Agroforestry Systems

, Volume 93, Issue 3, pp 1093–1105 | Cite as

Embryogenesis followed by enhanced micro-multiplication and eco-restoration of Calamus thwaitesii Becc.: an economic non-wood forest produce for strengthening agroforestry system

  • Achuthan Sudarsanan Hemanthakumar
  • Thankappan Suvarna PreethaEmail author
  • Padmesh Pandaram Pillai
  • Sooriamuthu Seeni


The present study is focussed on development of a high-frequency micro-multiplication system in Calamus thwaitesii, through somatic embryogenesis from immature zygotic embryos cultured in Murashige and Skoog (MS) medium supplemented with 31.68 µM 2,4-dichlorophenoxyacetic acid (2,4-D). The semi-friable calli when cultured in the same medium augmented with 2.22 µM 6-benzylaminopurine (BAP) and 1.07 µM α-naphthalene acetic acid (NAA) induced ~ 12 discrete globular embryoids in 6 weeks. The isolated embryoids in hormone-free media yielded 65% plantlets. Furthermore, embryoids and axenic shoots exhibited maximum shoot induction in medium supplemented with 0.45 µM Thidiazuron (TDZ). The shoot initials after subculture in media supplemented with 1.78 µM BAP and 0.45 µM TDZ produced shoot proliferation followed by elongation in basal medium. The elongated shoots produced roots in media supplemented with 16.11 µM NAA. With this established protocol, ~ 5940 rooted plantlets could be harvested after 40 weeks from a single embryoid. Genetic stability analysis of the plantlets using inter simple sequence repeat markers recorded only 0.05% genetic polymorphism. The plantlets were hardened in a mist house for 8 weeks, and then to 50% shade house for another 16 weeks, and the well-established 6-month-old nursery plants, reintroduced to selected forest segments, exhibited 86% field establishment even after 3 years of observation. Thus, the mass multiplication system developed could be a breakthrough for large-scale multiplication of C. thwaitesii to ensure continuous supply of quality planting material to the cottage industry through the development of agroforestry systems. Furthermore, the in vitro culture system developed here can be replicated for research activities related to the long-term–short-term conservation, micro-multiplication and sustainable utilization of rare, endangered, endemic, monopodial/single stemmed rattan palms.


Calamus thwaitesii Rattan palm Genetic variability Embryoids Eco-restoration 





2,4-Dichlorophenoxyacetic acid


Indole-3-butyric acid


Inter simple sequence repeat


Murashige and Skoog


α-Naphthaleneacetic acid


Plant growth regulator





We thankfully acknowledge the Department of Biotechnology (DBT), Government of India, for providing the financial assistance through the research grant (No BT/R&D/08/04/95), the Director, Jawaharlal Nehru Tropical Botanic Garden and Research Institute (JNTBGRI) for providing laboratory facilities and the Department of Forests, Government of Kerala extended support for eco-restoration activities.


  1. Aminuddin M, Nur Surpadi, Noor Md, Woon WC (1992) Economics of cultivation of large diameter rattan. In: Wan Mohd., Wan R, Dransfield J, Manokaron N (eds) A guide to the cultivation of rattan. Malayan For.Rec. p 35Google Scholar
  2. Arditti J, Ernst R (1993) Micropropagation of Orchids. Wiley, New York, pp 1–682Google Scholar
  3. Barba RC, Patena LJ, Mercado MM, Lorico L (1985) Tissue culture of rattan (Calamus manillensis. Wendl.) In: Proceedings of the Second National Symposium on Tissue Culture of Rattan, University Partanian MalaysiaGoogle Scholar
  4. Bhatia R, Singh KP, Jhang T, Sharma TR (2008) Assessment of clonal fidelity of micropropagated gerbera plants by ISSR markers. Sci Hortic 119:208–2111CrossRefGoogle Scholar
  5. Bingshan Zeng (1997) Tissue culture of Calamus egregius. J. Cent South For Univ 17(4):563–569Google Scholar
  6. Blake J, Eeuwens CJ (1982) Culture of coconut palm tissues with a view to vegetative propagation. In: Rao AN (ed) Proceedings of COSTE\symposium on Tissue Culture of Economically Important Plants, Singapore, pp 145–148Google Scholar
  7. Chengji Zhuang, Jiankui Zhou (1991) Plant regeneration in tissue culture of rattan. Acta Bot Yunnaica 13(1):97–100Google Scholar
  8. Corley RHV, Barrett JN, Jones LH (1976) Vegetative propagation of oil palm via tissue culture. Malaysian International Agricultural Oil Palm Conference, pp 1–8Google Scholar
  9. Cuenca B, Ballester A, Vieitez AM (2000) In vitro adventitious bud regeneration from internode segments of beech. Plant Cell Tiss Org Cult 60:213–220CrossRefGoogle Scholar
  10. Damasco OP, Graham GC, Henry RJ, Adkins SW, Smith MK, Godwin ID (1996) Random amplified polymorphic DNA (RAPD) detection of dwarf off-types in micropropagated Cavendish (Musa spp. AAA) bananas. Plant Cell Rep 16:118–123CrossRefGoogle Scholar
  11. Debnath SC (2005) A two-step procedure for adventitious shoot regeneration from in vitro-derived lingonberry leaves: shoot induction with TDZ and shoot elongation using zeatin. Hortic Sci 40:189–192Google Scholar
  12. Eeuwens CJ (1976) Mineral requirements for growth and callus initiation of tissue explants excised from mature coconut palms (Cocos nucifera) and cultured in vitro. Physiol Plant 36:23–28CrossRefGoogle Scholar
  13. Eeuwens CJ (1978) Effects of organic nutrients and hormones on growth and development of tissue explants from coconut (Cocos nucifera) and date palms (Phoenix dactylifera) cultured in vitro. Physiol Plant 42:173–178CrossRefGoogle Scholar
  14. Eshraghi P, Zarghami R, Mirabdulbaghi M (2005) Somatic embryogenesis in two Iranian date palm cultivars. Afr J Biotechnol 4(11):1309–1312., ISSN 1684–5315
  15. Fangqiu Zhang (1993) A study on Rattan tissue culture. J For Res 6(5):486–492Google Scholar
  16. Fasolo F, Zimmerman RH, Fordham I (1989) Adventitious shoot formation on excised leaves of in vitro grown shoots of apple cultivars. Plant Cell Tiss Org Cult 16:75–87CrossRefGoogle Scholar
  17. Goto S, Thakur RC, Ishii K (1998) Determination of genetic stability in long-term micropropagated shoots of Pinus thunbergii Parl. using RAPD markers. Plant Cell Rep 18:193–197CrossRefGoogle Scholar
  18. Hazarika BN (2003) Acclimatization of tissue-cultured plants. Curr Sci 85:1704–1712Google Scholar
  19. Hemanthakumar AS (2010) Studies on embryo and tissue culture of three economically important rattan sps. Thesis submitted to Kerala UniversityGoogle Scholar
  20. Hemanthakumar AS, Preetha TS, Krishnan PN, Seeni S (2013) Utilization of zygotic embryos of an economic rattan palm Calamus thwaitesii Becc. (Arecaceae) for somaplant regeneration and cryobanking. 3 Biotech 3:195–203CrossRefGoogle Scholar
  21. Joshi P, Dhawan V (2007) Assessment of genetic fidelity of micropropagated Swertia chirayita plantlets by ISSR marker assay. Biol Plant 51(1):22–26CrossRefGoogle Scholar
  22. Kundu M, Sett R (1999) Regeneration through organogenesis in rattan. Plant Cell Tissue Org Cult 59:219–222CrossRefGoogle Scholar
  23. Lopez-Villalobos A, Dodds PF, Hornung R (2001) Changes in fatty acid composition during development of tissues of coconut (Cocos nucifera L.) embryos in the intact nut and in vitro. J Exp Bot 52:933–942CrossRefGoogle Scholar
  24. Lyyra S, Lima A, Merkle S (2006) In vitro regeneration of Salix nigra from adventitious shoots. Tree Physiol 26:969–975CrossRefGoogle Scholar
  25. Magioli C, Rocha APM, de Oliveria DE, Mansur E (1998) Efficient shoot organogenesis of eggplant (Solanum melongena L.) induced by thidiazuron. Plant Cell Rep 17:661–663CrossRefGoogle Scholar
  26. Martin JP, Rabechault H (1976) Procede de multiplication vegetative de vegetaux et plants ainsi obtenus, French Patent No. 7628361Google Scholar
  27. Martin KP, Pachathundkandi SK, Zhang CL, Slater A (2006) RAPD analysis of a variant of Banana (Musa sp.) cv. Grande Naine and its propagation via shoot tip culture. In Vitro Cell Dev Biol 42:188–192CrossRefGoogle Scholar
  28. Merillon JM, Rideau M, Chenieux JC (1984) Influence of sucrose on levels of ajmalicine, serpentine and triptamine in Catharanthas roseus cells in vitro. Plant Med 50:497–501CrossRefGoogle Scholar
  29. Mojarabi M, Nasr SMH, Jalilvand H, Kooch Y (2011) Effect of activated charcoal, growth supplements and storage on removing dormancy, germination indices and vigour of Ash (Fraxinus excelsior L.). Ann Biol Res 2(5):203–212Google Scholar
  30. Murashige T, Skoog F (1962) A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol Plant 15:473–497CrossRefGoogle Scholar
  31. Murray HG, Thompson WF (1980) Rapid isolation of high molecular weight DNA. Nucl Acids Res 8:4321–4325CrossRefGoogle Scholar
  32. Murthy BNS, Murch SJ, Saxena PK (1998) Thidiazuron: a potent regulator of in vitro morphogenesis. In Vitro Cell Dev Biol 34:267–275CrossRefGoogle Scholar
  33. Nair LG, Seeni S (2002) Rapid clonal multiplication of Morinda umbellate Linn. (Rubiaceae), a medicinal liana, through cultures of nodes and shoot tips from mature plant. Phytomorphology 52:77–81Google Scholar
  34. Padmanabhan D, Ilangovan R (1993) Surgical induction of multiple shoots in embryo cultures of Calamus gamblei Becc. RIC Bull 12:8–12Google Scholar
  35. Ramanayake SMSD (1999) Viability of excised embryos, shoot proliferation and in vitro flowering in a species of rattan Calamus thwaitesii Becc. J Hortic Sci Biotechnol 74(5):594–601CrossRefGoogle Scholar
  36. Reynolds JF, Murashige T (1979) Asexual embryogenesis in callus cultures of palms. In Vitro 15:383–387CrossRefGoogle Scholar
  37. Rout GR, Das P, Goel S, Raina SN (1998) Genetic stability of micropropagated plants of ginger using random amplified polymorphic DNA (RAPD) markers. Bot Bull Acad Sin 39:23–27Google Scholar
  38. Sáenz L, Azpeitia A, Chuc-Armendariz B, Chan JL, Verdeil JL, Hocher V, Oropeza C (2006) Morphological and histological changes during somatic embryo formation from coconut plumule explants. In Vitro Cell Dev Biol 42:19–25. CrossRefGoogle Scholar
  39. Sane D, Aberlenc-Bertossi F, Gassama-Dia YK, Sagna M, Trouslot MF, Duval Y, Borgel A (2006) Histocytological analysis of callogenesis and somatic embryogenesis from cell suspensions of date palm (Phoenix dactylifera). Ann Bot 98(2):301–308CrossRefGoogle Scholar
  40. Satheeshkumar K, Seeni S (2000) In vitro multiplication of Nothapodites foetida (Wight.) Sleumer through seedling explant cultures. Indian J Exp Biol 38:273–277Google Scholar
  41. Scherwinski-Pereira JE, da Guedes RS, Fermino PCP Jr, Silva TL, Costa FHS (2010) Somatic embryogenesis and plant regeneration in oil palm using the thin cell layer technique. In Vitro Cell Dev Biol 46:378–385. CrossRefGoogle Scholar
  42. Seeni S, Latha PG (2000) In vitro multiplication and ecorehabilitation of the endangered Blue Vanda. Plant Cell Tiss Org Cult 61:1–8CrossRefGoogle Scholar
  43. Sett R, Kundu M, Sharma P (2002) Regeneration of plantlets through organogenesis in Calamus tenuis. Indian J Plant Physiol 7:358–361Google Scholar
  44. Singha S, Bhatia SK (1988) Shoot proliferation of pear cultures on medium containing thidiazuron and benzylamino purine. Hortic Sci 23:803Google Scholar
  45. SPSS 9 (version 10.0) (1999) Manuals SPSS Inc., ChicagoGoogle Scholar
  46. Thomas TD (2008) The role of activated charcoal in plant tissue culture. Biotechnol Adv 26(6):618–631CrossRefGoogle Scholar
  47. Tisserat B (1983) Tissue culture of date palms: a new method to propagate an ancient crop-and a short discussion of the california date industry. Principes 27(3):105–117Google Scholar
  48. Victório CP, Lage CLS, Sato A (2012) Tissue culture techniques in the proliferation of shoots and roots of Calendula officinalis. Rev Ciênc Agron 43(3):539–545CrossRefGoogle Scholar
  49. Vieitez AM, San-Jose MC (1996) Adventitious shoot regeneration from Fagus sylvatica leaf explants in vitro. In Vitro Cell Dev Biol 32:140–147CrossRefGoogle Scholar
  50. Wainwright H, Scrace J (1989) Influence of in vitro preconditioning with carbohydrates during the rooting of microcuttings on in vivo establishment. Sci Hortic 38:261–267CrossRefGoogle Scholar
  51. Wang H-C, Chen J-T, Wu S-P, Lin M-C, Chang W-C (2003) Plant regeneration through shoot formation from callus of Areca catechu. Plant Cell Tiss Org Cult 75:95–98CrossRefGoogle Scholar
  52. Yang H, Tabei Y, Kamada H, Kayamo T, Takaiwa F (1999) Detection of somaclonal variation in cultured rice cells using digoxygenin-based random amplified polymorphic DNA. Plant Cell Rep 18:520–526CrossRefGoogle Scholar
  53. Yap IV, Nelson RJ (1996) Win Boot: a program for performing bootstrap analysis of binary data to determine the confidence limits of UPGMA-based dendrograms. IRRI Discussion Paper Series No. 14, International Rice Research Institute, Manila, PhilippinesGoogle Scholar
  54. Yusoff AM (1989) Shoot formation in Calamus manan under in vitro. In: Rao AN, Aziah Mohd. Yusoff (eds) Proceedings of the seminar on Tissue Culture of Forest Species, Forest Research Institute Malaysia and International Development Research Centre, Singapore, pp 45–49Google Scholar

Copyright information

© Springer Science+Business Media B.V., part of Springer Nature 2018

Authors and Affiliations

  1. 1.Biotechnology and Bioinformatics DivisionJawaharlal Nehru Tropical Botanic Garden and Research InstituteThiruvananthapuramIndia
  2. 2.Department of BotanyUniversity CollegeThiruvananthapuramIndia
  3. 3.Department of Genomic ScienceCentral University of KeralaKasaragodIndia
  4. 4.School of BiosciencesMar Athanasios College for Advanced StudiesTiruvallaIndia

Personalised recommendations