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Introduction
The castor-oil plant, Ricinus communis L., is an oleaginous plant belonging to the Euphorbiaceae family, which comprises 280 genera. This species has been cultivated for more than 6000 years on the Asian continent, and more recently on the African and American continents (Govaerts et al. 2000; Salihu et al. 2014). R. communis is a non-edible plant, mainly used in chemical, pharmaceutical, and automobile industries, where it has numerous applications (Savy 2005; Barnes et al. 2009; Severino et al. 2010). All parts of this plant contain lectin ricin - one of the most potent lethal natural poisons known - but is particularly concentrated in the seeds and pods (Audi et al. 2005).
In recent years, R. communis oil has acquired importance as a biofuel, due to the possibility of its use in producing biodiesel (Baldwin and Cossar 2009; César and Batalha 2010). R. communis is distributed in tropical and subtropical regions and is also adaptable to temperate zones (Lima et al. 2011). The principal producer countries of R. communis seeds are India, China, and Mozambique; whereas the countries with the highest consumption of the products of this plant are Holland, Japan, and Italy (Faostat 2015). India, China, and Brazil contribute approximately 95 % of the world production of seeds (Sailaja et al. 2008).
Ricinus communis seeds are outstanding for their high oil content, between 40 and 60 %, compared with sunflower (Helianthus annuus L.) seeds with 38 to 48 %, soybean (Glycine max (L.) Merr.) between 18 and 19 %, moringa (Moringa oleifera L.) with 14 to 24 %, neem [Azadirachta indica (Juss)] between 17 and 39 %, and cotton (Gossypium hirsutum L.) with 15 to 19 % (Kittock and Williams 1970; Severino et al. 2006; Nass et al. 2007; Baldwin and Cossar 2009; Martín et al. 2010), a characteristic that makes this plant very attractive as a source of biofuel, particularly biodiesel.
The extensive cultivation of varieties and hybrids of R. communis under different management practices has made the plant vulnerable to biotic and abiotic factors. R. communis plants may lose leaves, seeds and pods for different reasons: damage by pests, diseases, wind, hail, traffic of machinery, and inappropriate use of herbicides and defoliation (Severino et al. 2010). Even though a castor-oil plant can recover from severe defoliation, the damage suffered by the leaves may reduce the production. It is estimated that for 1 m2 of lost leaf area, seed production diminishes by 37.8 g and oil production by 24.4 g (Lakshmamma et al. 2009; Lakshmi et al. 2010; Severino et al. 2010). Continuous sowing of R. communis in the same areas, as well as the lack of intercropping has increased the occurrence of pests and diseases. There are reports that more than 100 species of insects in different parts of the world feed on R. communis and can cause serious damage (Barteneva 1986; Kolte 1995). In India, for example, insect pests caused losses in seed production from 35 to 50 % (Kolte 1995). Integrated pest management programs are therefore important to prevent losses that can affect the economy of producer-countries.
The present literature reviewed focuses on the phytophagous arthropods associated with R. communis in different parts of the world, as well as, their natural enemies and floral visitors. The information was obtained through extense search of scientific literature on these subjects published in the Web of Science database, Ebsco database and Google Scholar, using appropriate key words (e.g. ‘insects on Ricinus communis’ ‘arthropods on Ricinus communis’, ‘pests of Ricinus communis or castor-oil’); the search was conducted until January 2019. Afterwards, the information collected was analyzed from the perspective of the co-evolutionary hypothesis following the approach of literature review analysis of arthropod herbivory on physic nut (Jatropha curcas L.) conducted by Lama et al. (2015) . Specifically, we set out to answer the following questions regarding the arthropods associated with R. communis: (1) What is the diversity of arthropod taxa associated with this plant? (2) In what geographic area does the greatest richness of associated arthropod species occur? (3) What are the parts of the plant most preferred by the herbivorous arthropods? and (4) What mouthpart classes of the arthropods associated with R. communis can be identified? According to the co-evolutionary hypothesis, it would be expected to find greater richness of native arthropod species in Asia and Africa, the origin area of R. communis, in comparison with those areas where this plant has been introduced or cultivated more recently.
Phytophagous arthropods associated with R. communis
Ricinus communis has been considered tolerant and/or resistant to pest attack due to the toxic compounds present in different parts of the plant. Some of the most common compounds found in this plant species are ricin, ricinine, N-demethylricinine, flavonoids, gallic acid, gentisic acid, coumaric acid, syringic acid, cinnamic acid, vanillic acid and rutin, and allergen proteins such as Ric c1 and Ric c3 (Usha Rani et al. 2006; Gahukar 2010; Vandenborre et al. 2011; Usha Rani and Pratyusha 2014). Some of these are toxic compounds that may even have insecticidal or antifeedant properties against insect pests of other crops (Rossi et al. 2012; Amoabeng et al. 2014; Dinesh et al. 2014). Despite the insecticidal properties of R. communis, there are reports of arthropods that feed on several parts of this plant. Ricinine, for example, one of its main alkaloids that has shown insecticidal effect on some insect pests of other plants (Bigi et al. 2004; Liu and Li 2006; Rossi et al. 2012) does not have any detrimental effect on certain specialist phytophagous insects that are common pests of R. communis, such as Achaea janata (L., 1758) (Lepidoptera: Noctuidae), Spodoptera litura (F., 1775) (Lepidoptera: Noctuidae) and others (Prabhakar et al. 2003; Usha Rani and Pratyusha 2014). This is due to the presence of enzymes in the midgut of these insects that are able to degrade toxins and thus breakdown the plants’ natural defenses (Yasur et al. 2009; Usha Rani and Pratyusha 2014).
The arthropod pests of R. communis damage all parts of the plant, including the seeds, where some toxic compounds such as lipases, the alkaloid ricinine (including the protein ricin) and glycosides of ricinoleic, isoricinoleic, stearic and dihydroxystearic acids are even more concentrated (Jena and Gupta 2012). The type of pest and damage varies from place to place; some pests of R. communis can be present in different regions. Table 1 presents information published in the literature on arthropods that attack R. communis.
According to Table 1, 59 % of the arthropod species feed on foliage, 20 % on roots and seedlings, 17 % on flowers, fruits and seeds, and 5 % on stems and branches. The low percentage of arthropods feeding on seeds and roots can be explained in part by the high concentration of ricinine in these parts of the plant (Salihu et al. 2014). To feed on seeds and roots, these arthropods have had to develop highly efficient mechanisms of detoxification (Yasur et al. 2009).
Table 1 Order, family and geographical distribution of the phytophagous arthropod species that attack cultivated Ricinus comunis.
| Order | Family | Species | Geographical distribution | References |
|---|---|---|---|---|
| Roots and seedlings | ||||
| Coleoptera | Curculionidae | Protostropus spp. | Africa | Salihu et al. (2014) |
| Elateridae | Agriotes sp. | Costa Rica | Anónimo (1991) | |
| Scarabeidae | Amphimallon solstitialis (Linnaeus, 1758) | Russia | Arkhangel’Skii and Romanova (1930) | |
| Holotrichia consanguinea Blanchard, 1850 | India | Gahukar (2018) | ||
| Phyllophaga sp. | Colombia and Costa Rica | Anónimo (1991); Londoño-Zuluaga (2008) | ||
| Holochelus aequinoctialis (Herbst, 1790) [= Rhizotrogus aequinoctialis (Herbst, 1790)] | Russia | Arkhangel’Skii and Romanova (1930) | ||
| Diptera | Agromyzidae | Liriomyza trifolii (Burgess, 1880) | India | Anjani et al. (2007) |
| Lepidoptera | Noctuidae | Agrotis ipsilon (Hüfnagel, 1766) | Colombia and Egypt | Mona et al. (2005); Saldarriaga Cardona et al. (2011) |
| Helicoverpa zea (Boddie, 1850) | USA | Wene (1933) | ||
| Spodoptera frugiperda (J. E. Smith, 1797) | Colombia | Saldarriaga Cardona et al. (2011) | ||
| Spodoptera marima (Schaus, 1904) | Brazil | Ribeiro and Costa (2008) | ||
| Sphingidae | Erinnyis ello (Linnaeus, 1758) | Brazil | Ribeiro and Costa (2008) | |
| Orthoptera | Gryllidae | Brachytrupes spp. | Africa | Salihu et al. (2014) |
| Pyrgomorphidae | Chrotogonus spp. | Africa | Salihu et al. (2014) | |
| Zonocerus variegatus (Linnaeus, 1758) | Africa | Salihu et al. (2014) | ||
| Isoptera | Termitidae | Odontotermes obesus (Rambur, 1842) | India | Gahukar (2018) |
| Leaves | ||||
| Coleoptera | Curculionidae | Naupactus glaucus Perty, 1832 [= Pantomorus glaucus (Perty, 1830)] | Brazil | Cavalcante et al. (1974) |
| Diptera | Agromyzidae | Liriomyza sativae Blanchard, 1938 | China | Zhang et al. (2006) |
| Liriomyza subpusilla Frost, 1943 | USA | Wene (1933); Parkman et al. (1989) | ||
| Liriomyza trifolii (Burgess, 1880) | India | Galande et al. (2005) | ||
| Hemiptera | Aleyrodidae | Bemisia tabaci (Gennadius, 1889) | Costa Rica and Africa | Anónimo (1991); Salihu et al. (2014) |
| Trialeurodes ricini (Misra, 1924) (= Trialeurodes rara Singh, 1931) | India and Egypt | Idriss et al. (1997); Sarma et al. (2005); Abdullah and Martin (2007); Raghavaiah (2011) | ||
| Aphrophoridae | Ptyelus grossus (Fabricius, 1781) | Uganda | Darling (1946) | |
| Cicadellidae | Amrasca (Amrasca) biguttula (Ishida, 1913) [= Amrasca biguttula biguttula (Ishida, 1912)] | India | Sharma and Singh (2002); Raghavaiah (2011) | |
| Agallia sp. | Spain | Durán et al. (2010) | ||
| Edwardsiana flavescens (Fabricius, 1794) [= Empoasca flavescens ((Fabricius, 1794)] | India | Jayaraj (1964); Sarma et al. (2005); Lakshmi et al. (2005); Jyothsna et al. (2009) | ||
| Empoasca (Empoasca) solana Delong, 1931 (= Empoasca solana Delong, 1931) | USA | Wene (1933) | ||
| Empoasca sp. | Costa Rica | Anónimo (1991) | ||
| Empoasca sp. | Africa | Salihu et al. (2014) | ||
| Jacobiasca furcostylus (Ramakrishnan y Menon, 1972) | India | Parmar et al. (2006) | ||
| Miridae | Falconia antioquiana Carvalho, 1987 | Colombia | Saldarriaga Cardona et al. (2011) | |
| Polymerus cognatus (Fieber, 1858) (= Poeciloscytus cognatus Fieber, 1858) | Russia | Arkhangel’Skii and Romanova (1930) | ||
| Pentatomidae | Acrosternum pallidoconspersum (Stål, 1858) | Egypt | Jannone (1952) | |
| Nezara viridula (Linnaeus, 1758) | Costa Rica and Egypt | Jannone (1952); Anónimo (1991) | ||
| Pseudococcidae | Paracoccus marginatus Williams and Granara de Willink, 1992 | Cuba | Martínez et al. (2005) | |
| Tingidae | Corythucha gossypii (Fabricius, 1794) | USA, Colombia, Mexico, and Cuba | Miller and Nagamine (2005); Londoño-Zuluaga (2008); Saldarriaga Cardona et al. (2011), López-Guillén et al. (2012) | |
| Lepidoptera | Arctiidae | Amsacta moorei Butler, 1876 | India | Sarma et al. (2005) |
| Amsacta albistriga Walker, 1864 | India | Sarma et al. (2005) | ||
| Pericallia ricini (Fabricius, 1775) | India | Mathur et al. (1994); Neelanarayanan and Indira (2010) | ||
| Spilosoma obliqua Walker, 1855 | India and Pakistan | Singh and Grewal (1982); Khattak et al. (1991); Sarma et al. (2005) | ||
| Dalceridae | Anacraga citrinopsis Dyar, 1927 | Brazil | Lourenção et al. (1989) | |
| Limacodidae | Parasa lepida Cramer, 1799 | India | Raghavaiah (2011) | |
| Lymantriidae | Dasychira sp. | Africa | Salihu et al. (2014) | |
| Euproctis fraterna Moore, 1883 | India | Paul et al. (2000); Suganthy (2010) | ||
| Noctuidae | Achaea janata (Linnaeus, 1758) | India, USA, and China | Hua (1984); Delaya et al. (1985); Basappa and Lingappa (2001); Mau and Kessing (2007) | |
| Helicoverpa armigera (Hübner, 1803-1808) | India and USA | Wene (1933); Ribeiro and Costa (2008) | ||
| Spodoptera cosmioides (Walker, 1858) | Brazil | Bavaresco et al. (2003) | ||
| Spodoptera exigua (Hübner, 1808) | Egypt | Ribeiro and Costa (2008) | ||
| Spodoptera litura (Fabricius, 1775) | India and Pakistan | Lohar et al. (1997); Usha Rani and Rajasekharreddy (2009) | ||
| Spodoptera ornithogalli (Guenée, 1852) [= Spodoptera marima (Schaus, 1904)] | Brazil | Ribeiro and Costa (2008) | ||
| Spodoptera sp. | Costa Rica | Anónimo (1991) | ||
| Nymphalidae | Ariadne merione Cramer, 1779 (= Ergolis merione Cramer, 1779) | India | Ghosh (1914); Sarma et al. (2005) | |
| Saturniidae | Samia ricini (Drury, 1773) | Egypt, India, and Brazil | El-Shaarawy et al. (1975); Negreiros et al. (1998) | |
| Rothschildia jacobaeae Walker, 1855 | Brazil | Ribeiro and Costa (2008) | ||
| Orthoptera | Acrididae | Chrotogonus (Chrotogonus) trachypterus robertsi Kirby & W. F., 1914 (= Chrotogonus robertsi Kirby & W. F., 1914) | India | Sarma et al. (2005) |
| Thysanoptera | Thripidae | Retithrips syriacus (Mayet, 1890) | India | Sarma et al. (2005) |
| Scirtothrips dorsalis Hood, 1919 | India | Patel et al. (2009) | ||
| Zaniothrips ricini Bhatti, 1967 | India | Daniel et al. (1983) | ||
| Acarina | Tetranychidae | Eutetranychus orientalis (Klein, 1936) | India | Ahuja (1994) |
| Eutetranychus sp. | India | Raghavaiah (2011) | ||
| Tetranychus piercei McGregor, 1950 | China | Lui and Lui (1986) | ||
| Tetranychus urticae Koch, 1836 [= Tetranychus telarius (Linnaeus, 1758)] | Morocco and India | Cangardel (1954); Rajasekhar et al. (1999); Raghavaiah (2011) | ||
| Tarsonemidae | Polyphagotarsonemus latus (Banks, 1904) | Belgium | Heungens and Degheele (1986) | |
| Stems and branches | ||||
| Coleoptera | Buprestidae | Sphenoptera sp. | Africa | Salihu et al. (2014) |
| Tenebrionidae | Blapstinus sp. | USA | De Ong (1918) | |
| Hemiptera | Membracidae | Oxyrhachis taranda (Fabricius, 1798) | India | Ali et al. (2006) |
| Lepidoptera | Cossidae | Strigocossus capensis (Walker, 1856) [= Xyleutes capensis (Walker, 1856)] | Africa | Salihu et al. (2014) |
| Flowers, fruits and seeds | ||||
| Coleoptera | Anobiidae | Lasioderma serricorne (Fabricius, 1792) | India and Africa | Hussain and Khan (1966); Salihu et al. (2014) |
| Tenebrionidae | Tribolium castaneum (Herbst, 1797) | Africa | Salihu et al. (2014) | |
| Hemiptera | Cicadellidae | Empoasca sp. | Costa Rica | Anónimo (1991) |
| Miridae | Eurystylus sp. | Africa | Salihu et al. (2014) | |
| Helopeltis sp. | Africa | Salihu et al. (2014) | ||
| Pentatomidae | Nezara viridula (Linnaeus, 1758) | Costa Rica, and USA | Anónimo (1991); Golden and Follett (2006) | |
| Scutelleridae | Calidea sp. | Africa | Salihu et al. (2014) | |
| Lepidoptera | Crambidae | Conogethes punctiferalis (Guenée, 1854) [= Dichocrocis punctiferalis (Guenée, 1854)] | India and Australia | Anonymous (1913); Sharma et al. (1995); Jyothsna et al. (2009); Patel and Patel (2009); Hedge et al. (2009) |
| Noctuidae | Achaea janata (Linnaeus, 1758) | India, USA, and China | Hua (1984); Delaya et al. (1985); Basappa and Lingappa (2001); Mau and Kessing (2007) | |
| Heliothis sp. | Costa Rica | Anónimo (1991) | ||
| Helicoverpa armigera (Hübner, 1803-1808) | India, and USA | Wene (1933); Geetha et al. (2003); Satyanarayana and Sing (2003) | ||
| Spodoptera sp. | Costa Rica | Anónimo (1991) | ||
| Pyralidae | Cadra cautella (Walker, 1863) [= Ephestia cautella (Walker, 1863)] | Africa | Salihu et al. (2014) | |
| Tortricidae | Thaumatotibia leucotreta (Meyrick, 1913) (= Cryptophlebia leucotreta Meyrick, 1913) | Africa | Salihu et al. (2014) | |
A total of 76 species of phytophagous arthropods associated to cultivated plants of R. communis is found worldwide (Table 1). Before the present literature review, the report was of 60 species (Raoof et al. 2003). The arthropods reported in Table 1 belong to eight orders and 38 families; 40 % of these species belong to Lepidoptera, 27 % to Hemiptera, 14 % to Coleoptera and 19 % to other orders. The species that belong to Lepidoptera, Hemiptera and Coleoptera represent 81 % of the total. These phytophagous arthropods are distributed geographically in Asia (39 %), America (34 %), Africa (25 %) and Europe (2 %). As it was supposed, it was not uncommon to find that the greatest richness of arthropods associated to R. communis occurred in Asia and Africa, continents considered as the center of origin of this plant (Govaerts et al. 2000). 63 % of the species had mandibulate mouthparts (Lepidoptera, Coleoptera, Orthoptera, Isoptera and Diptera) and 37 % were piercing-and-sucking mouthpart classes (Hemiptera, Thysanoptera and Acarina).
Of the pests listed in Table 1, the castor semilooper A. janata, the tobacco caterpillar S. litura, the green leafhopper Edwardsiana flavescens (F., 1794) [= Empoasca flavescens (F., 1794)] (Hemiptera: Cicadellidae), the serpentine leafminer Liriomyza trifolii Burgess, 1880, the vegetable leafminer L. sativae Blanchard, 1938 (Diptera: Agromyzidae), the Bihar hairy caterpillar Spilosoma obliqua Walker, 1855 (Lepidoptera: Arctiidae), the shoot and capsule borer Conogethes punctiferalis (Guenée, 1854) [= Dichocrocis punctiferalis (Guenée, 1854)] (Lepidoptera: Crambidae), the cowbug Oxyrhachis taranda (F., 1798) (Hemiptera: Membracidae), and the cotton bullworm Helicoverpa armigera (Hübner, 1803-1808) (Lepidoptera: Noctuidae), among others, are the most devastating pests in Asia. In Africa, the black cutworm Agrotis ipsilon (Hüfnagel, 1776), the armyworm S. exigua (Hübner, 1808) (Lepidoptera: Noctuidae), the stink bug Nezara viridula (L., 1758) (Hemiptera: Pentatomidae), the castor bean whitefly Trialeurodes ricini (Misra, 1924) (= Trialeurodes rara Singh, 1931) (Hemiptera: Aleyrodidae), the red spider mite Tetranychus urticae Koch, 1836 [= Tetranychus telarius (Linnaeus, 1758)] (Acarina: Tetranychidae), among others, are mentioned as the most important. In Central and South-America, the white grub Phyllophaga sp. (Coleoptera: Scarabaeidae), Agrietes sp., Erinnyis ello (F., 1794) (Lepidoptera: Sphingidae), N. viridula, the cotton lace bug Corythucha gossypii (Fabricius, 1794) (Hemiptera: Tingidae), the sucking bug Falconia antioquiana Carvalho, 1987 (Hemiptera: Miridae), S. marima (Schaus, 1904) (Lepidoptera: Noctuidae) and others, are reported as pests of economic importance for R. communis (Varón et al. 2010; Saldarriaga Cardona et al. 2011; López-Guillén et al. 2012).
The principal pests in Brazil are N. viridula, the leafhopper Empoasca spp., some defoliator larvae including S. frugiperda Smith (Lepidoptera: Noctuidae), A. janata, and A. ipsilon, and mites such as T. urticae, and T. ludeni Zacher, 1913 (Acarina: Tetranychidae) (Soares et al. 2001; Ribeiro and Costa 2008). In Colombia, C. gossypii is mentioned as the pest of greatest economic importance in R. communis crops (Varón et al. 2010). In Mexico, C. gossypii, N. viridula, and Tetranychus spp. are reported as the main potential pests of R. communis (López-Guillén et al. 2012) (Table 1).
Phytophagous arthropods found on noncultivated R. communis
There are reports of phytophagous insects mostly found in noncultivated R. communis plants, isolated plants, as well as, in more or less clustered plants or plants growing in urban and suburban areas, and in disturbed landscapes of Egypt, India, Spain, Uganda, USA and other countries (Oshaibah et al. 1986; Singh et al. 1991; Jacob et al. 2000; Pons et al. 2002; Ylla et al. 2008; Boland 2016; Egonyu et al. 2017).
Table 2 presents a total of 20 species of phytophagous arthropods associated with non-cultivated plants of R. communis in the world. These species belong to five orders and 16 families. 60 % of the arthropod species belong to Lepidoptera (30 %) and Hemiptera (30 %), while 40% belong to Coleoptera (20 %) and other orders (20 %). The species that belong to Lepidoptera, Hemiptera and Coleoptera represent 80 % of the total. 59 % of the arthropod species registered in Table 2 are distributed geographically in Asia (32 %) and Africa (27 %), while 41 % are registered in America (32 %) and Europe (9 %). 60 % of the species of arthropods are mandibulate mouthpart (Lepidoptera, Coleoptera, and Orthoptera) and 40 % are piercing-and-sucking mouthpart (Hemiptera and Acarina).
Table 2 Order, family and geographical distribution of the phytophagous arthropod species found on non-cultivated Ricinus communis.
| Order | Family | Species | Geographical distribution | References |
|---|---|---|---|---|
| Leaves | ||||
| Coleoptera | Bostrichidae | Prostephanus truncatus (Horn, 1878) | Mexico | Bourne-Murrieta et al. (2014) |
| Chrysomelidae | Diabrotica graminea Baly, 1886 | Puerto Rico | Woloott (1917) | |
| Scarabaeidae | Lepadoretus sinicus Burmeister, 1855 (= Adoretus sinicus Burmeister, 1855) | USA | McQuate y Jameson (2011) | |
| Scolytidae | Euwallacea sp. | Uganda and USA | Boland (2016); Egonyu et al. (2017) | |
| Hemiptera | Aleyrodidae | Aleurodicus dispersus Russell, 1965 | Cape Verde | Monteiro et al. (2005) |
| Cicadellidae | Amrasca (Amrasca) biguttula (Ishida, 1913) [= Amrasca devastans (Distant, 1918)] | India | Jacob et al. (2000) | |
| Empoasca (Empoasca) kerri Singh-Pruthi, 1940 (= Empoasca kerri Pruthi, 1940) | India | Singh et al. (1991); Jacob et al. (2000) | ||
| Empoasca (Empoasca) motti Singh-Pruthi, 1940 (= Empoasca motti Singh-Pruthi, 1940) | India | Jacob et al. (2000) | ||
| Flatidae | Metcalfa pruinosa (Say, 1830) | Spain | Pons et al. (2002) | |
| Miridae | Apolygus lucorum (Meyer-Dür, 1843) | China | Lu et al. (2010) | |
| Lepidoptera | Arctiidae | Amsacta moorei Butler, 1876 | India | Singh et al. (1989) |
| Cosmopterigidae | Pyroderces rileyi (Walsingham, 1882) (= Sathrobrota rileyi Walsingham, 1882) | Egypt | Oshaibah et al. (1986) | |
| Lymantriidae | Euproctis lunata Walker, 1855 | Bangladesh | Islam et al. (1988) | |
| Noctuidae | Agrotis ipsilon (Hüfnagel, 1766) | Egypt | Younis (1992) | |
| Pyralidae | Phycita diaphana (Staudinger, 1870) | Spain | Huertas Dionisio (2002); Ylla et al. (2008) | |
| Tortricidae | Thaumatotibia leucotreta (Meyrick, 1913) (= Cryptophlebia leucotreta Meyrick, 1913) | South Africa | Kirkman and Moore (2007) | |
| Orthoptera | Acrididae | Melanoplus differentialis (Thomas, 1865) | USA | Spain (1940) |
| Acarina | Tetranychidae | Eutetranychus banksi (McGregor, 1914) | USA | McGregor (1914) |
| Eutetranychus orientalis (Klein, 1936) | Palestine and Egypt | Klein (1936) | ||
| Tetranychus gloveri Banks, 1900 (= Tetranychus quinquenychus McGregor, 1914) | USA | McGregor (1914) | ||
Such insects were observed feeding on leaves of R. communis plants, and even though some species have been reported as pests of R. communis in other countries, most of them cause no considerable damage. However, they have the potential of becoming pests of R. communis if it is cultivated as a monoculture or, R. communis could be a plant host for important pests as the invasive ambrosia beetle Euwallacea sp. (Coleoptera: Curculionidae) (Boland 2016; Egonyu et al. 2017). Among these potential pests are insect and mite species of various families of Lepidoptera, Hemiptera, Orthoptera, and others (Table 2).
Pollinator insects and floral visitors in R. communis
Ricinus communis is a monoecious cross-pollinating plant, cultivated as a hybrid in India, Brazil, China, and other countries because they produce better yields than pure lines or varieties (Moll et al. 1962; Birchler et al. 2003; Reif et al. 2007). Several studies demonstrate that certain species of pollinator insects may improve seed production of R. communis. For example, it is mentioned that Apis mellifera (L., 1758) (Hymenoptera: Apidae) contributes to increasing R. communis crop productivity by incrementing fruit numbers as well as oil content in seeds (Freitas and Cruz 2010).
Among the pollinator insects of R. communis, A. mellifera is recorded as the main pollinating insect. It is also mentioned that this insect feeds on the nectar produced by the plant’s extrafloral nectar glands (Rizzardo et al. 2012; Waters et al. 2014). A. mellifera is the principal pollinating insect of R. communis, and laboratory work has demonstrated that the pollen of this plant reduces bee survival (Junior et al. 2011). According to these studies, expansion of R. communis as a crop in the semiarid region of Brazil for biodiesel production represents a risk for the native and domestic bees used for honey production.
As shown in Table 3, a total of 36 species of pollinator insects and floral visitors of non-cultivated plants of R. communis is found in the world. These species belong to four orders and 16 families. 25 % of the species belong to Lepidoptera (19 %) and Hemiptera (6 %), while 75 % belong to Hymenoptera (67 %) and Diptera (8 %). 55 % of the arthropod species registered in Table 3 are distributed geographically in Asia (33 %) and Africa (22 %), while 45 % are registered in America; no records were found for Europe.
In Mexico, Cameroon, USA, India, and Brazil, entomophagous Hymenoptera, as well as several species of Lepidoptera, Diptera, and Hemiptera have been reported to feed on nectaries and flowers of R. communis; however, only A. mellifera has been reported as a pollinator. Therefore, it is necessary to carry out studies on pollination and floral ecology in order to determine if there are other insect pollinators of R. communis that should be protected or may be used to increase crop yield (Table 3).
Table 3 Order, family and geographical distribution of the pollinators and floral visitors, reported on Ricinus communis plants.
| Order | Family | Species | Geographical distribution | References |
|---|---|---|---|---|
| Diptera | Muscidae | Musca domestica Linnaeus, 1758 | Cameroon | Douka and Tchuenguem (2014) |
| Richardiidae | Sepsisoma sp. | Brazil | Souza-Silva et al. (2001) | |
| Syrphidae | Ischiodon scutellaris (Fabricius, 1805) | India | Navatha and Sreedevi (2012) | |
| Hemiptera | Coreidae | Anoplocnemis curvipes (Fabricius, 1871) | Cameroon | Douka and Tchuenguem (2014) |
| Pentatomidae | Nezara viridula Linnaeus, 1758 | India | Navatha and Sreedevi (2012) | |
| Lepidoptera | Nymphalidae | Acraea acerata (Hewitson, 1874) | Cameroon | Douka and Tchuenguem (2014) |
| Acraea terpsicore (Linnaeus, 1758) | India | Navatha and Sreedevi (2012) | ||
| Hypolimnas misippus (Linnaeus, 1764) | India | Navatha and Sreedevi (2012) | ||
| Pieridae | Catopsilia florella (Fabricius, 1775) | Cameroon | Douka and Tchuenguem (2014) | |
| Eurema blanda (Boisduval, 1836) | India | Navatha and Sreedevi (2012) | ||
| Eurema sp. | Cameroon | Douka and Tchuenguem (2014) | ||
| Pieris brassicae (Linnaeus, 1758) | India | Navatha and Sreedevi (2012) | ||
| Hymenoptera | Apidae | Apis mellifera Linnaeus, 1758 | Brazil | Freitas et al. (2009); Freitas and Cruz (2010) |
| Apis florea Fabricius, 1973 | India | Navatha and Sreedevi (2012) | ||
| Ceratina sp. | India | Navatha and Sreedevi (2012) | ||
| Scaptotrigona sp. | Brazil | Freitas et al. (2009) | ||
| Trigona sp. | India | Navatha and Sreedevi (2012) | ||
| Xylocopa fenestrata (Fabricius, 1798) | India | Navatha and Sreedevi (2012) | ||
| Braconidae | Bracon spp. | Mexico | Álvarez and Reyes (1987) | |
| Ephiaulax sp. | Mexico | Álvarez and Reyes (1987) | ||
| Chalcididae | Conura igneoides Kirby, 1883 [= Spilochalcis igneoides (Kirby, 1883)] | Mexico | Álvarez and Reyes (1987) | |
| Conura maria Riley, 1870 [= Spilochalcis mariae (Riley, 1872)] | Mexico | Álvarez and Reyes (1987) | ||
| Eurytomidae | Neorileya sp. | Mexico | Álvarez and Reyes (1987) | |
| Formicidae | Camponotus compressus (Fabricius, 1787) | India | Navatha and Sreedevi (2012) | |
| Linepithema humile (Mayr, 1868) | USA | Line et al. (2013) | ||
| Polyrachis sp. | Cameroon | Douka and Tchuenguem (2014) | ||
| Halictidae | Halictus sp. | India | Navatha and Sreedevi (2012) | |
| Sphecidae | Liris sp. | Mexico | Álvarez and Reyes (1987) | |
| Sceliphron assimile (Dahlbom, 1843) | Mexico | Álvarez and Reyes (1987) | ||
| Tachysphex sp. | Mexico | Álvarez and Reyes (1987) | ||
| Tachytes sp. | Mexico | Álvarez and Reyes (1987) | ||
| Trypoxilon sp. | Mexico | Álvarez and Reyes (1987) | ||
| Torymidae | Torymus capillaceus (Huber, 1927) | Mexico | Álvarez and Reyes (1987) | |
| Vespidae | Synagris cornuta (Linnaeus, 1758) | Cameroon | Douka and Tchuenguem (2014) | |
| Delta sp. | Cameroon | Douka and Tchuenguem (2014) | ||
| Polistes sp. | Mexico | Álvarez and Reyes (1987) |
Some pests can affect pollinators through herbivory. In the case of R. communis, Wäckers et al. (2001) showed that plants damaged by larvae of Spodoptera littoralis (Boisd., 1833) (Lepidoptera: Noctuidae) increased the total amount of nectar produced by extrafloral nectaries compared to undamaged plants. De Sibio and Rossi (2016) found a similar result for the herbivory of S. frugiperda on R. communis. The secretion of carbohydrates through extrafloral nectaries is considered an indirect strategy of plant defense because it serves to attract parasitoids and predators (Heil 2008). Unlike floral nectaries, extrafloral nectaries do not participate in pollination, however, in plants pollinated by insects, extrafloral nectaries can negatively affect the effectiveness of pollination by distracting pollinators away from floral nectaries or when the ants that are attracted by the nectar attack the floral visitors (Wäckers et al. 2001; Turlings and Wäckers 2004).
Natural enemies of the pests of R. communis
Among the natural enemies of the key pests of cultivated R. communis, there are parasitoids, predators, and entomopathogens such as fungi, bacteria, nematodes, and viruses, which are used as biological control agents or have been found parasitizing, depredating, or naturally infecting some pests of the crop. An extensive list of natural enemies of phytophagous arthropods of R. communis grouped by taxa with information of their host or prey and geographical distribution is shown in Table 4; as it can appreciate in this table, the most commonly reported natural enemies in countries like India, Brazil, China, and USA, are Bacillus spp., Trichogramma spp., Microplitis spp., Telenomus spp., Stethorus spp., and other species attacking pests such as A. janata, S. litura, Anacraga citrinopsis Dyar, 1927, S. obliqua, Phyllophaga sp., Eutetranychus banksi (McGregor, 1914), Tetranychus piercei McGregor, 1950, Zaniothrips ricini Bhatti, 1967, and other species. Table 4 shows a total of 61 natural enemies of phytophagous insects of R. communis. Three species are bacteria belonging to the same genus; four species are nematodes of different genera; two species are fungi of different genera; two reports are viruses; 36 species are parasitoids of eight families of Hymenoptera and one family of Diptera; and 14 species are predators of six different families and order 74 % of the species is distributed geographically in Asia, 24 % in America, 2 % in Africa and 0 % in Europe.
An example of natural enemies of pest of R. communis is presented by Basappa (2009) . According to this author, parasitoids, insect predators, spiders, insectivorous birds and some microbial organisms are important natural enemies of the pest complex of R. communis ecosystem in India. In the case of A. janata, Trichogramma chilonis Ishii, 1941, Trichogramma achaeae Nagaraja and Nagarkatti, 1970, Telenomus sp. and Trissolcus sp. were recorded from eggs; Microplitis maculipennis (Szepligeti, 1900), Euplectrus maternus Bhatnagar, 1952, Rhogas spp. and Apanteles hyposidrae Wilkinson, 1928 were found among larval parasitoids; and pupae were found to be parasitised by Tetrastichus howardi (Olliff, 1893) (= Tetrastichus ayyari Rohwer, 1921) and Phorocera sp. Among insect predators of A. janata, Chrysoperla sp. and Cheilomenes sexmaculata (Fabricius, 1781) were found feeding on the eggs and neonate larvae; other general insect predators like mantids, paper wasps, and sphecid digger wasps were also found predating on larvae; spiders like green lynx spiders, jumping spiders and crab spiders were found feeding on early instar larvae. Among the entomopathogens of A. janata, Spodoptera litura nucleopolyhedrovirus and granulosis virus were isolated from dead larvae and the fungi Metarhizium rileyi (Farl.) Kepler, S.A.Rehner & Humber, 2014 [= Nomuraea rileyi (Farlow) Samson, 1974] and Beauveria bassiana (Balsamo) Vuillemin were found infecting larvae (Basappa 2009). Many other examples of natural enemies of pests of R. communis are shown in Table 4
Table 4 Natural enemies of phytophagous arthropods of Ricinus communis.
| Species | Host and/or prey | Geographical distribution | References |
|---|---|---|---|
| Entomopathogens | |||
| Bacteria | |||
| Bacillus thuringiensis var. kurstaki (Berliner, 1915) | Larvae of Achaea janata | India | Vimala Devi and Sudhakar (2006) |
| Bacillus cereus (Manson, Pollock & Tridgell, 1954) | Larvae of Achaea janata | India | Kattegoudar et al. (1994) |
| Bacillus popilliae Dutky, 1940 | Larvae of Phyllophaga sp. | Colombia | Saldarriaga Cardona et al. (2011) |
| Nematodes | |||
| Hexamermis dactylocercus Poinar and Linares, 1985 | Larvae of Amsacta albistriga | India | Prabhakar et al. (2010) |
| Steinernema carpocapsae (Weiser, 1955) | Larvae of Spodoptera litura | India | Raveendranath et al. (2008) |
| Heterorhabditis indica Poinar, Karunaka y David, 1992 | Larvae of Spodoptera litura | India | Raveendranath et al. (2008) |
| Mermis sp. | Larvae of Achaea janata | India | Sujatha et al. (2011) |
| Fungi | |||
| Metarhizium rileyi (Farl.) Kepler, S.A.Rehner & Humber, 2014 [= Nomuraea rileyi (Farlow) Samson, 1974] | Larvae of Spodoptera litura | India and USA | Mau and Kessing (2007) |
| Beauveria bassiana (Balsamo) Vuillemin, 1912 | Larvae of Achaea janata and Cogenethes punctiferalis | India | Duraimurugan et al. (2015) |
| Virus | |||
| Nucleopolyhedrovirus | Larvae of Spodoptera litura | India | Basappa (2009) |
| Granulovirus | Larvae of Achaea janata and Spodoptera litura | India | Naveen Kumar et al. (2013) |
| Parasitoids | |||
| INSECTA | |||
| Hymenoptera | |||
| Aphelinidae | |||
| Encarsia formosa Gahan, 1924 | Nymphs of Trialeurodes ricini | China | Wang et al. (2016) |
| Braconidae | |||
| Habrobracon hebetor (Say, 1836) | Larvae of Cogenethes punctiferalis | India | Basappa (2003) |
| Apanteles hyposidrae Wilkinson, 1928 | Larvae of Achaea janata | India | Basappa (2009) |
| Apanteles ricini Bhatnagar, 1948 | Larvae of Cogenethes punctiferalis | India | Basappa (2003) |
| Cotesia flavipes (Cameron, 1891) [Apanteles flavipes (Cameron, 1891)] | Larvae of Spilosoma obliqua and Spodoptera litura | India | Yadav et al. (2010); Basappa (2009) |
| Glyptapanteles dalosoma de Santis, 1987 | Larvae of Anacraga citrinopsis | Brazil | Lourenção et al. (1989) |
| Microplitis (= Microgaster) rufiventris Kokujev, 1914 | Larvae of Spodoptera litoralis | Egypt | Shalaby et al. (1988) |
| Microplitis maculipennis (Szepligeti, 1900) (= Microplitis ophiusae Aiyar, 1921) | Larvae of Achaea janata | India | Suganthy (2010); Naik et al. (2010) |
| Chalcididae | |||
| Brachymeria euploeae (Westwood, 1837) | Pupae of Cogenethes punctiferalis | India | Sujatha et al. (2011) |
| Eulophidae | |||
| Ceranisus menes (Walker, 1839) | 2º instar nymph of Zaniothrips ricini | India | Daniel et al. (1983) |
| Euplectrus maternus Bhatnagar, 1952 | Larave of Achaea janata | India | Basappa (2009) |
| Tetrastichus howardi (Olliff, 1893) (= Tetrastichus ayyari Rohwer, 1921) | Pupae of Spodoptera litura and Achaea janata | India | Basappa (2009) |
| Trichospilus pupivorus Ferrière, 1930 | Pupae of Spodoptera litura and Achaea janata | India | Basappa (2009) |
| Trichogrammatidae | |||
| Trichogramma achaeae Nagaraja and Nagarkatti, 1970 | Eggs of Achaea janata | India | Basappa (2009) |
| Trichogramma chilonis Ishii, 1941 | Eggs of Achaea janata and Spodoptera litura | India | Singh et al. (2008); Suganthy (2010); Naik et al. (2010) |
| Trichogramma minutum Riley, 1879 | Eggs of Achaea janata | USA | Mau and Kessing (2007) |
| Trichogramma australicum Girault, 1912 | Eggs of Achaea janata | China | Hua (1984) |
| Trichogramma dendrolimi Matsumura, 1926 | Eggs of Achaea janata | China | Hua (1984) |
| Trichogramma pretiosum Riley, 1879 | Eggs of S. cosmioides | Brazil | Cabezas et al. (2013) |
| Trichogramma evanescens Westwood, 1833 | Eggs of Achaea janata | India | Basappa (2009) |
| Scelionidae | |||
| Telenomus remus Nixon, 1937 | Eggs of Spodoptera litura, Spodoptera cosmioides and Spodoptera frugiperda | India | Satyanarayana et al. (2005); Pomari et al. (2013) |
| Telenomus proditor Nixon, 1937 | Eggs of Lepidoptera | USA | Mau and Kessing (2007) |
| Telenomus sp. | Eggs of Achaea janata | India | Basappa (2009) |
| Trissolcus sp. | Eggs of Achaea janata | India | Basappa (2009) |
| Vespidae | |||
| Polistes sp. | Larvae of Phyllophaga sp., Agrotis sp. and Spodoptera spp. | Costa Rica | Anónimo (1991) |
| Ichneumonidae | |||
| Campoletis chlorideae Uchida, 1957 | Larvae of Spodoptera litura | India | Satyanarayana et al. (2005) |
| Charops obtusus Morley, 1913 | Larvae of Spilosoma obliqua and Achaea janata | India | Basappa (2009) |
| Hyposoter exiguae (Viereck,1912) | Larvae of Achaea janata | USA | Mau and Kessing (2007) |
| Diadegma ricini Row & Kurian,1950 | Larvae of Cogenethes punctiferalis | India | Basappa (2003) |
| Theronia sp. | Larvae of Cogenethes punctiferalis | India | Basappa (2003) |
| Isdromas monterai (Costa Lima, 1948) | Larvae of Anacraga citrinopsis | Brazil | Lourenção et al. (1989) |
| Tachinidae | |||
| Palexorista parachrysops Bezzi, 1925 | Larvae of Cogenethes punctiferalis | India | Kalra (1984) |
| Eucelatoria armigera (Coquillett, 1889) | Larvae and pupae of Achaea janata | USA | Mau and Kessing (2007) |
| Chaetogaedia monticola (Bigot, 1887) | Larvae and pupae of Achaea janata | USA | Mau and Kessing (2007) |
| Predators | |||
| Coleoptera | |||
| Carabidae | |||
| Calosoma sp. | Larvae of Phyllophaga sp., Agrotis sp. and Spodoptera spp. | Costa Rica | Anónimo (1991) |
| Coccinellidae | |||
| Cheilomenes sexmaculata (Fabricius, 1781) | Eggs larvae of Achaea janata and Spodoptera litura | India | Basappa (2009) |
| Micraspis cardoni (Weise, 1892) | Zaniothrips ricini | India | Daniel et al. (1983) |
| Scymnus sp. | Eutetranychus orientalis | Palestine and Egypt | Klein (1936) |
| Stethorus sp. | Eutetranychus banksi | USA | McGregor (1914) |
| Stethorus siphonulus Kapur, 1948 | Tetranychus piercei | China | Lui and Lui (1986) |
| Stethorus histrio Chazeau, 1974 | Tetranychus urticae | Chile | Aguilera (1987) |
| Hemiptera | |||
| Pentatomidae | |||
| Eocanthecona furcellata (Wolff, 1811) | Achaea janata | India | Rao (1977); Usha Rani (2009) |
| Reduviidae | |||
| Rhynocoris kumarii Ambrose and Livingstone, 1986 | Eggs larvae of Achaea janata and Spodoptera litura | India | Basappa (2009) |
| Thysanoptera | |||
| Aeolothripidae | |||
| Franklinothrips megalops (Trybom, 1912) | Zaniothrips ricini | India | Daniel et al. (1983) |
| Mymarothrips garuda Ramakrishna and Margabandhu, 1931 | Zaniothrips icini | India | Daniel et al. (1983) |
| Neuroptera | |||
| Chrysopidae | |||
| Chrysoperla carnea (Stephens, 1836) | Tetranychus urticae | India | Rajasekhar et al. (1999) |
| Chrysoperla sp. | Eggs and larvae of Achaea janata and Spodoptera litura | India | Basappa (2009) |
| Mantodea | |||
| Mantidae | |||
| Haldwania lilliputana Beier, 1930 | Zaniothrips ricini | India | Daniel et al. (1983) |
| ARACHNIDA / Acari | |||
| Phytoseiidae | |||
| Sciulus sp. | Eutetranychus banksi | USA | McGregor (1914) |
Pest management of phytophagous arthropods in R. communis
Pest management methods used to control the principal arthropod pests of R. communis include cultural, genetic, ethological, biological, and chemical control.
Cultural control is the use of agronomical practices designed to reduce the presence of pests in crops of R. communis. Intercropping is a type of cultural control recommended to diminish the damage caused by insect pests in R. communis.Srinivasa Rao et al. (2012) found that plants such as Cyamopsis tetragonoloba (L.) Taub., 1891, Vigna unguiculata (L.) Walp., 1845, Vigna mungo (L.) Hepper, 1956, and Arachis hypogaea L., 1753, intercropped with R. communis in a 1:2 proportion, decreased the incidence of insect pests such as A. janata, E. flavescens, and C. punctiferalis. Moreover, a more considerable presence of natural enemies of these pests was observed in these intercropping systems. Patel and Patel (2009) recommended intercropping R. communis with Vigna radiata (L.) Wilczek, 1954, Sesamum indicum L., 1753, Vigna aconitifolia (Jacq.) Marechal, 1969, and V. unguiculata, to reduce damage by C. punctiferalis. When R. communis was monocropped, C. punctiferalis caused 53 % damage, but when intercropped with the above mentioned species, the damage was between 35 and 53 %. Sowing date is another cultural method for reducing damage and the presence of pests. Salihu et al. (2014) suggest that the correct time for planting R. communis crop must be related to the rainy season, which is more important than any other pest control measure in Africa, since the rains decrease the presence of certain pests.
Genetic control includes the use of cultivars resistant to insect pests, however, according to Singh et al. (2015) , breeding R. communis is complicated by limited sources of pest resistance. In India, there are R. communis varieties that are tolerant or resistant to attack by pests of greater economic importance, such as E. flavescens, T. ricini, S. litura, A. janata, C. punctiferalis, and L. trifolii (Anjani et al. 2010; Anjani 2012). Resistant or tolerant plants have high oil content (between 40 and 49 %) and yields that oscillate between 540 and 1,580 kg/ha (Lavanya et al. 2012). It is mentioned that the cultivars having purple leaves are resistant to the attack of L. trifolii, while those with green leaves are susceptible (Sarma et al. 2006; Anjani et al. 2007). Sarma et al. (2006) mention that purple-leaf varieties have high levels of anthocyanin, which make the plant more tolerant to L. trifolii attack, and the epicuticular wax on their leaves reduces infestation and defoliation by A. janata and S. litura. There are hybrids, such as GCH4, that are resistant to the attack by E. flavescens due to the high wax content on the plant stems and leaves (Lakshmi et al. 2005). Five accessions viz., RG-43, RG-631, RG-1621, RG-3037 and RG-3067, among 165 core set accessions representing diversity in the entire collection maintained at ICAR-Indian Institute of Oilseeds Research, Hyderabad, India, exhibited resistance reaction against E. flavescens; oil content of these accessions was 46, 51, 51, 51, 52 %, respectively (Anjani et al. 2018). On the other hand, Severino et al. (2012) recommended parallel research to determine the increased potential susceptibility to pests in breeding programs to develop low-ricin, low-ricinine, and low-allergen cultivars to reduce hazardous chemical products found in R. communis.
Research is being carried out on the use of transgenic plants of R. communis. In India, two transgenic varieties of R. communis, Jyothi and VP1, developed by genetic engineering induce A. janata mortality above 88 % due to the Bacillus thuringiensis gene CryAb (Malathi et al. 2006).
A little explored method for monitoring and massive trapping of R. communis pests has been the use of pheromones, kairomonal attractants and light traps. In India, the pheromone compounds of some pests of economic importance have been identified and used for monitoring and massive trapping of C. punctiferalis, S. litura, Amsacta albistriga Walker, 1864 (Lepidoptera: Arctiidae), A. janata, and S. obliqua (Cork and Hall 1998). In this country, an important prerequisite for successful management of S. litura, the most destructive insect pest of R. communis damaging the crop from July- October during the south-west monsoon (kharif season), has been the implementation of an intensive monitoring program of S. litura population using sex pheromone traps (Satyagopal et al. 2014). Setting twelve traps baited with pheromone compounds per hectare for massive trapping of S. litura is recommended (Nandagopal and Rathod 2007; Raghavaiah 2011). In Brazil, researchers are now taking the first steps toward identifying the pheromone compounds of C. gossypii (Fregadolli et al. 2012) with the aim of developing a commercial pheromone. In India, the kairomonal compounds of the most destructive lepidopteran insect pest of R. communis, such as S. litura, A. janata, and C. punctiferalis have been identified for trapping. In field experiment, water trap baited with phenyl acetaldehyde + 2-phenyl ethanol recorded significantly higher moth catches of S. litura (6.8 moths/trap/wk) and C. punctiferalis (5.8 moths/trap/wk) (Duraimurugan et al. 2017). Recently, Duraimurugan and Alivelu (2018), determined the relationship of pheromone trap catches corresponding to the economic threshold level of 25 % defoliation of S. litura on R. communis, which was estimated to be 81.4 moths/trap/week. Light traps using ultraviolet black-blue spectrum have also been suggested to capture Phyllophaga sp. adults as a measure of ethological control Saldarriaga Cardona et al. (2011).
Biological control (spraying entomopathogenic microorganisms and releasing entomophagous insects) has been implemented in the control of key R. communis pests in countries such as India and Colombia. In India, for example, parasitism rates between 10.4 and 28.7 % of M. maculipennis and Cotesia sp. were recorded on larvae of A. janata and S. litura, respectively (Suganthy 2007), while 9.5 % parasitism rates of Cotesia flavipes (Cameron, 1891) against S. obliqua larvae have been observed (Yadav et al. 2010). Also, in India, Rajasekhar et al. (1999) mention that when Chrysoperla carnea (Stephens, 1836) were released, the T. urticae (= T. telarius) mite populations diminished by 75 %. In the case of S. litura, the release of 150 adults of the parasitoid Telenomus remus (Nixon, 1937) per egg mass and the release of the larval parasitoid Campoletis chlorideae Uchida, 1957 in a parasitoid: host ratio of 1:15 achieved parasitism rates above 96 % (Satyanarayana et al. 2005).
Biological control through entomopathogenic nematodes, bacteria, fungi, and virus exposed to the principal pests of R. communis has been evaluated in India. For instance, mortality of S. litura pupae was evaluated with two nematode species: Heterorhabditis indica Poinar, Karunaka and David, 1992, and Steinernema carpocapsae (Weiser, 1955) (Raveendranath et al. 2008). Other studies evaluated the mortality of A. janata and Samia ricini (Drury, 1773) larvae exposed to two species of bacteria: Bacillus thuringiensis Berliner, 1915 and Bacillus cereus Frankland & Frankland 1887 (Manson et al. 1954; Kattegoudar et al. 1994; Mathur et al. 1994; Vimala Devi and Sudhakar 2006). Duraimurugan et al. (2015) conducted research to determine the mortality of A. janata larvae and C. punctiferalis adults with Beauveria bassiana (Balsamo) Vuillemin, 1912 fungus. Mortality of A. janata larvae exposed to granulovirus was also evaluated (Naveen Kumar et al. 2013). In Colombia, Saldarriaga Cardona et al. (2011) reported that the control of Phyllophaga sp. larvae was achieved by applying the B. popilliae Dutky bacterium at a concentration of 24,000.00 billion spores/ha a year, during five consecutive years.
The use of secondary metabolites derived from plants and other organisms, as well as methods of chemical control, have been assessed for controlling some R. communis pests. In laboratory studies, it has been found that methanol extracts of Clathria longitoxa (Hentschel, 1912) and Callyspongia diffusa (Ridley, 1884), two marine sponges, have insecticidal effect on A. janata and P. recini larvae (Joseph et al. 2010). Furthermore, Calotropis gigantea (L.) W. T. Aiton, 1811, leaf extracts demonstrated strong feeding deterrent activity against larvae of Pericallia ricini (Fabricius, 1775) at high concentrations (Neelanarayanan and Indira 2010). Likewise, toxic effects and strong antifeedant activity of raw acetonic extracts of Mormodica charantia L., 1753, Tectona grandis L. f., 1790, and Madhuca indica J. F. Gmel., (1791) against S. litura and A. janata larvae (Devanand and Usha Rani 2008) were found. Neem, Azadirachta indica (Juss, 1830), also evaluated for the control of S. litura, induces mortality of larvae at high concentrations (Choudhury and Aizur Rahman 2008).
Chemical control through insecticides is one of the most common practices for control of R. communis pests (Gahukar 2018). In Colombia, six pesticides for the control of C. gossypii were evaluated. The results of insect control efficacy three days after pesticide application were as follows: (from the least to the most effective) thiamethoxam + lambda-cihalotrin (0.00 %), spinetoram (0.00 %), malathion (20.35 %), thiamethoxam (38.62 %), dimethoate (86.94 %), and imidacloprid (87.33 %); whereas after seven days the following results were obtained: thiamethoxam + lambda-cihalotrin (0.00 %), spinetoram (21.46 %), malathion (38.77 %), thiamethoxam (50.84 %), dimethoate (86.14 %), and imidacloprid (90.37 %) (Varón et al. 2010). Mead (1989) suggested the use of carbaryl or malathion for controlling C. gossypii in Florida, USA. In Colombia, Saldarriaga Cardona et al. (2011) recommended application of baits poisoned with carbaryl at a dose of 2 to 3 g/L for the control of A. ipsilon and S. frugiperda; the same authors recommended application of liquid chlorpyrifos at the base of the plants at a dose of 1.5 - 2.0 cc/L.
The most recommendable strategy of R. communis pest control is Integrated Pest Management (IPM). Most of the IPM programs have been directed against key pests of R. communis, such as S. litura, C. punctiferalis, and A. janata (Prabhakar et al. 2003; Singh et al. 2006; Basappa 2009). In India, the growers increased seed production of R. communis up to 28 %, by implementing IPM programs with insecticides, crop rotation, insect traps, application of neem extract, and intercropping (Basappa 2007). The results of research in India demonstrate that IPM is an efficient strategy for the control of A. janata and S. litura, two of the key pests of R. communis. It is possible to decrease populations of these pests by using the recommended IPM program, which includes the use of bird perches for predatory birds to rest and to look for preys, foliar applications of 5 % neem seed extracts, biological insecticide consisting of nuclear polyhedrosis virus (S. litura NPV 100 LE/ha), monocrotophos at 0.5 %, and manual removal of larvae (Suganthy 2010). The pest control effectiveness of carbaryl 50W 0.2 %, endosulfan 35 EC 0.05 %, triazophos 40 EC 0.05 %, spinosad 45 SC 0.018 %, fipronil SSC 0.01 %, extract of neem seeds 5 % (weight/volume), B. thuringiensis 0.1 %, and a control without applying the dose of 500 L/ha, was evaluated under field conditions 30 and 45 days after establishing a plantation of a R. communis variety susceptible to leafminer L. trifolii. The results showed that the least damage (lowest number of insect mines) was found when spinosad and triazophos were applied and, at the same time, the best yield was obtained with both treatments (883 and 835 kg seed/ha, respectively) (Akashe et al. 2009). On the other hand, natural enemy impact has been proven to be greatest at sites adopting biointensive IPM (BIPM); par example, studies conducted by Basappa (2009) shown that BIPM modules were safer to A. janata eggs (T. chilonis) and larvae (M. maculipennis) parasitoids with 16.1 and 66.1 % average field parasitism, compared to chemical pesticide intensive integrated pest management modules with 6.9 and 21.2 % parasitism, respectively.
Conclusions
There is a wide range of arthropods that damage R. communis in different parts of the world where this plant is cultivated; many of these are considered pests of economic importance. Likewise, there are reports of a great variety of natural enemies, which have been used in biological control programs. According to the coevolutive hypothesis, it was found that the greatest richness and abundance of arthropods associated with R. communis is in Asia and Africa, considered as the center of origin of this plant. Most phytophagous arthropods feed on leaves. The natural enemies with more abundance and richness are the parasitoids that mostly attack the larvae of phytophagous arthropods. With respect to pollinators, A. mellifera is the principal pollinating insect, however, more research on pollination and floral ecology in R. communis is needed, in order to determine what other floral visitors may act as pollinators, and how they can be protected or manipulated to increase crop yield. The pest management programs of phytophagous arthropods of R. communis must be directed toward promoting and preserving natural enemies and pollinating insects by means of environment-friendly pest management techniques, for which use of wide-spectrum insecticides must be avoided.
