, Volume 50, Issue 4, pp 529–540 | Cite as

Light-dependent and light-independent protochlorophyllide oxidoreductases share similar sequence motifs —in silico studies

  • M. Gabruk
  • J. Grzyb
  • J. Kruk
  • B. Mysliwa-Kurdziel


In the present studies, we have found a fragment of amino acid sequence, called TFT motif, both in light-dependent protochlorophyllide oxidoreductase (LPOR) and in the L subunit of dark-operative (light-independent) protochlorophyllide oxidoreductases (DPOR). Amino acid residues of this motif shared similar physicochemical properties in both types of the enzymes. In the present paper, physicochemical properties of amino acid residues of this common motif, its spatial arrangement and a possible physiological role are being discussed. This is the first report when similarity between LPOR and DPOR, phylogenetically unrelated, but functionally redundant enzymes, is described.

Additional key words

chlorophyll biosynthesis homology modeling protochlorophyllide protochlorophyllide oxidoreductase sequence analysis 


BchB, BchL and BchN

B, L and N subunits of bacterial DPOR, respectively


L subunit of plant DPOR




dark-operative (light-independent) protochlorophyllide oxidoreductase


light-dependent protochlorophyllide oxidoreductase


National Center for Biotechnology Information


a protomer of Fe protein of nitrogenase




Protein Data Bank


one of the isoforms of LPOR


short chain dehydrogenases/reductases


Unable to display preview. Download preview PDF.

Unable to display preview. Download preview PDF.


  1. Arnold, K., Bordoli, L., Kopp, J., Schwede, T.: The SWISSMODEL workspace: A web-based environment for protein structure homology modeling. —Bioinformatics 22: 195–201, 2006.PubMedCrossRefGoogle Scholar
  2. Belyaeva, O.B., Litvin, F.F.: Photoactive pigment-enzyme complexes of chlorophyll precursor in plant leaves. —Biochemistry Moscow 72: 1458–1477, 2007.PubMedCrossRefGoogle Scholar
  3. Benach, J., Atrian, S., Ladenstein, R., Gonzàlez-Duarte, R.: Genesis of Drosophila ADH: the shaping of the enzymatic activity from a SDR ancestor. —Chem-Biol Interactions 130–132: 405–415, 2001.CrossRefGoogle Scholar
  4. Bröcker, M.J., Schomburg, S., Heinz, D.W., Jahn, D., Schubert, W.D., Moser, J.: Crystal structure of the nitrogenase-like dark operative protochlorophyllide oxidoreductase catalytic complex (ChlN/ChlB)2. J. Biol. Chem. 285: 27336–27345, 2010a.PubMedCrossRefGoogle Scholar
  5. Bröcker, M.J., Virus, S., Ganskow, S., Heathcote, P., Heinz, D.W., Schubert, W.D., Jahn, D., Moser, J.: ATP-driven reduction by dark-operative protochlorophyllide oxidoreductase from Chlorobium tepidum mechanistically resembles nitrogenase catalysis. —J. Biol. Chem. 283: 10559–10567, 2008.PubMedCrossRefGoogle Scholar
  6. Bröcker, M.J., Wätzlich, D., Saggu, M., Lendzian, F., Moser, J., Jahn, D.: Biosynthesis of (Bacterio)chlorophylls. ATP-dependent transient subunit interaction and electron transfer of dark operative protochlorophyllide oxidoreductase. —J. Biol. Chem. 285: 8268–8277, 2010b.PubMedCrossRefGoogle Scholar
  7. Buhr, F., El Bakkouri, M., Valdez, O., Pollmann, S., Lebedev, N., Reinbothe, S., Reinbothe, C.: Photoprotective role of NADPH:protochlorophyllide oxidoreductase A. —Proc. Nat. Acad. Sci. USA 105: 12629–12634, 2008.PubMedCrossRefGoogle Scholar
  8. Dahlin, C., Aronsson, H., Wilks, H., Lebedev, N., Sundqvist, C., Timko, M.P.: The role of protein surface charge in catalytic activity and chloroplast membrane association of the pea NADPH: protochlorophyllide oxidoreductase (POR) as revealed by alanine scanning mutagenesis. —Plant Mol. Biol. 39: 309–323, 1999.PubMedCrossRefGoogle Scholar
  9. Dietzek, B., Kiefer, W., Popp, J., Hermann, G., Schmitt, M.: Solvent effects on the excited-state processes of protochlorophyllide: A femtosecond time-resolved absorption study. —J. Phys. Chem. 110: 4399–4406, 2006.CrossRefGoogle Scholar
  10. Dietzek, B., Tschierlei, S., Hermann, G., Yartsev, A., Pascher, T., Sundstrom, V., Schmitt, M., Popp, J.: Protochlorophyllide a: a comprehensive photophysical picture. —Chem. Phys. Phys. Chem. 10: 144–150, 2009.Google Scholar
  11. Dietzek, B., Tschierlei, S., Hanf, R., Seidel, S., Yartsev, A., Schmitt, M., Hermann, G., Popp, J.: Dynamics of charge separation in the excited-state chemistry of protochlorophyllide. —Chem. Phys. Lett. 492: 157–163, 2010.CrossRefGoogle Scholar
  12. Filling, C., Berndt, K.D., Benach, J., Knapp, S., Prozorowski, T., Nordling, E., Ladenstein, R., Jornvall, H., Oppermann, U.: Critical residues for structure and catalysis in short-chain dehydrogenases/reductases. —J. Biol. Chem. 277: 25677–25684, 2002.PubMedCrossRefGoogle Scholar
  13. Garnier, J., Gibrat, J.F., Robson, B.: GOR method for predicting protein secondary structure from amino acid sequence. —In: Doolittle, R.F. (ed.): Methods in Enzymology 266: 540–553, 1996.Google Scholar
  14. Heyes, D.J., Hunter, C.N.: Site-directed mutagenesis of Tyr-189 and Lys-193 in NADPH: protochlorophyllide oxidoreductase from Synechocystis. —Biochem. Soc. Trans. 30: 601–604, 2002.PubMedCrossRefGoogle Scholar
  15. Heyes, D.J., Hunter, C.N.: Making light work of enzyme catalysis: protochlorophyllide oxidoreductase. —Trends Plant Sci. 30: 642–649, 2005.Google Scholar
  16. Heyes, D.J., Martin, G.E., Reid, R.J., Hunter, C.N., Wilks, H.M.: NADPH:protochlorophyllide oxidoreductase from Synechocystis: overexpression, purification and preliminary characterisation. —FEBS Lett. 483: 47–51, 2000.PubMedCrossRefGoogle Scholar
  17. Heyes, D.J., Scrutton, N.S.: Conformational changes in the catalytic cycle of protochlorophyllide oxidoreductase: what lessons can be learnt from dihydrofolate reductase? —Biochem. Soc. Trans. 37: 354–357, 2009.PubMedCrossRefGoogle Scholar
  18. Igarashi, R.Y., Seefeldt, L.C.: Nitrogen fixation: the mechanism of the Mo-dependent nitrogenase. —Crit. Rev. Biochem. Mol. Biol. 38: 351–381, 2003.PubMedCrossRefGoogle Scholar
  19. John, B., Šali, A.: Comparative protein structure modeling by iterative alignment, model building and model assessment. —Nucl. Acids Res. 31: 3982–3992, 2003.PubMedCrossRefGoogle Scholar
  20. Lebedev, N., Karginova, O., McIvor, W., Timko, M.P.: Tyr275 and Lys279 stabilize NADPH within the catalytic site of NADPH:protochlorophyllide oxidoreductase and are involved in the formation of the enzyme photoactive state. —Biochemistry 40: 12562–12574, 2001.PubMedCrossRefGoogle Scholar
  21. Martin, G.E., Timko, M.P., Wilks, H.M.: Purification and kinetic analysis of pea (Pisum sativum L.) NADPH:protochlorophyllide oxidoreductase expressed as a fusion with maltose-binding protein in Escherichia coli. —Biochem. J. 325: 139–145, 1997.PubMedGoogle Scholar
  22. Masuda, T.: Recent overview of the Mg branch of the tetrapyrrole biosynthesis leading to chlorophylls. —Photosynth. Res. 96: 121–143, 2008.PubMedCrossRefGoogle Scholar
  23. Meiler, J., Baker, D.: Coupled prediction of protein secondary and tertiary structure. —Proc. Nat. Acad. Sci. USA 100: 12105–12110, 2003.PubMedCrossRefGoogle Scholar
  24. Muraki, N., Nomata, J., Ebata, K., Mizoguchi, T., Shiba, T., Tamiaki, H., Kurisu, G., Fujita, Y.: X-ray crystal structure of the light-independent protochlorophyllide reductase. —Nature 465: 110–114, 2010.PubMedCrossRefGoogle Scholar
  25. Mysliwa-Kurdziel, B., Franck, F., Strzałka, K.: Analysis of fluorescence lifetime of protochlorophyllide and chlorophyllide in isolated etioplast membranes measured from multifrequency cross-correlation phase fluorometry. —Photochem. Photobiol. 70: 616–623, 1999.Google Scholar
  26. Mysliwa-Kurdziel, B., Kruk, J., Strzałka, K.: Fluorescence lifetimes and spectral properties of protochlorophyllide in organic solvents and their relations to the respective parameters in vivo. —Photochem. Photobiol. 79: 62–67, 2004.PubMedGoogle Scholar
  27. Mysliwa-Kurdziel, B., Solymosi, K., Kruk, J., Böddi, B., Strzałka, K.: Solvent effects on fluorescence properties of protochlorophyll and its derivatives with various porphyrin side chains. —Eur. Biophys. J. 37: 1185–1193, 2008.PubMedCrossRefGoogle Scholar
  28. Nomata, J., Ogawa, T., Kitashima, M., Inoue, K., Fujita, Y.: NB-protein (BchN-BchB) of dark-operative protochlorophyllide reductase is the catalytic component containing oxygen-tolerant Fe-S clusters. —FEBS Lett. 582: 1346–1350, 2008.PubMedCrossRefGoogle Scholar
  29. Oppermann, U., Filling, C., Hult, M. et al.: Short-chain dehydrogenases/ reductases (SDR): the 2002 update. —Chem-Biol. Interactions 143/144: 247–253, 2003.CrossRefGoogle Scholar
  30. Ouazzani-Chahdi, M.A., Schoefs, B., Franck, F.: Isolation and characterization of photoactive complexes of NADPH:protochlorophyllide oxidoreductase from wheat. —Planta 206: 673–680, 1998.CrossRefGoogle Scholar
  31. Pruitt, K.D., Tatusova, T., Maglott, D.R.: NCBI reference sequences (RefSeq): a curated non-redundant sequence database of genomes, transcripts and proteins. —Nucl. Acids Res. 35: D61–D65, 2006.PubMedCrossRefGoogle Scholar
  32. Reinbothe, C., El Bakkouri, M., Buhr, F., Muraki, N., Nomata, J., Kurisu, G., Fujita, Y., Reinbothe, S.: Chlorophyll biosynthesis: spotlight on protochlorophyllide reduction. —Trends Plant Sci. 15: 614–624, 2010.PubMedCrossRefGoogle Scholar
  33. Sarma, R., Barney, B.M., Hamilton, T.L., Jones, A., Seefeldt, L.C., Peters, J.W.: Crystal structure of the L protein of Rhodobacter sphaeroides light-independent protochlorophyllide reductase with MgADP bound: a homologue of the nitrogenase Fe protein. —Biochemistry 47: 13004–13015, 2008.PubMedCrossRefGoogle Scholar
  34. Schoefs, B.: Protochlorophyllide reduction —what is new in 2005? —Photosynthetica 43: 329–343, 2005.CrossRefGoogle Scholar
  35. Schoefs, B., Franck, F.: Protochlorophyllide reduction: mechanisms and evolution. —Photochem. Photobiol. 78: 543–557, 2003.PubMedCrossRefGoogle Scholar
  36. Solymosi, K., Schoefs, B.: Etioplast and etio-chloroplast formation under natural conditions:the dark side of chlorophyll biosynthesis in angiosperms. —Photosynth. Res. 105: 143–166, 2010.PubMedCrossRefGoogle Scholar
  37. Solymosi, K., Smeller, L., Böddi, B., Fidy, J.: Activation volumes of processes linked to the phototransformation of protochlorophyllide determined by fluorescence spectroscopy at high pressure. —Biochim. Biophys. Acta 1554: 1–4, 2002.PubMedCrossRefGoogle Scholar
  38. Sytina, O.A., Alexandre, M.T., Heyes, D.J., Hunter, C.N., Robert, B., van Grondelle, R., Groot, M.L.: Enzyme activation and catalysis: characterisation of the vibrational modes of substrate and product in protochlorophyllide oxidoreductase. —Phys. Chem. Chem. Phys. 13: 2307–2313, 2011.PubMedCrossRefGoogle Scholar
  39. Sytina, O.A., Heyes, D.J., Hunter, C.N., Alexandre, M.T., van Stokkum, I.H.M., van Grondelle, R., Groot, M.L.: Conformational changes in an ultrafast light-driven enzyme determine catalytic activity. —Nature 456: 1001–1004, 2008.PubMedCrossRefGoogle Scholar
  40. Sytina, O.A., Heyes, D.J., Hunter, C.N., Groot, M.L:. Ultrafast catalytic processes and conformational changes in the lightdriven enzyme protochlorophyllide oxidoreductase (POR). —Biochem. Soc. Trans. 37: 387–391, 2009.PubMedCrossRefGoogle Scholar
  41. Sytina, O.A., van Stokkum, I.H.M., Heyes, D.J., Hunter, C.N., van Grondelle, R., Groot, M.L.: Protochlorophyllide excitedstate dynamics in organic solvents studied by time-resolved visible and mid-infrared spectroscopy. —J. Phys. Chem. 114: 4335–4344, 2010.CrossRefGoogle Scholar
  42. Thompson, J.D., Higgins, D.G., Gibson, T.J.: CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. —Nucl. Acids Res. 22: 4673–4680, 1994.PubMedCrossRefGoogle Scholar
  43. Townley, H.E., Sessions, R.B., Clarke, A.R., Dafforn, T.R., Griffiths, W.T.: Protochlorophyllide oxidoreductase: a homology model examined by site directed mutagenesis. —Proteins 44: 329–335, 2001.PubMedCrossRefGoogle Scholar
  44. Wätzlich, D., Bröcker, M.J., Uliczka, F., Ribbe, M., Virus, S., Jahn, D., Moser, J.: Chimeric nitrogenase-like enzymes of (bacterio)chlorophyll biosynthesis. —J. Biol. Chem. 284: 15530–15540, 2009.PubMedCrossRefGoogle Scholar
  45. Wiktorsson, B., Ryberg, M., Gough, S., Sundqvist, C.: Isoelectric focusing of pigment-protein complexes solubilized from non-irradiated and irradiated prolamellar bodies. —Physiol. Plant. 82: 659–669, 1992.CrossRefGoogle Scholar
  46. Wilks, H.M., Timko, M.P.: A light-dependent complementation system for analysis of NADPH:protochlorophyllide oxidoreductase: Identification and mutagenesis of two conserved residues that are essential for enzyme activity. —Proc. Nat. Acad. Sci. USA 92: 724–728, 1995.PubMedCrossRefGoogle Scholar
  47. Yang, J., Cheng, Q.: Origin and evolution of the lightdependent protochlorophyllide oxidoreductase (LPOR) genes. —Plant Biol. 6: 537–544, 2004.PubMedCrossRefGoogle Scholar
  48. Zhao, G.J., Han, K.L.: Site-specific solvation of the photoexcited protochlorophyllide a in methanol: formation of the hydrogen-bonded intermediate state induced by hydrogen bond strengthening. —Biophys. J. 94: 38–46, 2008.PubMedCrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media B.V. 2012

Authors and Affiliations

  • M. Gabruk
    • 1
  • J. Grzyb
    • 2
  • J. Kruk
    • 1
  • B. Mysliwa-Kurdziel
    • 1
  1. 1.Department of Plant Physiology and Biochemistry, Faculty of Biochemistry, Biophysics and BiotechnologyJagiellonian UniversityKrakówPoland
  2. 2.Laboratory of Biological PhysicsInstitute of Physics PASWarsawPoland

Personalised recommendations