TSitologiya i Genetika 2021, vol. 55, no. 6, 72-74
Cytology and Genetics 2021, vol. 55, no. 6, 558–565, doi: https://www.doi.org/10.3103/S009545272106013X

The feasibility of microalgae Dunaliella identification based on conserved regions of mitochondrial cytochrome b and cytochrome oxidase genes

Razeghi J., Pishtab P.A., Fathi P., Panahi B., Hejazi M.A.

  1. Department of Plant Sciences, Faculty of Natural Sciences, University of Tabriz, Tabriz 51666­16471, Iran
  2. Department of Genomics, Branch for Northwest and West Region, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Tabriz 5156915­598, Iran
  3. Department of Food Biotechnology, Branch for Northwest and West Region, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Tabriz 5156915­598, Iran

Classification of unicellular algae at species level has not been successfully characterized with common morphological, physiological, and molecular approaches. In this study, the efficiency of two mitochondrial cob and cox1 genes as new molecular targets for the study of the phylogeny relationships was investigated among twenty isolated Dunaliella species from different regions of Iran. First, specific primers were designed based on the conserved regions of the cob and cox1 sequences in Dunaliella species and other microalgae, followed by analysis of PCR products. Based on the analysis of amplification products, some isolates were selected for subsequent RFLP and sequencing processes. Findings revealed that cob gene was not amplified in the isolates, whereas cox1 gene was amplified in B60, M12, G3, and CCAP19/18 isolates. RFLP results showed that B60 with 19/18 and G3 with M1.2 had a similar pattern and were grouped in the same clade. Interestingly, the findings on cox1 gene sequencing demonstrated complete congruence with RFLP results. Although the results of this study highlighted the efficiency of cox1 gene in determining the phylogeny relationships between Dunaliella species, further cox1 gene sequences and subsequently PCR and RFLP analyses are required to approve the results. Our results open a new avenue on growing bodies of knowledge regarding the phylogeny relationships of Dunaliella species and could be suitable in taxonomical studies of other microalgae.

Keywords: cob, cox1, Dunaliella, mitochondrial genes, molecular phylogeny

TSitologiya i Genetika
2021, vol. 55, no. 6, 72-74

Current Issue
Cytology and Genetics
2021, vol. 55, no. 6, 558–565,
doi: 10.3103/S009545272106013X

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1. Endo, H., Park, E.J., Sato, Y., Mizuta, H., and Saga, N., 2009) Intraspecific diversity of Undaria pinnatifida (Harvey) Suringar (Laminariales, Phaeophyta) from Japan, China and Korea, based on the cox1 gene and ITS2 sequences, Fish Sci., 2012, vol. 75, no. 2, pp. 393–400.

2. Evans, K.M., Wortley, A.H., and Mann, D.G., An assessment of potential diatom “barcode” genes (cox1, rbcL, 18S and ITS rDNA) and their effectiveness in determining relationships in Sellaphora (Bacillariophyta), Protist, 2007, vol. 158, no. 3, pp. 349–364.

3. Ghorbani, S., Manaffar, R., Taee, A., and Malekzadeh, R., A study on molecular diversity of Dunaliella algae species in some of Urmia Lake’s stations, Iran. J. Plant Biol., 2007, vol. 5, no. 17, pp. 1–8.

4. Ghorbanzadeh Naghab, M. and Panahi, B., Molecular characterization of Iranian black cumin (Nigella sativa L.) accessions using RAPD marker, Biotechnologia, 2017, vol. 98, pp. 2, pp. 97–102.

5. Gomez, P.I. and Gonzalez, M.A., Genetic variation among seven strains of Dunaliella salina (Chlorophyta) with industrial potential, based on RAPD banding patterns and on nuclear ITS rDNA sequences, Aquaculture, 2004, vol. 233, pp. 149–162.

6. Hejazi, M.A., Barzegari, A., Gharajeh, N.H., and Hejazi, M.S., Introduction of a novel 18S rDNA gene arrangement along with distinct ITS region in the saline water microalga Dunaliella, Saline Syst., 2010, vol. 6, no. 1, pp. 11–41.

7. Hejazi, M.A., Khoshrouy, R., Hosseinzadeh Gharajeh, N., Etemadi, M.R., Madayen, L., and Javanmard, A., Conservation and biodiversity analysis of the microalga Dunaliella in shrinking highly saline Urmia Lake based on intron-sizing method, J. Agr. Sci. Tech., 2016, vol. 18, pp. 1693–1703.

8. Jolodar, A., Molecular characterization and phylogeny analysis based on sequences of cytochrome oxidase gene from Hemiscorpius lepturus of Iran, Iran. J. Vet. Med., 2019, vol. 13, no. 1, pp. 59–67.

9. Kamikawa, R., Nagai, S., Hosoi-Tanabe, S., Itakura, S., Uchida, Y., Baba, T., and Sako, Y., Application of real-time PCR assay for detection and quantification of Alexandrium tamarense and Alexandrium catenella cysts from marine sediments, Harmful Algae, 2007, vol. 6, pp. 413–420.

10. Le Gall, L. and Saunders, G.W., DNA barcoding is a powerful tool to uncover algal diversity: a case study of the Phyllophoraceae (Gigartinales, Rhodophyta) in the Canadian flora, J. Phycol., 2010, vol. 46, no. 2, pp. 374–389.

11. Moniz, M.B.J. and Kaczmarska, I., Barcoding of diatoms: nuclear encoded ITS revisited, Protist, 2010, vol. 161, no. 1, pp. 7–34.

12. Olmos, J., Paniagua, J., and Contreras, R., Molecular identification of Dunaliella sp. utilizing the 18S rDNA gene, Lett. Appl. Microbiol., 2000, vol. 30, pp. 80–84.

13. Panahi, B., Afzal, R., Ghorbanzadeh Neghab, M., Mahmoodnia, M., and Paymard, B., Relationship among AFLP, RAPD marker diversity and agromorphological traits in safflower (Carthamus tinctorius L.), Progr. Biol. Sci., 2013, vol. 3, no. 1, pp. 90–99.

14. Panahi, B., Frahadinan, M., Dumas, J., and Hejazi, M., Integration of cross species RNA-seq Meta- analysis and Machine Learning Models identifies the most important salt stress responsive pathways in microalga Dunaliella, Front. Genet., 2019, vol. 10, p. 752.

15. Preetha, K., John, L., Subin, C.S., and Vijayan, K.K., Phenotypic and genetic characterization of Dunaliella (Chlorophyta) from Indian salinas and their diversity, Aquat. Biosyst., 2012, vol. 8, no. 1, p. 27.

16. Quispe-Tintaya, W., White, R.R., Popov, V.N., Vijg, J., and Maslov, A.Y., Fast mitochondrial DNA isolation from mammalian cells for next-generation sequencing, BioTechniques, 2013, vol. 55, pp. 133–136.

17. Raho, N., Rodríguez, F., Reguera, B., and Marín, I., Are the mitochondrial cox1 and cob genes suitable markers for species of Dinophysis Ehrenberg?, Harmful Algae, 2013, vol. 28, pp. 64–70.

18. Robba, L., Russell, S.J., Barker, G.L., and Brodie, J., Assessing the use of the mitochondrial cox1 marker for use in DNA barcoding of red algae (Rhodophyta), Am. J. Bot., 2006, vol. 93, no. 8, pp. 1101–1108.

19. San, M.D., Gower, D.G., Zardoya, R., and Wilkinson, M., A hotspot of gene order rearrangement by tandem duplication and random loss in the vertebrate mitochondrial genome, Mol. Biol. Evol., 2006, vol. 23, pp. 227–234.

20. Sathasivam, R., Praiboon, J., Chirapart, A., Trakulnaleamsai, S., Kermanee, P., Roytrakul, S., and Juntawong, N., Screening, phenotypic and genotypic identification of β-carotene producing strains of Dunaliella salina from Thailand, Ind. J. Geo-Mar. Sci., 2014, vol. 43, no. 12, pp. 2198–2216.

21. Saunders, G.W., Applying DNA barcoding to red macroalgae: a preliminary appraisal holds promise for future applications, Philos. Trans. R. Soc., B, 2005, vol. 360, no. 1462, pp. 1879–1888.

22. Smith, D.R., Lee, R.W., Cushman, J.C., Magnuson, J.K., Tran, D., and Polle, E.J., The Dunaliella salina organelle genomes: large sequences, inflated with intronic and intergenic DNA, BMC Plant Biol., 2010, vol. 10, pp. 83–96.

23. Tamura, K., Peterson, D., Peterson, N., Steecher, G., and NeiMand Kumar, S., MEGA: Molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods, Mol. Biol. Evol., 2011, vol. 28, pp. 2731–2739.

24. Tempesta, S., Paoletti, M., and Pasqualetti, M., Morphological and molecular identification of a strain of the unicellular green algae Dunaliella sp. isolated from Tarquinia salterns, Transit. Water Bull., 2010, vol. 4, pp. 60–70.

25. Thompson, J.D., Gibson, T.J., and Higgins, D.G., Multiple sequence alignment using ClustalW and ClustalX, Curr. Protoc. Bioinformatics, 2002, Chapter 2, Unit2.

26. Turmel, M., Otis, C., and Lemieux, C., The chloroplast and mitochondrial genome sequences of the charophyte Chaetosphaeridium globosum: insights into the timing of the events that restructured organelle DNAs within the green algal lineage the led to land plants, Proc. Natl. Acad. Sci. U. S. A., 2002, vol. 99, no. 17, pp. 11275–11280.

27. Turmel, M., Otis, C., and Lemieux, C., The mitochondrial genome of Chara vulgaris: insights into the mitochondrial DNA architecture of the last common ancestor of green algae and land plants, Plant Cell, 2003, vol. 15, no. 8, pp. 1888–1903.

28. Wahrmund, U., Quandt, D., and Knoop, V., The phylogeny of mosses—addressing open issues with a new mitochondrial locus: group I intron cobi420, Mol. Phylogenet. Evol., 2010, vol. 54, pp. 417–426.

29. Wang, F., Jia, F., Jie, W., Bo, L., and Shulian, X., Phylogenetic and morphological investigation of a Dunaliella strain isolated from Yuncheng Salt Lake, China, Acta Geol. Sin. (Engl. Ed.), 2014, vol. 88, no. 1, pp. 106–107.

30. Zhang, H., Bhattacharya, D., Maranda, L., and Lin, S.,Mitochondrial cob and cox1 genes and editing of the corresponding mRNAs in Dinophysis acuminata from Narragansett Bay, with special reference to the phylogenetic position of the genus Dinophysis, Appl. Environ. Microbiol., 2008, vol. 74, no. 5, pp. 1546–1554.