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In Vitro Culture of Croftonweed (Ageratina adenophora): Considerable Potential for Fast and Convenient Plantlet Production

Published online by Cambridge University Press:  20 January 2017

Jinbo Shen
Affiliation:
College of Chemistry and Life Science, Zhejiang Normal University, Jinhua 321004, China
Xia Li
Affiliation:
College of Chemistry and Life Science, Zhejiang Normal University, Jinhua 321004, China
Dandan Wang
Affiliation:
College of Chemistry and Life Science, Zhejiang Normal University, Jinhua 321004, China
Hongfei Lu*
Affiliation:
College of Chemistry and Life Science, Zhejiang Normal University, Jinhua 321004, China
*Corresponding
Corresponding author's E-mail: luhongfei0164@sina.com

Abstract

A callus induction and plantlet regeneration system for croftonweed was developed by studying the influence of explant type (leaf, stem, and nodal segment) and different concentrations of plant growth regulators. The leaf was a better explant for callogenesis compared to the stem. The highest callus induction frequency (87.2%) was obtained from leaf segments on Murashige and Skoog's medium (MS medium) supplemented with 0.5 mg/L (2,4-dichlorophenoxy)acetic acid and 2.0 mg/L 6-benzylaminopurine (BA), and 71.6% differentiation along with a multiplication rate of 4.1 adventitious shoots per callus was achieved with a combination of 0.5 mg/L 1-naphthaleneacetic acid (NAA) and 1.0 mg/L BA. In addition, MS medium supplemented with 0.5 mg/L NAA and 1.0 mg/L BA was the best medium for axillary shoot regeneration from nodal segments. Rhizogenesis of cultured shoots was satisfactorily obtained in half-strength MS without any growth regulators. The regenerated rooted plantlets were successfully acclimatized in soil where they grew normally without showing any morphological variation. These studies provide the prerequisite system for the development of genetic engineering in the future and propagating croftonweed rapidly for further study.

Type
Research
Copyright
Copyright © Weed Science Society of America 

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References

Arockiasamy, S. and Ignacimuthu, S. 1998. Plant regeneration from mature leaves and roots of Eryngium foetidum L., a food flavouring agent. Curr. Sci. 75:644666.Google Scholar
Arya, M. P. S. and Singh, R. V. 1998. Evaluation of herbicides for the control of croften weed (Eupatorium adenophorum Sprengel) for different land use. Indian J. Weed Sci. 30:179181.Google Scholar
Auld, B. A. 1969. The distribution of Eupatorium adenophorum Spreng. On the far north coast of New South Wales. J. Proc. Royal Soc. New South Wales 102:159161.Google Scholar
Auld, B. A. 1972. Chemical control of Eupatorium adenophorum, crofton weed. Trop. Grass. 6:5560.Google Scholar
Auld, B. A. and Martin, P. M. 1975. The autecology of Eupatorium adenophorum Spreng. in Australia. Weed Res. 15:2731.CrossRefGoogle Scholar
Baker, M. C., Munoz-Fernandez, N., and Carter, C. D. 1999. Improved sheet development and rooting from mature cotyledons of sunflower. Plant Cell Tiss. Org. Cult. 58:3949.CrossRefGoogle Scholar
Bess, H. A. and Haramoto, F. H. 1958. Biological control of Pamakani Eupatorium adenophorum in Hawaii by a Tephritid gall fly. Procecidochares utilis. 1. The life history of the fly and its effectiveness in the control of the weed. Pages 543548. in Proceedings of the 10th International Congress of Entomology. Montreal.Google Scholar
Borthakur, M., Dutta, K., Nath, S. C., and Sing, R. S. 2000. Micropropagation of Eclipta alba and Eupatorium adenophorum using a single-step nodal cutting technique. Plant Cell Tiss. Org. Cult. 62:239242.CrossRefGoogle Scholar
Chengalrayan, K. and Gallo-Meagher, M. 2001. Effect of various growth regulators on shoot regeneration of sugarcane. In Vitro Cell. Dev. Biol. Plant. 37:434439.CrossRefGoogle Scholar
Chraibi, K. M. B., Castelle, J. C., Latché, A., Roustan, P., and Fallot, F. 1992. A genotype-independent system of regeneration from cotyledons of sunflower (Helianthus annuus L.): the role of ethylene. Plant Sci. 86:215221.CrossRefGoogle Scholar
Christopher, T. and Rajam, M. V. 1996. Effect of genotype, explant and medium on in vitro regeneration of red pepper. Plant Cell Tiss. Org. Cult. 46:245250.CrossRefGoogle Scholar
Cuenca, S. and Amo-Marco, J. B. 2000. In vitro propagation of two spanish endemic species of salvia through bud proliferation. In Vitro Cell. Dev. Biol. Plant. 36:225229.CrossRefGoogle Scholar
Deglene, L., Lesignes, P., Alibert, G., and Saraffi, A. 1997. Genetic control of organogenesis in cotyledons of sunflower (Helienthus annuus). Plant Cell Tiss. Org. Cult. 48:127130.CrossRefGoogle Scholar
Dhar, A. C., Kishor, P. B. K., and Rao, A. M. 1989. In vitro propagation of guayule (Parthenium argentatum)—a rubber-yielding shrub. Plant Cell Rep. 8:489492.CrossRefGoogle ScholarPubMed
Dhar, U. and Joshi, M. 2005. Efficient plant regeneration protocol through callus for Saussurea obvallata (DC.) Edgew. (Asteraceae): effect of explant type, age and plant growth regulators. Plant Cell Rep. 24:195200.CrossRefGoogle ScholarPubMed
Dodd, A. P. 1961. Biology control of Eupatorium adenophorum in Queensland. Aust. J. Sci. 23:356365.Google Scholar
Echeverrigaray, S., Fracaro, F., Andrade, L. B., Biasio, S., and Atti-Serafini, L. 2000. In vitro shoot regeneration from leaf explants of Roman Chamomile. Plant Cell Tiss. Org. Cult. 60:14.CrossRefGoogle Scholar
Ferreira, C. M. and Handro, W. 1988. Production, maintenance and plant regeneration from cell suspension cultures of Stevia rebaudiana (Bert.) Bertoni. Plant Cell Rep. 7:123126.CrossRefGoogle ScholarPubMed
Heirwegh, K. M. G., Banerjee, N., Nerum, K., and Langhe, E. 1985. Somatic embryogenesis and plant regeneration in Cichorium intybus L. (Witloof, Compositae). Plant Cell Rep. 4:108111.CrossRefGoogle Scholar
Henson, S. E., Skroch, W. A., Burton, J. D., and Worsham, A. D. 2003. Herbicide efficacy using a wet-blade application system. Weed Technol. 2:320324.CrossRefGoogle Scholar
Hoy, J. M. 1960. Establishment of Procecidochares utilis Stone (Diptera: Trypetidae) on Eupatorium adenophorum Spreng. in New Zealand. N Z J. Sci. 3:200208.Google Scholar
Janick, J. and Whipkey, A. 2002. Trends in new crops and new uses. Pages 522526. in Koroch, A., Kapteyn, J., Juliani, H.R. and Simon, J.E. eds. In Vitro Regeneration and Agrobacterium Transformation of Echinacea purpurea Leaf Explants. Alexandria ASHS.Google Scholar
Kluge, R. L. 1991. Biological control of crofton weed, Ageratina adenophora (Asteraceae) in South Africa. Agric. Ecosystems Environ. 37:187191.CrossRefGoogle Scholar
Knittel, N., Escandon, A. S., and Hahne, G. 1991. Plant regeneration at high frequency from mature sunflower cotyledons. Plant Sci. 73:219226.CrossRefGoogle Scholar
Koroch, A. R., Kapteyn, J., Juliani, H. R., and Simon, J. E. 2003. In vitro regeneration of echinacea pallida from leaf explants. In Vitro Cell. Dev. Biol. Plant. 39:415418.CrossRefGoogle Scholar
Laparra, H., Stoeva, P., Ivanov, P., and Hahne, G. 1997. Plant regeneration from different explants in Helianthus smithii Heiser. Plant Cell Rep. 16:692695.CrossRefGoogle Scholar
Li, H., Qiang, S., and Cui, J. 2005. Eupatorium adenophorum micropropagation by bud and callus culture. Acta Bot. Boreal. Occident. Sin. 25:14581462.Google Scholar
Lisowska, K. and Wysokinska, H. 2000. In vitro propagation of Catalpa ovata G. Don. Plant Cell Tiss. Org. Cult. 60:171176.CrossRefGoogle Scholar
Liu, L. H., Xie, S. C., and Zhang, J. H. 1985. Studies on the distribution, harmfulness and control of Eupatorium adenophorum Spreng. Acta Ecol. Sin. 5:16.Google Scholar
MacDonald, G. E., Brecke, B. J., Colvin, D. L., and Shilling, D. G. 1994. Chemical and mechanical control of dogfennel (Eupatorium capillifolium). Weed Technol. 3:483487.CrossRefGoogle Scholar
Mechanda, S. M., Baum, B. R., Johnson, D. A., and Arnason, J. T. 2003. Direct shoot regeneration from leaf segments of mature plants of Echinacea Purpurea (L.) Moench. In Vitro Cell. Dev. Biol. Plant. 39:505509.CrossRefGoogle Scholar
Misra, P. and Datta, S. K. 2001. Direct differentiation of shoot buds in leaf segments of white marigold (Tagetes erecta L). In Vitro Cell. Dev. Biol. Plant. 37:466470.CrossRefGoogle Scholar
Murashige, T. and Skoog, F. 1962. A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol. Plant. 15:473497.CrossRefGoogle Scholar
Nikam, T. D. and Shitole, M. G. 1998. In vitro culture of Safflower L. cv. Bhima: initiation, growth optimization and organogenesis. Plant Cell Tiss. Org. Cult. 55:1522.CrossRefGoogle Scholar
Nin, S., Morosi, E., Sahiff, S., and Bennici, A. 1996. Callus cultures of Artemisia absinthium L.: initiation, growth optimization and organogenesis. Plant Cell Tiss. Org. Cult. 45:307320.CrossRefGoogle Scholar
O'Sullivan, B. M. 1985. Investigations into crofton weed (Eupatorium adenophorum) toxicity in horses. Aust. Vet. J. 62:3032.CrossRefGoogle ScholarPubMed
Pereira, A. M. S., Bertoni, B. W., Gloria, B. A., Araiyo, A. R. B., Janauario, A. H., Loureno, M. V., and Franca, S. C. 2000. Micropropagation of Pathomorphe umbellate via direct organogenesis from leaf explants. Plant Cell Tiss. Org. Cult. 60:4753.CrossRefGoogle Scholar
Saini, J. P. 2002. Chemical control of Eupatorium (Chromolaena adenophorum L.) in Himachal Pradesh. Indian J. Weed Sci. 34:156157.Google Scholar
Sharma, O. P., Dawra, R. K., Kurade, N. P., and Sharma, P. D. 1998. A review of the toxicosis and biological properities of genus Eupatorium . Nat. Toxins. 6:114.3.0.CO;2-E>CrossRefGoogle Scholar
Ślusarkiewicz–Jarzina, A., Ponitka, A., and Kaczmarek, Z. 2005. Influence of cultivar, explant source and plant growth regulator on callus induction and plant regeneration of Cannabis sativa L. Acta Biol. Cracov. Bot. 47:145151.Google Scholar
Staden, J. and Bennett, P. H. 1991. Gall formation in crofton weed. Differences between normal stem tissue and gall tissue with respect to cytokinin levels and requirements for in vitro culture. S. Afr. J. Bot. 57:246248.CrossRefGoogle Scholar
Sudarshana, M. S. and Shanthamma, C. 1991. In vitro regeneration from excised leaves of Flaveria trinervia (Sprengel) C Mohr. Plant Cell Tiss. Org. Cult. 27:297302.CrossRefGoogle Scholar
Swanson, S. M., Mahady, G. B., and Beecher, C. W. W. 1992. Stevioside biosynthesis by callus, root, shoot and rooted-shoot cultures in vitro. Plant Cell Tiss. Org. Cult. 28:151157.CrossRefGoogle Scholar
Vanegas–Espinoza, P. E., Cruz–Hernández, A., Valverde, M. E., and Paredes-López, O. 2002. Plant regeneration via organogenesis in marigold. Plant Cell Tiss. Org. Cult. 3:279283.CrossRefGoogle Scholar
Wan, Z. X., Zhu, J. J., and Qiang, S. 2001. The pathogenic mechanism of toxin of Alternaria alternata (Fr.) Keissler to Eupatorium adenophorum Spreng. J. Plant Res. Environ. 10:4750.Google Scholar
Xu, Z. Q. and Jia, J. F. 1996. Callus formation from protoplasts of Artemisia sphaerocephala Krasch and some factors influencing protoplast division. Plant Cell Tiss. Org. Cult. 44:129134.CrossRefGoogle Scholar
Zhao, D., Xing, J., Li, M., Lu, D., and Zhao, Q. 2001. Optimization of growth and jaceosidin production in callus and cell suspension cultures of Saussurea medusa . Plant Cell Tiss. Org. Cult. 67:227234.CrossRefGoogle Scholar
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