Submitted Successfully!
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Ver. Summary Created by Modification Content Size Created at Operation
1 Therefore, due to the development of insecticides’ resistance by some of the insect pests of Brassica oleracea, there is a need for search of the alternative products which can effectively be used to control those insect pests in the field. + 6457 word(s) 6457 2020-05-15 10:45:48 |
2 update layout and reference -1976 word(s) 4481 2020-05-27 08:45:02 | |
3 update layout -25 word(s) 4456 2020-10-28 09:54:05 |

Video Upload Options

Do you have a full video?


Are you sure to Delete?
If you have any further questions, please contact Encyclopedia Editorial Office.
Mpumi, N.; Machunda, R.S.; Mtei, K.M.; Ndakidemi, P.A. Insect Pests of Brassica oleracea. Encyclopedia. Available online: (accessed on 05 December 2023).
Mpumi N, Machunda RS, Mtei KM, Ndakidemi PA. Insect Pests of Brassica oleracea. Encyclopedia. Available at: Accessed December 05, 2023.
Mpumi, Nelson, Revocatus S. Machunda, Kelvin M. Mtei, Patrick A. Ndakidemi. "Insect Pests of Brassica oleracea" Encyclopedia, (accessed December 05, 2023).
Mpumi, N., Machunda, R.S., Mtei, K.M., & Ndakidemi, P.A.(2020, May 22). Insect Pests of Brassica oleracea. In Encyclopedia.
Mpumi, Nelson, et al. "Insect Pests of Brassica oleracea." Encyclopedia. Web. 22 May, 2020.
Insect Pests of Brassica oleracea

The propagation and regeneration of Brassica species has been successful using seeds and different explants like petioles, cotyledons, stems and shoot tips. Shoot regeneration and rooting of Brassica species are successfully obtained from cotyledons and hypocotyl explants. The biological cycle length of Brassica species may either differ slightly or may not differ from one species to another. For instance, the seeds of Brassica oleracea take five days to germinate after sowing at 20–25 °C while the seeds of Brassica campestries take about three to five days to germinate after sowing at 20–25 °C. The most common insect pests of economic importance to Brassica oleracea in African smallholder farmers include Plutella xylostella, Helula undalis, Pieris brassicae, Brevycoryne brassicae, Trichoplusia ni and Myzus persicae. Those insect pests infest cabbages at different stages of growth, causing huge damage and resulting into huge yield losses. The African smallholder farmers use cultural and synthetic pesticides to control those insect pests and minimize infestations. The cultural practices are environmental friendly but are ineffective to control the insect pests. Due to ineffectiveness of cultural practices, African smallholder famers use broad-spectrum synthetic pesticides to effectively control the Brassica species insect pests. The improper and misuse of synthetic pesticides result into insect pests resistance towards the insecticides applied, environmental pollution and human health threats. Insect pests such as Plutella xylostella, Hellula undalis, Brevicoryne brassicae and Myzus persicae have developed resistance to a wide range of pesticides used such as cypermethrin, parathion, decamethrin, quinalphos and lamda-cyhalothrin. Therefore, that calls for search of the alternative products which can effectively be used to control those insect pests in the field.

Plutella xylostella Helula undalis biological cycle length, and regeneration

1. The Biological Cycle of Brassica Species and Their Common Insect Pests in Africa

1.1. Propagation and Biological Cycle Length of Selected Brassica Species

The propagation and regeneration of Brassica species has been successful using seeds and different explants like petioles, cotyledons, stems and shoot tips [1] (Table 1). Shoot regeneration and rooting of Brassica species are successfully obtained from cotyledons and hypocotyl explants [2]. The shoot tip explants of Brassica species are reported to be effective for initiating shoots and roots [3]. Table 1 shows the Brassica species propagation and biological cycle length.

Table 1. Propagation and biological cycle length of Brassica species.

1.2. The Common Insect Pests Affecting Cabbages in Africa

Many insect pests such as diamondback moth (Plutella xylostella), cabbage webworm (Helula undalis), cabbage white butterfly (Pieris brassicae), the cabbage aphids (Brevycoryne brassicae), green peach aphids (Myzus persicae) and cabbage loopers (Trichloplusia ni) [10][11] (Table 2) hinder the proper cabbage crop production on the field in Africa. Those insect pests (Table 2) infest the cabbage crops at different stages of growth, causing significant damage to the crop [12] and resulting into huge cabbage yield losses. Krishnamoorthy [13] showed that, cabbage insect pests all together can cause 52% yield loss on cabbage. Severe infestation by Plutella xylostella usually causes huge economic crop losses and may result in 100% yield loss of the Brassica oleracea [14]. Due to heavy infestations which result into huge losses, the African smallholder farmers of cabbages spray four or more than four times in a month and two or more than two mixed insecticides into the field for strongly and effectively control of the cabbage insect pests [15][16]. The consequence of that scenario is environmental pollution especially water and soil, detrimental effects to non-target organisms and endanger the health of the human being [17]. This section reviews the major insect pests of economic importance infesting cabbage crop at different stages of growth in African countries and how the control measures are potential water and soil pollution threat.

Table 2. Common insect pests of cabbages [10][11].

1.2.1. Cabbage Looper (Trichoplusia ni)

The cabbage looper (Figure 1A) (Trichoplusia ni) is a moth found in the family noctuidae a family which is commonly referred to as owlet moths [18]. Its common name comes from its preferred host plants and distinctive crawling behavior. The members of noctuidae are brown or gray night-flying moths whereby the larvae infest the growth of cruciferous vegetables [19]. Cruciferous vegetables like cabbages, bok choy and broccoli are the main host plants to cabbage lopper and hence, the reference to cabbage in its common name [20]. The larvae is called a looper since it arches its back into a loop when it crawls [18]. While crucifers are preferred, however, over 160 plants can serve as hosts of cabbage looper larvae [18]. The adult cabbage looper is a migratory moth and its migratory behavior can be found in a wide range of host plants and this contribute to its wide range of distribution [19].

Figure 1. (A) Mature larva of the cabbage looper, Trichoplusia ni. (B) The cabbage plant damaged by Cabbage looper larva. Photograph by Nelson Mpumi, NM-AIST-Arusha, Tanzania.

The cabbage looper larvae is a vegetable pest for crucifers and has been reported to damage broccoli, cabbages, cauliflowers, chinese cabbages, collards, kale, mustards, radish, turnip and watercress [19]. The cabbage looper larvae interfere with plant growth and marketability by making irregular holes of variable shapes (Figure 1A,B) while feeding on the leaves of the host cabbage plants [21]. Although it is not extremely destructive, but it is becoming difficult to control and manage due to its broad distribution and resistance to many insecticides [20][21]. Therefore, African smallholder farmers rely intensively on the application of synthetic pesticides to control the cabbage insect pests. However, synthetic pesticides result into environmental pollution, insect pest resistance and contaminate the foods which consequently threaten the human health [21]. Therefore, environmental benign, the botanical pesticides from T. vogelii, S. aromaticum and C. dichogamus can be utilized to control the insect pests in the field instead of synthetic pesticides [22]. Although the potentialities are ignored, but botanical pesticides have been in use for centuries by smallholder farmers in developing countries to control insect pests of both field and stored products [23][24]. Therefore, they could be used to control cabbage insect pests in the field to minimize the infestation.

1.2.2. Cabbage Webworm (Hellula undalis)

Among the most destructive insect pests which attack cruciferous vegetables is the cabbage webworm (Figure 2A) (Hellula undalis) (Lepidoptera: Pyralidae) [25]. The cabbage webworm (Hellula undalis) is a major pest of cruciferous crops in the tropics and subtropics [26]. It is a widespread species in the world especially in Europe across Asia to the Pacific and also, in African countries [27][28]. Shine et al. [29] reported that H. undalis is distributed mostly in tropical and subtropical regions but can similarly be found in countries with moderate climates.

Figure 2. (A) Mature larva of the cabbage webworm, Hellula undalis. Photograph by Lyle Buss, Entomology and Nematology Department, University of Florida (March, 2016). (B) The cabbage plant head damaged by larva of H. undalis. Photograph by Nelson Mpumi, NM-AIST Arusha, Tanzania.

Ebenebe et al. [27] reported that, Helulla. undalis larva causes a serious and severe damage to the leaves and the heads of cabbages (Figure 2B) in the field. According to Waterhouse and Norris [30] H. undalis feeds on a variety of plants especially the Brassicaceae family members. Waterhouse and Norris [30] revealed that, H. undalis larva can cause a huge yield loses of up to 100% to crucifers crops in the field and its management is not well taken into account. The larvae feed on leaves, petioles, growing points and stems [28]. According to Sivapragasam and Aziz [25] and Waterhouse and Sands [28], the plants in which H. undalis larvae feed include broccoli, head cabbage, chinese cabbage, spoon cabbage, daikon radish, horseradish, mustard, radish and turnip. Shine et al. [29] revealed that, H. undalis is a very serious agricultural pest to crucifer crops grown by the African smallholder famers. The incidence of H. undalis did not depend on the number of insecticide applications, but depend highly to host crop abundance and the temperature of the area [31]. The larvae make mines in the leaves and bore into the stem and later, they tunnel into the heart of the plant, destroying the bud causing the leaves to become distorted and stunted [10]. A study done by Sivapragasam and Aziz [25] indicated that, a single larva of cabbage webworm, can either cause a number of deaths to the young plant or lead to the formation of unmarketable multiple heads on relatively older plant. On the field, a low population of larvae can cause very huge significant losses to the cabbage crop and in untreated cabbages, losses could go as high as 99% [27]. Although, the larva can be present throughout the cropping season, it is severe only during the period between transplanting and the heading stage of cabbage [32].

Currently, African smallholder farmers rely intensively on the application synthetic pesticides as the only effective control method to the cabbage webworm on the field [25]. The effective insecticides which are used to control cabbage webworm worldwide and Africa particularly, include permethrin, abamectin, teflubenzuron, chlorfluazuron, triflumuron, phenthoate, exthofenprox and Lamda-cyhalothrin and among those insecticides, abamectin is found to be the most effective of the other insecticides [25]. However, some are reported hazardous and therefore unwise and overuse of those insecticides can result into severe environmental pollution especially water and soil, development of insect resistance to some of insecticides and health problems to human being. Therefore, there is a need of searching and utilizing benign and environmental friendly botanicals from pesticidal plants as cabbage insect pests control strategy.

1.2.3. Diamondback month (Plutella xyllostela)

The diamondback moth (Figure 3A) (Plutella xylostella), sometimes called the cabbage moth, is a moth species belonging to the family Plutellidae and genus Plutella [32][33]. Badenes-Perez et al. [34] reported that Plutella xylostella is believed to have originated in Europe, South Africa, or the Mediterranean region, but it has now spread worldwide. The diamondback moth is the dominant and most destructive insect pest of crucifer crops worldwide [23]. Justus and Mitchell [35] reported that, Plutella xylostella larva feeds on the leaves between the large veins and midribs of cruciferous crops and the plants which produce glucosinolates. Plutella xylostella larva prefers to feed on the lower leaf surface, leaving the upper epidermis intact creating a “window-paning” effect (Figure 3B) [23]. Timbilla and Nyarko [12] showed that, severe feeding damage (Figure 3B) stunts and destroys the cabbage heads and can cause heads to abort leading to huge yield depression and total crop loss. The most cabbage plant damage is caused by larval feeding resulting in a complete removal of foliar tissues and disrupt head formation in cabbages, broccoli and cauliflower [23]. The destruction of cruciferous crops by diamondback moth larva reduces the quality and the marketability of the cabbage crops and hence yield losses due to P. xylostella can go up to 100% [14][36] and vary widely depending on the season and severity of pest infestation [37].

Figure 3. (A) Mature larva of the Diamond back moth, Plutella xylostella. (B) The cabbage plant damaged by larva of P. xylostella. Photograph by Nelson Mpumi. NM-AIST, Arusha, Tanzania.

Generally, it is estimated that, diamondback moth causes an annual loss of about 16 million dollar on the basis of 2.5 per cent damage even on the protected crop [38]. Also, in the tropics, diamondback moth causes threat of great loss of 90% and above to crucifer production crop [39]. Therefore, there is a need to conduct a research to determine the cabbage losses due to infestation of diamondback moth in various parts of Africa.

The diamondback moth and its larvae control in cabbage by African smallholder farmers is still deeply dependent on chemical insecticides although their use is connected with many adverse and lethal consequences. Inappropriate and excessive application of chemical insecticides result into environmental pollution especially water and soil pollution [40][41]. Pedigo and Rice [42] indicated that, extreme use of insecticides also induces resistance development in target pests as well as killing beneficial organisms like pollinators such as bees and other natural enemies such as spiders, lacewings and ladybird beetle. Therefore, the benign, environmental friendly botanicals have to be searched to control this pest instead of relying on the synthetic pesticides which have many negative impacts and problems to the environment. The benign and environmental friendly control measures with broad spectrum of the activities are the botanicals (phytochemicals), the chemicals from pesticidal plants [22]. Those alternatives with antifeedant, repellency and insect growth regulators of their natural origin having non-neurotoxic modes of action to human being and low environmental persistence can be applied.

Botanical pesticides are not only effective against crop pests but remain safe to the environment and to natural enemies [43]. In developing countries, botanicals have been in use for centuries by smallholder farmers to control insect pests both in field and storage [23]. For instance nicotine, rotenone and pyrethrum were famous and among the botanical insecticides used in those days [24]. Those chemicals from pesticidal plants possess one or more useful properties like repellency, anti-feeding, fast knock down, flushing action, biodegradability, broad spectrum of activity and ability to reduce insect resistance [24][44]. Therefore, there is a need to use the environmental friendly products for instance, the botanicals/phytochemicals from Tephrosia vogelii, Syzygium aromaticum and Croton dichogamus to control cabbage insect pests in the field.

1.2.4. The Cabbage Aphids, (Brevicoryne brassicae)

Cabbage aphid (Figure 4A) (Brevicoryne brassicae) belongs to the family Aphididae of the order Hemiptera [45] and the genus Brevicoryne [46]. The name is derived from two Latin words “brevi” and “coryne” and which means “small pipes” [45]. In those aphids, there are two small pipes called cornicles or siphunculi at the posterior end which can be observed when using hand lens during the observation [47]. The cornicles of the cabbage aphid are comparatively shorter than the cornicles of other aphids except those of the turnip aphid, Lipaphis erysimi [47]. The short cornicles and the waxy coating present on cabbage aphids differentiate cabbage aphids from other aphids which can attack the same host plants [47][48]. The cabbage aphid is native to Europe, but now has a world wide distribution [48][49] and can be found in Africa, Asia, Canada, Australia [50], America, India, China and Netherland [46] and also in African countries.

Figure 4. (A) Cabbage aphids, Brevicoryne brassicae. (B) The damaged plant cabbage by cabbage Aphids. Photograph by Nelson Mpumi. NM-AIST, Arusha, Tanzania.

Jahan et al. [51] and Moharramipour et al. [52] indicated that, cabbage aphids are serious plant sap sucking pests worldwide. Those aphids are the most common damaging species causing significant yield loss to many crops of Brassicaceae, like the mustards and crucifers [52][53]. Blackman and Eastop [54] insisted that, cabbage aphids mostly attack growing parts of the host plants such as tips, flowers, developing pods, leaves and eventually cover the whole plants (Figure 4B) at high population. According to Elwakil and Mossler [55] and Lashkari et al. [56] cabbage aphids (Figure 4A) have direct and indirect damaging effects to cabbage crops. The direct damage caused by this pest is by sucking cell sap, secrete honey dew which result into sooty mold formation on leaves and shoots and indirect damaging effect is as a vector of 20 plant viral diseases in a wide range of plants. According to Valenzuela and Hoffmann [57], the damaging viruses transmitted by cabbage aphids are such as potato leafroll virus, potyviruses, beet western yellows, beet yellows, cauliflower mosaic, cucumber mosaic, lettuce mosaic, turnip mosaic and watermelon mosaic. High population and feeding of cabbage aphids result into curling, distortion and yellowing of leaves, stunting plant growth, deforming developing heads, damaging of flowers and green pods and discoloration of any growth stage and part of plants [47][50]. Feeding by cabbage aphids can stop terminal growth resulting into reduced plant size and yield [58].

Eliminating weeds in Brassicaceae field borders is one of the cultural methods which may help to reduce the population and damaging of the cabbage aphids [56][59]. However, cultural methods alone are less effective to completely control the cabbage aphids from the farmers’ field [45]. So, biological control can play a major role in the natural suppression of aphids. Among the biological controls which can be applied to control the aphids are the natural enemies such as ladybird beetles adult and larvae, lacewing larvae, syrphid fly larvae, predatory bugs and lacewing larvae [56][60]. Other biological control agents are entomopathogenic fungi, which particularly can be applied during the periods of high humidity and precipitation [55][59]. However, natural enemies alone and other biological controls are also insufficient to prevent economic damage by a rapidly increasing population of cabbage aphids [61].

Due to high pest pressure and damaging caused by those aphids on cabbages in African countries, growers resort to excessive and intensive chemical pesticides application for aphids and other insect pest management [45][60]. Chemical pesticides are intensively, excessively and doubly rated for insect pest management [45][60]. However, intensive and heavily reliance on the application of the synthetic pesticides results into extreme soil and water pollution and pose serious threats to the non-target organisms including human beings [45]. For instance, Bami [62] reported that, every year, one million people are suffering from pesticide poisoning in India. The pesticides poisoning threatens the health of human being and the natural enemies. Also, the soil pollution threatened the soil ecosystem. Decomposers are also in danger due to soil pollution through excessive and intensive application of synthetic pesticides [17].

Due to those problems associated with the application of synthetic pesticides, there is a need of assessing the potential of botanical pesticides from various plants such as T. vogelli, S. aromaticum and C. dichogamus for cabbage aphid control and management in the field. Botanicals from different pesticidal plants have many advantages over synthetic pesticides such as local availability and inexpensive pest control agents [45][63].

1.2.5. The Green Peach Aphids (Myzus persicae)

The green peach aphid, (Myzus persicae) (Figure 5A), is found throughout the world and can be present at any time throughout the year [64]. Generally, its color is pale green, and there are two forms of green peach aphids; winged and wingless forms [65]. The green peach aphid have prominent cornicles on the abdomen that are markedly swollen and club like in appearance [54]. The frontal tubercles at the base of the antennae are very prominent and are convergent [65]. Winged forms of the green peach aphid have a distinct dark patch near the tip of the abdomen; wingless forms lack this dark patch [66]. The green peach aphid is adapted to high environmental temperatures [64].

Figure 5. (A) Green peach aphids, Myzus persicae. (B) The cabbage affected by green peach aphids. Photograph by Nelson Mpumi. NM-AIST, Arusha, Tanzania.

Blackman and Eastop [54] and Gu et al. [64] showed that, over 40 plant families are hosts of green peach aphids. According to them, those plants include woody and herbaceous plants including vegetable crops in the family Solanaceae, Chenopodiaceae, Compositaceae, Brassicaceae, and Cucurbitaceae. Some of the host plants which support the growth and development of green peach aphids include cabbages, spinach, asparagus, bean, beets, broccoli, Brussels sprouts, carrot, cauliflower, cantaloupe, celery, corn, cucumber, fennel, kale, turnip, eggplant, lettuce, mustard, okra, parsley, parsnip, pea, pepper, potato, radish, squash, tomato, turnip, watercress and watermelon [54]. Moreover, Gu et al. [64] added that, many flower crops and ornamental plants are also suitable for growth and development of green peach aphids. Those all crops differ in their vulnerability to green peach aphids, but the actively growing plants and plants’ parts, or the youngest plant tissues often are affected by large aphid populations [66]. Broadleaf vegetables are particularly very suitable host plants for green peach aphids. Therefore, the broadleaf vegetables create pest infestation problems in nearby crops [64]. The green peach aphids can achieve very high densities on young plant tissues, causing water stress, wilting and reduced growth rate of the plant [65].

Anstead et al. [67] and Umina et al. [66] indicated that, adults and nymphs of aphids can damage the crops in three ways: - firstly, they feed directly on young tender plant tissues and causes drying out of shoots, wilting and distortions of the plants’ parts (Figure 5B); secondly, they produce honeydew which falls onto foliage and becomes blackened by sooty mould fungi; and thirdly, they spread more than 100 viruses. According to Anstead et al. [67], de Little and Umina [68] and Valenzuela and Hoffmann [69], the damaging viruses transmitted by green peach aphids are such as potato leafroll, potyviruses in pepper, beet western yellows, beet yellows, cauliflower mosaic, cucumber mosaic, lettuce mosaic, papaya ringspot, turnip mosaic and watermelon mosaic. These viruses affect the proper growth and development of the crops and reduce the marketability. The damaging levels caused by green peach aphids are characterized by large numbers of aphids found on the underside of leaves sucking the plant saps [67][68]. In addition to attacking plants in the field, the green peach aphid can readily infest vegetables and ornamental plants grown in glasshouses [64]. Umina et al. [66] reported that, the aphids feed by sucking sap from leaves and flower buds, but the entire crop foliage may be covered when populations are large resulting in reduced or stunted growth of young plants. The extensive feeding of green peach aphids on crops enforces plants to turn yellow and the leaves to curl downward and inward from the edges resulting into wilting, stunted growth and finally death of the crops [68]. When young plants are infested in glasshouses and then transplanted into the field, the fields will not only be inoculated with aphids but insecticide resistance may be introduced [64]. de Little and Umina [68] insisted that, the green peach aphid is considered the most important vector of plant viruses in the world. Also, contamination of harvestable plant material with aphids, or aphid honeydew, causes the loss of the food quality and quantity [70]. Therefore, prolonged aphid infestation of crops can reduce the yield of crop products.

The green peach aphid is attacked by a number of common predators such as lacewings, lady beetles, syrphid flies and parasites, including the parasitic wasps (Lysiphlebus testaceipes, Aphidius matricariae, Aphelinus semiflavus, and Diaeretiella rapae), and is susceptible to the fungus disease, Entomophthora spp. All those natural enemies together with field sanitation helps to control the incidence and spread of viruses transmitted by green peach aphid, but it does little to control the aphid itself. So, the smallholder farmers rely on the application of chemical insecticides to control the green peach aphids in the field. The use of chemical pesticides to control M. persicae on the food crops is increasing globally [71]. For instance in African countries like Tanzania, M. persicae are now extensively controlled with insecticides in oilseeds, pulses, and vegetable crops [65]. However, heavy reliance on insecticides to manage aphid populations result into strong insect pest resistance and M. persicae has probably developed resistance to more insecticides than any other insect species [71][72]. Therefore, broad spectrum insect pest control strategies are needed to ensure the aphids are controlled.

The severe damaged caused by insect pests in various parts of B. oleracea (Table 3 and Figure 5) compel the African smallholder farmers to increases the doses of the synthetic pesticides during the application.

Table 4. The parts of Brassica oleracea damaged by insect pests, signs and their effects [10][36][44][73].

1.2.6. Brassica oleracea Insect Pests with Insecticides' Resistance

Some of the important pests of Brassica oleracea such as the Diamond Back Moth (Plutella xylostella), cabbage webworms (Hellula undalis), whiteflies (Bemicia tabaci) and aphids (Brevicoryne brassicae and Myzus persicae) have developed resistance to a wide range of commonly used pesticides [74]. For instance, Plutella xylostella is documented to have developed resistance to a number of insecticides [75]. The tests done in four regions in New Zeland between 1999 and 2000 reported that, P. xylostella developed resistance to synthetic pyrethroids [76]. The resistance of P. xylostella to pyrethroids is based on the oxidative detoxification of monooxygenase enzymes [77]. The level of resistance of P. xylostella to cypermethrin can be 1096 fold [78]. However, the resistance of P. xylostella to pyrethroids insecticides can be even 2880 fold [75]. Varma and Sandhu [79] reported the resistance of P. xylostella to DDT and parathion organochlorine insecticides in India. Also, P. xylostella is reported to developed resistance to fenitrothion and malathion [80], cypermethrin, decamethrin and quinalphos [81], cartap hydrochloride, diafenthiuron and flufenexuron [75][80]. The major reasons for P. xylostella to develop resistance to insecticides includes: the increase in number of sprays, misuse of pesticides, inappropriate dosages used by farmers and frequency of applications [82]. Apart from the insecticides resistance developed by P. xylostella, also cabbage webworms (Hellula undalis), whiteflies (Bemicia tabaci), aphids (Brevicoryne brassicae and Myzus persicae) have developed resistance to cypermethrin, decamethrin, chlorpyrfors, malathion and lamda-cyhalothrine [76]. Therefore, Brassica oleracea insect pest resistance to synthetic pesticides calls for search of the alternatives products which can effectively control those insect pests in the field.


  1. Basak, H.; Biswas, B.; Azad, M.; Arifuzzaman, M.; Sharmeen, F. Micropropagation of Mustard (Brassica spp.) from Leaf Explants. Thai J. Agric. Sci. 2012, 45, 75–81.
  2. Singh, S.; Cheema, G.; Bhatia, D. Evaluation of Plant Regeneration and Somaclonal Variation in Brassica Juncea L (Coss and Czern); World Vegetable Center: Tainan, Taiwan, 2001.
  3. Zhang, F.; Takahata, Y.; Xu, J. Plant regeneration from cotyledonary explants of Chinese cabbage cultured in vitro. Hort. Sin. 2002, 29, 348–352.
  4. W. Msikita; H.T. Wilkinson; R.M. Skirvin; Propagation of Tronchuda (Brassica oleracea var. tronchuda Bailey) from Cuttings. HortScience 1992, 27, 1036-1038, 10.21273/hortsci.27.9.1036.
  5. S. Eapen; V. Abraham; M. Gerdemann; O. Schieder; Direct Somatic Embryogenesis, Plant Regeneration and Evaluation of Plants Obtained from Mesophyll Protoplasts of Brassica juncea). Annals of Botany 1989, 63, 369-372, 10.1093/oxfordjournals.aob.a087753.
  6. Sharma, M.; Gupta, S. Effective Callus Induction and Plant Regeneration in Brassica napus (L.) Var Dgs-1. J. Cell Tissue Res. 2012, 12, 3229–3234.
  7. Dubey, S.K.; Gupta, A.K. Callus induction and shoot proliferation from seedling explants of different mustard genotypes. Int. J. Curr. Microbiol. App. Sci. 2014, 3, 858–864.
  8. JohnE. Hachey; KiranK. Sharma; MauriceM. Moloney; Efficient shoot regeneration of Brassica campestris using cotyledon explants cultured in vitro. Plant Cell Reports 1991, 9, 549–554, 10.1007/bf00232329.
  9. Smartt, J.; Simmonds, N.W. Evolution of Crop Plants, 2nd ed.; Longman Scientific & Technical: Harlow, UK, 1995; pp. 82–86.
  10. Baidoo, P.; Mochiah, M. Comparing the effectiveness of garlic (Allium sativum L.) and hot pepper (Capsicum frutescens L.) in the management of the major pests of cabbage Brassica oleracea (L.). Sustain. Agric. Res. 2016, 5, 83–91.
  11. P. K. Baidoo; J. I. Adam; The Effects of Extracts of Lantana camara (L.) and Azadirachta indica (A. Juss) on the Population Dynamics of Plutella xylostella, Brevicoryne brassicae and Hellula undalis on Cabbage. Sustainable Agriculture Research 2012, 1, 229–234, 10.5539/sar.v1n2p229.
  12. Ja Timbilla; Ko Nyarko; A survey of cabbage production and constraints in Ghana. Ghana Journal of Agricultural Science 2004, 37, 93–101, 10.4314/gjas.v37i1.2084.
  13. Krishnamoorthy, A. Biological control of diamondback moth Plutella xylostella (L.), an Indian scenario with reference to past and future strategies. In Proceedings of the International Symposium, Montpellier, France, 21–24 October 2002; Kirk, A.A., Bordat, D., Eds.; CIRAD: Montpellier, France, 2004.
  14. Waiganjo, M.; Waturu, C.; Mureithi, J.; Muriuki, J.; Kamau, J.; Munene, R. Use of entomopathogenic fungi and neem bio-pesticides for Brassica pests control and conservation of their natural enemies. East Afric. Agric. For. J. 2011, 77, 545–549.
  15. Claude Ahouangninou; Benjamin E. Fayomi; Thibaud Martin; Assessing health and environmental risks as regards pesticide practices of vegetable growers in the rural city of Tori-Bossito in southern Benin. Cahiers Agricultures 2011, 20, 216-222, 10.1684/agr.2011.0485.
  16. A.V.F. Ngowi; T.J. Mbise; A.S.M. Ijani; L. London; O. C. Ajayi; Smallholder vegetable farmers in Northern Tanzania: Pesticides use practices, perceptions, cost and health effects. Crop Protection 2007, 26, 1617-1624, 10.1016/j.cropro.2007.01.008.
  17. John J. Ondieki; The current state of pesticide management in Sub-Saharan Africa. Science of The Total Environment 1996, 188, 30–34, 10.1016/s0048-9697(96)90506-9.
  18. Lingren, P.; Green, G. Suppression and Management of Cabbage Looper Populations; Agricultural Research Service: Washington, DC, USA, 1984.
  19. Chomchalow, N; Protection of stored products with special reference to Thailand. AU J. 2003, 7, 34–47.
  20. Capinera, J. Handbook of Vegetable Pests; Academic Press: Cambridge, MA, USA, 2001.
  21. K. O. Fening; B. W. Amoabeng; I. Adama; M. B. Mochiah; H. Braimah; M. Owusu-Akyaw; E. Narveh; S. O. Ekyem; Sustainable management of two key pests of cabbage, Brassica oleracea var. capitata L. (Brassicaceae), using homemade extracts from garlic and hot pepper. Organic Agriculture 2013, 3, 163-173, 10.1007/s13165-014-0058-2.
  22. Blankson W. Amoabeng; Geoff M. Gurr; Catherine W. Gitau; Helen I. Nicol; Louis Munyakazi; Philip C. Stevenson; Correction: Tri-Trophic Insecticidal Effects of African Plants against Cabbage Pests. PLOS ONE 2013, 8, 1–10, 10.1371/annotation/f0351003-b6f8-4249-ace5-bcd84dead916.
  23. Begna, F.; Damtew, T. Evaluation of four botanical insecticides against Diamondback Moth, Plutella Xylostella L.(Lepidoptera: Plutellidae) on head cabbage in the central rift valley of Ethiopia. Sky J. Agric. Res. 2015, 4, 97–105.
  24. Murray Isman; BOTANICAL INSECTICIDES, DETERRENTS, AND REPELLENTS IN MODERN AGRICULTURE AND AN INCREASINGLY REGULATED WORLD. Annual Review of Entomology 2006, 51, 45-66, 10.1146/annurev.ento.51.110104.151146.
  25. Sivapragasam, A.; Aziz, A. Cabbage webworm on crucifers in Malaysia. In Diamondback Moth and Other Crucifer Pests, Proceedings of the Second International Workshop, Tainan, Taiwan, 10–14 December 1990; Talekar, N.S., Ed.; Asian Vegetable Research and Development Center: Taipei, Taiwan, 1992.
  26. Pérez-Lucas, G.; Vela, N.; El Aatik, A.; Navarro, S. Environmental Risk of Groundwater Pollution by Pesticide Leaching through the Soil Profile. In Pesticides, Anthropogenic Activities and the Health of Our Environment; IntechOpen: London, UK, 2018.
  27. Adama A. Ebenebe; Saidi R. Achari; Nitesh Chand; Annas A. Krishna; Saula Baleisuva; The cabbage webworm (Hellula undalis) on tickweed (Cleome viscosa) in Samoa. The South Pacific Journal of Natural and Applied Sciences 2011, 29, 1-6, 10.1071/sp11001.
  28. Waterhouse, D.F.; Sands, D.P.A. Classical Biological Control of Arthropods in Australia; Australian Centre for International Agricultural Research: Bruce Town, Australia, 2001.
  29. Shine, C.; Reaser, J.; Gutierrez, A. Invasive Alien Species in the Australa; Global Invasive Species Programme; National Reports & Directory of Resources: Cape Town, South Africa, 2003.
  30. Waterhouse, D.F.; Norris, K.R. Biological Control: Pacific Prospects; Inkata Press: Melbourne, Australia, 1987.
  31. Babacar Labou; Thierry Brévault; Serigne Sylla; Mamadou Diatte; Dominique Bordat; Karamoko Diarra; Spatial and temporal incidence of insect pests in farmers’ cabbage fields in Senegal. International Journal of Tropical Insect Science 2017, 37, 225-233, 10.1017/s1742758417000200.
  32. Zhenyu Li; X. Feng; Shu-Sheng Liu; Minsheng You; Michael J. Furlong; Biology, Ecology, and Management of the Diamondback Moth in China. Annual Review of Entomology 2016, 61, 277-296, 10.1146/annurev-ento-010715-023622.
  33. Z. Li; Myron P Zalucki; T. Yonow; Darren J. Kriticos; H. Bao; H. Chen; Z. Hu; X. Feng; Michael J. Furlong; Population dynamics and management of diamondback moth (Plutella xylostella) in China: the relative contributions of climate, natural enemies and cropping patterns. Bulletin of Entomological Research 2016, 106, 197-214, 10.1017/s0007485315001017.
  34. Francisco Rubén Badenes-Pérez; Brian A. Nault; Anthony M. Shelton; Dynamics of diamondback moth oviposition in the presence of a highly preferred non-suitable host. Entomologia Experimentalis et Applicata 2006, 120, 23-31, 10.1111/j.1570-7458.2006.00416.x.
  35. Justus, K.; Mitchell, B. Oviposition site selection by the diamondback moth, Plutella xylostella (L.)(Lepidoptera: Plutellidae). J. Insect Behav. 1996, 9, 887–898.
  36. Katinka Weinberger; R. Srinivasan; Farmers' management of cabbage and cauliflower pests in India and their approaches to crop protection. Journal of Asia-Pacific Entomology 2009, 12, 253-259, 10.1016/j.aspen.2009.08.003.
  37. G. Ayalew; Comparison of yield loss on cabbage from Diamondback moth, Plutella xylostella L. (Lepidoptera: Plutellidae) using two insecticides. Crop Protection 2006, 25, 915-919, 10.1016/j.cropro.2005.12.001.
  38. M Mohan; G.T. Gujar; Local variation in susceptibility of the diamondback moth, Plutella xylostella (Linnaeus) to insecticides and role of detoxification enzymes. Crop Protection 2003, 22, 495-504, 10.1016/s0261-2194(02)00201-6.
  39. Charleston, D.S.; Gols, R.; Hordijk, K.A.; Kfir, R.; Vet, L.E.; Dicke, M; Impact of botanical pesticides derived from Melia azedarach and Azadirachta indica plants on the emission of volatiles that attract parasitoids of the diamondback moth to cabbage plants. Journal of Chemical Ecology 2006, 39, 105–114.
  40. Ralf Schulz; Sue Kc Peall; James Dabrowski; Adriaan J Reinecke; Current-use insecticides, phosphates and suspended solids in the Lourens River, Western Cape, during the first rainfall event of the wet season. Water SA 2001, 27, 65-70, 10.4314/wsa.v27i1.5012.
  41. Dalvie, M.A.; Cairncross, E.; Solomon, A.; London, L; Contamination of rural surface and ground water by endosulfan in farming areas of the Western Cape, South Africa. Environmental Health 2003, 2, 1–15.
  42. Pedigo, L.P.; Rice, M.E. Entomology and Pest Management; Waveland Press: Long Grove, IL, USA, 2014.
  43. Patel, P.; Shukla, N.; Patel, G. Enhancing insecticidal properties of cow urine against sucking pests of cotton. In Proceedings of the National Symposium on Fronterier Areas of Entomological Research, New Delhi, India, 5–7 November 2003.
  44. Mochiah, M.; Banful, B.; Fening, K.; Amoabeng, B.; Ekyem, S.; Braimah, H.; Owusu-Akyaw, M. Botanicals for the management of insect pests in organic vegetable production. J. Entomol. Nematol. 2011, 3, 85–97.
  45. Mersha, W.; Ayele, N.; Fentahun, G.; Getinet, M.; Kassu, K.; Nagappan, R. Repellent and insecticidal activity of Mentha piperita (L.) plant extracts against cabbage aphid [Brevicoryne brassicae Linn. (Homoptera: Aphididae)]. Am. Eurasian J. Sci. Res. 2014, 9, 150–156.
  46. Gill, H.K.; Garg, H.; Gillett-Kaufman, J.L. Cabbage aphid, Brevicoryne brassicae Linnaeus (Insecta: Hemiptera: Aphididae); IFAS Extension University of Florida: Gainesville, FL, USA, 2013.
  47. Carter, C.; Sorensen, K. Insect and related pests of vegetables. In Cabbage and Turnip Aphid; Center for Integrated Pest Management; North Carolina State University: Raleigh, NC, USA, 2013.
  48. Opfer, P.; McGranth, D. Oregon Vegetables, Cabbage Aphid and Green Peach Aphid; Department of Horticulture Oregon State University Corvallis: Corvallis, OR, USA, 2013.
  49. Beata Gabryś; H. J. Gadomski; Z. Klukowski; J. A. Pickett; G. T. Sobota; L. J. Wadhams; C. M. Woodcock; Sex Pheromone of Cabbage Aphid Brevicoryne brassicae: Identification and Field Trapping of Male Aphids and Parasitoids. Journal of Chemical Ecology 1997, 23, 1881-1890, 10.1023/b:joec.0000006457.28372.48.
  50. Mushtaq Ahmad; Shamim Akhtar; Development of insecticide resistance in field populations of Brevicoryne brassicae (Hemiptera: Aphididae) in Pakistan. Journal of Economic Entomology 2013, 106, 954-958, 10.1603/EC12233.
  51. Fatemeh Jahan; Habib Abbasipour; Alireza Askarianzade; Gholamhossein Hassanshahi; AyatAllah Saeedizadeh; Effect of eight cauliflower cultivars on biological parameters of the cabbage aphid,Brevicoryne brassicae(L.) (Hem: Aphididae) in laboratory conditions. Archives of Phytopathology and Plant Protection 2013, 46, 636-642, 10.1080/03235408.2012.749697.
  52. Moharramipour, S.; Monfared, A.; Fathipour, Y. Comparison of intrinsic rate of increase and mean relative growth rate of cabbage aphid (Brevicoryne brassicae L.) on four rapeseed (Brassica napus L.) varieties in growth room. Agric. Sci. 2003, 13, 79–89.
  53. Mudzingwa, S.; Muzemu, S.; Chitamba, J. Pesticidal efficacy of crude aqueous extracts of Tephrosia vogelii L., Allium sativum L. and Solanum incanum L. in controlling aphids (Brevicoryne brassicae L.) in rape (Brassica napus L.) n controlling aphids (Brevicoryne brassicae L.) in rape (Brassica napus L.). J. Agric. Res. 2013, 2, 157–163.
  54. Blackman, R.L.; Eastop, V.F. Aphids on the World’s Crops: An Identification and Information Guide; John Wiley & Sons Ltd.: Hoboken, NJ, USA, 2000.
  55. Elwakil, W.M.; Mossler, M. Florida Crop/Pest Management Profile: Cabbage; University of Florida: Gainesville, FL, USA, 2016; Volume 1256, p. 18.
  56. Mohammadreza Lashkari; Ahad Sahragard; Mohammad Ghadamyari; Sublethal effects of imidacloprid and pymetrozine on population growth parameters of cabbage aphid, Brevicoryne brassicae on rapeseed, Brassica napus L. Insect Science 2007, 14, 207-212, 10.1111/j.1744-7917.2007.00145.x.
  57. Mudzingwa, S.; Muzemu, S.; Chitamba, J; Pesticidal efficacy of crude aqueous extracts of Tephrosia vogelii L., Allium sativum L. and Solanum incanum L. in controlling aphids (Brevicoryne brassicae L.) in rape (Brassica napus L.) n controlling aphids (Brevicoryne brassicae L.) in rape (Brassica napus L.). J. Agric. Res. 2013, 2, 157–163.
  58. Liu, T.X.; Sparks, A.N., Jr. Aphids on Cruciferous Crops: Identification and Management; Texas A&M University: College Station, TX, USA, 2001.
  59. Liu, Y.Q.; Shi, Z.-H.; Zalucki, M.P.; Liu, S.-S; Conservation biological control and IPM practices in Brassica vegetable crops in China. Biol. Control. 2014, 68, 37–46.
  60. Hines, R.; Hutchison, W. Cabbage Aphids. VegEdge, Vegetable IPM Resource for the Midwest; University of Minnesota: Minneapolis, MN, USA, 2013.
  61. Zaki, F; Field application of plant extracts against the aphid, B. brassicae and the whitefly, B. abaci and their side effects on their predators and parasites. Arch. Phytopathol. Pflanzenschutz. 2008, 41, 462–466.
  62. Bami, H. Pesticide use in India-Ten questions. Chem. Wkly. Bombay 1997, 42, 137–146.
  63. Mkenda, P.; Mwanauta, R.; Stevenson, P.C.; Ndakidemi, P.; Mtei, K.; Belmain, S.R; Extracts from field margin weeds provide economically viable and environmentally benign pest control compared to synthetic pesticides. PLoS ONE 2015, 10, 1-14.
  64. Gu, H.; Fitt, G.P.; Baker, G.H; Invertebrate pests of canola and their management in Australia: A review. Aust. J. Entomol. 2007, 46, 231–243.
  65. Edwards, O.R.; Franzmann, B.; Thackray, D.; Micic, S; Insecticide resistance and implications for future aphid management in Australian grains and pastures: A review. Aust. J. Exp. Agric. 2008, 48, 1523–1530.
  66. Umina, P.A.; Edwards, O.; Carson, P.; Van Rooyen, A.; Anderson, A; High levels of resistance to carbamate and pyrethroid chemicals widespread in Australian Myzus persicae (Hemiptera: Aphididae) populations. J. Econ. Entomol. 2014, 107, 1626–1638.
  67. Anstead, J.; Mallet, J.; Denholm, I; Temporal and spatial incidence of alleles conferring knockdown resistance to pyrethroids in the peach–potato aphid, Myzus persicae (Hemiptera: Aphididae), and their association with other insecticide resistance mechanisms. Bull. Entomol. Res. 2007, 97, 243–252.
  68. De Little, S.C.; Umina, P.A; Susceptibility of Australian Myzus persicae (Hemiptera: Aphididae) to three recently registered insecticides: Spirotetramat, cyantraniliprole, and sulfoxaflor. J. Econ. Entomol. 2017, 110, 1764–1769.
  69. Stewart, J.K.; Aharoni, Y.; Hartsell, P.L.; Young, D.K; Acetaldehyde fumigation at reduced pressures to control the green peach aphid on wrapped and packed head lettuce. J. Econ. Entomol. 1980, 73, 149–152.
  70. Stewart, J.K.; Aharoni, Y.; Hartsell, P.L.; Young, D.K; Acetaldehyde fumigation at reduced pressures to control the green peach aphid on wrapped and packed head lettuce. J. Econ. Entomol. 1980, 73, 149–152.
  71. Sparks, T.C.; Nauen, R; IRAC: Mode of action classification and insecticide resistance management. Pestic. Biochem. Physiol. 2015, 121, 122–128.
  72. Whalon, M.; Mota-Sanchez, D.; Hollingworth, R. Analysis of Global Pesticide Resistance in Arthropods; CABI Publishing: Wallingford, UK, 2008; pp. 5–31.
  73. Kasina, M.A.; Kraemer, M.; Holm-Mueller, K; Economic Benefit of Crop Pollination by Bees: A case of Kakamega Small-holder Farming in Western Kenya. J. Econ. Entomol. 2009, 102, 467–473.
  74. Seif, A.; Nyambo, B. Integrated Pest. Management for Brassica Production in East Africa; ICIPE Science Press: Addis Ababa, Ethiopia, 2013.
  75. Vuković, S.; Inđić, D.; Gvozdenac, S.; Červenski, J; Efficacy of Insecticides in the Control of Cabbage Pests. Res. J. Agric. Sci. 2014, 46, 421–425.
  76. Walker, G.; Cameron, P.; Berry, N. Implementing an IPM programme for vegetable brassicas in New Zealand. In The management of diamondback moth and other crucifer pests. In Proceedings of the Fourth International Workshop, Melbourne, Australia, 26–29 November 2001.
  77. Sun, C.; Tsai, Y.; Chiang, F. Resistance in the Diamondback Moth to Pyrethroids and Benzoylphenylureas; ACS Publications: Washington, DC, USA, 1992.
  78. Takahashi, H.; Mitsui, J.; Takakusa, N.; Matsuda, M.; Yoneda, H.; Suzuki, J.; Ishimitsu, K.; Kishimoto, T. NI-25, a new type of systemic and broad spectrum insecticide. In Proceedings of the Brighton Crop Protection Conference, Pests and Diseases, Brighton, UK, 23–26 November 1992.
  79. Verma, A.; Sandhu, G; Chemical control of diamondback moth, Plutella maculipennis (Curtis). J. Res. Punjab Agric. Univer. 1968, 5, 420–423..
  80. Joia, B.; Chawla, R.; Udeaan, A. Present insecticide use practices on cole crops in Punjab and strategies for managing multiple insecticide resistance in diamondback moth. In Proceedings of the XX International Congress of Entomology, Florence, Italy, 25–31 August 1996.
  81. Saxena, J.; Rai, S.; Srivastava, K.; Sinha, S; Resistance in the field populations of the diamondback moth to some commonly used synthetic pyrethroids. Ind. J. Entomol. 1989, 51, 265–268.
  82. Imran, M.; Kanwal Hanif, M.A.; Nasir, M.; Sheikh, U.A.A; Comparative Toxicity of Insecticides against Two Important Insect Pests of Cauliflower Crop. Asian J. Agric. Biol. 2017, 5, 88–98.
Subjects: Entomology
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to : , , ,
View Times: 2708
Revisions: 3 times (View History)
Update Date: 28 Oct 2020