Journal of
Medicinal Plants Research

  • Abbreviation: J. Med. Plants Res.
  • Language: English
  • ISSN: 1996-0875
  • DOI: 10.5897/JMPR
  • Start Year: 2007
  • Published Articles: 3707

Full Length Research Paper

The effect of growth regulators on two different in vitro-cultured explants of Carapa guianensis

Larisse Lobo de Oliveira
  • Larisse Lobo de Oliveira
  • Laboratório Integrado de Biologia Vegetal, Departamento de Botânica, Centro de Ciências Biológicas e da Saúde, Universidade Federal do Estado do Rio de Janeiro, Brazil.
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Anaíze Borges Henriques
  • Anaíze Borges Henriques
  • Laboratório de Fisiologia do Desenvolvimento Vegetal, Departamento de Botânica, Centro de Ciências e da Saúde, Instituto de Biologia, Universidade Federal do Rio de Janeiro, Brazil.
  • Google Scholar
Andrea Furtado Macedo*
  • Andrea Furtado Macedo*
  • Laboratório Integrado de Biologia Vegetal, Departamento de Botânica, Centro de Ciências Biológicas e da Saúde, Universidade Federal do Estado do Rio de Janeiro, Brazil. Centro de Inovação em Espectrometria de Massas do Laboratório de Bioquímica de Proteínas, Centro de Ciências Biológicas e da Saúde, Universidade Federal do Estado do Rio de Janeiro, Brazil.
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  •  Received: 10 October 2014
  •  Accepted: 14 January 2015
  •  Published: 10 February 2015


Carapa guianensis Aubl. (Meliaceae), known locally as andiroba, is a multi-use species from Amazonia. Andiroba oil is considered an important natural product in the Brazilian market, and international demand is increasing due to its cosmetic and pharmaceutical potential. C. guianensis trees produce seed irregularly over different harvest periods, leading to inconsistent oil production and difficulties with supply. No management plans or protocols have been developed for in vitro or clonal production of Carapa seedlings and the maintenance of genetic resources. The objective of this study was to assess the effect of growth regulators on explants (young leaves, old leaves and apical buds). Explants consisting of leaf segments 1 cm on a side were cultivated in MS medium with and without growth regulators. Evaluation was based on fresh and dry weight of the explants after 20 days. In the media with 2,4-dichlorophenoxyacetic acid (5, 15, 35 or 45 µM), changes were observed in weight and explant appearance (callus). Bud breakage and development of shoots were achieved using 5 µM of 6-benzylaminopurine. Overall, the results showed that 2,4-dichlorophenoxyacetic acid stimulates callus formation on andiroba foliar explants, while 6-benzylaminopurine was superior to thidiazuron for the initial development of shoots.


Key words: Growth regulators, Carapa guianensis, in vitro, tissue culture, organogenesis.


Carapa guianensis Aubl. (Melicaceae), commonly known as andiroba, is a neotropical tree distributed throughout South and Central America, as well as the Caribbean Islands (Cloutier et al., 2007). It is a multi-use species, the main product being seed oil used for medicinal purposes due to its significant limonoid content (Mendonça and Ferraz, 2007; Henriques and Penido, 2014).     The    bio-oils    obtained    from    C. guiannesis seed have physical and chemical properties that make them acceptable renewable diesel fuels (Iha et al., 2014). Andiroba oil is used as an insect repellent (Freire et al., 2006) and in the manufacture of cosmetics, due to its high level of unsaturated triacylglycerols (TAG) (Cabral et al., 2013). Additionally, its wood is valued for the construction of buildings and furniture (Guariguata et al., 2002), and cultivated andiroba trees have the potential to recover degraded land.

The exploitation of C. guianensis is inevitable and is intensifying, mainly in central Amazonia where seed extraction leads to population reduction, as seed dispersal is the main reproductive mode. Evidence indicates that the size of felled logs has been decreasing for decades (Fortini and Zarin, 2011).

Andiroba trees produce seed irregularly over different harvest periods (Tonini et al., 2008). This variability, which does not allow continuous oil production, generates management difficulties, resulting in periods with low seed and oil production (Frankie et al., 1974; McHargue and Hartshorn, 1983). Industries that use C. guianensis need a constant source of homogeneous raw plant material.

Plant regeneration by tissue culture, through either organogenesis or somatic embryogenesis, is a prerequisite for potential clone propagation, genetic transformation and in vitro preservation for germplasm from timber trees, including andiroba (Handley, 1995; Park et al., 1998; Minocha and Jain, 2000). Clone propagation in aseptic conditions is an alternative method of propagation for some medicinal plant species with large-scale production issues, accelerating the conventional propagation process and producing genetically identical plants (Zhou and Wu, 2006).

This study evaluated the effects of growth regulators on C. guianensis foliar explants and apical buds, with a view toward shoot induction and providing guidelines for optimizing andiroba cultivation. 



Seeds of C. guianensis Aublet (andiroba) were collected in the city of Rio de Janeiro (Jardim Botânico do Rio de Janeiro) beneath identified parent trees, with previous authorization from the institution. Seeds, weighing around 20 to 30 g were washed, soaked in water for 24 h, placed in 200 ml plastic bottles with equal volumes of sterile soil fertilized with plant humus, and watered twice a week. Three kinds of explants were used: (a) young leaf explants (less than 1 week old), when they were pink-colored; (b) old leaf explants, when each leaf was at most 2 weeks old and green; and (c) shoot apical buds. The younger leaves for explants were simply cut into three parts (apex, middle and base). Leaf fragments (1 cm2) were obtained from the older green leaves. Apical buds were collected from the same seedlings when the apical segment was still green and soft.

Surface sterilization of explants

The surfaces of the explants from young and old leaves were sterilized with a 50% (v/v) commercial bleach solution for 1.5 min and then washed three times for 1 min each in sterile distilled water. Surface-sterilized explants were placed with their adaxial or abaxial surfaces firmly in contact with the medium in culture flasks.

The apical bud explants were surface-sterilized with a 50% (v/v) commercial bleach solution, for 2 min, followed by a quick dip in 70% (v/v) ethanol solution, and then washed 3 times with sterile distilled water. The exposed ends of the explant were trimmed aseptically and then inoculated on the medium.

Tissue culture

To assay the effectiveness of growth regulators (GR), surface-sterilized leaf explants were inoculated on sterile MS medium (Murashige and Skoog, 1962). The medium was supplemented with 30 g/L sucrose, 7 g/L agar and vitamins, with or without growth regulators (MS 0) (Macedo et al., 1999).

To determine if foliar explants would respond to medium supplemented with one GR at a time, the following GRs were used: indoleacetic acid (IAA) (1, 5, 15, 35 or 45 µM), 6-benzylaminopurine (BA) (1, 5, 15, 35 or 45 µM), 2,4-dichlorophenoxyacetic acid (2,4-D) (1, 5, 15, 35 or 45 µM), and thidiazuron (TDZ) (0.5, 1 or 5 µM) (Figure 1).

Combinations of BA and IAA (1 + 1; 1 + 5 or 5 + 5 µM) were also tested. The plant material was observed for 2 months. For each treatment, 12 explants were used, and the experiments were repeated three times. The explants were placed on the medium with either the abaxial or the adaxial surface turned up. In order to determine if callus obtained from foliar explants would undergo indirect organogenesis, callus explants were subcultivated on control, IAA, BA and TDZ media. Then, calli developed on the lowest and the highest (5 and 45 µM) 2,4-D medium concentrations were subcultivated (Figure 1). Explants cultured for 4 weeks in 2,4-D supplemented medium with callus formation were transferred to fresh MS 0 medium or to medium supplemented with IAA (1 or 5 µM), BA (1 or 5 µM), 2,4-D (1 or 5 µM) or TDZ (0.5, 1 or 5 µM). Explants alive after 4 weeks were subcultivated on MS 0 supplemented with 3 g/L charcoal (Figure 1).

For bud growth experiments, the explants were inoculated in MS medium with IAA, BA, TDZ (1 or 5 µM) or no GR, and with or without 3 g/L charcoal (Figure 1). After 4 weeks, the bud explants were transferred to MS medium supplemented with cytokinins and auxins, in an attempt to achieve organogenesis: IAA (0.5, 1 or 5 µM), BA (0.5, 1 or 5 µM), TDZ (0.5, 1 or 5 µM) and combinations of IAA and BA (1 + 5 µM; 5 + 1 µM; 1 + 1 µM) (Fig. 1). Assays performed on a small number of samples with higher concentrations of IAA and BA (15, 35 or 45 µM) and a combination of IAA and BA (5 + 5 µM) produced brown and dry explants. Therefore, these GR combinations were not tested for bud explants.

 All experiments were performed in a climate-controlled room equipped with white fluorescent lamps (Osram F20T12/CW) (approximately 20 µmol m2 s1 photosynthetically active radiation, PAR). For all treatments, a 16-h photoperiod was used. Cultures were maintained at 25±1°C.



Assessment of leaf explant development

After the culture periods detailed above, the effect of each treatment was evaluated by dry and fresh explant mass. For dry weight, the leaf explants were individually  oven-dried  in  aluminum vessels at 40°C to constant mass and then weighed. To measure fresh weight, the material was removed from the culture flasks and immediately weighed to prevent dehydration.

Assessment of apical bud explants development

After the culture period, the height of explants and leaf length were measured weekly with a ruler. Fresh weight was determined immediately after each explant was removed from the culture flask after 8 weeks of culture; the dry weight was determined after the explant was oven-dried at 40°C to constant mass.


The results were analyzed by Analysis of Variance (ANOVA), followed by Tukey’s test with a significance level set at a = 0.05, using Statistica 7 software for Windows. Means ± standard error (SE) are presented.


A method for organogenesis, either direct or indirect, was developed. Indirect organogenesis involves the production of organs by callus stage, whilst direct organogenesis is related to the formation of organs directly on the surface of cultured intact explants (Us-Camas et al., 2014). The purpose of this study was to produce in vitro shoots as an alternative to sexual propagation for C. guianensis. Sexual propagation is limited by the tendency of andiroba seeds to lose their power of germination soon after harvest, as a result of dehydration. Micropropagation of selected phenotypes of C. guianensis is also desirable since propagation by seed yields high levels of genetic variability, a limiting factor for its commercial use.

Old leaf explants

No callus formation was observed from old leaf-tissue explants cultured in media supplemented with IAA, BA, TDZ or IAA-BA combinations (Figure 2). However, friable whitish callus was formed in all explants cultured in 2,4-D media, except the medium supplemented with  1  µM 2,4-D (Figures 2 and 3). After 4 weeks of observation, a significant difference, according to Tukey’s test, in explant fresh and dry weight was observed when comparing explants cultured in 2,4-D supplemented medium with others cultured in MS 0 or with IAA, BA, TDZ or IAA-BA supplemented medium (Figure 2). No difference was observed between the orientations of explants (abaxial or adaxial surface turned up) in differently supplemented MS medium or MS 0 for 4 weeks.




After calluses were obtained on 2,4-D supplemented medium, the effects of different subculture medium conditions on callus development were investigated (Figure 1). Therefore, calluses obtained from old leaf explants, after 4 weeks of culture in 2,4-D medium (5 or 45 µM of 2,4-D), were transferred to fresh medium. Calluses obtained with 5 µM of 2,4-D and then transferred to medium with 1 or 5 µM of 2,4-D or 0.5, 1 or 5 µM of TDZ survived and showed increase in callus mass (Figures 4 and 5.). These calli acquired a brighter green color (Figure 5). The calli that were subcultivated on MS 0 or MS supplemented with IAA or BA did not survive after 4 weeks. These explants turned brown and became dry. Calli subcultivated on 2,4-D and TDZ that showed no or very few signs of brown parts after 4 weeks were transferred to MS 0 supplemented with 3 g/L of charcoal (Figure 1). Only a few calli (30%) that came from 1 µM 2,4-D produced very small thin roots after 4 weeks (Figure 6). The calli that came from TDZ in all concentrations merely maintained their green color.





Callus obtained with 45 µM of 2,4-D and then transferred to medium with 1 or 5 µM of 2,4-D or 0.5, 1 or 5 µM of TDZ also showed increases in callus mass (Figure 7). The calli that were transferred to MS 0 or MS supplemented with IAA or BA did not survive after 4 weeks. These explants turned brown and became dry, as shown subsequently. Rhizogenesis was not observed in any calli that were first subjected to 2,4-D 45 µM supplemented medium and then transferred to MS 0 supplemented with charcoal, as observed with the callus from 2,4-D. They merely maintained the green callus mass.



Young leaf explants

Young, pink-colored leaf explants were tested to compare the   results   obtained   using  old  leaf  explants,  and  to determine if they would respond to GR better than older leaves from the same plants (Figure 1). However, these explants did not survive more than 4 weeks on any culture medium. After the first week of incubation in all media, explants turned from pink, to a pale yellow to green, and then to brown. There was no difference in response among the three parts of the leaf (apex, middle and base).

Apical shoot bud explants

Apical bud explants were cultured in MS 0 and with cytokinins and auxins to check their development (Figure 1). Bud breakage was 100% successful only in MS 0 supplemented with charcoal, but shoots did not grow longer than 1 cm. A mean of 3 to 4 small shoots were obtained per bud explant (Figure 8). On the other media, no bud explant development was observed.

Shoots developed only on BA supplemented medium (Figure 8), after the initial growth on MS 0. On the other GR supplemented media, the explants did not develop and the shoots maintained the size that they had reached on the first medium (MS 0). No signs of rooting were observed. When the medium was not supplemented with 3 g/L of charcoal, all the explants turned brown and died. The plantlets reached 2.55 ± 0.36 cm in height (mean ± standard error) at the end of 2 months. The leaves reached 3.66 ± 0.38 cm in length (mean ± standard error).


The present results for callus culture contrast with those obtained by Da Costa Nunes et al. (2002) and Rocha and Quoirin (2004). Using a cotyledonary node culture, Da Costa Nunes et al. (2002) found that callus formation in Cedrela fissilis Vell., a woody species of Meliaceae, occurred with the growth regulators naphthalene acetic acid (NAA) and BA. These authors obtained the largest increase of fresh weight in treatments  with  combinations of 6-BA at 1.25, 2.5 and 5.0 µM with 2.5, 1.25 to 5.0 or 5.0 μM of NAA, respectively. Rocha and Quoirin (2004) observed callus formation in mahogany (Swietenia macrophylla King), using BA. However, Vila et al. (2009) reported that 2,4-D induces callus formation in C. fissilis and somatic embryos were formed after 6 months, reducing the concentration of GR in the medium. In this present study, although the entire plant did not regenerate, there was a morphogenic response with the appearance of roots. In general, the absence or reduction of plant growth regulators led to the development and differentiation of somatic embryos or their conversion into plantlets (Merkle, 1995; Hu et al., 2008; Kumar et al., 2008; Yang et al., 2008).

In the present study, C. guianensis leaf explants developed in vitro in different 2,4-D concentrations and showed callus formation. In agreement with our results, Vila et al. (2007) noted that callus mass in zygotic embryo cultures of Melia azedarach L. (Meliaceae) was induced by high concentrations of 2,4-D and Picloran. To differentiate embryos from calli originating from hypocotyls or immature cotyledons in Azadirachta indica A. juss. (Su et al., 1997), it was necessary to use medium supplemented with IAA. Thus, it is evident that callus production in different woody species of Meliaceae is induced by different growth regulators, in varied concentrations. Furthermore, the type of auxins and cytokinins used in the culture media was shown to strongly influence callus formation.

Cytokinins, principally BA, have been reported to be a positive influence to break dormancy from buds and increase its development, as seen on Husain and Anis (2009), where MS medium with 5 µM of BA was the best condition for multiple shoots growth and the increase of length. BA is naturally present in plant tissues, plus its stability in comparison to other cytokinins (Letham and Palni, 1983), may be an explanation for the better response from explants using BA.

In the present work, rhizogenesis was only obtained in callus and not in plantlets. Rhizogenesis was observed when GR was reduced in MS 0 medium. These results agree with that of Basto et al. (2012). It was not possible in the same medium to develop roots from plantlets. A different response was also observed by Da Costa  et  al. (2002), who reported rooting rates of over 87% of C. fissilis node cuttings without growth regulators, and with Milla?n-Orozco et al. (2011) regarding C. odorata shoots from seeds germinated in vitro.

The success of in vitro regeneration relies on the rooting percentage and survival of the plantlets in field conditions. Future studies can focus on achieving rhizogenesis by using media with indole-3- butyric acid (IBA). The IBA improved rooting efficiency and the superiority of IBA in rhizogenesis was also envisaged by other workers (Chiruvella et al., 2011). Rooted plantlets with 4 to 6 fully expanded leaflets will be transferred into plastic cups containing sterilized soil, sand and water to acclimatization tests.

In conclusion, although the induction of callus in C. guianensis has been achieved and bud breakage was inducted, further research is required to confirm the efficiency of embryogenic tissue or bud induction. However, the protocol described here may be suitable for clonal   propagation   and   genetic   transformation  of  C. guianensis. 


The authors are indebted to the Universidade Federal do Estado do Rio de Janeiro (UNIRIO) and to the Instituto de Pesquisas Jardim Botânico do Rio de Janeiro.


We have no conflicting or competing financial interests.


Basto S, Serrano C, Hodson de Jaramillo E (2012). Effects of donor plant age and explants on in vitro culture of Cedrela montana Moritz ex Turcz. Univ. Sci. 17(3):263-271.
Cabral EC, da Cruz GF, Simas RC, Sanvido GB, Gonçalves LDV, Leal RV, Eberlin MN (2013). Typification and quality control of the Andiroba (Carapa guianensis) oil via mass spectrometry fingerprinting. Anal. Methods 5(6):1385-1391.
Chiruvella KK, Mohammed A, Dampuri G, Ghanta RG (2011). In vitro shoot regeneration and control of shoot tip necrosis in tissue cultures of Soymida febrifuga (Roxb.) A. Juss. Plant Cell Tiss. Org. Cult. 21(1):11-25.
Cloutier D, Kanashiro M, Ciampi AY (2007). Impact of selective logging on inbreeding and gene dispersal in an Amazonian tree population of Carapa guianensis Aubl. Mol. Ecol. 16:797-809.
Da Costa Nunes E, Volkmer de Castilho C, Moreno FN, Viana AM (2002). In vitro culture of Cedrela fissilis Vellozo (Meliaceae). Plant Cell Tiss. Org. Cult. 70:259-268.
Fortini LB, Zarin DJ (2011). Population dynamics and management of Amazon tidal floodplain forests: Links to the past, present and future. Forest Ecol. Manag. 261:551-561.
Frankie GW, Baker HG, Opler PA (1974). Comparative phenological studies of trees in tropical wet and dry forest in the lowlands Costa Rica. J. Ecol. 62:881-919.
Freire DCB, Brito-Filha CRC, Carvalho-Zilse GA (2006). Efeito dos óleos vegetais de andiroba (Carapa sp.) e Copaíba (Copaifera sp.) sobre forídeo, pragas de colméias, (Diptera: Phoridae) na Amazônia Central. Acta Amaz. 36:365-368.
Guariguata MR, Arias-LeClaire H, Jones G (2002). Tree seed fate in a logged and fragmented forest landscape, Northeastern Costa Rica. Biotropica 34:405-415.
Handley LW (1995). Future uses of somatic embryogenesis in woody plantation species. In: Jain S, Gupta P, Newton R (eds.), Somatic embryogenesis in woody plants, Vol. 1. Kluwer Academic Publishers, Dordrecht, The Netherlands. pp. 415-434.
Henriques MG, Penido C (2014). The Therapeutic Properties of Carapa guianensis. Curr. Pharm. Des. 20(6):850-856.
Hu Z, Hu Y, Gao HH, Guan XQ, Zhuang DH (2008). Callus production, somatic embryogenesis and plant regeneration of Lycium barbarum root explants. Biol. Plant 52:93-96.
Husain MK, Anis M (2009). Rapid in vitro multiplication of Melia azedarach L. (a multipurpose woody tree). Acta Physiol. Plant 31:765-772.
Iha OK, Alves FC, Suarez PA, Silva CR, Meneghetti MR, Meneghetti SM (2014). Potential application of Terminalia catappa L. and Carapa guianensis oils for biofuel production: Physical-chemical properties of neat vegetable oils, their methyl-esters and bio-oils (hydrocarbons). Ind. Crop Prod. 52:95-98.
Letham DS, Palni LMS (1983). The biosynthesis and metabolism of cytokinins. Ann. Rev. Plant Physiol. 34:163-97.
Macedo AF, Barbosa NC, Esquibel MA, Cechinel Filho V (1999). Pharmacological and phytochemical studies of callus culture extracts from Alternanthera brasiliana. Pharmazie (Berlin) 54(10):776-777.
McHargue LA, Hartshorn GS (1983). Seed and seedling ecology of Carapa guianensis. Turrialba 33:39-404.
Mendonça AP, Ferraz IDK (2007). Óleo de andiroba: processo tradicional da extração, uso e aspectos sociais no estado do Amazonas, Brasil. Acta Amaz. 37:353-364.
Merkle SA (1995). Strategies for dealing with limitations of somatic embryogenesis in hardwood trees. Plant Cell Tiss. Org. Cult. 1:112-121.
Minocha R, Jain SM (2000). Tissue culture of woody plants and its relevance to molecular biology. In: Jain SM, Minocha SC (eds.), Molecular biology of woody plants. Kluwer Academic Publishers, Dordrecht, The Netherlands. pp. 315-339.
Murashige T, Skoog F (1962). A Revised Medium for Rapid Growth and Bio Assays with Tobacco Tissue Cultures. Physiol. Plant 15:473-497.
Park YS, Barrett JD, Bonga JM (1998). Application of somatic embryogenesis in high-value clonal forestry: deployment, genetic control, and stability of cryopreserved clones. In Vitro Cell Dev. Biol. Plant 34:231-239.
Rocha SC, Quoirin M (2004). Calogênese e rizogênese em explantes de mogno (Swietenia macrophylla King) cultivados in vitro. Sci Flor. 14:91-101.
Su WW, Hwang WI, Kim SY, Sagawa Y (1997). Induction of somatic embryogenesis in Azadirachta indica. Plant Cell Tissue Org. Cult. 50:91-95.
Tonini H, Kaminski PE, Costa P, Schwengber LAM (2008). Estrutura populacional e produção de Castanha-do-Brasil (Bertholletia excelsa Bonpl.) e andiroba (Carapa sp.) no Sul do Estado de Roraima. In: Wadt LHO (ed.), Anais/1º. Seminário do Projeto Kamukaia Manejo Sustentável de produtos florestais não-madereiros na Amazônia, Embrapa Acre, Rio Branco, AC. P 182.
Us-Camas R, Rivera-Solís G, Duarte-Aké F, De-la-Pe-a C (2014). In vitro culture: an epigenetic challenge for plants. Plant Cell Tissue Org. Cult. 1:1-15.
Vila SA, Rey HY, Mroginski LA (2007). Factors affecting somatic embryogenesis induction and conversion in "paradise tree" (Melia azedarach L.). J. Plant Growth Regul. 26:268-277.
Yang XM, An LZ, Xiong YC, Zhang JP, Li Y, Xu SJ (2008). Somatic embryogenesis from immature zygotic embryos and monitoring the genetic fidelity of regenerated plants in grapevine. Biol. Plant 52:209-214.
Zhou LG, Wu JY (2006). Development and application of medicinal plant tissue cultures for production of drugs and herbal medicinals in China. Nat. Prod. Rep. 23:789-810.