Skip to main content
PMC home page
Biochemical Journal logoLink to Biochemical Journal
. 2002 Mar 1;362(Pt 2):423–432. doi: 10.1042/0264-6021:3620423

Measurement of ferrochelatase activity using a novel assay suggests that plastids are the major site of haem biosynthesis in both photosynthetic and non-photosynthetic cells of pea (Pisum sativum L.).

Johanna E Cornah 1, Jennifer M Roper 1, Davinder Pal Singh 1, Alison G Smith 1
PMCID: PMC1222403  PMID: 11853551

Abstract

Ferrochelatase is the terminal enzyme of haem biosynthesis, catalysing the insertion of ferrous iron into the macrocycle of protoporphyrin IX, the last common intermediate of haem and chlorophyll synthesis. Its activity has been reported in both plastids and mitochondria of higher plants, but the relative amounts of the enzyme in the two organelles are unknown. Ferrochelatase is difficult to assay since ferrous iron requires strict anaerobic conditions to prevent oxidation, and in photosynthetic tissues chlorophyll interferes with the quantification of the product. Accordingly, we developed a sensitive fluorimetric assay for ferrochelatase that employs Co(2+) and deuteroporphyrin in place of the natural substrates, and measures the decrease in deuteroporphyrin fluorescence. A hexane-extraction step to remove chlorophyll is included for green tissue. The assay is linear over a range of chloroplast protein concentrations, with an average specific activity of 0.68 nmol x min(-1) x mg of protein(-1), the highest yet reported. The corresponding value for mitochondria is 0.19 nmol x min(-1) x mg of protein(-1). The enzyme is inhibited by N-methylprotoporphyrin, with an estimated IC(50) value of approximately 1 nM. Using this assay we have quantified ferrochelatase activity in plastids and mitochondria from green pea leaves, etiolated pea leaves and pea roots to determine the relative amounts in the two organelles. We found that, in all three tissues, greater than 90% of the activity was associated with plastids, but ferrochelatase was reproducibly detected in mitochondria, at levels greater than the contaminating plastid marker enzyme, and was latent. Our results indicate that plastids are the major site of haem biosynthesis in higher plant cells, but that mitochondria also have the capacity for haem production.

Full Text

The Full Text of this article is available as a PDF (170.7 KB).

Selected References

These references are in PubMed. This may not be the complete list of references from this article.

  1. Arnon D. I. COPPER ENZYMES IN ISOLATED CHLOROPLASTS. POLYPHENOLOXIDASE IN BETA VULGARIS. Plant Physiol. 1949 Jan;24(1):1–15. doi: 10.1104/pp.24.1.1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Brown S. B., Holroyd J. A., Vernon D. I., Jones O. T. Ferrochelatase activity in the photosynthetic alga Cyanidium caldarium. Development of the enzyme during biosynthesis of photosynthetic pigments. Biochem J. 1984 Jun 15;220(3):861–863. doi: 10.1042/bj2200861. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Camadro J. M., Labbe P. Purification and properties of ferrochelatase from the yeast Saccharomyces cerevisiae. Evidence for a precursor form of the protein. J Biol Chem. 1988 Aug 25;263(24):11675–11682. [PubMed] [Google Scholar]
  4. Camadro J. M., Matringe M., Scalla R., Labbe P. Kinetic studies on protoporphyrinogen oxidase inhibition by diphenyl ether herbicides. Biochem J. 1991 Jul 1;277(Pt 1):17–21. doi: 10.1042/bj2770017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Chow K. S., Singh D. P., Roper J. M., Smith A. G. A single precursor protein for ferrochelatase-I from Arabidopsis is imported in vitro into both chloroplasts and mitochondria. J Biol Chem. 1997 Oct 31;272(44):27565–27571. doi: 10.1074/jbc.272.44.27565. [DOI] [PubMed] [Google Scholar]
  6. Chow K. S., Singh D. P., Walker A. R., Smith A. G. Two different genes encode ferrochelatase in Arabidopsis: mapping, expression and subcellular targeting of the precursor proteins. Plant J. 1998 Aug;15(4):531–541. doi: 10.1046/j.1365-313x.1998.00235.x. [DOI] [PubMed] [Google Scholar]
  7. De Matteis F., Gibbs A. H. Drug-induced conversion of liver haem into modified porphyrins. Evidence for two classes of products. Biochem J. 1980 Apr 1;187(1):285–288. doi: 10.1042/bj1870285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Huang J., Struck F., Matzinger D. F., Levings C. S., 3rd Flower-enhanced expression of a nuclear-encoded mitochondrial respiratory protein is associated with changes in mitochondrion number. Plant Cell. 1994 Mar;6(3):439–448. doi: 10.1105/tpc.6.3.439. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Jacobs J. M., Jacobs N. J. Oxidation of protoporphyrinogen to protoporphyrin, a step in chlorophyll and haem biosynthesis. Purification and partial characterization of the enzyme from barley organelles. Biochem J. 1987 May 15;244(1):219–224. doi: 10.1042/bj2440219. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Jacobs J. M., Jacobs N. J. Porphyrin Accumulation and Export by Isolated Barley (Hordeum vulgare) Plastids (Effect of Diphenyl Ether Herbicides). Plant Physiol. 1993 Apr;101(4):1181–1187. doi: 10.1104/pp.101.4.1181. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Jacobs J. M., Jacobs N. J., Sherman T. D., Duke S. O. Effect of diphenyl ether herbicides on oxidation of protoporphyrinogen to protoporphyrin in organellar and plasma membrane enriched fractions of barley. Plant Physiol. 1991 Sep;97(1):197–203. doi: 10.1104/pp.97.1.197. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Jacobs J. M., Jacobs N. J. Terminal enzymes of heme biosynthesis in the plant plasma membrane. Arch Biochem Biophys. 1995 Nov 10;323(2):274–278. doi: 10.1006/abbi.1995.9964. [DOI] [PubMed] [Google Scholar]
  13. Jones M. S., Jones O. T. Ferrochelatase of Rhodopseudomonas spheroides. Biochem J. 1970 Sep;119(3):453–462. doi: 10.1042/bj1190453. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Jones M. S., Jones O. T. The structural organization of haem synthesis in rat liver mitochondria. Biochem J. 1969 Jul;113(3):507–514. doi: 10.1042/bj1130507. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Jones O. T. Ferrochelatase of spinach chloroplasts. Biochem J. 1968 Mar;107(1):113–119. doi: 10.1042/bj1070113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Jones O. T. Haem synthesis by isolated chloroplasts. Biochem Biophys Res Commun. 1967 Sep 7;28(5):671–674. doi: 10.1016/0006-291x(67)90367-1. [DOI] [PubMed] [Google Scholar]
  17. Karger G. A., Reid J. D., Hunter C. N. Characterization of the binding of deuteroporphyrin IX to the magnesium chelatase H subunit and spectroscopic properties of the complex. Biochemistry. 2001 Aug 7;40(31):9291–9299. doi: 10.1021/bi010562a. [DOI] [PubMed] [Google Scholar]
  18. Lee H. J., Duke M. V., Duke S. O. Cellular Localization of Protoporphyrinogen-Oxidizing Activities of Etiolated Barley (Hordeum vulgare L.) Leaves (Relationship to Mechanism of Action of Protoporphyrinogen Oxidase-Inhibiting Herbicides). Plant Physiol. 1993 Jul;102(3):881–889. doi: 10.1104/pp.102.3.881. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Lermontova I., Kruse E., Mock H. P., Grimm B. Cloning and characterization of a plastidal and a mitochondrial isoform of tobacco protoporphyrinogen IX oxidase. Proc Natl Acad Sci U S A. 1997 Aug 5;94(16):8895–8900. doi: 10.1073/pnas.94.16.8895. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Lister R., Chew O., Rudhe C., Lee M. N., Whelan J. Arabidopsis thaliana ferrochelatase-I and -II are not imported into Arabidopsis mitochondria. FEBS Lett. 2001 Oct 12;506(3):291–295. doi: 10.1016/s0014-5793(01)02925-8. [DOI] [PubMed] [Google Scholar]
  21. Little H. N., Jones O. T. The subcellular loclization and properties of the ferrochelatase of etiolated barley. Biochem J. 1976 May 15;156(2):309–314. doi: 10.1042/bj1560309. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Margalit R., Shaklai N., Cohen S. Fluorimetric studies on the dimerization equilibrium of protoporphyrin IX and its haemato derivative. Biochem J. 1983 Feb 1;209(2):547–552. doi: 10.1042/bj2090547. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Matringe M., Camadro J. M., Joyard J., Douce R. Localization of ferrochelatase activity within mature pea chloroplasts. J Biol Chem. 1994 May 27;269(21):15010–15015. [PubMed] [Google Scholar]
  24. Papenbrock J., Mishra S., Mock H. P., Kruse E., Schmidt E. K., Petersmann A., Braun H. P., Grimm B. Impaired expression of the plastidic ferrochelatase by antisense RNA synthesis leads to a necrotic phenotype of transformed tobacco plants. Plant J. 2001 Oct;28(1):41–50. doi: 10.1046/j.1365-313x.2001.01126.x. [DOI] [PubMed] [Google Scholar]
  25. Porra R. J., Lascelles J. Studies on ferrochelatase. The enzymic formation of haem in proplastids, chloroplasts and plant mitochondria. Biochem J. 1968 Jun;108(2):343–348. doi: 10.1042/bj1080343. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Smith A. G., Marsh O., Elder G. H. Investigation of the subcellular location of the tetrapyrrole-biosynthesis enzyme coproporphyrinogen oxidase in higher plants. Biochem J. 1993 Jun 1;292(Pt 2):503–508. doi: 10.1042/bj2920503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Smith A. G., Santana M. A., Wallace-Cook A. D., Roper J. M., Labbe-Bois R. Isolation of a cDNA encoding chloroplast ferrochelatase from Arabidopsis thaliana by functional complementation of a yeast mutant. J Biol Chem. 1994 May 6;269(18):13405–13413. [PubMed] [Google Scholar]
  28. Smith A. G. Subcellular localization of two porphyrin-synthesis enzymes in Pisum sativum (pea) and Arum (cuckoo-pint) species. Biochem J. 1988 Jan 15;249(2):423–428. doi: 10.1042/bj2490423. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Suzuki T., Masuda T., Inokuchi H., Shimada H., Ohta H., Takamiya K. Overexpression, enzymatic properties and tissue localization of a ferrochelatase of cucumber. Plant Cell Physiol. 2000 Feb;41(2):192–199. doi: 10.1093/pcp/41.2.192. [DOI] [PubMed] [Google Scholar]
  30. Thomas J., Weinstein J. D. Measurement of heme efflux and heme content in isolated developing chloroplasts. Plant Physiol. 1990 Nov;94(3):1414–1423. doi: 10.1104/pp.94.3.1414. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Walker C. J., Weinstein J. D. Further characterization of the magnesium chelatase in isolated developing cucumber chloroplasts : substrate specificity, regulation, intactness, and ATP requirements. Plant Physiol. 1991 Apr;95(4):1189–1196. doi: 10.1104/pp.95.4.1189. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Walker C. J., Willows R. D. Mechanism and regulation of Mg-chelatase. Biochem J. 1997 Oct 15;327(Pt 2):321–333. doi: 10.1042/bj3270321. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Witty M., Jones R. M., Robb M. S., Jordan P. M., Smith A. G. Subcellular location of the tetrapyrrole synthesis enzyme porphobilinogen deaminase in higher plants: an immunological investigation. Planta. 1996;199(4):557–564. doi: 10.1007/BF00195187. [DOI] [PubMed] [Google Scholar]
  34. ap Rees T., Bryce J. H., Wilson P. M., Green J. H. Role and location of NAD malic enzyme in thermogenic tissues of Araceae. Arch Biochem Biophys. 1983 Dec;227(2):511–521. doi: 10.1016/0003-9861(83)90480-0. [DOI] [PubMed] [Google Scholar]

Articles from Biochemical Journal are provided here courtesy of The Biochemical Society

RESOURCES