2006;27:473–84. Stauffer WR, Cui XT. Polypyrrole doped with 2 peptide sequences from laminin. Biomaterials 2006;27: 2405–13. Gomez N, Lee JY, Nickels JD, Schmidt CE. Micropatterned polypyrrole: a combination of electrical and topographical characteristics for the stimulation of cells. Adv Funct Mater 2007;17:1645–53.

[134] Ateh DD, Vadgama P, Navsaria HA. Culture of human keratinocytes on polypyrrole-based conducting polymers. Tissue Eng 2006;12:645–55. [135] Castano H, O’Rear EA, McFetridge PS, Sikavitsas VI. Polypyrrole thin films formed by admicellar polymerization support the osteogenic differentiation of mesenchymal stem cells. Macromol Biosci 2004;4:785–94. [136] Ateh DD, Navsaria HA, Vadgama P. Polypyrrole-based conducting polymers and interactions with biological tissues. J R Soc Interface 2006;3:741–52. [137] Williams RL, Doherty PJ. A preliminary assessment of poly(pyrrole) in nerve guide studies. J Mater Sci: Mater Med 1994;5:429–33. [138] Shastri VR, Schmidt CE, Kim T-H, Vacanti JP, Langer R. Polypyrrole—a potential candidate for stimulated nerve regeneration. Mater Res Soc Symp Proc 1996;414:113–8. [139] Chen SJ, Wang DY, Yuan CW, Wang XD, Zhang P, Gu XS. Template synthesis of the polypyrrole tube and its bridging in vivo sciatic nerve regeneration. J Mater Sci Lett 2000;19:2157–9. [140] Kotwal A, Schmidt CE. Electrical stimulation alters protein adsorption and nerve cell interactions with electrically conducting biomaterials. Biomaterials 2001;22: 1055–64. [141] Li Y, Neoh KG, Cen L, Kang ET. Porous and electrically conducting Ppy–poly(vinyl alcohol) composite and its applications in biomaterials. Langmuir 2005;21:10702–9. [142] Boyle A, Genie`s E, Fouletier M. Electrochemical behavior of polypyrrole films doped with ATP anions. J Electroanal Chem 1990;279:179–86. [143] Pyo M, Maeder G, Kennedy RT, Reynolds JR. Controlled release of biological molecules from conducting polymer modified electrodes: the potential dependent release of adenosine 50 -triphosphate from poly(pyrrole adenosine 50 triphosphate) films. J Electroanal Chem 1994;368:329–32. [144] Pyo M, Reynolds JR. Electrochemically stimulated adenosine 50 -triphosphate (ATP) release through redox switching of conducting polypyrrole films and bilayers. J Chem Mater 1996;8:128–33. [145] Richardson RT, Thompson B, Moulton S, Newbold C, Lum M, Cameron A, et al. The effect of polypyrrole with incorporated neurotrophin-3 on the promotion of neurite outgrowth from auditory neurons. Biomaterials 2007; 28:512–23. [146] Sanghvi AB, Miller KP-H, Belcher AM, Schmidt CE. Biomaterials functionalization using a novel peptide that selectively binds to a conducing polymer. Nat Mater 2005;4:496–502. [147] De Giglio E, Sabbatini L, Zambonin PG. Development and analytical characterization of cysteine-grafted polypyrrole films electrosynthesized on Pt- and Ti-substrates as precursors of bioactive interfaces. J Biomater Sci Polym Ed 1999;10:845–58. [148] De Giglio E, Sabbatini L, Colucci S, Zambonin G. Synthesis, analytical characterization, and osteoblast adhesion properties on RGD-grafted polypyrrole coating on titanium substrates. J Biomater Sci Polym Ed 2000;11: 1073–83. [149] Patel N, Davies MC, Hartshorne M, Heaton RJ, Roberts CJ, Tendler SJB, et al. Immobilization of protein molecules onto homogeneous and mixed carboxylate-terminated selfassembled monolayers. Langmuir 1997;13:6485–90.

ARTICLE IN PRESS N.K. Guimard et al. / Prog. Polym. Sci. 32 (2007) 876–921 [150] Chehimi MM, Azioune A, Bousalem S, Slimane AB, Yassar A. Synthesis, characterizationo, and biomedical applications of conducting polymer particles. Surf Sci Ser 2003;115:245–84. [151] Lee JW, Serna F, Schmidt CE. Carboxy-endcapped conductive polypyrrole: biomimetic conducting polymer for cell scaffolds and electrodes. Langmuir 2006;22: 9816–9. [152] Gomez N, Schmidt CE. Nerve growth factor-immobilized polypyrrole: bioactive electrically conducting polymer for enhanced neurite extension. J Biomed Mater Res A 2007; 81:135–49. [153] Cen L, Neoh KG, Kang ET. Surface functionalization of electrically conductive polypyrrole film with hyaluronic acid. Langmuir 2002;18:8633–40. [154] Cen L, Neoh KG, Li Y, Kang ET. Assessment of in vitro bioactivity of hyaluronic acid and sulfated hyaluronic acid functionalized electroactive polymer. Biomacromolecules 2004;5:2238–46. [155] Thomas CA, Zong K, Schottland P, Reynolds JR. Poly(3,4-alkylenedioxypyrrole)s as highly stable aqueouscompatible conducting polymers with biomedical implications. Adv Mater 2000;12:222–5. [156] Hakansson E, Kaynak A, Lin T, Nahavandi S, Jones T, Hu E. Characterization of conducting polymer coated synthetic fabrics for heat generation. Synth Met 2004;144:21–8. [157] Bhadani SN, Kumari M, Sengupta SK, Sahu GC. Preparation of conducting fibers via the electrochemical polymerization of pyrrole. J Appl Polym Sci 1997;64: 1073–7. [158] Tessier D, Dao LH, Zhang Z, King MW, Guidoin R. Polymerization and surface analysis of electrically conductive polypyrrole on surface-activated polyester fabrics for biomedical applications. J Biomater Sci Polym Ed 2000;11:87–99. [159] Jakubiec B, Marois Y, Zhang Z, Roy R, Sigot-Luizard MF, Dugre´ FJ, et al. In vitro cellular response to polypyrrolecoated woven polyester fabris: potential benefits of electrical conductivity. J Biomed Mater Res 1998;41: 519–26. [160] Zhang Z, Roy R, Dugre´ FJ, Tessier D, Dao LH. In vitro biocompatibility study of electrically conductive polypyrrole-coated polyester fabrics. J Biomed Mater Res 2001; 57:63–71. [161] Jiang X, Marois Y, Traore´ A, Tessier D, Dao LH, Guidoin R, et al. Tissue reaction to polypyrrole-coated polyester fabrics: an in vivo study in rats. Tissue Eng 2002;8:635–47. [162] Zelikin AN, Lynn DM, Farhadi J, Martin I, Shastri V, Langer R. Erodible conducting polymers for potential biomedical applications. Angew Chem Int Ed 2002;41: 141–5. [163] Wang Z, Roberge C, Wan Y, Dao LH, Guidoin R, Zhang Z. A biodegradable electrical bioconductor made of polypyrrole nanoparticle/poly(D,L-lactide) composite: a preliminary in vitro biostability study. J Biomed Mater Res 2003;66A:738–46. [164] Wang Z, Roberge C, Dao LH, Wan Y, Shi G, Rouabhia M, et al. In vivo evaluation of a novel electrically conductive polypyrrole/poly(D,L-lactide) composite and polypyrrole coated poly(D,L-lactide-co-glycolide) membranes. J Biomed Mater Res 2004;70A:28–38.

919

[165] Huang J, Ichinose I, Kunitake T. Nanocoating of natural cellulose fibers with conjugated polymer: hierarchical polypyrrole composite materials. R Soc Chem 2005:1717–9. [166] Bartlett PN, Birkin PR, Ghanem MA, Toh C-S. Electrochemical syntheses of highly ordered macroporous conducting polymers grown around self-assembled colloidal templates. J Mater Chem 2001;11:849–53. [167] Cassagneau T, Caruso F. Semiconducting polymer inverse opals prepared by electropolymerization. Adv Mater 2002;14:34–8. [168] LaVan DA, George PM, Langer R. Simple, three-dimensional microfabrication of electrodeposited structures. Angew Chem Int Ed 2003;42:1262–5. [169] Weiss Z, Mandler D, Shustak G, Domb AJ. Pyrrole derivatives for electrochemical coating of metallic medical devices. J Polym Sci, Part A: Polym Chem 2004;42: 1658–67. [170] Bidez PR, Li S, MacDiarmid AG, Venancio EC, Wei Y, Lelkes PI. Polyaniline, an electroactive polymer, supports adhesion and proliferation of cardiac myoblasts. J Biomater Sci Polym Ed 2006;17:199–212. [171] Wang CH, Dong YQ, Sengothi K, Tan KL, Kang ET. Invivo tissue response to polyaniline. Synth Met 1999;102: 1313–4. [172] Guterman E, Cheng S, Palouian K, Bidez P, Lelkes P, Wei Y. Peptide-modified electroactive polymers for tissue engineering applications. Polym Preprints (Am Chem Soc, Div Polym Chem) 2002;43:766–7. [173] Li ZF, Ruckenstein E. Two liquid adsorptive entrapment of a pluronic polymer into ths surface of polyaniline films. J Colloid Interface Sci 2003;264:370–7. [174] Cheng D, Xia H, Chan HS. Synthesis and characterization of surface-functionalized conducting polyaniline–chitosan nanocomposite. J Nanosci Nanotechnol 2005;5:466–73. [175] Li M, Guo Y, Wei Y, MacDiarmid AG, Lelkes PI. Electrospinning polyaniline-containing gelatin nanofibers for tissue engineering applications. Biomaterials 2006;27:2705–15. [176] Waugaman M, Sannigrahi B, McGeady P, Khan IM. Synthesis, characterization and biocompatibility studies of oligosiloxane modified polythiophenes. Eur Polym J 2003; 39:1405–12. [177] Rivers TJ, Hudson TW, Schmidt CE. Synthesis of a novel, biodegradable electrically conducting polymer for biomedical applications. Adv Funct Mater 2002;12:33–7. [178] George PM, Lyckman AW, LaVan DA, Hegde A, Leung Y, Rupali A, et al. Fabrication and biocompatibility of polypyrrole implants suitable for neural prosthetics. Biomaterials 2005;26:3511–9. [179] Cui X, Lee VA, Raphael Y, Wiler JA, Hetke JF, Anderson DJ, et al. Surface modification of neural recording electrodes with conducting polymer/biomolecule blends. J Biomed Mater Res 2001;56:261–72. [180] Cui X, Wiler J, Dzaman M, Altschuler RA, Martin DC. In vivo studies of polypyrrole/peptide coated neural probes. Biomaterials 2003;24:777–87. [181] Kim D-H, Abidian M, Martin DC. Conducting polymers grown in hydrogel scaffolds coated on neural prosthetic devices. J Biomed Mater Res 2004;71A:577–85. [182] Yamato H, Ohwa M, Wernet W. Stability of polypyrrole and poly(3,4-ethylenedioxythiophene) for biosensor application. J Electroanal Chem 1995;397:163–70.



[183] Cui XY, Martin DC. Electrochemical deposition and characterization of poly(3,4-ethylenedioxythiophene) on neural microelectrode arrays. Sensors Actuators B 2003; 89:92–102. [184] Xiao YH, Cui XY, Hancock JM, Bouguettaya M, Reynolds JR, Martin DC. Electrochemical polymerization of poly(hydroxymethylated-3,4-ethylenedioxythiophene) (PEDOT–MeOH) on multichannel neural probes. Sensors Actuators B 2004;99:437–43. [185] Yang J, Kim D-H, Hendricks JL, Leach M, Northey R, Martin DC. Ordered surfactant-templated poly(3,4ethylenedioxythiophene) (PEDOT) conducting polymer on microfabricated neural probes. Acta Biomater 2005; 1:125–36. [186] Ludwig KA, Uram JD, Yang J, Martin DC, Kipke DR. Chronic neural recordings using silicon microelectrode arrays electrochemically deposited with a poly(3,4-ethylenedioxythiophen) (PEDOT) film. J Neural Eng 2006;3: 59–70. [187] Entezami AA, Massoumi B. Artificial muscles, biosensors and drug delivery systems based on conducting polymers: a review. Iranian Polym J 2006;15:13–30. [188] Wadhwa R, Lagenaur CF, Cui XT. Electrochemically controlled release of dexamethasone from conducting polymer polypyrrole coated electrode. J Control Release 2006;110:531–41. [189] Abidian MR, Kim DH, Martin DC. Conducting polymer nanotubes for controlled drug release. Adv Mater 2006;18:405–9. [190] Li Y, Neoh KG, Kang ET. Controlled release of heparin from polypyrrole–poly(vinyl alcohol) assembly by electrical stimulation. J Biomed Mater Res A 2005;73A:171–81. [191] George PM, LaVan DA, Burdick JA, Chen CY, Liang E, Langer R. Electrically controlled drug delivery from biotindoped conductive polypyrrole. Adv Mater 2006;18:577–81. [192] Kulinsky L, Xu H, Tsai H-KA, Madou M. System-based approach for an advanced drug delivery platform. Proc SPIE Int Soc Opt Eng 2006;6173 (Smart Structures and Integrated Systems, 61730M/1-61730M/6). [193] Otero TF, Sansinena JM. Bilayer dimensions and movement in artificial muscles. Bioelectrochem Bioenergy 1997;42:117–22. [194] Otero TF, Cortes MT. A sensing muscle. Sensors Actuators B 2003;96:152–6. [195] Gandhi MR, Murray P, Spinks GM, Wallace GG. Mechanism of electromechanical actuation in polypyrrole. Synth Met 1995;73:247–56. [196] Tahhan M, Truong V-T, Spinks GM, Wallace GG. Carbon nanotube and polyaniline composite actuators. Smart Mater Struct 2003;12:626–32. [197] Spinks GM, Campbell TE, Wallace GG. Force generation from polypyrrole actuators. Smart Mater Struct 2005;14: 406–12. [198] Spinks GM, Xi B, Troung V-T, Wallace GG. Actuation behavior of layered composites of polyaniline, carbon nanotubes and polypyrrole. Synth Met 2005;151:85–91. [199] Spinks GM, Mottaghitalab V, Bahrami-Samani M, Whitten PG, Wallace GG. Carbon nanotube reinforced polyaniline fibres for high strength artificial muscles. Adv Mater 2006;18:637–40. [200] Mottaghitalab V, Xi B, Spinks GM, Wallace GG. Polyanline fibres containing single walled carbon nanotubes:

[201]

[202]

[203]

[204] [205]

[206]

[207]

[208]

[209]

[210]

[211]

[212]

[213]

[214]

[215]

[216]

[217]

[218]

enhanced performance artificial muscles. Synth Met 2006; 156:796–803. Mazzoldi A, De Rossi D. Conductive-polymer-based structures for a steerable catheter. Proc SPIE Int Soc Opt Eng 2000;3987:273–80. Smela E, Gadegaard N. Surprising volume change in PPy(DBS): an atomic force microscopy study. Adv Mater 1999;11:953–7. Low LM, Seetharaman S, He KQ, Madou MJ. Microactuators toward microvalves for responsive controlled drug delivery. Sensors Actuators B 2000;67:149–60. Smela E. Conjugated polymer actuators for biomedical applications. Adv Mater 2003;15:481–94. Gizdavic-Nikolaidis M, Travas-Sjdic J, Bowmaker GA, Cooney RP, Kilmartin PA. Conducting polymers as free radical scavengers. Synth Met 2004;140:225–32. Ouerghi O, Touhami A, Jaffrezic-Renault N, Martelet C, Ben Ouada H, Cosnier S. Impedimetric immunosensor using avidin–biotin for antibody immobilization. Bioelectrochemistry 2002;56:131–3. Marks RS, Novoa A, Konry T, Krais R, Cosnier S. Indium tin oxide-coated optical fiber tips for affinity electropolymerization. Mater Sci Eng C 2002;21:189–94. Ho HA, Dore K, Biossinot M, Bergeron MG, Tanguay RM, Boudreau D, et al. Direct molecular detection of nucleic acids by fluorescence signal amplification. J Am Chem Soc 2005;127:12673–6. Dubus S, Gravevl J-F, Le Drogoff B, Nobert P, Veres T, Boudreau D. PCR-free DNA detection using a magnetic bead-supported polymeric transducer and microelectromagnetic traps. Anal Chem 2006;78:4457–64. Richardson-Burns SM, Hendricks JL, Foster B, Povlich LK, Kim D-H, Martin DC. Polymerization of the conducting polymer poly(3,4-ethylenedioxythiophene) (PEDOT) around living neural cells. Biomaterials 2007;28:1539–52. Brahim S, Narinesingh D, Guiseppi-Elie A. Polypyrrole– hydrogel composites for the construction of clinically important biosensors. Biosensors Bioelectron 2002;17:53–9. Brahim S, Guiseppi-Elie A. Electroconductive hydrogels: electrical and electrochemical properties of polypyrrole– poly(HEMA) composites. Electroanalysis 2005;17:556–70. Chronakis IS, Grapenson S, Jakob A. Conductive polypyrrole via electrospinning: electrical and morphological properties. Polymer 2006;47:1597–603. Zhang X, Manohar SK. Bulk synthesis of polypyrrole nanofibers by a seeding approach. J Am Chem Soc 2004; 126:12714–5. Zhang X, MacDiarmid AG, Manohar SK. Chemical synthesis of PEDOT nanofibers. Chem Commun (Cambridge, United Kingdom) 2005;42:5328–30. Ramanathan K, Pandey SS, Kumar R, Gulati A, Murthy AS. Covalent immobilization of glucose oxidase to poly (o-amino benzoic acid) for application to glucose biosensor. J Appl Polym Sci 2000;78:662–7. Ramanathan K, Ram MK, Malhotra BD, Murthy AS. Appication of polyaniline—langmuir—blodgett films as a glucose biosensor. Mater Sci Eng C 1995;3:159–63. Kumar A, Rajesh, Chaubey A, Grover SK, Malhotra BD. Immobilization of cholesterol oxidase and potassium ferricyanide on dodecylbenzene sulfonate ion doped polypyrrole film. J Appl Polym Sci 2001;82:3486–91.

ARTICLE IN PRESS N.K. Guimard et al. / Prog. Polym. Sci. 32 (2007) 876–921 [219] Singh S, Chaubey A, Malhotra BD. Preparation and characterization of an enzyme electrode based on cholesterol esterase and cholesterol oxidase immobilized onto conducting polypyrrole films. J Appl Polym Sci 2004; 91:3769–73. [220] Chaubey A, Gerard M, Singhal R, Singh VS, Malhotra BD. Immobilization of lactate dehydrogenase on electrochemically prepared polypyrrole–polyvinyl sulphonate composite films for application to lactate biosensors. Electrochim Acta 2000;46:723–9.

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[221] Chaubey A, Pande KK, Singh VS, Malhotra BD. Coimmobilization of lactate oxidase and lactate dehydrogenase on conducting polyaniline films. Anal Chim Acta 2000;407:97–103. [222] Gerard M, Ramanathan K, Chaubey A, Malhotra BD. Immobilization of lactate dehydrogenase on electrochemically prepared polyaniline films. Electroanalysis 1999;12:450–2. [223] Adeloju SB, Shaw SJ, Wallace GG. Polypyrrole-based amperometric flow injection biosensor for urea. Anal Chim Acta 1996;323:107–13.