Decoration of amyloid fibrils with luminescent conjugated polymers

Abstract

In this work we report the coating of a biological template with a polar, but uncharged, luminescent conjugated polymer, soluble in organic solvents but not in water, to produce a nanowire. Amyloid fibrils from bovine insulin were decorated with an alternating polyfluorene derivative. Decorated fibrils were partially aligned on hydrophobic surfaces as separate and bundled fibrils, by means of molecular combing. The single molecule spectroscopy technique utilizing excitation by rotating linearly polarized light and fluorescence detection through a rotating polarizer showed a high degree of anisotropy of the polymer chains on the individual fibrils. The high degree of polarization indicated highly oriented polymer chains with the preferential orientation of the polymer backbone along the fibrils. The anisotropy ratios are comparable with those of well-oriented polymer chains in films.

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Introduction

Nanoscale organization of electroactive, metallic and semi-conducting, materials into 1D objects is a key task in nanotechnology. There is an interest in materials and components with inherent properties to self-assemble into a functional organization. This bottom-up approach offers, unlike conventional top-down techniques, the self-assembly possibilities to control the material on the single molecule level.¹ Fibrillar or tubular biological molecules have inherent 1D nanoscale geometry and specific biochemical structures, making them suitable as templates for organization of organic as well as inorganic materials.^2,3 There are a number of nanoscale electronic devices generated through assembly based on the well-defined structures of biomolecular materials.^4–6 However, biomolecules are in many cases incompatible with the processing conditions of electrically or optically active materials. Harsh conditions, which normally deviate from physiological conditions in terms of temperature and access to water, result in loss of biochemical functionality or degradation of the material. The processing may also lead to loss of geometry in the desired structure. Metallization of fibrillar biological molecules give thicker fibrils, widening 10 to 50 times for DNA,⁷ and at least 5 times for amyloid fibrils.⁸ Here we focus on nanostructures made with electronic polymers, wirelike macromolecules capable of electronic transport and optical absorption and emission. Water-soluble conjugated polymers (CPs) can form complexes with biological molecules and through changes in the optical properties of the CP specific interactions with a secondary biomolecule have been shown, e.g.DNA hybridization,⁹protein–protein¹⁰ and peptide–peptide interaction.¹¹ Similar CPs have also been shown to decorate fibrillar biomolecules with very small coarsening of the structures.^12,13 Well-performing CPs for device applications, such as solar cells and organic light emitting diodes, are normally molecules of low polarity and only soluble in organic solvents. The mismatch in solubility makes the combination of these CPs and biomolecules difficult. Here we demonstrate one route to such objects.

An amyloid fibril is a protein fibril, which can be formed from proteins,^14–16 specifically designed peptides^17–19 and peptide–polymer complexes²⁰ under conditions that destabilize the native state. The high stability together with the geometry, a diameter of ∼10 nm²¹ for mature fibrils and ∼1.2 nm for protofibrils²² and lengths up to over 10 μm, make amyloid fibrils suitable for self-assembly of nanowires. Metallization of amyloid fibrilsvia binding of gold nanoparticles followed by successive deposition of silver and gold in order to form conducting nanowires have been demonstrated by Scheibel et al.⁸ We have reported that water-soluble, semiconducting conjugated oligoelectrolytes (CPE) can be integrated within amyloid fibrils of bovine insulin (BI) through self-assembly.²³ The fibrillar material demonstrated the electro-optical activity of the conjugated oligoelectrolyte. In another study we showed that an anionic water-soluble CP can decorate insulin fibrils with the polymer chains along the fibril axis.¹³ The decorated fibrils were aligned on surfaces by the means of molecular combing.

In this report we show that amyloid fibrils can be decorated with a water-insoluble CP, APFO-12 (see Fig. 1), without major degradation of the biological material. APFO-12 is a low bandgap, random, alternating polyfluorene co-polymer with m = n = 50%. It has two absorption peaks at λ_max= 398, and 543 nm when spincoated from chloroform.²⁴ Luminescence is observed with a peak at 650 nm. Alternating polyfluorenes with similar structure have shown good performance in organic solar cells.^25–27 In conjugated materials the supramolecular order is crucial for the performance in opto-electronic processes.^28,29 Through fluorescence polarization we can conclude very good alignment of the APFO-12 chains along the fibril axis and polarization modulation in the range 79 to 97%.

Fig. 1 Chemical structure of APFO-12.

Experimental

The synthesis of APFO-12 has been reported elsewhere.²⁴M_n = 1900 and M_w = 3600, n and m in Fig. 1 is 50%. Insulin amyloid fibrils were formed through incubation of 320 μM native insulin (Sigma Aldrich) in 25 mM HCl at 65 °C for 6–10 h as earlier described by us.¹³ The insulin amyloid fibrils were diluted 50 times in MilliQ water. Tetrahydrofuran (THF) was added 1 : 1 to the diluted insulin fibrils and the solution was incubated for ∼2 min. APFO-12 dissolved in THF was added to a concentration of 40 ng ml⁻¹ (Conc 1), 200 ng ml⁻¹ (Conc 2) or 8 ng ml⁻¹ (Conc 3). Glass and Si/SiO₂ substrates were cleaned and silanized with dichlorodimethylsilane to yield a hydrophobic surface. The complexes between amyloid fibrils and APFO-12 were deposited/aligned on the surfaces with the method reported for amyloid fibrils/PTAA (poly(thiophene acetic acid)) complexes.¹³ Droplets of the amyloid fibril/APFO-12 solution was deposited on the hydrophobic surface, incubated for 30 s and moved over the surface with a nitrogen gas flow. The fibrils were partially aligned along the direction of movement. Fluorescence images of the fibrils on glass were recorded with an epifluorescence microscope (Zeiss Axiovert inverted microscope A200 Mot) equipped with a CCD camera (Axiocam HRc 3-chip), using a 546/12 nm filter (LP590) and a 100× oil immersion objective. Atomic force microscopy (AFM) imaging was performed in ambient conditions by standard procedures in tapping mode on a SFM-Nanoscope III, Digital Instruments, with a J scanner. Cantilevers for tapping mode were obtained from NT-MDT. For studies of fluorescence of single complexes a home- built wide-field fluorescence microscope based on a commercial Olympus IX-71 inverted microscope was used. A sketch of the setup is shown in ESI† Fig. S1. Excitation was carried out using the 514 nm line of a cw argon-ion laser. Fluorescence light was collected by an objective lens (Olympus LUCPlanFl 40×, NA 0.6) and projected onto a CCD camera (Photometrics, Cascade 512B). Spatial resolution of the microscope was 0.6 μm corresponding to 3 pixels on the CCD chip. During all experiments the sample was kept at 10⁻² Torr to increase photostability. Simultaneous imaging and recording of spectra was achieved by placing a holographic grating (Edmund Optics) in front of the CCD camera. The dispersion of the grating was horizontal giving 2 nm per pixel of the CCD chip. The distance between the CCD chip and the grating was such that zero-order (image) and first-order (spectrum) of the diffraction appeared on the CCD chip at the same time. Selection of a particular place on the sample was done by using a vertical slit with adjustable width placed in an intermediate image plane. Calibration of the overall spectral sensitivity of the setup (including all optics and CCD camera) was done by using a special calibrated tungsten lamp with known spectrum. Exposure time was up to 20 s depending on the brightness of the complexes. The instrumental response function had a FWHM ≈ 6 nm. The spectra were smoothed over 2 data points before analysis. To record fluorescence polarization sensitive microscopy in excitation and emission, stepmotor driven rotating polarizers were placed in front of the CCD camera and in the exciting beam, operated at two different frequencies (see ESI† Fig. S4). Time dependence of the fluorescence intensity of a chosen “spot” on the image was obtained by taking the average intensity over 5 × 5 pixels for all consecutive frames in the “movie” and then subtracting the background fluorescence for each frame. The background signal was obtained in the same way but from an empty position on the sample.

Results and discussion

We incubated bovine insulin in acidic conditions (25 mM HCl in H₂O, pH 1.6) at 65 °C for 6–10 h to form amyloid-like fibrils. Chloroform is a commonly used solvent for conjugated polymers, but dispersion of amyloid fibrils in chloroform or the addition of smaller volumes of chloroform to an aqueous fibril solution resulted in visible sedimentation of the biological material. Tetrahydrofuran (THF) is a polar organic solvent, which is miscible with water, and also a solvent for APFO-12. Sedimentation of fibrils can not be seen in a 1 : 1 mixture between aqueous fibril solution and THF. This result suggest that water–THF mixtures can be a suitable environment for decorating amyloid fibrils with a water-insoluble CP. The polar ethylene oxide side chain enables a good solubility of APFO-12 in THF, making it a good model compound for studies of CP decoration of fibrils. It is known that certain molecules, such as Thioflavine T³⁰ and Congo Red,³¹ interact specifically, and that water-soluble thiophene derivatives can interact, with amyloid fibrils in solution^32,33 and in tissue samples.^34,35

To evaluate the possibility to decorate insulin fibrils with APFO-12, insulin fibrils were diluted in water to a concentration of 6.4 μM, followed by the addition of THF in a 1 : 1 volume ratio. After a few minutes of incubation APFO-12 in THF was added to a concentration of Conc 1 = 40 ng ml⁻¹, Conc 2 = 200 ng ml⁻¹ and Conc 3 = 8 ng ml⁻¹. The solution was studied using a fluorescence microscope and fluorescing wire-like objects were observed. To facilitate the studies of the decorated fibrils, they were deposited with molecular combing on glass surfaces made hydrophobic with dichlorodimethylsilane. In molecular combing techniques a receding air/water meniscus is used to stretch and align wire-like molecules on surfaces.³⁶ The orientation of the objects will be perpendicular to the receding meniscus. Different classes of wire/fibril-like objects have been subjected to molecular combing: biomolecules (DNA,³⁶ fibrillin,³⁷ titin,³⁸ and amyloid fibrils¹³) but also carbon nanotubes³⁹ and inorganic nanowires.^40,41 The process is dependent on moderate interaction strength between the fibrillar molecule and the surface, and orientation/stretching will be controlled by viscous drag and surface tension. Our studies of molecular combing of the PTAA/insulin fibril complex have shown that alignment can be achieved on hydrophobic surfaces formed by PDMS printing.¹³

The APFO-coated fibrils can be partially aligned on surfaces with a molecular combing method, similar to what we earlier have demonstrated with PTAA/insulin fibrils (Fig. 2, Fig. 3 and ESI† Fig. S2). The APFO-12/insulin fibril solution described above was deposited in small droplets on a surface, incubated for ∼1 min and blown over the surface with a nitrogen gas stream. The surface was thoroughly rinsed with milli-Q water and examined in a fluorescence microscope. Clean hydrophilic glass resulted in poor deposition of decorated fibrils, but hydrophobic surfaces, either silanized or PDMS printed, gave better results. The observed fibrils are less aligned and more small fluorescent fragments can be seen, if compared with fibrils decorated with PTAA.

Fig. 2 Fluorescence micrographs of partially aligned insulin fibrils decorated with APFO-12 at Conc 1. Scale bar represents 15 μm.

Fig. 3 Atomic force micrograph of partially aligned insulin fibrils decorated with APFO-12 at Conc 1. (ESI† Fig. S2 shows fluorescence micrographs from the same sample.) The white arrows point at examples of fibrils with height of ∼4 nm and the black arrows point at fibrils with height ∼1.5 nm. Scale bar represents 4 μm.

These fragments are also visible in AFM (see Fig. 3) and can be fragments of fibrils or aggregated, non-fibrillar, insulin. A dilution of APFO-12 in THF–water does not give similar aggregates. The mixing with THF might be the cause of degradation as well as making alignment more difficult. It is also likely that the coating of APFO-12 affects the fibril–surface interaction. Interestingly fibrils in U-shapes can be found, which often is the case when applying molecular combing to DNA,¹² but has not been observed by us for PTAA/insulin. The interactions between hydrophobic surfaces and APFO-12/insulin fibril complexes are strong enough to withstand extensive rinsing with water as well as THF. Using AFM the thicknesses (d) of the fibrils have been determined and two groups of fibrils could be found, d₁ = 1.4 ± 0.4 nm (based on measurements on 15 individual fibrils, examples of such fibrils are marked with black arrows in Fig. 3) and d₂ = 4.2 ± 1.1 nm (based on measurements on 20 individual fibrils, examples of such fibrils are marked with white arrows in Fig. 3). These values are similar to what Jansen et al.²² reported for protofilaments (1.2 nm) and fibrils (3–7 nm) of insulin. The heterogeneity in fibril thickness makes it difficult to estimate the thickness of the APFO-12-coating on the fibrils. Furthermore it is not possible to tell if all fibrils in Fig. 3 are covered with APFO-12 or if some fibrils are only partly covered. In our studies of decoration of fibrils we have used fibrils formed during various times of heat incubation. We have seen no difference in the degree of coverage on fibrils from shorter incubation times, presumably containing more protofilaments, compared to the more mature fibrils from longer incubation times.

To further analyze the decoration of APFO-12 on insulin fibrils, the emission spectra and the fluorescence polarization of single fibrils were studied in single molecule spectroscopy set-up. No significant difference could be found in the emission spectra within the same decorated fibril or when different fibrils are compared. Typical spectra can be found in ESI† (Fig. S2). To evaluate the organization and degree of order of the APFO-12 polymer chains on the fibrils, polarization sensitive spectroscopy was used, a powerful method to analyze the orientation of the transition dipoles of the polymer backbone.⁴² The intensity of the emitted light from APFO-12/insulin fibril complexes was recorded while rotating polarizers in the exciting and emitting beam simultaneously at two different frequencies. The intensity trajectory follows two superimposed sinusoidal curves (see ESI† Fig. S4). After background subtraction, the fluorescence intensity can be fitted to the function:

I = I₀(1 −m_exsin²(ω_ext−φ_ex))(1 −m_emsin²(ω_emt−φ_em)),  m_ex,m_em∈ [0,1] (1)

where I₀ is the maximal intensity, m_ex and m_em are modulations or polarization degrees in excitation and emission respectively.

Note that this expression is not a general one and, therefore, it does not represent the emission intensity of an arbitrary molecular system. We found that this particular model can describe the fluorescence of decorated fibrils very well. The model will be valid for some systems, for example for very highly oriented systems (then the polarization is simply given by the geometry) and for systems showing perfect energy transfer from initially excited states to emitting states having identical orientations of transition dipole moments.

In Fig. 4 the modulation in excitation (m_ex) is plotted against the modulation in emission (m_em) for parts of fibrils decorated with three different concentrations of APFO-12. Modulation in emission from these fibrils spans from 0.79 to 0.97 (average = 0.89) and modulation in excitation from 0.79 to 0.93 (average = 0.87) (see Fig. 4 and 5). In Fig. 4 it can be seen that the modulation in emission is slightly higher than in excitation. No clear relation between the concentration of APFO-12 and the modulation can be seen in the concentration range used in this study (8 ng ml⁻¹ to 200 ng ml⁻¹).

Fig. 4 Polarization modulation of emitted light from fibrils decorated with APFO-12 at concentration Conc 1 = 40 ng ml⁻¹, Conc 2 = 200 ng ml⁻¹ and Conc 3 = 8 ng ml⁻¹. The line y = x is added as a guideline for the eye.

Fig. 5 (left) m_em and m_ex are plotted against emission intensity of fibrils decorated with APFO-12. (right) The same data set where G and LD are plotted against emission intensity of fibrils decorated with APFO-12.

Many reports on single molecule and ensemble molecule polarization evaluate data using such a parameter as modulation depth M = (I_max−I_min)/(I_max + I_min), where I_max and I_min are maximal and minimal fluorescence intensities obtained upon rotating a polarizer.

In order to correlated the modulations depths m_ex and m_em obtained in our experiments with two rotating polarizers to the literature data obtained in experiments with single rotating polarizer one can express Mviam_ex and m_em using eqn (1):

M = m_em/(2 −m_em)

when the sample is excited by light and its fluorescence is measured through a rotated polarizer, then M reflects fluorescence states anisotropy; or

M = m_ex/(2 −m_ex)

when the sample is excited by linearly polarized rotated light, and overall emission is detected, then M has the meaning of anisotropy of light absorbers.

For example in the experiment with excitation by circular polarized light M = 1 corresponds to fully polarized fluorescence coming from a single dipole or several perfectly aligned dipoles; M = 0 corresponds to totally unpolarized (e.g. from randomly oriented dipoles) or circularly polarized fluorescence. M < 1 indicates partially polarized emission e.g. from several dipoles with different orientations. We can correlate the amplitudes of excitation and emission in our experiments with 2 polarizers to related experiments done on oriented conjugated polymer films (vide infra) and note that the linear dichroism (LD) of the sample is LD = A_max/A_min = 1/(1 −m_ex) and the often used fluorescence polarisation anisotropy is G = (1 + M)/(1 −M) = 1/(1 −m_em). However, our model (1) does not account for dependence between emission and absorption processes and by this underestimates the G value, compared to studies of polymers in films (vide infra) where G values are determined with excitation linearly polarized parallel to the axis of the polymer chain orientation.

The modulation values can be compared with studies on single molecules studies of conjugated polymers, where the CPs normally are studied in a matrix of another polymer. Forster et al. recently showed that single polymer chains of a polythiophene derivative, PDOPT, had an average degree of polarization (modulation) in emission of 0.52, but were in the wide range from 0.06 to 0.96.⁴³ The cause of the relatively low polarization was attributed to emission from several differently oriented dipoles during the exposure time. The lack of correlation between emission intensity and polarization modulation was also seen by Forster et al. Becker and Lupton studied the fluorescence polarization of the backbone of a dye-endcapped polyindenofluorene derivative and concluded that the polarization modulation was significantly larger in the emission compared to excitation, but in the range from 0.1 to 0.9.⁴² This can be explained by a funneling of the excitation energy to a few or possibly only one emitting chromophore on the chain, leading to linearly polarized emission. In two studies of single OPV (oligo(phenylenevinylene)) molecules with rotating polarization of excitation the depth of modulation (DOM = (I_max−I_min)/I_average) was varying between 0.1 to 1, where DOM = 1 was considered to come from elongated molecules.^44,45

The intensity modulation from the decorated fibrils is not from single polymer chains, but every value in Fig. 3 and 4 is rather from a population of many emitting dipoles. All APFO-12 decorated fibrils showed a continuous fluorescence, with no discrete intensity fluctuations, “blinking”, as seen in a discontinuous decoration of conjugated polyelectrolyte on DNA.¹² Continuous fluorescence was also seen in the case of PTAA decorated insulin fibrils. This observation indicates that the APFO-12 clusters on the fibrils are larger than 10 nm, the upper limit of Förster energy transfer distance towards a fluorescence quencher in conjugated polymers.^46–48 Summers et al. have shown that single OPV molecules can be in a state of continuous fluorescence when present in a rigid environment.⁴⁴ Possibly the APFO-12 are locked in a non-blinking state when decorated on the fibrils due to various interactions with the environment.

If the bundles of APFO-12 chains are relatively large, compared to the single polymer chain, the polarization values can be compared with those of oriented polymer chains in films. The degrees of absorption and emission polarization for aligned CP chains in films are often presented with linear dichroism of absorption (LD) and the photoluminescence anisotropy of emission G = I_max/I_min. Stretch-oriented gel films of poly(2-methoxy)-5-(2′-hexoxy)-p-phenylenevinylene (MEHPPV)²⁹ and poly(3-octylthiophene)⁴⁹ in polyethylene showed a polarization direction along the draw axis and G≈ 60 and G≈ 40, respectively. Gustavsson et al. demonstrated stretch-oriented films of pure poly(3-octylthiophene) with an LD ≈ 7.⁵⁰ Recently Zheng et al. oriented a liquid-crystalline conjugated polymer, poly(9,9-dioctylfluorene-co-benzothiadiazole) (F8BT) by means of nanoconfinement during nanoimprinting, with a maximum G value of 66,‡, and LD = 7.⁵¹ These values should be compared with our fibril data, where G values range from 5 to 38 (average 10.5). Our measured excitation amplitude does reflect the orientation of the polymer chain, and the range of 0.8 < m_ex < 0.93 indicates a linear dichroism of the order LD ≈ 5–14.4 (average 8), depending on the absorber. These numbers also contain errors, which are presently difficult to quantify, but should be high for high LD, G. The problem of accurately determining the orientation function at very high orientation is also found in other methods. We conclude that for alignment of conjugated polymer chains, alignment along amyloid nanowires is comparable to stretch aligning of dilute polymer gels and to nanoimprinting in liquid crystalline polymers, and superior to stretch alignment in the bulk.

In Fig. 5 (right) the G and LD values from the analyzed fibrils are plotted against I₀. No clear correlation can be seen between emission intensity and G or LD value. As m_ex < m_em, transport of excited states leads to enhanced polarisation of emission. We conclude that the subset of absorbers found among the emitting species is less oriented. In the case of polymer films, the transport of the excited state might lead to a selection of better-aligned dipoles giving a higher emission anisotropy. We have measured the decay time (∼2 ns) of photoluminescence from APFO-12 in films as well as decorated on fibrils: the values do not differ significantly. Therefore excited state pathways and energy transfer are similar for APFO-12 in film and coated on fibrils. The thickness of the coating is not known.

The G values of PTAA decorated fibrils in our previous study, recorded with circularly polarized excitation and a rotating polarizer in emission, ranged from 2.5 to 17.3 (average 7). In the case of PTAA decorated fibrils there is a correlation between low emission intensity and high G value. The cause of the higher G values in the APFO-12 may be caused by the greater stiffness of the polymer backbone. A quantitative value of the stiffness is the persistence length, which for poly-3-hexylthiophene in THF-d₈ was measured as 33 Å,⁵² while for a polyfluorene derivative, poly{2,7-[9,9-bis((S)-3,7-dimethyloctyl)]fluorene} (PDMOF) in THF it was measured as 95 Å.⁵³ The low solubility of APFO-12 in water might also increase the π-stacking of the polymer chain onto the fibrils. The lower polarization anisotropy in some cases can be attributed to a considerable variation of the direction of the polymer chains, with respect to the direction defined by the protein fibril. There may also be a bundling of several, undistinguishable in the microscope, decorated fibrils. The variation in the degree of order of the fibrils will not affect the degree of polarization in our measurements where we collect light from only parts of one emitting fibril.

The angle of the polarizer when the maximum intensity is recorded from a part of the decorated fibrils indicates the average orientation of the emitting dipoles. When comparing this angle with the orientation of the fibrils on the surface it can be concluded that the polymer chains are preferentially aligned along the fibrils. This result was also seen for insulin fibrils decorated with PTAA. In fluorescence polarization of single molecules of rigid-rod ladder type CPs Müller et al. observed ultrafast rotation of the plane of polarization in absorption and emission.⁵⁴ This rotation of the plane of polarization has also been observed for dye-endcapped CP , which is attributed to energy transfer between absorbing and emitting dipoles of different orientation.⁴² In our studies of APFO-12 decorated fibrils only very small phase shifts, maximum up to six degrees, between polarization in emission and excitation were observed. We can therefore conclude that the absorbing as well as the emitting dipoles are preferentially aligned along the fibrillar axis.

The nature of the interaction between the CP and the amyloid fibril is an interesting subject for bioimaging. Water-soluble polythiophenes with various side-chains has been shown to interact with amyloid both in vitro^32,35 and in tissue samples.^34,35 Also here the exact nature of the interaction is unknown, but both backbone and sidechain interactions are discussed. In the case of APFO-12 the side-chain is uncharged, but polar, which could in the slightly acidic solution mediate interactions with the insulin fibrils by hosting cationic charges. Calamai et al. have recently shown that amyloid fibrils can interact strongly with oppositely charged polyelectrolytes.⁵⁵ The backbone of APFO-12 could also be dominant in the interaction with the fibrils. Nesterov et al. suggests that the presence of a hydrophobic planarized, or easily planarizable, π system is an important design feature to obtain high binding specificity between optical probes and amyloid.⁵⁶ For the well-established amyloid probe Congo Red,⁵⁷ parallel as well as perpendicular orientation³¹ of the probe has been discussed, however both electrostatic and aromatic interactions have been pointed out as the source of specificity.³¹ Hydrophobic interactions are important in fibril nucleation and elongation for insulin fibrils.⁵⁸ The areas of the growing amyloid fiber showing hydrophobic protein–protein interactions might also interact with a hydrophobic conjugated backbone.

Organized conjugated polymers in stable nanowire geometries, such as the APFO-12 decorated fibrils, may be very interesting both for use as separated nanowires and in oriented film assemblies. In this work we have demonstrated the polarization anisotropy of absorption and emission, but assuming that charge injection and transport can be achieved into the APFO-12 on the fibrils polarized electroluminescence could also be realized. Organization of decorated fibrils in device structures would then give electro-optic components on the nanoscale. Alternating polyfluorenes have (vide supra) been used in well performing organic solar cells, and another potential use of decorated fibrils is to structure the conjugated polymer in bulk heterojunction cells. The formation of an interpenetrating network of decorated fibrils could enhance the charge extraction and thereby the efficiency of the cell. It should also be emphasized that the use of protein based fibrils as a structural element gives the possibility to vary the functionality of the fibrils, since a large number of protein and peptides are known to form amyloid fibrils. Several routes, including peptide synthesis, recombinant protein synthesis and covalent modification of proteins, can be used to achieve fibrils with affinities for certain materials or surfaces.

Conclusions

Amyloid fibrils are biological nanowires of unusually high stability. In this article we demonstrate the first example of a biological nanowire coated with a polar, non-charged conjugated polymer. The interaction between the CP and the amyloid fibrils was made in a water–THF mixture. The CP-decorated fibrils could be partially aligned on hydrophobic surfaces with molecular combing. Fluorescence polarization showed that the polymer chains are preferentially aligned along the fibrils. Furthermore the degree of polarization was varying, but for all probed fibril like objects it was high (>0.7) for polarization in both excitation and emission. In some cases the polarization degree was higher than 0.9 signifying very high orientation of the polymer chains along the fibrils. The corresponding I_max/I_min values for these decorated fibrils were 10.5 (for emission) on average, with extreme values up to 38, which is similar to values found for highly oriented polymer chains in gel films. The polarization degree was slightly higher for emission than excitation, possibly indicating that the emission occurred from dipoles of higher orientation than the absorbing dipoles. No significant rotation of the plane of polarization for excitation and emission was observed for the decorated fibrils. These nanowires with highly oriented CP could demonstrate very interesting anisotropic properties in optical and electrical applications.

Acknowledgements

We thank Per Björk, IFM, for helpful discussions, and Wendimagnegn Mammo and Mats Andersson, Chalmers, for polymer samples. These studies were financially supported by VINNOVA within a Bionanoit project named MEM, by the Science Council (VR), Crafoord foundation, Kungliga Fysiografiska Sällskapet in Lund and KAW foundation. Studies of nanostructuring and nanopatterning are also supported by the EC-funded project NaPa (Contract no. NMP4-CT-2003-500120), which is gratefully acknowledged. The content of this work is the sole responsibility of the authors.

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Footnotes

† Electronic supplementary information (ESI) available: Fluorescence micrograph, spectra of APFO-12 decorated fibrils and fluorescence intensity trace. See DOI: 10.1039/b712829k

‡ I_max is with polarizers both in excitation and emission parallel to the polymer chain orientation, whereas I_min is with both polarizers perpendicular to this orientation.

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