Advances in the Micropropagation and Genetic Transformation of Abelmoschus esculentus (L.) Moench for Insect Resistance

Introduction

 Agricultural biotechnology has the potential to enhance crop yield¹. Researchers have developed various techniques to boost crop production and make agriculture more sustainable in the environment². Through genetic engineering, several crop varieties for drought resistance, insect resistance, herbicide tolerance and tolerance to salinity have been developed³. Crops that have been genetically modified (GM) have substantially contributed to improving global food security as well as poverty alleviation⁴. Plant transformation technology has grown into a flexible platform in the area of plant biology, with Bacillus thuringiensis (Bt)-producing transgenic crops being commercialized throughout the world to replace insect pesticides for pest control⁵. Many studies employing B. thuringiensis (Bt) formulations for the biocontrol of the insect pests of A. esculentus were published and have confirmed their effective control along with a significant reduction in the plant population. The major pests that caused damage are Pectinophora gossypiella, Amrasca biguttula, Syllepte derogata, Earias vittella, Acontia intersepta, Aphis gossypii, Mylabris, Odontotermes obesus, Nezara viridula, Bagrada cruciferarum, Helicoverpa armigera, Spodoptera litura, Tetranychus urticae, Tetranychus cinnabarinus and Dysdercus koenigii^6,7,8,9,10. Based on their structure of protein and genetic variation, B. thuringiensis genes were divided into three different classes: CryI, CryII, and CryIII¹¹. CryI toxins are found to be deadly against the Lepidoptera insects, whereas CryIII acts against Coleoptera insects¹². An attempt has been made to transfer the Bt Cry1Ac gene into A. esculentus plant to confer a high level of resistance against natural pests and improve the crop yield without affecting the environment^13,14,15.

A. esculentus (L.) (Lady’s finger) is an annual or perennial herb that grows in the tropics, sub-tropics and milder climates, belonging to the family Malvaceae. The plant grows to approximately 2 m tall with heart-shaped leaves that are 10–20 cm in length and breadth and palmately lobed with 5–7 lobes. The petals range in colour from white to yellow, and the blooms are big. Seed pods range in length from 3 to 10 inches and taper towards the tip¹⁶. Immature seed pods of A. esculentus are grown and consumed worldwide as a source of vital nutrients and fibre in the human diet. A. esculentus is commonly known by a variety of names across the world, such as Lady’s Finger, ochro, quiabo and okra. In addition, it is called quimbombó in Spain, gombo in French, bamia or bamya in the eastern Mediterranean, bamies in Arab countries and bhindi in India¹⁷. This crop was first cultivated in Ethiopia, Sudan and other north-eastern African countries. In terms of pod consumption, India is at the top. At present, it is widely grown in many countries with its distribution spread throughout Africa, Egypt, Asia, America and southern Europe^{18, 19}. Protein-rich fruits and seeds are low in calories and abundant in vitamins A and C, and iron, glucose, calcium and enzymes^{20, 21}. A. esculentus is a plant of high medicinal importance whose fruits, leaves, roots and seeds are used for several treatments²². Soluble fibres in the form of gums and pectin, which help reduce serum cholesterol, lower risks of cardiovascular diseases and are also used as a substitute for plasma for expanding the volume of blood; the pharmaceutical applications include thickeners in oral liquids, disintegrates in tablets, binders, protective colloids in suspensions and gelling agents in gel²³. The mucilaginous texture offers healing properties for ulcer and may also help reduce acid reflux^{24, 25}. It also helps in cardiovascular and gastrointestinal health, and has antioxidant and anticancer properties and also the potential benefit against SARS-CoV-2^{26, 27, 28, 29}. A. esculentus fibre has high cellulose content and may be utilised as a raw cellulosic resource in the cellulose-based sectors. Yellowing and photochemical deterioration is caused by the low lignin content. It exhibits dyeing fastness, ultimate strength, and other characteristics due to its large-molecular-weight components³⁰.

Insecticidal sprays, the most popular approach for controlling insect pests, are hard to implement at the initial stages of plant development due to insufficient area and poor efficiency in the growing leaves³¹. Synthetic insecticides, without a doubt, are effective in controlling insect pests, but, when used on a regular basis, can pollute the environment, cause pests to develop resistance to these over a period of long exposure and also lead to the emergence of secondary pests³². Pest management solutions that are environmentally friendly are in high demand across the world. Pest control using varietal resistance has been identified as one of the most economical and ecologically safe methods. Insect-resistant cultivars can be developed and utilized as sole control strategies working in tandem with biological, chemical and cultural control methods to reduce the spreading of pest insects^{33, 32, 34}.

Farmers greatly rely on chemical pesticides to combat the pests of A. esculentus. Synthetic pesticides used in vegetable crops are causing enormous problems to the environment. Precisely vegetable crops use around 13–14% of all pesticides used in India³⁵. As the young pods of A. esculentus are picked on consecutive days, many environmental issues with spraying can result from the reliance on the use of synthetic insecticides. The pesticide residues from vegetable market have been confirmed in A. esculentus that contains toxic pesticides such as acetamiprid and thiamethoxam³⁶. Therefore, there is an urgent need to develop an alternative strategy of eco-friendly pest control in A. esculentus⁷.

Effects caused by the chemical pesticides

Non-target species, such as pests, predators, and parasites, can be harmed by chemical application, and the loss of these useful organisms can disrupt natural biological balances³⁷. Pollinating insects such as honeybees and others can be affected by these pesticides. Sprays and vapour drifts during application along with careless discharge can also cause severe damage and residual problems in crops, livestock, waterways and the environment, resulting in the extinction of animals and fish³⁸. Residues in foods of humans and livestock can be a result of the direct application of a chemical pesticide on the food source, and the presence of pollutants in the environment or their transmission and bio-magnification along the food chain³⁹. Pesticides that have leached into the ground can also contaminate the water supply. Overuse and improper use of the chemical pesticides might result in the pests’ species susceptibility, while its excessive exposure can result in poisoning and other health risks for operators if proper handling protocols are not followed.

Insecticides, herbicides, fungicides, nematicides, rodenticides, and miticides are some of the chemical pesticides used in the chemical approach in okra. The most efficient pesticides for okra shoot borer (Earias vitella) control were determined to be fenvalerate (0.01%) and thiodicarb (0.15%)⁹⁶. Several pesticides pose risks to people, plants, and the environment⁹⁷. The length of chemical exposure, toxicity, dosage, and route of entry into the body are some variables that affect pesticide toxicity in humans⁹⁸. India has investigated more than 400 genotypes for resistance to the shoot and fruit borer using chemical pesticide, however, none of these lines could be categorized as highly resistant or immune⁹⁹.  In okra, traditional and mutation breeding strategies to breed for disease resistance have actually rarely been successful. Because okra germplasm lacks sources of resistance to insect pests and diseases, genetic improvement through traditional plant breeding takes a long time¹⁰⁰. However, with the development of genetic engineering techniques, it is now possible to add a variety of genes for disease resistance, insect pest resistance, and nutritional enrichment to this crop. Transgenic plants produced by inserting the genes of insecticidal proteins are showing promising results for effective pest control while causing no harm to beneficial insects.

Biotechnological studies on A. esculentus

Tissue culture studies

Plant tissue culture is an important field of study for the current transformation procedures for efficient gene transfer. Successful tissue culture protocols are required for Agrobacterium-mediated transformation. Tissue culture helps in the selection of explants since each responds differently due to differences in their endogenous hormone level and the efficiency with which transformed plants regenerate. It is a crucial parameter for selecting appropriate plantlets⁴⁰. A critical function is played by the plant growth regulators (PGR) in the development of in vitro plants⁴¹. Their advancement and growth depend upon the type of PGRs, and the concentration varies according to the plant genotype, as one genotype’s growing condition may not favour another⁴²; so, it is a must to standardise the protocol. The exposure to these PGRs also varies according to the genotype that is used, and media composition determines the regeneration efficiency. The addition of exogenous nutrients such as vitamins, amino acids, inorganic nutrients, casein hydrolysate, proline and glutamine^{43, 44} exhibit a greater influence on somatic embryogenesis and in shoot regeneration. Temperature and light and dark exposure time also assume a major function in the in vitro propagation of tissue culture plants. Standardisation of these parameters paves the way for the development of the different transformation techniques and the development of a successful genetically transformed plant.

A successful transformation of A. esculentus is a primary requirement for the establishment of an effective tissue culture procedure^{45, 46}. But, only a few tissue culture reports on it are available so far. The association of auxin [indole-3-acetic acid (IAA)] and GA₃ on A. esculentus for the induction of adventitious root by stem cuttings was studied in a medium supplemented with both IAA and GA⁴⁷. For callus formation and seedling establishment, different explants (hypocotyl, cotyledon, cotyledonary node and leaf segment) were employed. In the presence of naphthalene acetic acid (NAA) or indoleacetic acid, callus and root differentiation were developed. On the cotyledon and cotyledonary node explants grown in a media enriched with benzyl adenine (BAP) and naphthalene adenine (NAA), shoots were generated⁵¹. In another study on A. esculentus, callus was induced from hypocotyl and cotyledonary axil explants on kinetin-supplemented MS medium and rejuvenation of plants by supplementing benzyl adenine was observed⁴⁸. Following this, Haider et al. (1993) also regenerated the whole plant from the hypocotyl callus culture with a combination of BAP and NAA treatment⁴⁹.

Ganesan et al. (2007) have also reported a tissue culture study on A. esculentus; they devised a method for generating somatic embryos and regenerating plants using hypocotyl explants⁵⁰. They used 2,4-dichlorophenoxy acetic acid and NAA acid for somatic embryogenesis; on MS medium enriched with BAP and gibberellic acid (GA₃), plants were regenerated. Kabir et al. (2008) reported that the hypocotyl and leaf disc showed 95% callus induction with the combination of BAP and NAA followed by 60.82% highest regenerated shoots from the callus⁵¹. Another study using hypocotyl and leaf disc was conducted in the same year, and it was discovered that the combination of NAA and thidiazuron (TDZ) produced the most morphogenic callus, with 80% regeneration from callus in the medium containing BAP and IBA (indole-3-butyric acid)⁵². TDZ showed better multiple shoot regeneration through direct regeneration method⁵³. Direct regeneration of apical shoot showed better response with the combination of IBA and NAA⁵⁴.

A recent study utilizing A. esculentus cotyledonary leaf explants revealed a highly efficient procedure for somatic embryogenesis and rejuvenation.  When L-glutamine and casein hydrolysate were added to the MS medium, the somatic embryo maturation and plant regeneration frequency were enhanced, along with successful somatic embryogenesis from A. esculentus cotyledonary leaf explants and the genomic integrity of regenerated plants being affirmed using ISSR markers⁴³. A summary of the reports on A. esculentus regeneration in vitro is provided in Table 1.

Table 1: Plant tissue culture and Biotechnological studies reported in A. esculentus.

S.No.	Explant	Type of culture	Medium	Phytohormones	References
1.	Hypocotyl and leaf segment	Callus-induced organogenesis	MS basal medium	0.5 mg/l BAP and 2.0 mg/l NAA for callus induction and 2.0 mg/l BAP + 0.1 mg/l IAA and 2.0 mg/l BAP + 0.5 mg/l NAA gave the most effective for plant regeneration from callus	(51)
2.	Hypocotyl and leaf disc	Callus-induced organogenesis	MS basal medium	2.0 mg/l NAA plus 0.5 mg/l TDZ for callus and 2.0 mg/l BAP plus 0.1 mg/l IBA for shoot regeneration	(52)
3.	Hypocotyl	Somatic embryogenesis through suspension cultures	MS basal medium	2.0 mg dm–3 2,4-D and 1.0 mg dm–3 kinetin in suspension culture and 0.2 mg dm–3 BAP and 0.2 mg dm–3 gibberellic GA3 on half strength MS medium for regeneration	(50)
4.	Apical shoot	Direct regeneration	MS basal medium	1.0 mg/l IBA and 0.5 mg/l NAA were found to be most effective	(54)
5.	Hypocotyl and cotyledonary axil	Callus culture	MS basal medium	Benzyl adenine (BA) (1.0 mg/l) showed rapid callus induction and presence of 10 and 20 mg/l of silver nitrate to BA (1.0 mg/l) and NAA (1.0 mg/l) significantly increased calli producing multiple shoots	(48)
6.	Stem cutting	Root formation	indole-3-acetic acid, (IAA) and GA3	At concentrations (10 mg/l IAA + 5 mg/l GA3) the number of roots formed was greater than the sum of roots formed in the individual treatments	(47)
7.	Hypocotyl, cotyledon, cotyledonary node and leaf segment	Callus formation and root differentiation	MS basal medium	Shoot, root and callus development on cotyledonary node explant cultured on MS medium supplemented with NAA (1.0 mg/l) and BA (1.0 mg/l)	(45)
8.	Hypocotyl explants	Direct and indirect organogenesis	MS basal medium	1–3 mg/l benzyl adenine (BA) and 0.1–0.3 mg/l NAA induced direct shoot organogenesis and indirect shoot organogenesis 0.1–0.3 mg/l BA and 1–3 mg/l NAA supported callus growth followed by addition of 1 mg/l BA developed shoots from callus	(49)
9.	Cotyledonary Node	Direct organogenesis	MS basal medium	0.01 mg/l TDZ found to be best for multiple shoot induction and 0.5 mg/l IBA showed successful root induction in excised micro-shoots	(53)
10.	Cotyledonary leaf	Somatic embryogenesis	MS basal medium	1.5 mg/l 2,4-D and 1 mg/l NAA along with 400 mg/l l-glutamine, 300 mg/l casein hydrolysate and half strength MS medium 1 mg/l BAP and 0.5 mg/l GA3 were used for regeneration and shoot elongation	(43)
11.	Seed embryo	Agrobacterium-mediated transformation	MS basal medium	MS salts, B5 vitamins, zeatin 2 mg/l, agar 0.8%, sucrose 3%, kanamycin 50 mg/l, cefotaxime 500 mg/l and cry1Ac gene used for Agrobacterium transformation	(13)
12.	Seeds	Agrobacterium-mediated transformation	MS basal medium	MS basal medium containing 250 mg/l cefotaxime and 15 mg/l BASTA. Agrobacterium tumefaciens strain EHA105 harbouring pCAMBIA 1301 were used for transformation	(78)
13.	Seed embryo	Agrobacterium-Mediated Genetic Transformation	MS basal medium	Agrobacterium tumefaciens strain EHA105 was carrying cry1Ac gene against fruit and shoot borer, CaMV 35S as promoter and nptII as plant selectable marker gene	(92)
14.	Pre-cultured seeds	Agrobacterium-Mediated Genetic Transformation	MS basal medium	Agrobacterium-mediated transformation was performed using LBA4404 strain harbouring the binary vector pBinAR carrying cry3a gene under the control of CaMV35s promoter and npt II gene as a selectable marker	(93)
15.	Seed embryo	Agrobacterium-Mediated Genetic Transformation	MS basal medium	cry1Ac gene was borne on the T-DNA of one plasmid while nptII and uidA (GUS) marker genes	(14)

Bacillus thuringiensis (Bt) crystal protein gene

Bacillus thuringiensis (Bt) is the bacteria that produces Bt toxin. Cloning and transformation of the Bt toxin genes were done and introduced into host plants, where they were expressed and provided resistance to insects without the need for insecticides⁵⁵. B. thuringiensis (Bt) crystal proteins have a low toxicity to vertebrates towards non-target organisms, making them an environment friendly alternative to conventional insecticides. They have an essential part in the development of pest resistance being an important tool in modern biotechnology for creating transgenic plants for integrated pest management strategies, where these techniques are utilized with maximum efficiency^{56, 57, 58}. The initial case reported for pesticide resistance was in 1948, and that synthetic insecticide (DDT) was used against insect pests, and six years into its implementation, the population of insecticide-resistant bugs had skyrocketed⁵⁹. Using Agrobacterium tumefaciens strain (C58Clrif) bearing the pGV3850: pAK1003 Ti plasmid, Feldmann and Marks (1987) successfully established the first Agrobacterium-mediated transformation in Arabidopsis thaliana⁶⁰. In order to eliminate specific insect pests as well as reduce the reliance on chemical pesticides, transgenic crops are developed by transforming B. thuringiensis (Bt) toxins into the host plant⁶¹. Several transformation works have been carried out by researchers by developing various transgenic plants, including crops of vegetables such as brinjal, tomato, cotton, radish, snake gourd and soybean^{62, 63, 64, 65, 66, 67, 68, 69}. The role of Bt toxins (δ-endotoxins) inside the insect is as follows: solubilization in the midgut once ingested and activation of the gut protease enzymes, cleaving the proteins into a lesser-polypeptides; then, the epithelial cells in the mid gut surface get bound in specific target sites by these toxins and kill the pest^{70, 71, 72, 73, 74}. The most often utilised Bt toxins belong to the Cry1A family, especially Cry1Ac in transgenic Bt cotton and Cry1Ab in transgenic Bt corn⁷⁵.

Agrobacterium tumefaciens-mediated transformation of A. esculentus

It is critical to develop Agrobacterium-mediated transformation in A. esculentus because it may be utilised to introduce Bt genes into this crop for pest resistance. A. esculentus has a significant set of chromosomes (2n=130) in its genome^{76, 77}. Both A. esculentus and cotton (Gossypium sp.) are members of the Malvaceae family; however, only very few transformation works have been done on A. esculentus⁷⁸, whereas many have been carried out on genetic transformation of cotton^{79, 80, 81, 82, 83, 84, 85, 86, 87, 88}. The protocols are genotype-dependent throughout the time periods for the varying transgenic plant’s regeneration. Transformation mediated by A. tumefaciens can aid in DNA transfer to organisms on a wide range alongside several monocot and dicot taxa^{89, 90, 91}.

The first report on Agrobacterium-mediated transformation method for A. esculentus using transgenic Bt plants exhibited tolerance to Earias vittella, the target pest that was a fruit and shoot borer of A. esculentus (Narendran et al. 2013) (table 1).

Narendran et al. (2013) in their study used CAMBIA 2300, a plasmid containing the Cry1Ac gene that was regulated by an improved CaMV 35S promoter along with a selectable marker for plants in the T-DNA, the gene nptII. Plasmid was introduced into the A. tumefaciens strain EHA105.  Embryos were isolated from A. esculentus and punctured for 2–3 times on the plumule area with a syringe needle (23G, 100, 0.6 9 25 mm), followed by inoculation into a suspension of Agrobacterium (EHA105 carrying the Cry1Ac and nptII genes), and 20 embryos per petri dish were transferred at a time.  Monoclonal antibodies specific to the Cry1Ac protein, coated on an ELISA plate, were used to assess the transgenic plants for Cry1Ac protein expression. The Bt Cry1Ac protein was effectively altered, according to Molecular and Genetic Analysis utilising polymerase chain reaction (PCR) and southern hybridization experiments. The insect bioassay study showed mortality of about 83.33 and 100.00 % in the transgenic fruits¹³.

In another study, successful transformation was done on A. esculentus genotype Arka Anamika. For this experiment, A. tumefaciens EHA 105 was utilised, which carried the binary vector pCAMBIA 1301–bar. Genomic DNA was extracted from 45-day-old transformed A. esculentus leaves. Basta resistance was checked on the transformed and non-transformed control A. esculentus for herbicide tolerance in transgenic plants⁷⁸. Agrobacterium-mediated genetic transformation was used to establish a method for genetic transformation in okra. Explants for transformation were imbibed seed embryos pierced from the meristematic area of the plumule. Cry1Ac gene against fruit and shoot borer, CaMV 35S promoter, and nptII plant selectable marker gene were all carried out by A. tumefaciens strain EHA105. The potential transgenic plants were validated by polymerase chain reaction to amplify the transgene⁹². A study used LBA4404 strain to undergo Agrobacterium-mediated transformations, with the binary vector pBinAR carrying the cry3a gene under the control of the CaMV35s promoter and the npt II gene as a selectable marker. The transformation event, which included sonicating the explants for 3 minutes, vacuum infiltration (750 mm Hg) for 2 minutes in Agrobacterium (pBinAR-cry3a) and co-cultivation in MS medium with acetosyringone (100M) for 3 days, yielded a 12.5-percent transformation efficiency. Polymerase chain reaction (PCR) was used to confirm the presence and integration of the npt II and cry3a transgenes into the A. esculentus genome⁹³.

Agrobacterium-mediated co-transformation was used to create insect-resistant transgenic okra plants expressing the cry1Ac gene that was free of markers. The cry1Ac gene was found on one plasmid’s T-DNA, while the nptII and uidA (GUS) marker genes were found on the T-DNA of second plasmid. The larvae of the shoot and fruit borer, Earias vittella, a significant okra pest, were used to rigorously screen the plants from selected transgenic events in whole plant insect bioassays. Insect bioassays revealed 100 per cent larval mortality without infestation in five of the transgenic events, and 5 to 10% infestation in two others, demonstrating the transgenic plants’ insect resistance¹⁴.

These plant transformation methods hold much promise to develop insect-resistant A. esculentus for target resistance against insect pests. Commercial insecticides will not be as effective as a well-designed genetic transformation method for A. esculentus.

CRISPR/Cas9 transformation

When it comes to plants, the majority of the early CRISPR/Cas9 findings focused on genome editing. Feng et al., reported the CRISPR/Cas9 investigation with targeted site analysis in different plant taxa⁹⁴. Research using Bombyx mori as a model organism for CRISPR/Cas9 technology is driving the evolution of Lepidoptera⁹⁵. These studies provided strong evidence that CRISPR/Cas9 system could be employed in A. esculentus plants for insect resistance. The conventional transgenic technique may lack the inheritance of stability due to the introduced genes may get lost or silent within a few generations, CRISPR/Cas9 system will likely be a promising alternative to the conventional transgenic approach and will deliver good results in the field.

Figure 1: Figure shows the Agrobacterium-mediated gene transfer for insect resistance, confirmation and insect bioassay of A. esculentus in our laboratory Click here to view Figure

Conclusion and Future Perspective

This review gives a detailed report on the works carried out for insect resistance in okra using different genes and the various explants and their response to different auxins and cytokinins for the rapid growth and development, and rejuvenation and transformation of okra through Agrobacterium provide valuable information to the scientific community for further studies in this important vegetable crop. Screening of different genotypes of A. esculentus using in vitro studies will help isolate elite genotypes for future transformation studies. Further, the Agrobacterium-mediated transformation system will be extremely beneficial in introducing useful genes into A. esculentus for improving the yield by providing biotic and abiotic stress resistance. The insect-resistant A. esculentus plant will also significantly minimize synthetic pesticides usage as well as enhance small-scale farmer’s economy who largely depend on this important vegetable crop for their livelihood.

In recent years, and still today, transgenic plants have garnered a lot of media interest. Despite this, the general public still doesn’t fully understand what a GM plant is or the benefits of the technology. Many government agencies have strict regulations in place for transgenic crops. The specifications for a thorough risk assessment of transgenic plants and any related food and feed have been outlined by the European Food Safety Authority¹⁰¹. For more than 15 years, hundreds of millions of people have consumed foods made from transgenic crops without any negative side effects being noted.

Acknowledgement

The authors wish to thank the Entomology Research Institute, Loyola College Chennai – 34, Tamil Nadu, India, for extending necessary support and guidance.

Author’s contribution

All the authors contributed to the study conception and design. Material preparation, data collection and analysis were performed by Dr Melvin A Daniel, Dr V. Duraipandiyan and Dr S. Maria Packiam. The first draft of the manuscript was written by Dr Melvin A Daniel, and all the authors commented on previous versions of the manuscript. All the authors read and approved the final manuscript.

Conflict of Interest

There is no conflict of interest

Funding source

The authors declare that no funds, grants, or other support were received during the preparation of this manuscript.

References

1. Kader AA. Increasing food availability by reducing postharvest losses of fresh produce. Proceedings of the V International Postharvest Symposium 682; 2004. https://doi.org/10.17660/ActaHortic.2005.682.296
2. Hansson SO, Joelsson K. 2013. Crop biotechnology for the environment. J Agric Environ Ethics.26:759-770. https://doi.org/10.1007/s10806-012-9405-z
3. Björnberg KE, Jonas E, Marstorp H, Tidåker P. 2015. The role of biotechnology in sustainable agriculture: Views and perceptions among key actors in the Swedish food supply chain. Sustainability.7:7512-7529. https://doi.org/10.3390/su7067512
4. Qaim M. 2009. The economics of genetically modified crops. Annu Rev Resour Econ.1:665-694. https://doi.org/10.1146/annurev.resource.050708.144203
5. James C. 2015. Global status of commercialized biotech/GM crops: 2014. ISAAA brief.49.
6. Obeng-Ofori D, Sackey J. 2003. Field evaluation of non-synthetic insecticides for the management of insect pests of okra Abelmoschus esculentus (L.) Moench in Ghana. SINET: Ethiop J Sci.26:145-150. https://doi.org/10.4314/sinet.v26i2.18210
7. Praveen P, Dhandapani N. Development of biocontrol-based pest management in tomato, Lycopersicon esculentum (Mill.). Proceedings of the Biological control of Lepidopteran pests: proceedings of the symposium on biological control of Lepidopteran pests; 2002.
8. Aarwe, R., Mishra, Y. K., Patidar, S., Shukla, A., & Sharma, A. K. (2021). Population trend and bioefficacy of different insecticides against major insect pests in okra, Abelmoschus esculentus (L.) Moench: A review.
9. Chauhan, V., Vashisht, S. D., Sharma, L., & Gandhi, V. (2021). Assessment of leaf hopper, management of Earias vittella and population dynamics of sucking pests of okra: A review.
10. Sapkal, S. D., Mehendale, S. K., Shinde, B. D., Sanap, P. B., & Chavan, S. S. (2022). Seasonal incidence of major sucking pests on okra.
11. Jouanin L, Bonadé-Bottino M, Girard C, Morrot G, Giband M. 1998. Transgenic plants for insect resistance. Plant Sci.131:1-11. https://doi.org/10.1016/S0168-9452(97)00239-2
12. Carozzi NB. 1997. Advances in insect control: the role of transgenic plants: CRC Press. https://doi.org/10.4324/9780203211731
13. Narendran M, Deole SG, Harkude S, Shirale D, Nanote A, Bihani P, Parimi S, Char BR, Zehr UB. 2013. Efficient genetic transformation of okra (Abelmoschus esculentus (L.) Moench) and generation of insect-resistant transgenic plants expressing the cry1Ac gene. Plant Cell Rep.32:1191-1198. https://doi.org/10.1007/s00299-013-1415-4
14. Deole, S., Padakipatil, S., Sandhya, S. R., Nanote, A., Jadhav, M., Bihani, & Char, B. R. (2021). Development of marker-free insect-resistant transgenic okra (Abelmoschus esculentus Moench) expressing the cry1Ac gene and identification of vector backbone-free events. Physiology and Molecular Biology of Plants, 27(10), 2379-2387. https://doi.org/10.1007/s12298-021-01074-3
15. Chen, Y., Pan, L., Ren, M., Li, J., Guan, X., & Tao, J. (2022). Comparison of genetically modified insect-resistant maize and non-transgenic maize revealed changes in soil metabolomes but not in rhizosphere bacterial community. GM crops & food, 13(1), 1-14. https://doi.org/10.1080/21645698.2022.2025725
16. Jain P, Parkhe G, Jain N, Jain R. 2017. Antidiabetic potential of Abelmoschus esculentus In alloxan-induced diabetic rats. International Interdisciplinary Conference on Science Technology Engineering Management Pharmacy and Humanities.522-528
17. Manohar MS. 1969. Pod development and germination of Bhindi (Abelmoschus esculentus). Exp Agric.5:249-255. https://doi.org/10.1017/S0014479700004506
18. Camciuc M, Deplagne M, Vilarem G, Gaset A. 1998. Okra—Abelmoschus esculentus (Moench.) a crop with economic potential for set aside acreage in France. Ind Crop Prod.7:257-264. https://doi.org/10.1016/S0926-6690(97)00056-3
19. Kumar DS, Tony DE, Kumar AP, Kumar KA, Rao DBS, Nadendla R. 2013b. A review on Abelmoschus esculentus (Okra). Int Res J Pharm App3:129-132.
20. Siemonsma J, Kouame C. 2004. In plant resources of tropical Africa Vegetable. J Nutr.7:21-29.
21. Dantas, T. L., Alonso Buriti, F. C., & Florentino, E. R. (2021). Okra (Abelmoschus esculentus) as a potential functional food source of mucilage and bioactive compounds with technological applications and health benefits. Plants, 10(8), 1683. https://doi.org/10.3390/plants10081683
22. Amin IM. Nutritional Properties of Abelmoschus Esculentus as Remedy to Manage Diabetes Mellitus: A Literature Review. Proceedings of the Proceedings of International Conference on Biomedical Engineering and Technology (ICBET 2011); 2011.
23. Singh K, Kumar A, Langyan N, Ahuja M. 2009. Evaluation of Mimosa pudica seed mucilage as sustained-release excipient. AAPS PharmSciTech.10:1121-1127. https://doi.org/10.1208/s12249-009-9307-1
24. Ameena K, Dilip C, Saraswathi R, Krishnan P, Sankar C, Simi S. 2010. Isolation of the mucilage’s from Hibiscus rosasinensis and Okra (Abelmoschus esculentus linn.) and studies of the binding effects of the mucilage’s. Asian Pac J Trop Biomed.3:539-543. https://doi.org/10.1016/S1995-7645(10)60130-7
25. Collins EM. 2010. An AZ Guide to Healing Foods: A Shopper’s Reference: Conari Press.
26. Adelakun O, Oyelade O, Ade-Omowaye B, Adeyemi I, Van de Venter M. 2009. Chemical composition and the antioxidative properties of Nigerian Okra Seed (Abelmoschus esculentus Moench) Flour. ‎Food Chem. Toxicol.47:1123-1126. https://doi.org/10.1016/j.fct.2009.01.036
27. Mollick MMR, Bhowmick B, Mondal D, Maity D, Rana D, Dash SK, Chattopadhyay S, Roy S, Sarkar J, Acharya K. 2014. Anticancer (in vitro) and antimicrobial effect of gold nanoparticles synthesized using Abelmoschus esculentus (L.) pulp extract via a green route. RSC Advances.4:37838-37848. https://doi.org/10.1039/c4ra07285e
28. Subrahmanyam G, Sushma M, Alekya A, Neeraja C, Harsha HS, Ravindra J. 2011. Antidiabetic activity of Abelmoschus esculentus fruit extract. Int J Res Pharm Chem.1:17-20.
29. Ansori, A. N. M., & Nur, A. (2021). A mini-review of the medicinal properties of okra (Abelmoschus esculentus) and potential benefit against SARS-CoV-2. Indian J Forensic Med Toxicol, 15(1), 852-856.
30. Kumar DS, Tony DE, Kumar AP, Kumar KA, Rao DBS, Nadendla R. 2013a. A review on Abelmoschus esculentus (Okra). Int Res J Pharm App Sci.3:129-132.
31. Nault BA, Taylor AG, Urwiler M, Rabaey T, Hutchison WD. 2004. Neonicotinoid seed treatments for managing potato leafhopper infestations in snap bean. Crop Prot.23:147-154. https://doi.org/10.1016/j.cropro.2003.08.002
32. Wiseman B. 1990. Plant resistance to insects in the southeastern United States: an overview. Fla Entomol.351-358.
33. Javed H, Aziz M, Leghari R. 2009. Resistance in different okra cultivars (Abelmoschus esculentus) against American bollworm (Helicoverpa armigera Hub.). J AHgric Res.47:433-438.
34. Zhou, B., & Li, X. (2021). The monitoring of chemical pesticides pollution on ecological environment by GIS. Environmental Technology & Innovation, 23, 101506. https://doi.org/10.1016/j.eti.2021.101506
35. Agnihotri N. 1999. Supervised trials of pesticides on crops. Pesticide, Safety Evaluation and Monitoring.71.
36. Singh S, Kulshrestha G. 2005. Residues of Thiamethoxam and Acetamiprid, Two Neonicotinoid Insecticides, in/on Okra Fruits (Abelmoschus esculentus L). Bull Environ Contam Toxicol.75:945-951. https://doi.org/10.1007/s00128-005-0841-6
37. Rani, L., Thapa, K., Kanojia, N., Sharma, N., Singh, S., Grewal, A. S. & Kaushal, J. (2021). An extensive review on the consequences of chemical pesticides on human health and environment. Journal of Cleaner Production, 283, 124657. https://doi.org/10.1016/j.jclepro.2020.124657
38. Kumar, R., Sankhla, M. S., Kumar, R., & Sonone, S. S. (2021). Impact of pesticide toxicity in aquatic environment. Biointerface Research in Applied Chemistry, 11(3), 10131-10140.
39. Zhou, S., & Jander, G. (2021). Engineering insect resistance using plant specialized metabolites. Current Opinion in Biotechnology, 70, 115-121. https://doi.org/10.1016/j.copbio.2021.03.005
40. Pal AK, Acharya K, Ahuja PS. 2012. Endogenous auxin level is a critical determinant for in vitro adventitious shoot regeneration in potato (Solanum tuberosum). J Plant Biochem Biot.21:205-212. https://doi.org/10.1007/s13562-011-0092-z
41. Zhou, J., Liu, Y., Wu, L., Zhao, Y., Zhang, W., Yang, G., & Xu, Z. (2021). Effects of Plant Growth Regulators on the Rapid Propagation System of Broussonetia papyrifera Vent Explants. Forests, 12(7), 874. https://doi.org/10.3390/f12070874
42. Bhaskaran S, Smith RH. 1990. Regeneration in cereal tissue culture: a review. Crop Sci.30:1328-1337. https://doi.org/10.2135/cropsci1990.0011183X003000060034x
43. Daniel MA, David RHA, Caesar SA, Ramakrishnan M, Duraipandiyan V, Ignacimuthu S, Al-Dhabi N. 2018. Effect of l-glutamine and casein hydrolysate in the development of somatic embryos from cotyledonary leaf explants in okra (Abelmoschus esculentus moench). S Afr J Bot.114:223-231. https://doi.org/10.1016/j.sajb.2017.11.014
44. Satish L, Rency AS, Rathinapriya P, Ceasar SA, Pandian S, Rameshkumar R, Rao TB, Balachandran S, Ramesh M. 2016. Influence of plant growth regulators and spermidine on somatic embryogenesis and plant regeneration in four Indian genotypes of finger millet (Eleusine coracana (L.) Gaertn). Plant Cell, Tiss Org.124:15-31. https://doi.org/10.1007/s11240-015-0870-8
45. Mangat B, Roy M. 1986. Tissue culture and plant regeneration of okra (Abelmoshus esculentus). Plant Sci.47:57-61. https://doi.org/10.1016/0168-9452(86)90010-5
46. Fufa, N. (2019). Propagation methods of okra (Abelmoschus esculentus) and its application used in vitro plant regeneration”. Acta Scientific Agriculture, 3(2019), 125-130.
47. Bhattacharya S, Bhattacharya N, Malik C. 1978. Synergistic effect of gibberellic acid and indole-3-acetic acid on rooting in stem cuttings of Abelmoschus esculentus Planta.138:111-112. https://doi.org/10.1007/BF00392926
48. Roy M, Mangat B. 1989. Regeneration of plants from callus tissue of okra (Abelmoschus esculentus). Plant Sci.60:77-81. https://doi.org/10.1016/0168-9452(89)90046-0
49. Haider S, Islam R, Kamal A, Rahman S, Joarder O. 1993. Direct and indirect organogenesis in cultured hypocotyl explants of Abelmoschus esculentus (L.) moench [in Bangladesh]. Plant Tissue Cult.3: 85-89
50. Ganesan, M., Chandrasekar, R., Ranjitha Kumari, B. D., & Jayabalan, N. (2007). Somatic embryogenesis and plant regeneration of Abelmoschus esculentus through suspension culture. Biologia plantarum, 51(3), 414-420. https://doi.org/10.1016/j.phymed.2010.05.005
51. Kabir, A. H., Sarker, K. K., Sharmin, S. A., Islam, M. S., & Alam, M. F. (2008). Callus induction and plantlet regeneration in Abelmoschus esculentus (L.) Moench. Agric. Technol, 4, 193-204.
52. Anisuzzaman M, Jarin S, Naher K, Akhtar M, Alam M, Khalekuzzaman M, Alam I, Alam M. 2008. Callus induced organogenesis in okra (Abelmoschus esculents Moench.). Asian J Plant Sci. 7: 677-681.
53. Sharma R, Shahzad A. 2008. Thidiazuran (TDZ) induced regeneration from cotyledonary node explant of Abelmoschus moschatus, A valuable medicinal plant. W J Agri Sci.4:449-452.
54. Dhande G, Patil VM, Raut R, Rajput J, Ingle A. 2012. Regeneration of okra (Abelmoschus esculentus) via apical shoot culture system. Afr J Biotechnol.11:15226-15230.
55. Estrada EAC. 2017. The role of biotechnology in agricultural production and food supply. Cienc Investig Agrar. (En línea).44:1-11.
56. Schnepf E, Crickmore N, Van Rie J, Lereclus D, Baum J, Feitelson J, Zeigler D, Dean D. 1998. Bacillus thuringiensis and its pesticidal crystal proteins. Microbiol Mol Biol reviews.62:775-806. https://doi.org/10.1128/MMBR.62.3.775-806.1998
57. Carriere Y, Ellers-Kirk C, Sisterson M, Antilla L, Whitlow M, Dennehy TJ, Tabashnik BE. 2003. Long-term regional suppression of pink bollworm by Bacillus thuringiensis Proc Natl Acad Sci.100:1519-1523. https://doi.org/10.1073/pnas.0436708100
58. Shelton AM, Zhao J-Z, Roush RT. 2002. Economic, ecological, food safety, and social consequences of the deployment of Bt transgenic plants. Annu Rev Entomol.47:845-881. https://doi.org/10.1146/annurev.ento.47.091201.145309
59. Denholm I, Devine G, Williamson M. 2002. Insecticide resistance on the move. Science.297:2222-2223. https://DOI: 10.1126/science.1077266
60. Feldmann KA, Marks MD. 1987. Agrobacterium-mediated transformation of germinating seeds of Arabidopsis thaliana: a non-tissue culture approach. Mol Gen Genet.208:1-9.
61. Wu K-M, Lu Y-H, Feng H-Q, Jiang Y-Y, Zhao J-Z. 2008. Suppression of cotton bollworm in multiple crops in China in areas with Bt toxin–containing cotton. 321:1676-1678. https://doi.org/10.1126/science.1160550
62. Curtis IS, Nam HG. 2001. Transgenic radish (Raphanus sativus longipinnatus Bailey) by floral-dip method–plant development and surfactant are important in optimizing transformation efficiency. Transgenic Res.10:363-371. https://doi.org/10.1023/A:1016600517293
63. Hu C-Y, Wang L. 1999. In planta soybean transformation technologies developed in China: procedure, confirmation and field performance. In Vitro Cell Dev Biol Plant.35:417-420. https://doi.org/10.1007/s11627-999-0058-1
64. Park B-J, Liu Z, Kanno A, Kameya T. 2005. Transformation of radish (Raphanus sativus) via sonication and vacuum infiltration of germinated seeds with Agrobacterium harboring a group 3 LEA gene from B. napus. Plant Cell Rep.24:494-500. https://doi.org/10.1007/s00299-005-0973-5
65. Yasmeen A, Mirza B, Inayatullah S, Safdar N, Jamil M, Ali S, Choudhry MF. 2009. In planta transformation of tomato. Plant Mol Biol Rep.27:20-28. https://doi.org/10.1007/s11105-008-0044-5
66. Subramanyam K, Arunachalam C, Thaneswari RM, Sulaiman AA, Manickavasagam M, Ganapathi A. 2015. Highly efficient Agrobacterium-mediated in planta genetic transformation of snake gourd (Tricosanthes cucumerina). Plant Cell, Tiss Org.123:133-142. https://doi.org/10.1007/s11240-015-0821-4
67. Eck, J. V., Keen, P., & Tjahjadi, M. (2019). Agrobacterium tumefaciens-mediated transformation of tomato. In Transgenic plants (pp. 225-234). Humana Press, New York, NY. https://doi.org/10.1007/978-1-4939-8778-8_16
68. Bhat, S. G., Arulananthu, G., Rajesh, G., & Ramesh, N. (2020). Agrobacterium-mediated transformation of brinjal (Solanum melongena L.) using fungal resistant gene. Electronic Journal of Plant Breeding, 11(01), 160-168.
69. Kesiraju, K., Mishra, P., Bajpai, A., Sharma, M., Rao, U., & Sreevathsa, R. (2020). Agrobacterium tumefaciens-mediated in planta transformation strategy for development of transgenics in cotton (Gossypium hirsutum L.) with GFP as a visual marker. Physiology and Molecular Biology of Plants, 26(11), 2319-2327. https://doi.org/10.1007/s12298-020-00887-y
70. Ferré J, Van Rie J. 2002. Biochemistry and Genetics of Insect Resistance to Bacillus thuringiensis. Annu Rev Entomol.47:501-533.
71. Knowles BH, Dow JA. 1993. The crystal δ‐endotoxins of Bacillus thuringiensis: Models for their mechanism of action on the insect gut. BioEssays.15:469-476. https://doi.org/10.1002/bies.950150706
72. Morin S, Biggs RW, Sisterson MS, Shriver L, Ellers-Kirk C, Higginson D, Holley D, Gahan LJ, Heckel DG, Carriere Y. 2003. Three cadherin alleles associated with resistance to Bacillus thuringiensis in pink bollworm. Proc Natl Acad Sci.100:5004-5009. https://doi.org/10.1073/pnas.0831036100
73. Azizoglu, U. (2019). Bacillus thuringiensis as a biofertilizer and biostimulator: a mini-review of the little-known plant growth-promoting properties of Bt. Current Microbiology, 76(11), 1379-1385. https://doi.org/10.1007/s00284-019-01705-9
74. Qaim, M. (2020). Bt cotton, yields and farmers’ benefits. Nature Plants, 6(11), 1318-1319. https://doi.org/10.1038/s41477-020-0615-5 (2020)
75. Tabashnik BE, Gould F. 2012. Delaying corn rootworm resistance to Bt corn. J Econ Entomol.105:767-776. https://doi.org/10.1603/EC12080
76. Joshi A, Hardas M. 1956. Alloploid Nature of Okra, Abelmoschus esculentus (L.) Moench. Nature.178:1190. https://doi.org/10.1038/1781190a0
77. Nwangburuka C, Kehinde O, Adegbite O, Denton O. 2011. Mitotic chromosomes in Abelmoschus esculentus (L.) Moench. Ann Biol Res.2:85-90.
78. Manickavasagam M, Subramanyam K, Ishwarya R, Elayaraja D, Ganapathi A. 2015. Assessment of factors influencing the tissue culture-independent Agrobacterium-mediated in planta genetic transformation of okra [Abelmoschus esculentus (L.) Moench]. Plant Cell, Tiss Org.123:309-320. https://doi.org/10.1007/s11240-015-0836-x
79. Finer JJ, McMullen MD. 1990. Transformation of cotton (Gossypium hirsutum) via particle bombardment. Plant Cell Rep. 8:586-589. https://doi.org/10.1007/BF00270059
80. Katageri I, Vamadevaiah H, Udikeri S, Khadi B, Kumar PA. 2007. Genetic transformation of an elite Indian genotype of cotton (Gossypium hirsutum) for insect resistance. Curr Sci.1843-1847.
81. Kumar S, Dhingra A, Daniell H. 2004. Stable transformation of the cotton plastid genome and maternal inheritance of transgenes. Plant Mol Biol.56:203-216. https://doi.org/10.1007/s11103-004-2907-y
82. Lei J, Li X, Wang D, Shao L, Wei X, Huang L. 2012. Agrobacterium-mediated Transformation of Cotton Shoot Apex with SNC1 Gene and Resistance to Cotton Fusarium Wilt in T 1 Generation. Cotton Genomics and Genetics.3.1-7(1) https://doi: 10.5376/cgg.2012.03.0001
83. Sunilkumar G, Rathore KS. 2001. Transgenic cotton: factors influencing Agrobacterium-mediated transformation and regeneration. Mol Breeding.8:37-52. https://doi.org/10.1023/A:1011906701925
84. Tohidfar M, Ghareyazie B, Mosavi M, Yazdani S, Golabchian R. 2008. Agrobacterium-mediated transformation of cotton (Gossypium hirsutum) using a synthetic cry1Ab gene for enhanced resistance against Heliothis armigera. Ir J Biotech.6:164-173. https://doi.org/10.1007/s11240-004-6155-2
85. Yuceer SU, Koc N. 2006. Agrobacterium-mediated transformation and regeneration of cotton plants. Russ J Plant Physiol.53:413-417.
86. Zhang B. 2013. Agrobacterium-mediated transformation of cotton. In: Transgenic Cotton. Springer. p. 31-45. https://doi.org/10.1007/978-1-62703-212-4_3
87. Blaise, D., Manikandan, A., Verma, P., Nalayini, P., Chakraborty, M., & Kranthi, K. R. (2020). Allelopathic intercrops and its mulch as an integrated weed management strategy for rainfed Bt-transgenic cotton hybrids. Crop Protection, 135, 105214. https://doi.org/10.1016/j.cropro.2020.105214
88. Liu, Z., Wang, G., Zhang, Z., Zhang, C., Li, H., Wu, T. & Chen, D. (2022). Recovery Characteristics of Cry1Ac Endotoxin Expression and Related Physiological Mechanisms in Bt Transgenic Cotton Squares after High-Temperature Stress Termination. Agronomy, 12(3), 668. https://doi.org/10.3390/agronomy12030668
89. Anderson A, Moore L. 1979. Host specificity in the genus Agrobacterium. Phytopathology.69:320-323.
90. Nester EW, Gordon MP, Amasino RM, Yanofsky MF. 1984. Crown gall: a molecular and physiological analysis. Annu Rev Plant Physiol.35:387-413. https://doi.org/10.1146/annurev.pp.35.060184.002131
91. Van Wordragen MF, Dons HJ. 1992. Agrobacterium tumefaciens-mediated transformation of recalcitrant crops. Plant Mol Biol Rep.10:12-36. https://doi.org/10.1007/BF02669262
92. Menon, R., Sarao, N. K., & Pathak, M. (2018). In planta Agrobacterium-Mediated Genetic Transformation in Okra (Abelmoschus esculentus (L.) Moench). Applied Biological Research, 20(3), 221-227. https://doi.org/10.5958/0974-4517.2018.00030.7
93. Anandan, R., Tohidfar, M., Kumar, P. S., Kamaraj, A., & Vijayakumar, N. (2019). Transgenic okra plants expressing a cry3a gene for fruit borer (Helicoverpa armigera) resistance-an in planta transformation approach. Journal of Pharmacognosy and Phytochemistry, 8(3), 79-83.
94. Feng Z, Mao Y, Xu N, Zhang B, Wei P, Yang DL, Wang Z, Zhang Z, Zheng R, Yang L, Zeng L, Liu X, Zhu JK (2014) Multigeneration analysis reveals the inheritance, specificity and patterns of CRISPR/cas-induced gene modification in Proceedings of the National Academy of Sciences of the United States of America 111(12):4632-4637. doi:10.1073/pnas.1400822111
95. Taning, C. N. T., Van Eynde, B., Yu, N., Ma, S., & Smagghe, G. (2017). CRISPR/Cas9 in insects: Applications, best practices and biosafety concerns. Journal of insect physiology, 98, 245-257.
96. Dhamdhere, S.V., J. Bahadur, and U.S. Mistra. 1984. Efficacy of some foliar insecticides against Earias vitella fabricius infesting okra, Abelmoschus esculentus (L.) Moench. Journal of Entomological Research (India). 8(2):128-131.
97. Delzell, E. and S. Grufferman. 1985. Mortality among white and non-white farmers in North Carolina, 1976-1978. American Journal of Epidemiology. 121:391-402.
98. Alimi, T. (2004). Use of cultural practices, economic impact of insecticide use, and awareness and practice of insecticide safety precaution on Okra production. Journal of vegetable crop production, 10(1), 23-35.
99. Dhankhar BS, Singh R, Kumar R, Kumar S (2009) Genetic improvement. In: Dhankhar BS, Singh R (eds) Okra handbook: global production, processing and crop improvement. HNB Publishing, New York, pp 125–158
100. Rajamony L, Chandran M, Rajmohan K (2006) In Vitro embryo rescues of interspecific crosses for transferring virus resistance in okra (Abelmoschus esculentus (L.) Moench). Acta Hort (ISHS) 725:235–240
101. Guidance document of the genetically modified organisms for the risk assessment of genetically modified plants and derived food and feed. EFSA J. 2004;99:1–94