Charge transfer between photosynthetic proteins and hematite in bio-hybrid photoelectrodes for solar water splitting cells
© Faccio et al.; licensee Springer. 2015
Received: 2 November 2014
Accepted: 7 November 2014
Published: 6 May 2015
Functionalization of the hematite photoanode with the photosynthetic light antenna protein C-phycocyanin (PC) can yield substantial enhancement of the photocurrent density. Photoelectrochemical cells with bio-hybrid electrodes from photosynthetic proteins and inorganic semiconductors have thus potential for the use in artificial photosynthesis. We investigate here processing routes for the functionalization of hematite photoanodes with PC, including in situ co-polymerization of PC with enzymatically-produced melanin, and using a recombinant PC genetically engineered to carry a hexa-histidine tag (αHisPC). First, the effect of the immobilisation of PC on the electrode morphology and photocurrent production is evaluated. Then, the electronic charge transfer in dark and light conditions is assessed with electrochemical impedance spectroscopy and valence band (VB) X-ray photoemission spectroscopy. The relative shift of the VB spectrum towards the Fermi energy EF upon illumination is smaller for the more complex processed coating and virtually disappears for αHisPC immobilised with a melanin film. Optimal conditions for protein immobilisation are determined and the dark currents benefit most from the most advanced protein coating processes.
KeywordsArtificial photosynthesis Antenna protein C-phycocyanin Hematite photoanode Impedance spectroscopy Valence band (VB) X-ray photoemission spectroscopy
Hematite, a ferric oxide with 2.1 eV optical band gap energy, is considered a low cost and environmentally benign photoanode material for solar hydrogen production by water splitting in photoelectrochemical cells (PEC). The research and development of photoelectrodes for PEC is focused on the tailoring of the electronic structure and the microstructure, and recently also on the design of hetero structures. The functionalization of a photoelectrode with light harvesting proteins was recently demonstrated for hematite using C-phycocyanin (PC) , which constitutes basically a dye sensitization.
Dye sensitization  is an established technology for the extension of the light absorption range of inorganic photoelectrodes for solar energy conversion. To be fully functional, the electric charge generated in the dye upon light absorption needs to be transferred to the photoelectrode, i.e. the interface between the dye and the photoelectrode must warrant proper charge transfer for electrons or holes.
We recently developed a process which enhances the photocurrent of hematite thin-film electrodes by immobilising light-harvesting proteins such as PC on the surface . So far, the functionalization has been performed with the commercially available PC from Spirulina sp. which contains the α- and β-subunits of the protein. In the present study, we evaluate the simpler α-subunit of PC from Synechocystis sp. PCC6803 which is recombinantly produced in E. coli and engineered to carry an N-terminal hexa-histidine tag (αHisPC) . The electronic charge transfer is investigated by electrochemical impedance spectroscopy (EIS) and with valence band X-ray photoemission spectroscopy (VB PES).
1,1′-Carbonyldiimidazol (CDI, >97%), 1,4-dioxane (99.8%), 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt (ABTS, >98%), monopotassium phosphate (1 mol l−1), hydrogen dipotassium phosphate (1 mol l−1), potassium hydroxide (KOH, 95%), iron nitrate nonahydrate (>98%), tetrahydrofuran (THF, >99.9%), sodium chloride (NaCl, 5 mol l−1), L-tyrosine (>99%), Tris–HCl (>99%) and C-phycocyanin isolated from Spirulina sp. (99.9%) were purchased from Sigma-Aldrich. Oleic acid (>60%) and L(+)-tartaric acid (>99.5%) were purchased from Fluka. Tyrosinase from Verrucomicrobium spinosum was produced in E. coli and proteolitically activated as described in .
2.2 Production and purification of C-phycocyanin from Synechocystis sp. PCC6803
The α-subunit of PC from Synechocystis sp. PCC6803 was produced in an N-terminally His-tagged form according to  and purified with a modified protocol from . Briefly, strain E. coli DH5α was co-transformed with plasmids pAT101 and pBD414V and transformants were selected for resistance to both kanamycin (50 μg ml−1) and spectinomycin (100 μg ml−1). Cultures were routinely set up in TB medium (500 ml volume), incubated at 37°C under constant shaking (180 rpm), and expression was induced by adding IPTG (final conc. 1 mM). Cells were harvested by centrifugation 17 h post-induction and stored at −20°C. Upon thawing and re-suspension in 100 mM potassium phosphate buffer with pH 7.5 (5 ml g−1 wet cell weight), lysozyme (final conc. 1 mg ml−1) and a protease inhibitor mix (Complete Protease Inhibitor Cocktail Tablets, Roche) were added. The cell suspension was then subjected to a freeze-thaw cycle at −20°C. Cells were further disrupted by sonication, and the cell-free extract, isolated by ultracentrifugation, was loaded onto a HisTrap column (1 ml, GE Healthcare Life Sciences) for purification by immobilized metal ion affinity chromatography (IMAC). αHisPC was eluted with imidazole (conc. 200 mM) in 100 mM potassium phosphate (pH 7.5). αHisPC-containing fractions were concentrated and desalted with a Vivaspin ultrafiltration device (10000 cut-off, Sartorius). The purified αHisPC was stored at −20°C for further use. Protein concentrations of αHisPC solutions were determined spectrophotometrically with a Synergy Mx spectrophotometer (Biotek, Switzerland) and using an extinction coefficient of 140000 M−1 cm−1 at 620 nm. Protein purity was assessed spectrophotometrically as the ratio of the absorbance at 280 nm and at 620 nm.
2.3 Hematite films and single crystals
Hematite films were prepared as described previously . Fluorine doped tin oxide (FTO) coated glass slides (12 mm × 30 mm × 2 mm, TEC-8 from Hartford Glass Inc.) were automatically dip coated (DipMasterTM-50, Chemat Technology Inc., USA) with the precursor solution and dried at 75°C for 5 min on a hot plate. The samples were then further annealed for 30 min at 500°C in an air vented muffle furnace. Dip coating and annealing were repeated three times so as to obtain a film of four layers of hematite, which has typically a thickness of 500 to 600 nm. Prevalence of the hematite phase was confirmed with X-ray diffraction (Siemens D5000).
The thickness of the hematite films was determined with a mechanical profilometer (XP-1 Stylus Profiler by Ambios Technology). The morphology of the pristine and protein coated hematite films was investigated with a Philips XL 30 scanning electron microscope (SEM, Oxford instruments).
Hematite single crystals with (0001) orientation were purchased from SurfaceNet GmbH, Rheine, Germany. They had a size of 5 mm × 5 mm × 1 mm size and were Epi surface polished.
2.4 Coating of hematite with proteins and melanin
Similarly to , the hematite thin films were first dip-coated with a 0.001% agarose solution, let dry, and second with ~100 μl cm−2 of a 30 mg ml−1 CDI solution prepared in dioxan. Protein immobilisation was carried out by coating the hematite surface with a melanin-forming mixture containing PC or αHisPC (concentration 0.2 mg ml−1), L-tyrosine (concentration 0.1 mg ml−1), and tyrosinase (50 μg ml−1 concentration) in 0.05 M phosphate buffer saline (PBS; 160 g l−1 NaCl, 4 g l−1 KCl, 28.88 gl−1 Na2HPO4, 4.4 g l−1 KH2PO4, pH 7.2-7.4) for 3 h. The surface was dried between all the steps, eventually rinsed with PBS and distilled water, and then stored at 4°C in the dark until the actual measurements were performed. Because the tyrosinase preparation contains traces of trypsin, the protease inhibitor phenylmethanesulfonyl fluoride (PMSF) was added to the coating mixture at a 1 mM final concentration. The thickness of the films treated with melanin alone, or with melanin plus PC or with melanin plus αHisPC was below 1000 nm. All samples were prepared in either duplicate or triplicate and average values are reported throughout this paper.
In addition to the aforementioned thin hematite films on FTO-coated glass slides, three (0001) hematite single crystals were coated with:
1) Melanin plus αHisPC: 50 μl solution containing αHisPC, L-tyrosine and tyrosinase in PBS (concentrations as above) after pre-treatment of the hematite single crystal with agarose and CDI solution (as described above).
2) PC only: 50 μl solution containing PC, after pre-treatment of the hematite single crystal with agarose and CDI solution (as described above).
3) Melanin plus PC: 50 μl solution containing PC and the melanin-forming mixture (as described above), after pre-treatment of the hematite single crystal with agarose and CDI solution (as described above).
2.5 Photoelectrochemical assessment of the films
For the assessment of the photoelectrochemical properties of the photoanode, we largely followed the recommendations provided in , when not otherwise stated in this manuscript. Linear voltammetry in the dark and under illumination was conducted using a specifically designed spectro-photoelectrochemical cell (known as EPFL cappuccino cell) made of polyether ether ketone (PEEK) and a potentiostat (Voltalab80 PGZ 402). The hematite photoanode was connected as the working electrode in a three electrode configuration. A platinum sheet of 5 mm × 5 mm was used as counter electrode and an Ag/AgCl (with saturated KCl) electrode was used as reference electrode. The electrolyte was 0.05 M PBS. Sunlight was simulated by a 1 Sun Oriel Lamp (L.O.T.–Oriel AG) with a quartz filter, corresponding to AM 1.5 global standard solar spectrum. The bias potential was scanned from 0 mV to 1000 mV. All shown results are the arithmetic average of data of at least three consecutive measurements. Electrochemical impedance spectra were recorded with the Voltalab80 PGZ 402 using the same cell in the frequency range from 100 kHz to 100 mHz with 20 data points per decade and a perturbation amplitude of 10 mV. Spectra of hematite films coated on FTO glass and of variously prepared PC films on hematite were recorded. Dark and light conditions as described above were applied during the impedance measurements.
2.6 Photoemission spectroscopy
X-ray photoemission (PES) spectra were recorded at beamline 8–1 at Stanford Synchrotron Radiation Lightsource, Menlo Park, CA. Specifically, the spectra were recorded under Fe 2p resonant condition with 53.2, 54.4 and 55.8 eV photon energy, so that the valence band spectra were particularly representing the Fe relevant features. The PES spectra were measured from the surface finished (0001) hematite single crystals that were coated with PC in the aforementioned different processing routes. The measurements of PES were carried out under ultrahigh vacuum condition, i.e. 10−10 mbar base pressure. At the time of experiment, the films could be considered dry by ambient conditions. A 1.5 AM solar simulator (HAL-302 Solar Simulator, 350–750 nm, Asahi Spectra, Japan) was used to illuminate the samples during the PES experiments.
3 Results and Discussion
We have recently observed that the photocurrent density of a hematite electrode increases when the light-harvesting protein phycocyanin (PC) is copolymerized with melanin on its surface. In the present work we investigate the influence of PC concentration, the time of incubation with the melanin-forming mixture on the photocurrent. In addition to the commercially available PC preparation, we test the effect of a recombinantly produced form of PC on the performance of the photoelectrochemical cell (PEC). Each sample is measured prior to and after the immobilization of proteins under pH-neutral conditions in PBS, if not mentioned otherwise.
3.1 Production of recombinant αHisPC
Glazer et al. reported that the peak of maximum absorbance of C-phycocyanin when composed of both α- and β-subunits results from the sum of the absorbance peaks of both subunits. The α-subunit absorbs at a longer wavelength than the β-subunit, e.g. the α-subunit has an absorbance maximum at 620 nm and the β-subunit at 608 nm in C-phycocyanin from Synechococcus sp. .
Similarly to what has been reported for PC , incubation under alkaline conditions, e.g. in the presence of 1 M KOH, absorbance in the 600–620 nm region is rapidly annulled and a novel absorbance peak at 750 nm is detected (Figure 1).
3.2 Effect of protein-melanin coating on the photocurrent
In order to establish optimal parameters for the coating process, three protein concentrations spanning two orders of magnitude (0.02, 0.2 and 2 mg ml−1) and three reaction times (1.5, 3 and 4.5 h) are tested. The highest enhancement in photocurrent density was achieved at concentrations between 0.2 mg ml−1 to 2 mg ml−1 of PC and a reaction time of 3 h to 4.5 h. For hematite thin films coated with PC, the highest relative enhancement in photocurrent density is 23% and is obtained at a concentration of 2 mg ml−1 and a reaction time of 3 h, when measured in KOH. Under identical conditions, a higher value is obtained with samples coated with αHisPC at a concentration of 0.2 mg ml−1 that give a 55% increase in photocurrent density.
We expect that the extreme pH conditions with the use of 1 M KOH electrolyte (~ pH 13–14) leads to melanin dissolution and structural changes in PC causing denaturation. Yet we measure in KOH a significant increase in photocurrent density (left panel, Figure 2a). A possible explanation might be found in the newly occurring absorbance feature at 750 nm for PC and αHisPC, when measured in the strongly alkaline environment (Figure 1). While the proteins may become denatured, the chromophores may still be intact and absorb light  and produce electron–hole pairs which add to the photocurrent. Moreover, the recombinant monomeric αHisPC appears to be a valid alternative to the commercial PC preparation. The 55% increase measured for αHisPC is significantly higher than the 8% increase measured for analogous samples prepared with PC (all measured in 1 M KOH). We thus speculate that PC and αHisPC couple differently with the surface, thus conferring different charge transfer. As a possible explanation, the presence of the Histag on αHisPC might provide a better conducting bonding to the hematite surface as histidine residues are known to coordinate iron ions and the Histag has been shown to bind iron in some proteins . For the evaluation of the effect of melanin alone, samples lacking any photosynthetic protein were prepared and analysed as a control experiment. Compared to the hematite films with coatings containing PC or αHisPC and melanin, no significant increase in photocurrent was observed. Therefore the key factor for the increase in photocurrent is the biliprotein, and not the melanin. Based on these results, a 0.2 mg ml−1 protein concentration and 3 h enzymatic reaction time was used for further experiments.
3.3 Surface morphology of phycocyanin-melanin coated samples
3.4 Charge transfer in PC-melanin coated hematite films
Moreover, upon illumination of the protein film, the impedance decreases dramatically (see the very small box in the lower left corner in the impedance spectrum in 7b) to far below 1 kOhm. Here it is sufficient to plot the impedance in an isometric window of 300 Ohm to present virtually the entire impedance spectra when measured under light condition. The lowest impedance is found for 800 mV bias. The curvature of the spectra in the inset of Figure 7b) shows the clear tendency towards semicircles as the direct current (DC) bias approaches the water splitting potential. We interpret this as increased charge transfer of the system during illumination and increased DC bias. The protein coating improves the conductivity and charge transfer between the hematite and the electrolyte also under illumination significantly. The impedance for pristine and protein coated hematite increases again in the transition from 800 mV to 1000 mV. This is, where typically considerable oxygen evolution by water oxidation takes place.
Charge transfer across interfaces depends on the electronic structure of these interfaces. A method for the assessment of the electronic structure of surfaces and interfaces is X-ray and photoelectron spectroscopy (XPS) and photoemission spectroscopy (PES). In Brizzolara et al.  the conclusion about covalent attachment of the proton pump protein bacteriorhodopsin via genetic substitution of cysteine for serine (S35C) was based partially on the XPS core level spectra of sulfur, i.e. via detection of the chemical shift of the sulphur core level spectrum. An early valence band (VB) XPS study on a protein (D-luciferin) is presented in Wada et al. , where XPS helped sketch a model for the luminescence. In this study, we find that the position of the VB spectrum changes depending on the type of phycocyanin coating process applied to hematite.
We observe that, under dark condition (8a), the intensity maximum of the spectrum from the PC film shifts from 12 eV to just below 11.5 eV towards the Fermi level, when PC was co-polymerized with melanin (Figure 8a). With αHisPC + Mel on hematite, the intensity maximum shifts to 10.3 eV. A similar trend is shown in the case of illumination with the solar simulator (Figure 8b). We notice that with increasing improvement of film processing technology, the maxima of the VB spectra move closer to the Fermi level EF, reflecting the changes in the electronic structure with increased DOS near EF. The charge transfer between hematite and phycocyanin might be improved in αHisPC as histidines are known to coordinate to iron ions. At this time it is not clear whether the spectral shift towards the EF is a result of hole doping originating from the PC + Mel in the films. In addition, the His-tag presumably improves entrapment of the His-tagged protein into the melanin network, as the formation of histidine-tyrosine bonds have been observed upon tyrosinase-catalysed crosslinking .
It is clear, however, that there is a correlation between improved electric transport, increased photocurrent and increased DOS in the VB near EF. Although literature on the electronic structure of the interfaces of biological macromolecules and inorganic materials is scarce, some pioneering studies have been published. A PES and NEXAFS study on the electronic structure of an ultrathin film of the surface layer of Bacillus sphaericus deposited on Si wafers showed how the highest occupied and lowest unoccupied molecular orbitals (HOMO and LUMO) constitute the band gap of the assembly . In an extension of that study, these authors showed with resonant PES how charge transport evolved from torsion effects in the protein .
We want to turn to another observation in our PES spectra. When subjecting the protein-hematite assembly to 1.5 AM simulated solar light, the spectral weight of the intensity range near the O2p bonding peak decreases and shifts by up to 0.7 eV toward the Fermi level (0 eV binding energy).
We interpret this shift as the hole-doped DOS resultant from the exposure of visible light. With the solar lighting on, the electron–hole pairs can be created in the conduction band (CB) and the holes stays near the surface generating a p-type DOS. A similar trend of binding energy shift is observed when silicon is doped . Our PES studies on proteins are in principal not different from studies on synthetic organic molecules and polymers which are frequently used in solar cells and light emitting diodes. A comprehensive review on such VB spectroscopy studies is presented by Koch et al. .
When we turn to the PC + Mel coated hematite (Figure 9b), the VB position shifts from 5.6 eV to 5 eV, i.e. the shift is only 0.6 eV upon illumination. In the case of the HisPC coating (Figure 9c), there is no shift of the VB spectrum when the light is switched on. Interestingly, for all films, the position of the VB in the illumination condition is 5 eV. Hence, it appears that light has no influence on the charge transfer as far as the interface of the protein film with the hematite is concerned.
In addition to the electric charge transfer, which is the scope of this paper, fluorescence resonance energy transfer (FRET) could hypothetically occur between phycocyanin and hematite and thus increase the photocurrent density. The efficiency of the FRET mechanism is inversely proportional to the sixth power of the distance between light donor and light acceptor, making it extremely sensitive to distances and virtually negligible for small distances. Due to the architectures of our bio-hybrid electrodes, where agarose, CDI and His tag virtually constitute “spacers” between PC and hematite, it is difficult to perceive that FRET could occur across these spacers.
The immobilization of photosynthetic proteins on hematite, in particular C-phycocyanin, yields a substantial increase of the photocurrent density which can be assigned to electron hole pair formation in the protein by light absorption in a wavelength range that exceeds the absorption range of the hematite. Refined immobilization strategies like co-polymerization of PC with melanin or genetic engineering to attach a His-Tag to PC successively increase the photocurrent. Analysis of the charge carrier dynamics of the bio-hybrid electrode assembly under photoelectrochemical conditions shows that upon illumination the enhanced charge transfer and DC bias are substantially amplified (factor 10) when the hematite film is coated with the light harvesting protein. This, however, seems to have not the same strong effect on the photocurrent density (only factor 2). A clear and systematic dependency of spectral shift of the valence band is observed in the dark and in the illuminated condition. While the His-tag strategy clearly improves the charge transfer between PC and hematite in the dark, no further enhancement is observed under illumination condition. This might be due to the fact that the illumination generates electron hole pairs, but this should have no effect on the charge transport characteristic of the covalent linkages between protein and hematite.
This study was funded at large by the VELUX Foundation under project no° 790 (Biomimetic photoelectrochemical cells for solar hydrogen generation: Bio-PEC) for K. G.-S. and G.F. G.F. was also financially supported by the TYROMAT project (COFUND action between Empa Postdocs programme and FP7: People) and COST TD1102 Action Project by SBFI C13.003. Financial support by the Strategic Korean-Swiss Cooperative Program in Science and Technology within the project “Spectroscopy on photoelectrochemical electrode materials: SOPEM” for A.B. and NRF-2013K1A3A1A14055158 for B.S.M. is gratefully acknowledged. Financial support by the Swiss National Science Foundation for F.B. (project # 200021–137868 Reaction–diffusion processes for the growth of patterned structures and architectures: bottom up approach for photoelectrochemical electrodes) and for Y.H. (project # 200021–132126 Defects in the bulk and on surfaces and interfaces of metal oxides with photoelectrochemical properties: In-situ photo-electro-chemical and resonant x-ray and electron spectroscopy studies) and R’Equip # 121306 is gratefully acknowledged.
Portions of this research were carried out at the Stanford Synchrotron Radiation Lightsource, a national user facility operated by Stanford University on behalf of the U.S. Department of Energy, Office of Basic Energy Sciences. The solar simulator was provided by Jinghua Guo, Advanced Light Source in Berkeley. Florian Häussler (Empa Dübendorf) assisted with the sample preparation at Empa. We are grateful to Prof. Wendy Schluchter from the Department of Biological Sciences of the University of New Orleans for providing the plasmids for the production of αHisPC in this study.
- Bora DK, Braun A, Constable EC: Energy Environ. Sci. 2013, 6(2):407–425.Google Scholar
- Anderson S, Constable EC, Dare Edwards MP, Goodenough JB, Hamnett A, Seddon KR, Wright RD: Nature 1979, 280(5723):571–573.View ArticleGoogle Scholar
- Tooley AJ, Cai YP, Glazer AN: Proc. Natl. Acad. Sci. 2001, 98(19):10560–10565.View ArticleGoogle Scholar
- Fairhead M, Thoeny-Meyer L: New Biotechnol 2012, 29(2):183–191.View ArticleGoogle Scholar
- Taurino I, Reiss R, Richter M, Fairhead M, Thoeny-Meyer L, De Micheli G, Carrara S: Electrochim. Acta. 2013, 93:72–79.View ArticleGoogle Scholar
- Hu Y, Bora DK, Boudoire F, Haeussler F, Graetzel M, Constable EC, Braun A: J. Renew. Sust. Ener. 2013, 5(4):043109.Google Scholar
- Chen Z, Jaramillo TF, Deutsch TG, Kleiman-Shwarsctein A, Forman AJ, Gaillard N, Garland R, Takanabe K, Heske C, Sunkara M, McFarland EW, Domen K, Miller EL, Turner JA, Dinh HN: J. Mater. Res. 2010, 25(1):3–16.View ArticleGoogle Scholar
- Ihssen J, Braun A, Faccio G, Gajda-Schrantz K, Thöny-Meyer L: Light harvesting proteins for solar fuel generation in bioengineered photoelectrochemical cells. Curr. Protein. Pept. Sci. 2014, 15: in press.Google Scholar
- Glazer AN, Fang S, Brown DM: J. Biol. Chem. 1973, 248:5679–5685.Google Scholar
- Berns DS, MacColl R: Chem. Rev. 1989, 89:807–825.View ArticleGoogle Scholar
- The Recombinant Protein Handbook: Protein amplification and simple purification. Amersham Biosciences 2002, 41–58. 18-1142-75.Google Scholar
- Watt AA, Bothma JP, Meredith P: Soft Matter 2009, 5(19):3754–3760.View ArticleGoogle Scholar
- Costa TG, Younger R, Poe C, Farmer PJ, Szpoganicz B: Bioinorg. Chem. Appl. 2012, 2012:712840.View ArticleGoogle Scholar
- Bridelli MG: Biophys. Chem. 1998, 73(3):227–239.View ArticleGoogle Scholar
- Frolov L, Rosenwaks Y, Richter S, Carmeli C, Carmeli I, Phys J: Chem. C. 2008, 112:13426–13430.Google Scholar
- Chi Q, Farver O, Ulstrup J: PNAS 2005, 102(45):16203–16208.View ArticleGoogle Scholar
- Deng S, Jian G, Lei J, Hu Z, Ju H: Biosens Bioelectron 2009, 25:373–377.View ArticleGoogle Scholar
- Omanovic S, Roscoe SG: Langmuir 1999, 15(23):8315–8321.View ArticleGoogle Scholar
- Owino IOK, Sadik OA: Electroanalysis 2005, 17(23):2101–2113.View ArticleGoogle Scholar
- Dheilly A, Linossier I, Darchen A, Hadjiev D, Corbel C, Alonso V: Appl. Microbiol. Biotechnol. 2008, 79(1):157–164.Google Scholar
- Bora DK, Rozhkova AA, Schrantz K, Wyss PP, Braun A, Graule T, Constable EC: Adv. Funct. Mater. 2012, 22(3):490–502.View ArticleGoogle Scholar
- Ben-Yoav H, Freeman A, Sternheim M, Shacham-Diamand Y: Electrochim. Acta. 2011, 56(23):7780–7786.View ArticleGoogle Scholar
- Zhang S, Xie J, Zhang J, Zhao J, Jiang L: Biochim. Biophys. Acta. 1999, 1426(1):205–211.Google Scholar
- Foote CS: Photochem. Photobiol. 1991, 54(5):659.Google Scholar
- Brizzolara RA, Boyd JL, Tate AE, Vac J: Sci. Technol. A. 1997, 15(3):773–778.Google Scholar
- Wada N, Shitaba R, Shibazaki M, Suzuki N: Photochem. Photobiol. 1989, 49(4):513–518.View ArticleGoogle Scholar
- Hellman M, Mattinen M-L, Fu B, Buchert J, Permi P: J. Biotechnol. 2011, 151(1):143–150.View ArticleGoogle Scholar
- Vyalikh DV, Danzenbacher S, Mertig M, Kirchner A, Pompe W, Dedkov YS: Phys. Rev. Lett. 2004, 93(23):238103.View ArticleGoogle Scholar
- Vyalikh DV, Maslyuk VV, Blueher A, Kade A, Kummer K, Dedkov YS: Phys. Rev. Lett. 2009, 102(9):098101.View ArticleGoogle Scholar
- Sezen H, Suzer S: J. Chem. Phys. 2011, 135:141102.View ArticleGoogle Scholar
- Koch N: phys. stat. sol. RRL. 2012, 6(7):277–293.Google Scholar
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