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Research on Chemical Intermediates

, Volume 28, Issue 7–9, pp 795–815 | Cite as

Photoinduced charge transfer in helical polypeptides

  • Valentine I. Vullev
  • Guilford Jones
Article

Abstract

Investigation of electron transfer in synthetic polypeptides provides an important probe of how charge entrainment is mediated in redox-active proteins, including photosynthetic reaction centers. Interest in this field has focused increasingly on experimental probes of photoinduced electron transfer kinetics and thermodynamics, and the influence of various features of polypeptide templates (e.g. the hydrogen bonding network, the permanent dipole moment for α-helices) that assemble redox groups for long range charge transfer. A review of the various approaches is presented here with attention to heliogenic peptides and the mediation of photoinduced charge entrainment.

Keywords

Polypeptide Electron Transfer Charge Transfer Dipole Moment Long Range 
These keywords were added by machine and not by the authors. This process is experimental and the keywords may be updated as the learning algorithm improves.

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REFERENCES

  1. 1.
    F. Rabanal, B. R. Gibney, W. F. DeGrado, C. C. Moser and P. L. Dutton, Engineering photosynthesis: synthetic redox proteins, Inorg. Chim. Acta 243, 213-218 (1996).Google Scholar
  2. 2.
    M. W. Mutz, J. F. Wishart and G. L. Mclendon, Electron transfer kinetics of bifunctional redox protein maquettes, Adv. Chem. Ser. 254, 145-159 (1998).Google Scholar
  3. 3.
    M. Y. Ogawa, Electron transfer within synthetic polypeptides and de novo designed proteins, Mol. Supramol. Photochem. 4, 113-150 (1999).Google Scholar
  4. 4.
    T. J. Meyer, Chemical Approaches to artificial photosynthesis, Acc. Chem. Res. 22, 163-170 (1989).Google Scholar
  5. 5.
    M. R. Wasielevski, Photoinduced electron transfer in supramolecular systems for arti. cial photosynthesis, Chem. Rev. 92, 435-461 (1992).Google Scholar
  6. 6.
    D. Gust, T. A. Moore, and A. L. Moore, Molecular mimicry of photosynthetic energy and electron transfer, Acc. Chem. Res. 26, 198-205 (1983).Google Scholar
  7. 7.
    R. B. Hill, D. P. Raleigh, A. Lombardi and W. F. DeGrado, De novo design of helical bundles as models for understanding protein folding and function, Acc. Chem. Res. 33, 745-754 (2000).Google Scholar
  8. 8.
    W. F. DeGrado, Protein design: Proteins from scratch, Science 278, 80-81 (1997).Google Scholar
  9. 9.
    M. D. Struthers, R. P. Cheng and B. Imperiali, Design of a monomeric 23-residue polypeptide with de. ned tertiary structure, Science 271, 342-345 (1996).Google Scholar
  10. 10.
    N. E. Zhou, C. M. Kay, B. D. Sykes and R. S. Hodges, A single-strandedamphipathic α-helix in aqueous solution: Design, structural characterization, and its application for determining α-helical propensities of amino acids, Biochemistry 32, 6190-6197 (1993).Google Scholar
  11. 11.
    D. L. Minor, Jr. and P. S. Kim, Context-dependent secondary structure formation of a designed protein sequence, Nature 380, 730-734 (1996).Google Scholar
  12. 12.
    M. A. Case and G. L. McLendon, A virtual library approach to investigate protein folding and internal packing, J. Am. Chem. Soc. 122, 8089-8090 (2000).Google Scholar
  13. 13.
    J. P. Allen, and J. C. Williams, Photosynthetic reaction centers, FEBS Lett. 438, 5-9 (1998).Google Scholar
  14. 14.
    J. Deisenhofer, R. Huber and H. Michel, in: Prediction of Protein Structure and the Principles of Protein Conformation, G. D. Fasman (Ed.), pp. 99-116. Plenum Press, New York, NY (1989).Google Scholar
  15. 15.
    J. Deisenhofer, O. Epp, K. Miki, R. Huber and H. Michel, Structure of the protein subunits in the photosynthetic reaction center of Rhodopseudomonas viridis at 3 Å resolution, Nature 318, 618-624 (1986).Google Scholar
  16. 16.
    J. Deisenhofer, O. Epp, K. Miki, R. Huber and H. Michel, X-ray structure analysis of a membrane protein complex. Electron density map at 3 Å resolution and a model of the chromophores of the photosynthetic reaction center from Rhodopseudomonas viridis, J. Mol. Biol. 180, 385-398 (1984).Google Scholar
  17. 17.
    P. Jordan, P. Fromme, H. T. Witt, O. Klukas, W. Saenger and N. Krauß, Three-dimensional structure of cyanobacterial photosystem I at 2.5 Å resolution, Nature 411, 909-917 (2001).Google Scholar
  18. 18.
    K.-H. Rhee, E. P. Morris, J. Barber and W. Kühlbrandt, Three-dimensional structureof the plant photosystem II reaction center at 8 Å resolution, Nature 396, 283 (1998).Google Scholar
  19. 19.
    P. K. Fyfe and M. R. Jones, Re-emerging structures: continuing crystallographyof the bacterial reaction center, Biochim. Biophys. Acta 1459, 413-421 (2000).Google Scholar
  20. 20.
    J. Deisenhofer and H. Michel, Structures of bacterial photosynthetic reaction centers, Annu. Rev. Cell Biol. 7, 1-23 (1991).Google Scholar
  21. 21.
    B. A. Diner, P. J. Nixon and J. W. Farchaus, Site-directed mutagenesis of photosynthetic reaction centers, Curr. Opin. Struct. Biol. 1, 546-554 (1991).Google Scholar
  22. 22.
    B. Heller, D. Holten and C. Kirmaier, Control of electron transfer between the L-and M-sides of photosynthetic reaction centers, Science 269, 940-945 (1995).Google Scholar
  23. 23.
    A. N. Webber, H. Su, S. E. Bingham, H. Käss, L. Krabben, M. Kuhn, R. Jordan, E. Schlodder and W. Lubitz, Site-directedmutations affecting the spectroscopic characteristics and midpoint potential of the primary donor in photosystem I, Biochemistry 35, 12857-12863 (1996).Google Scholar
  24. 24.
    V. Nagarajan, W. Parson, D. Davis and C. C. Schenck, Kinetics and free energy gaps of electrontransfer reactions in Rhodobacter sphaeroides reaction centers, Biochemistry 3212324-12326 (1993)Google Scholar
  25. 25.
    U. Finkele, C. Lauterwasser, W. Zinth, K. A. Gray and D. Oesterhelt, Role of tyrosine M210 in the initial charge separation of reaction centers of Rhodobacter sphaeroides, Biochemistry 29, 8517-8521 (1990).Google Scholar
  26. 26.
    B. Heller, D. Holten and C. Kirmaier, Characterization of bacterial reaction centers having mutations of aromatic residues in the binding site of the bacteriopheophytin intermediary electron carrier, Biochemistry 34, 5294-5302 (1995).Google Scholar
  27. 27.
    H. A. Murchison, J. P. Alden, J. M. Peloquin, A. K. W. Taguchi, N. W. Woodbury and J. C. Williams, Mutations designed to modify the environment of the primary electron donor of the reaction center from Rhodobacter sphaeroides: Phenylalanine to leucine at L167 and histidine to phenylalanine at L168, Biochemistry 32, 3498-3505 (1993).Google Scholar
  28. 28.
    C. Kirmaier, D. Gaul, R. DeBey, D. Holten and C. C. Schenck, Charge separation in a reaction center incorporating bacteriochlorophyll for photoactive bacteriopheophytin, Science 251, 992-927 (1991).Google Scholar
  29. 29.
    Y. Jia, T. J. DiMagno, C. K. Chan, Z. Wang, M. Du, D. K. Hanson, M. Schiffer, J. R. Norris, G. R. Fleming and M. Popov, Primary charge separation in mutant reaction centers of Rhodobacter capsulatus, J. Phys. Chem. 97, 13180-13191 (1993).Google Scholar
  30. 30.
    T. Nogi and K. Miki, Structural basis of bacterial photosynthetic reaction centers, J. Biochem. 130, 319-329 (2001).Google Scholar
  31. 31.
    A. N. Glazer and A. Melis, Photochemical reaction centers: structure, organization, and function, Annu. Rev. Plant Physiol. 38, 11-45 (1987).Google Scholar
  32. 32.
    R. J. Debus, G. Feher and M. Y. Okamura, Iron-depleted reaction centers from Rhodopseudomonas sphaeroides R-26.1: characterization and reconstitution with iron(2+), manganese(2+), cobalt(2+), nickel(2+), copper(2+), and zinc(2+), Biochemistry 25, 2276-2287 (1986).Google Scholar
  33. 33.
    R. J. Debus, G. Feher and M. Y. Okamura, LM complex of reaction centers from Rhodopseudomonas sphaeroides R-26: characterization and reconstitutionwith the H subunit, Biochemistry 24, 2488-2500 (1985).Google Scholar
  34. 34.
    T. Watanabe, M. Kobayashi, A. Hongu and T. Hiyama, Evidence that a chlorophyll a' dimer constitutes the photochemical reaction center 1 (P700) in photosynthetic apparatus, FEBS Lett. 235, 252-256 (1985).Google Scholar
  35. 35.
    H. Käss, P. Fromme, H. T. Witt and W. Lubitz, Orientation and electronic structure of the primary donor radical cation P700+ . in PhotosystemI; a single crystalEPR and ENDOR study, J. Phys. Chem. B 105, 1225-1239 (2001).Google Scholar
  36. 36.
    M. E. van Brederode and R. van Grondelle, New and unexpected routes for ultrafast electron transfer in photosynthetic reaction centers, FEBS Lett. 455, 1-7 (1999).Google Scholar
  37. 37.
    D. C. Arnett, C. C. Moser, P. L. Dutton and N. F. Scherer, The first events in photosynthesis: Electronic coupling and energy transfer dynamics in the photosynthetic reaction center from Rhodobacter sphaeroides, J. Phys. Chem. B 103, 2014-2032 (1999).Google Scholar
  38. 38.
    H. M. Visser, M. L. Groot, F. van Mourik, I. H. M. van Stokkum, J. P. Dekker and R. van Grondelle, Subpicosecond transient absorption difference spectroscopy on the reaction center of photosystem II: Radical pair formation at 77 K, J. Phys. Chem. 99, 15304-15309 (1995).Google Scholar
  39. 39.
    B. Heller, D. Holten and C. Kirmaier, Control of electron transfer between the L-and M-sides of photosynthetic reaction centers, Science 269, 940-945 (1995).Google Scholar
  40. 40.
    K. Brettel, Electron transfer and arrangement of the redox cofactors in photosystem I., Biochim. Biophys. Acta 1318, 322-373 (1997).Google Scholar
  41. 41.
    M. Guergova-Kuras, B. Boudreaux, A. Joliot, P. Joliot and K. Redding, Evidence for two active branches for electron transfer in photosystem I, Proc. Natl. Acad. Sci. USA 98, 4437-4442 (2001).Google Scholar
  42. 42.
    P. Joliot and A. Joliot, In vivo analysis of the electron transfer within photosystem I: Are the two phylloquinones involved? Biochemistry 38, 11130-11136 (1999).Google Scholar
  43. 43.
    R. A. Marcus and N. Sutin, Electron transfers in chemistry and biology, Biochim. Biophys. Acta 811, 265-322 (1985).Google Scholar
  44. 44.
    C. W. Hoganson and G. T. Tabcock, A metalloradicalmechanism for the generation of oxygen from water in photosynthesis, Science 277, 1953-1956 (1997).Google Scholar
  45. 45.
    D. Gust, T. A. Moore and A. L. Moore, Mimicking photosynthetic solar energy transduction, Acc. Chem. Res. 34, 40-48 (2001).Google Scholar
  46. 46.
    D. Gust, T. A. Moore and A. L. Moore, Mimicking bacterial photosynthesis, Pure Appl. Chem. 70, 2189-2200 (1998).Google Scholar
  47. 47.
    G. Steinberg-Yfrach, P. A. Liddell, S.-C. Hung, A. L. Moore, D. Gust and T. A. Moore, Conversion of light energy to proton potential in liposomes by arti. cial photosynthetic reaction centers, Nature 385, 239-241 (1997).Google Scholar
  48. 48.
    G. Steinberg-Yfrach, J.-L. Rigaud, E. N. Durantini, A. L. Moore, D. Gust and T. A. Moore, Light-driven production at ATP catalyzed by F0F1-ATP synthase in an artificial photosynthetic membrane, Nature 392, 479-482 (1998).Google Scholar
  49. 49.
    W. Parson, Z. T. Chu and A. Warshel, Electrostatic control of charge separation in bacterial photosynthesis, Biochim. Biophys. Acta 1017, 251-272 (1990).Google Scholar
  50. 50.
    M. A. Fox and E. Galoppini, Electric field effects on electron transfer rates in dichromophoric peptides: The effect of helix unfolding, J. Am. Chem. Soc. 119, 5277-5285 (1997).Google Scholar
  51. 51.
    E. Galoppini and M. A. Fox, Effect of the electric field generated by the helix dipole on photoinduced intramolecular electron transfer in dichromophoric α-helical peptides, J. Am. Chem. Soc. 118, 2299-2300 (1996).Google Scholar
  52. 52.
    A. Knorr, E. Galoppini and M. A. Fox, Photoinduced intramolecular electron transfer in dichromophore-appended α-helical peptides: spectroscopic properties and preferred conformations, J. Phys. Org. Chem. 10, 484 (1997).Google Scholar
  53. 53.
    W. G. J. Hol, The role of the α-helix dipole in protein function and structure, Prog. Biophys. Mol. Biol. 45, 149-195 (1985).Google Scholar
  54. 54.
    W. F. DeGrado, C. M. Summa, V. Pavone, F. Nastri and A. Lombardi, De novo design and structural characterization of proteins and metalloproteins, Annu. Rev. Biochem. 68, 779-819 (1999).Google Scholar
  55. 55.
    R. S. Hodges, Boehringer Mannheim Award Lecture 1995/La conference Boehringer Mannheim 1995. De novo design of α-helical proteins: basic research to medical applications, Biochem. Cell Biol. 74, 133-154 (1996).Google Scholar
  56. 56.
    J. W. Bryson, S. F. Betz, H. S. Lu, D. J. Suich, H. X. Zhou, K. T. O'Neil and W. F. DeGrado, Protein design: a hierarchic approach, Science 270, 935-941 (1995).Google Scholar
  57. 57.
    W. F. DeGrado, Z. R. Wasserman and J. D. Lear, Protein design, a minimalist approach, Science 243, 622-628 (1989).Google Scholar
  58. 58.
    P. B. Harbury, T. Zhang, P. S. Kim and T. Alber, A switch between two-, three-, and fourstranded coiled coils in GCN4 leucine zipper mutants, Science 262, 1401-1407 (1993).Google Scholar
  59. 59.
    D. L. Akey, V. N. Malashkevich and P. S. Kim, Buried polar residues in coiled-coil interfaces, Biochemistry 40, 6352-6360 (2001).Google Scholar
  60. 60.
    K. J. Lumb and P. S. Kim, Measurement of interhelical electrostatic interactions in the GCN4 leucine zipper, Science 268, 436-439 (1995).Google Scholar
  61. 61.
    E. K. O'shea, K. J. Lumb and P. S. Kim, Peptide ‘Velcro’: design of a heterodimeric coiled coil, Curr. Biol. 3, 658-667 (1993).Google Scholar
  62. 62.
    D. L. Daugherty and S. H. Gellman, A fluorescence assay for leucine zipper dimerization: Avoiding unintended consequences of fluorophore attachment, J. Am. Chem. Soc. 121, 4325-4333 (1999).Google Scholar
  63. 63.
    G. Jones, II and V. I. Vullev, Contribution of a pyrene fluorescence probe to the aggregation propensity of polypeptides, Org. Lett. 3, 2457-2460 (2001).Google Scholar
  64. 64.
    X. Chen, C. C. Moser, D. L. Pilloud, B. R. Gibney and P. L. Dutton, Engineering oriented heme protein maquette monolayers through surface residue charge distribution patterns, J. Phys. Chem. B 103, 9029-9037 (1999).Google Scholar
  65. 65.
    S. T. R. Walsh, H. Cheng, J. W. Bryson, H. Roder and W. F. DeGrado, Solution structure and dynamics of a de novo designed three-helix bundle protein, Proc. Natl. Acad. Sci. USA 96, 5486-5491 (1999).Google Scholar
  66. 66.
    J. S. Johansson, B. R. Gibney, J. J. Skalicky, A. J. Wand and P. L. Dutton, A native-like three-α-helix bundle protein from structure-basedredesign: A novel maquette scaffold, J. Am. Chem. Soc. 120, 3881-3886 (1998).Google Scholar
  67. 67.
    S. Roy, G. Ratnaswamy, J. A. Boice, R. Fairman, G. McLendon and M. H. Hecht, A protein designed by binary patterning of polar and nonpolar amino acids displays native-like properties, J. Am. Chem. Soc. 119, 5302-5306 (1997).Google Scholar
  68. 68.
    N. J. Greenfield and G. D. Fasman, Computed circular dichroism spectra for the evaluation of protein conformation, Biochemistry 8, 4108-4116 (1969).Google Scholar
  69. 69.
    Y.-H. Chen, J. T. Yang and K. H. Chau, Determination of the helix and β form of proteins in aqueous solution by circular dichroism, Biochemistry 13, 3350-3359 (1974).Google Scholar
  70. 70.
    W. W. Fish, Determination of the molecular weights of membrane proteins and polypeptides, Methods Membr. Biol. 4, 189-276 (1975).Google Scholar
  71. 71.
    L. Hagel and J. C. Janson, Size-exclusion chromatography, J. Chromatogr. Libr. 51A, A267-A307 (1992).Google Scholar
  72. 72.
    D. Rehm and A. Weller, Kinetics of fluorescence quenching by electron and hydrogen-atom transfer, Israel J. Chem. 8, 259-271 (1970).Google Scholar
  73. 73.
    P. Piotrowiak, Photoinduced electron transfer in molecular systems: recent developments, Chem. Soc. Rev. 28, 143-150 (1999).Google Scholar
  74. 74.
    M. Sisido, Elementary photoprocesses in designed chromophore sequence on α-helical polypeptides, Adv. Photochem. 22, 197-228 (1997).Google Scholar
  75. 75.
    M. Sisido, S. Hoshino, H. Kusano, M. Kuragaki, M. Makino, H. Sasaki, T. A. Smith and K. P. Ghiggino, Distance dependence of photoinduced electron transfer along α-helical polypeptides, J. Phys. Chem. B 105, 10407-10415 (2001).Google Scholar
  76. 76.
    B. Pispisa, A. Palleschi, M. Venanzi, G. Zanotti and L. Stella, A spectroscopic and structural investigation on model compounds of biomimetic systems in solution, Photochem. Photobiol. 3, 11-34 (1999).Google Scholar
  77. 77.
    G. Hungerford, F. Donald, D. J. S. Birch and B. D. Moore, Influence of secondary structure on the decay kinetics of fluorescent donor-acceptor labeled peptides, Biosens. Bioelectron. 12, 1183-1190 (1997).Google Scholar
  78. 78.
    B. I. Dahiyat, T. J. Meade and S. L. Mayo, Site-specific modification of α-helical peptideswith electron donors and acceptors, Inorg. Chim. Acta 243, 207-212 (1996).Google Scholar
  79. 79.
    S. S. Isied, I. Moreira, J. F. Wishart, M. Y. Ogawa, A. Vassilian, B. Arbo and J. Sun, New perspectives on long-range electron transfer in conformationally organized peptides and electron-transferproteins: an experimental approach, J. Photochem. Photobiol. A 82, 203-210 (1994).Google Scholar
  80. 80.
    Y. Inai, M. Sisido and Y. Imanishi, Photoinduced electron transfer in a single α-helical polypeptide chain: evidence of a through-space mechanism, J. Phys. Chem. 95, 3847-3851 (1991).Google Scholar
  81. 81.
    C. A. Slate, D. R. Striplin, J. A. Moss, P. Chen, B. W. Erickson and T. J. Meyer, Photochemical energy transduction in helical proline arrays, J. Am. Chem. Soc. 120, 4885-4886 (1998).Google Scholar
  82. 82.
    D. G. McCafferty, D. A. Friesen, E. Danielson, C. G. Wall, M. J. Saderholm, B. W. Erickson and T. J. Meyer, Photochemical energy conversion in a helical oligoproline assembly, Proc. Natl. Acad. Sci. USA 93, 8200-8204 (1996).Google Scholar
  83. 83.
    S. S. Isied, M. Y. Ogawa and J. F. Wishart, Peptide-mediated intramolecular electron transfer: Long-range distance dependence, Chem. Rev. 92, 381-394 (1992).Google Scholar
  84. 84.
    P. M. Cowan and S. McGavin, The structure of poly-L-proline, Nature 176, 501-503 (1955).Google Scholar
  85. 85.
    W. Traub and U. Shmueli, Structure of poly-L-proline I, Nature 198, 1165-1166 (1963).Google Scholar
  86. 86.
    A. R. Downie and A. A. Randall, The mutarotation of poly(L-proline), Trans. Faraday Soc. 55, 2132-2140 (1959).Google Scholar
  87. 87.
    M. W. Mutz, M. A. Case, J. F. Wishart, M. R. Ghadiri and G. L. McLendon, De novo design of protein function: Predictable structure-function relationshipsin synthetic redox protein, J. Am. Chem. Soc. 121, 858-859 (1999).Google Scholar
  88. 88.
    M. W. Mutz, G. L. McLendon, J. F. Wishart, E. R. Gaillard and A. F. Corn, Conformational dependence of electron transfer across de novo designed metalloproteins, Proc. Natl. Acad. Sci. USA 93, 9521-9526 (1996).Google Scholar
  89. 89.
    D. N. Beratan and S. S. Skourtis, Electron transfer mechanisms, Curr. Opin. Chem. Biol. 2, 235-243 (1998).Google Scholar
  90. 90.
    G. V. Kozlov and M. Y. Ogawa, Electron Transfer across a Peptide-Peptide interface within a designed metalloprotein, J. Am. Chem. Soc. 119, 8377-8378 (1997).Google Scholar
  91. 91.
    A. Y. Kornilova, J. F. Wishart, W. Xiao, R. C. Lasey, A. Fedorova, Y.-K. Shin and M. Y. Ogawa, Design and characterization of a synthetic electron-transfer protein, J. Am. Chem. Soc. 122, 7999-8006 (2000).Google Scholar
  92. 92.
    A. Y. Kornilova, J. F. Wishar and M. Y. Ogawa, Effect of surface charges on the rates of intermolecular electron-transfer between de novo designed metalloproteins, Biochemistry 40, 12186-12192 (2001).Google Scholar
  93. 93.
    A. Fedorova and M. Y. Ogawa, Site-specific modification of de novo designed coiled-coil polypeptides with inorganic redox complexes, Bioconjugate Chemistry 13, 150-154 (2002).Google Scholar
  94. 94.
    G. Jones, II, V. I. Vullev, E. Braswell and D. Zhu, Multistep photoinduced electron transfer in a de novo helix bundle. Multimer self-assembly of peptide chains including a chromophore special pair, J. Am. Chem. Soc. 122, 388-389 (2000).Google Scholar
  95. 95.
    G. Jones, II, V. I. Vullev, Ground-and excited-state aggregation properties of a pyrene derivative in aqueous media, J. Phys. Chem. A 105, 6402-6406 (2001).Google Scholar
  96. 96.
    S. Williams, T. P. Causgrove, R. Gilmanshin, K. S. Fang, R. H. Callender, W. H. Woodruff and R. B. Dyer, Fast events in protein folding: helix melting and formation in a small peptide, Biochemistry 35, 691-697 (1996).Google Scholar
  97. 97.
    V. I. Vullev, Towards artificial photosynthesis: photo-induced multiple-step electron transfer in supramolecular structures based on synthetic polypeptides, PhD Dissertation, Boston University, pp. 273-395 (2001).Google Scholar
  98. 98.
    F. Rabanal, W. F. DeGrado and P. L. Dutton, Toward the synthesis of a photosynthetic reaction center maquette: A cofacial porphyrin pair assembled between two subunits of a synthetic four-helix bundle multiheme protein, J. Am. Chem. Soc. 118, 473-474 (1996).Google Scholar
  99. 99.
    D. E. Robertson, R. S. Farid, C. C. Moser, J. F. Urbauer, S. E. Mulholland, R. Pidikiti, J. D. Lear, A. J. Wand, W. F. DeGrado and P. L. Dutton, Design and synthesis of multi-heme proteins, Nature 368, 425-432 (1994).Google Scholar
  100. 100.
    K. S. Akerfeldt, R. M. Kim, D. Camac, J. T. Groves, J. D. Lear and W. F. DeGrado, Tetraphilin: a four-helix proton channel built on a tetraphenylporphyrin framework, J. Am. Chem. Soc. 114, 9656-9657 (1992).Google Scholar
  101. 101.
    S. Kimura, K. Fujita, Y. Miura, G. Xu, Y. Muraji, T. Kidchob and Y. Imanishi, Peptide supramolecules, Curr. Top. Pept. Protein Res. 2, 137-143 (1997).Google Scholar

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© VSP 2002 2002

Authors and Affiliations

  • Valentine I. Vullev
  • Guilford Jones

There are no affiliations available

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