Oryctes rhinoceros (coconut rhinoceros beetle)
- Summary of Invasiveness
- Taxonomic Tree
- Distribution Table
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Links to Websites
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Oryctes rhinoceros (Linnaeus)
Preferred Common Name
- coconut rhinoceros beetle
Other Scientific Names
- Oryctes stentor Castelnau, 1840
- Scarabaeus rhinoceros Linnaeus
International Common Names
- English: Asiatic rhinoceros beetle; black beetle; coconut black beetle; coconut palm rhinoceros beetle; date palm beetle; dung beetle; rhinoceros beetle; scarab beetle
- Spanish: escarabajo rinoceronte Asiático
- French: oryctes du cocotier; rhinoceros du cocotier
Local Common Names
- Germany: Indischer nashornkaefer
- Indonesia: kumbang badak; kumbang tanduk
- Netherlands: klappertor
- ORYCRH (Oryctes rhinoceros)
Summary of InvasivenessTop of page
O. rhinoceros is included in the Global Invasive Species Database (ISSG, 2009).
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Arthropoda
- Subphylum: Uniramia
- Class: Insecta
- Order: Coleoptera
- Family: Scarabaeidae
- Subfamily: Dynastinae
- Genus: Oryctes
- Species: Oryctes rhinoceros
DescriptionTop of page
O. rhinoceros eggs are yellowish-white, measuring 3 mm in diameter and laid inside rotting vegetative matter. Initially oval in shape, they begin to swell about a week after laying and hatch within 11-13 days (Wood, 1968a).
The larval stages are usually yellowish-white in colour and may grow to about 60-100 mm long, or more (Wood, 1968a; Ooi, 1988). The maximum head capsule width is about 10.6-11.4 mm. The cranium is medium to dark brown, with numerous round pits, many of which bear minute setae. Thoracic spiracles are 1.85-2.23 mm long, 1.25-1.53 mm wide. Respiratory plate with a maximum of 40-80 or more small, round to oval holes along any diameter. Thoracic spiracles larger than abdominal spiracles; first abdominal spiracle somewhat smaller than succeeding spiracles (for detailed information, see Bedford (1974)). The larval stages may look similar to other species within the scarabeid family (e.g. Xylotrupes gideon, Scapanes australis, Trichogomphus fairmairei, Oryctes centaurus and Oryctoderus sp.). A key to differentiate the larval stages of these species has been provided by Bedford (1974). The presence of the organ of Herold on the ventral surface of the ninth abdominal segment indicates the male sex (Hurpin, 1953; Elliott, 1964). A description has been provided for third-instar larva of Chalcosoma atlas (Bedford, 1976a) which co-exists in Malaysia with O. rhinoceros and may be confused with it, while in the African region O. monoceros is widespread, for which a description of the larva is available (Bedford, 1979). Lucanid and cetoniine beetles, which often shared the same breeding sites, may also be confused with the early instars of O. rhinoceros (Wood, 1968a) and a key for separating some of these types of larvae has been provided for Papua New Guinea (Beaudoin-Ollivier et al., 1998, 2000).
The prepupa is somewhat similar in appearance to the larval stage, except that it is smaller than the final larval instar. Shrivelled in appearance, it shakes its body actively when disturbed.
The O. rhinoceros pupa is yellowish-brown in colour and measures up to 50 mm in length. It is segmented on the dorsal surface. The length of the horn-shaped protuberances on the anterior may indicate the sex of the adult.
Stout-looking adults, dark brown to black, shiny, 35-50 mm long and 20-23 mm wide, with a prominent horn on head (Wood, 1968a; Bedford, 1974). The males having a relatively longer horn than the female. The males can be differentiated more accurately by having a rounded, shiny terminal abdominal segment while the female has a relatively hairier 'tail' (Wood, 1968a). The strikingly similar but larger species of scarabeid, Oryctes gnu (=trituberculatus) may be confused with O. rhinoceros (Wood, 1968a). The difference is the three tubercles on the thoracic ridge for O. gnu, instead of two in O. rhinoceros. A key to Oryctes species is given by Endrödi (1985).
DistributionTop of page
O. rhinoceros is endemic to the coconut-growing regions of South and South-East Asia from Pakistan to the Philippines (CIE, 1967). It was accidentally introduced into parts of Papua New Guinea in the Bismarck Archipelago (New Britain, New Ireland, Manus Island); Western and American Samoas, Tonga, Fiji, Wallis Island, Micronesia, Mauritius and the Cocos Islands (Bedford, 1974, 1980). It has recently been found in Guam and Saipan (Moore, 2007) and Hawaii (Hawaii Department of Agriculture, 2014).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.Last updated: 23 Nov 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Mauritius||Present||EPPO (2020); CABI (Undated);|
|Réunion||Present||EPPO (2020); CABI (Undated)|
|Bangladesh||Present, Widespread||APPPC (1987); EPPO (2020); CABI (Undated)|
|British Indian Ocean Territory||Present||CABI (Undated a)|
|Brunei||Present||Waterhouse (1993); EPPO (2020); CABI (Undated)|
|Cambodia||Present||Waterhouse (1993); EPPO (2020); CABI (Undated)|
|China||Present, Localized||EPPO (2020)|
|-Hainan||Present||Zhang (1984); EPPO (2020); CABI (Undated)|
|Cocos Islands||Present||EPPO (2020); CABI (Undated);|
|Hong Kong||Present||EPPO (2020); CABI (Undated)|
|India||Present, Localized||EPPO (2020)|
|-Andaman and Nicobar Islands||Present||EPPO (2020); CABI (Undated)|
|-Andhra Pradesh||Present||EPPO (2020); CABI (Undated)|
|-Assam||Present||EPPO (2020); CABI (Undated)|
|-Bihar||Present||EPPO (2020); CABI (Undated)|
|-Goa||Present||EPPO (2020); CABI (Undated);|
|-Jammu and Kashmir||Present||EPPO (2020)|
|-Karnataka||Present||Dhileepan (1992); EPPO (2020); CABI (Undated)|
|-Kerala||Present||EPPO (2020); CABI (Undated)|
|-Lakshadweep||Present||EPPO (2020); CABI (Undated)|
|-Maharashtra||Present||EPPO (2020); CABI (Undated)|
|-Manipur||Present||Gope and Prasad (1983); EPPO (2020); CABI (Undated)|
|-Nagaland||Present||Sharma and Gupta (1988); EPPO (2020); CABI (Undated)|
|-Odisha||Present||EPPO (2020); CABI (Undated)|
|-Rajasthan||Present||Bhatnagar (1971); EPPO (2020); CABI (Undated)|
|-Tamil Nadu||Present||Rajamanickam et al. (1992); EPPO (2020); CABI (Undated)|
|-West Bengal||Present||EPPO (2020); CABI (Undated)|
|Indonesia||Present, Widespread||EPPO (2020); Hallett et al. (1995); CABI (Undated)|
|-Irian Jaya||Present||EPPO (2020); CABI (Undated)|
|-Java||Present||EPPO (2020); CABI (Undated)|
|-Lesser Sunda Islands||Present||EPPO (2014)|
|-Maluku Islands||Present||EPPO (2020)|
|-Sulawesi||Present||Zelazny et al. (1992); EPPO (2020); CABI (Undated)|
|-Sumatra||Present||EPPO (2020); CABI (Undated)|
|Iran||Present||EPPO (2020); CABI (Undated)|
|Japan||Present, Localized||EPPO (2020)|
|-Ryukyu Islands||Present||Hosoya (2011); EPPO (2020); CABI (Undated)|
|Laos||Present||Waterhouse (1993); EPPO (2020); CABI (Undated)|
|Malaysia||Present, Widespread||EPPO (2020); Huger (2005)|
|-Peninsular Malaysia||Present||EPPO (2020); CABI (Undated)|
|-Sabah||Present||EPPO (2020); CABI (Undated)|
|-Sarawak||Present||EPPO (2020); CABI (Undated)|
|Maldives||Present||EPPO (2020); CABI (Undated)|
|Myanmar||Present||Waterhouse (1993); EPPO (2020); CABI (Undated)|
|Oman||Present||Anon (1986); Kinawy (2004); EPPO (2020); CABI (Undated)|
|Pakistan||Present||EPPO (2020); CABI (Undated)|
|Philippines||Present||Zelazny and Alfiler (1987); Waterhouse (1993); EPPO (2020); CABI (Undated)|
|Singapore||Present||Waterhouse (1993); AVA (2001); EPPO (2020); CABI (Undated)|
|Sri Lanka||Present||Wettasinghe and Mahindapala (1986); EPPO (2020); CABI (Undated)|
|Taiwan||Present||EPPO (2020); CABI (Undated)|
|Thailand||Present||APPPC (1987); Waterhouse (1993); EPPO (2020); CABI (Undated)|
|Vietnam||Present||Waterhouse (1993); EPPO (2020); CABI (Undated)|
|Yemen||Present||Al-Habshi et al. (2006); EPPO (2020)|
|United States||Present, Transient under eradication||EPPO (2020); Ohlendorf (1916); HDOA (2014); HDOA (2014a); Marshall et al. (2015)|
|-Hawaii||Present, Transient under eradication||NAPPO (2014); Hawaii Department of Agriculture (2014); HDOA (2014); HDOA (2014a); Marshall et al. (2015); EPPO (2020)|
|American Samoa||Present||EPPO (2020); CABI (Undated);|
|Australia||Absent, Formerly present||EPPO (2020)|
|-Queensland||Absent, Formerly present||EPPO (2020); CABI (Undated)|
|Federated States of Micronesia||Present||Bedford (1974); EPPO (2020); CABI (Undated);|
|Fiji||Present||Bedford (1986); EPPO (2020); CABI (Undated);|
|Guam||Present, Localized||Molet (2013); Moore (2007); Mankin and Moore (2010); Marshall et al. (2015); EPPO (2020)|
|New Caledonia||Present, Localized||EPPO (2020); IPPC (2019)|
|Niue||Present||EPPO (2020); CABI (Undated)|
|Northern Mariana Islands||Present||Moore (2007)|
|Palau||Present||Marshall et al. (2015); EPPO (2020); CABI (Undated)|
|Papua New Guinea||Present||Bedford (1974); Marshall et al. (2015); EPPO (2020); CABI (Undated);|
|Samoa||Present||EPPO (2020); CABI (Undated);|
|Solomon Islands||Present||EPPO (2020); Marshall et al. (2015)|
|Tokelau||Present||Uili (1980); EPPO (2020); CABI (Undated)|
|Tonga||Present||EPPO (2020); CABI (Undated);|
|Wallis and Futuna||Present||EPPO (2020); CABI (Undated);|
Risk of IntroductionTop of page McKenna and Shroff (1911) reported that the pest first appeared in the extreme south of Myanmar, probably from Malaysia in 1895, and worked its way through to the north over the next 15 years. It was also introduced to Upolu island in Western Samoa in 1909 via potted rubber seedlings from Sri Lanka (Jepson, 1912).
Habitat ListTop of page
Hosts/Species AffectedTop of page
Primarily found attacking coconut and oil palm, O. rhinoceros has also occasionally been recorded on banana (Sharma and Gupta, 1988), sugarcane, papaya, sisal and pineapple (Khoo et al., 1991). In Mauritius, ornamentals such as the royal palm (Roystonea regia), the latanier palm (Livistona chinensis), the talipot palm (Corypha umbraculifera) and the raphia palm (Raphia ruffia) are attacked (Bedford, 1980).
Host Plants and Other Plants AffectedTop of page
|Agave sisalana (sisal hemp)||Agavaceae||Other|
|Ananas comosus (pineapple)||Bromeliaceae||Other|
|Areca catechu (betelnut palm)||Arecaceae||Main|
|Carica papaya (pawpaw)||Caricaceae||Other|
|Cocos nucifera (coconut)||Arecaceae||Main|
|Colocasia esculenta (taro)||Araceae||Main|
|Elaeis guineensis (African oil palm)||Arecaceae||Main|
|Metroxylon sagu (sago palm)||Arecaceae||Other|
|Musa x paradisiaca (plantain)||Musaceae||Other|
|Phoenix dactylifera (date-palm)||Arecaceae||Main|
|Saccharum officinarum (sugarcane)||Poaceae||Other|
|Wodyetia bifurcata (foxtail palm)||Arecaceae||Other|
Growth StagesTop of page Vegetative growing stage
SymptomsTop of page
O. rhinoceros adults feed in the crown region of both coconut and oil palm. On oil palms they bore through petiole bases into the central unopened leaves. This causes tissue maceration and the presence of a fibrous frass inside and at the entrance to the feeding hole is an indication of its activity within (Catley, 1969). A single attack may be followed by others on the same palm (Barlow and Chew, 1970; Young, 1975). These attacks subsequently produce fronds which have wedge-shaped gaps or the characteristic V-shaped cuts to fronds(Wood, 1968a; Sadakathulla and Ramachandran, 1990).
List of Symptoms/SignsTop of page
|Growing point / dead heart|
|Growing point / dieback|
|Growing point / distortion|
|Growing point / internal feeding; boring|
|Leaves / abnormal forms|
|Leaves / internal feeding|
Biology and EcologyTop of page
O. rhinoceros eggs are laid in rotting palm materials including trunks, tree stumps, compost heaps, dung hills, sawdust or garbage dumps. The eggs hatch after 8-12 days and the entire larval stage is spent inside the breeding medium. Development of the larva takes 80-200 days: first instar takes 10-21 days, second instar takes 12-21 days and third instar takes 60-125 days. The beetle then enters the prepupal stage of about 8-13 days, subsequently pupating within a pupal chamber made from the food substrate. The pupal stage lasts 17-30 days. Adults may live up to 6 months or more (Bedford, 1980; Khoo et al., 1991).
Bedford (1976c, 1980) reviewed the life-cycle of O. rhinoceros and noted a wide range in the duration of the third-instar larvae (60-165 days) compared to the other stages. This may be caused by differences in climatic and nutritional conditions of different habitats. Dry climate or low nutritional conditions often delayed larval development of O. rhinoceros, which can be extended to as long as 14 months, giving rise to smaller-sized adults (Catley, 1969). Catley (1969) also demonstrated that with ample food supply, the duration of the third-instar larvae is within 3-4 months. With superior and improved larval diet combined with suitable temperatures, the duration of the larval stages can be shortened from 6 to 5 months (Schipper, 1976). Similarly, Wood (1968a) demonstrated that the larvae required about 5-7 months to mature in oil palm log tissue, while a shorter maturity period of 4-5 months was observed in a habitat mixture of cowdung and sawdust. Conditions suitable for larval development are temperature (27-29°C) and RH 85-95% (Bedford, 1980).
The adult beetles feed by boring into the growing point or meristem of coconut palms and this is the primary cause of crop damage leading to loss of yield and death in in coconut palms of all ages. Similar attacks stunt growth or may kill young oil palms. The larvae feed on dead wood; dead palm logs and stumps and the tops of dead standing coconut poles are a primary breeding site.
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Alaus farinosus||Predator||Wallis and Futuna||Cocos nucifera|
|Alaus montravelii||Predator||Wallis and Futuna||Cocos nucifera|
|Alaus speciosus||Predator||Larvae||Western Samoa||Cocos nucifera|
|Bacillus thuringiensis kurstaki||Pathogen|
|Bacillus thuringiensis thuringiensis||Pathogen|
|Brachinus stenoderus||Predator||Fiji||Cocos nucifera|
|Campsomeris azurea||Parasite||Fiji||Cocos nucifera|
|Catascopus facialis||Predator||Western Samoa||Cocos nucifera|
|Elis romandi||Parasite||Western Samoa||Cocos nucifera|
|Hololepta marginipunctata||Predator||Fiji||Cocos nucifera|
|Hololepta quadridentata||Predator||Borneo; Cocos Islands; Fiji; Malaysia; Papua New Guinea; Sabah||Cocos nucifera|
|Hypoaspis dubia||Parasite||India; Gujarat||Cocos nucifera|
|Lanelater fuscipes||Predator||Larvae||Western Samoa||Cocos nucifera|
|Mecodema spinifer||Predator||Fiji||Cocos nucifera|
|Neochryopus savagei||Predator||Larvae||American Samoa; Fiji; Papua New Guinea; Tonga; Western Samoa||Cocos nucifera|
|Ochyropus gigas||Predator||Fiji||Cocos nucifera|
|Oryctes rhinoceros nudivirus||Pathogen||Adults/Larvae/Pupae||Andaman and Nicobar Islands|
|Oxylobus punctatosulcatus||Predator||Tonga||Cocos nucifera|
|Pachylister chinensis||Predator||Larvae||American Samoa; Belau; Papua New Guinea||Cocos nucifera|
|Pheropsophus consimilis||Predator||Mauritius||Cocos nucifera|
|Pheropsophus hilaris sobrinus||Predator||Fiji; Mauritius; Wallis and Futuna||Cocos nucifera|
|Pheropsophus lissoderus||Predator||Fiji; Mauritius||Cocos nucifera|
|Pheropsophus occipitalis||Predator||Fiji; Mauritius||Cocos nucifera|
|Pheropsophus stenoderus||Predator||Fiji||Cocos nucifera|
|Placodes ebenenus||Predator||Belau||Cocos nucifera|
|Platymeris laevicollis||Predator||Adults||American Samoa; Belau; Fiji; India; Laccadive, Minicoy and Amindivi Islands; Mauritius; New Britain; Papua New Guinea; Tonga; Wallis and Futuna; Western Samoa||Cocos nucifera|
|Scarites dubiosus||Predator||American Samoa; Tonga||Cocos nucifera|
|Scarites madagascariensis||Predator||Fiji; Mauritius; Wallis and Futuna||Cocos nucifera|
|Scolia oryctophaga||Parasite||Larvae||Fiji; Java; Mauritius; New Britain; Papua New Guinea; Western Samoa||Cocos nucifera|
|Scolia patricialis||Parasite||Larvae||Belau||Cocos nucifera|
|Scolia procer||Parasite||Larvae||Belau; Mauritius; Papua New Guinea||Cocos nucifera|
|Scolia quadripustulata||Parasite||Larvae||Fiji; Tonga||Cocos nucifera|
|Scolia ruficornis||Parasite||Larvae||Belau; Chagos Archipelogo; Fiji; Malaysia; Mauritius; New Britain; Papua New Guinea; Tokelau; Wallis and Futuna||Cocos nucifera|
Notes on Natural EnemiesTop of page
The entomopathogenic virus Oryctes rhinoceros nudivirus (OrNV), which was originally discovered in Malaysia (Huger, 1966), attacks both the larval and adult stages of O. rhinoceros. It has since been introduced into Fiji (Bedford, 1976d, 1985, 2013), the Maldives and other islands in the South Pacific to control O. rhinoceros in coconut (Marschall, 1970; Marschall and Ioane, 1982; Young and Longworth, 1981; Zelazny et al., 1992).
Oryctes larvae also succumb to attack by the green entomopathogenic fungus, Metarhizium anisopliae (Sundara Babu et al., 1983). However, tests have shown that O. rhinoceros larvae are susceptible only to strains isolated from the same species (Fargues, 1976). Latch (1976) tested 34 isolates of Beauveria bassiana and B. tenella on O. rhinoceros larvae and found 17 isolates causing lesions similar to those by M. anisopliae.
Bacterial pathogens which have the potential to control larval stages of Oryctes were identified as Acinetobacter calcoaceticus (Kannan et al., 1980), Pseudomonas alcaligenes (Gopal and Gupta, 2002), Bacillus thuringiensis and B. popilliae (Sundara Babu et al., 1971; Norman et al., 2005) and Protista flagellates Monocercomonoides oryctesae and M. qadrii (Krishnamurthy and Sultana, 1977).
A list of predators and parasitoids for O. rhinoceros tested in the Pacific area has been compiled by Swan (1974), and includes Coleoptera (Elateridae, Histeridae and Carabidae), Hemiptera (Reduviidae), centipedes and vertebrates (rats, mice, shrews, squirrels, lemurs, monkeys and pigs). Wood (1968a) also reported the barn owl (Tyto alba javanica) as a predator of the beetles. Other natural enemies affecting O. rhinoceros include hymenopteran (Scoliidae) and dipteran (Tachinidae) parasitoids.
Means of Movement and DispersalTop of page
O. rhinoceros is thought to have spread throughout the Pacific by increased sea traffic during World War II. The pest has also been found in air cargo and may be transported to new areas in decaying organic materials such as compost and sawdust, and in nursery pot plants. The larvae may also be transported to new areas in floating logs.
ImpactTop of page
On oil palms, O. rhinoceros bores into the base of the cluster of unopened leaves (spears), causing V- or wedge-shaped cuts in the unfolded fronds. Spears may collapse or emerged fronds may break off along the petiole or midrib (Wood, 1968). In young palms where the spears are narrower and penetration may occur lower down, the effects of damage can be much more severe than in older palms (Wood, 1968a). The young palms affected by the beetle damage are believed to have a delayed immaturity period (Liau and Ahmad, 1991). Thus, early oil palm yields could be considerably reduced after a prolonged and serious rhinoceros beetle attack. Although Wood et al. (1973) suggested that the damage to the immature palms results in relatively small crop losses, field experiments conducted by Liau and Ahmad (1991) revealed an average of 25% yield loss over the first 2 years of production. This was possibly caused by the reduction in the canopy size of more than 15% for moderately serious to higher damage levels (Samsudin et al., 1993). In India, the infestation in oil palm was more prevalent in mature plantations (10-15 year old) compared to immature or younger plantings (Dhileepan, 1988).
Similarly, on coconut the reduction in leaf area of the palms influences nut production (Zelazny, 1979a) but the attack was more towards the tall, mature trees, from about 5 years of age onwards (Bedford, 1976c). Serious attacks on coconut may be observed in areas adjacent to a breeding site with a high beetle population, especially in the coastal region of Peninsular Malaysia. Zelazny (1979a) reported an average of 10% of fronds damaged by cuts resulted in a 3% reduction of the leaf area and a 4-5% loss of nut production; similarly 30% of fronds damaged resulted in a 7% reduction of the leaf area and 13% nut yield reduction, i.e. loss.
Detection and InspectionTop of page
On both oil palms and coconuts, O. rhinoceros bores through the petiole bases into the central unopened leaves. This causes tissue maceration and the presence of a fibrous frass inside the feeding hole is an indication of its activity within (Catley, 1969). The adults may be forced out by 'winkling' with a hooked barbed wire into the feeding hole. Larval, pupal as well as adult population may be detected and inspected by digging into or breaking open its possible breeding sites its possible breeding grounds.
Similarities to Other Species/ConditionsTop of page
The strikingly similar but larger species of scarabeid, Oryctes gnu (=trituberculatus) may be confused with the adult O. rhinoceros (Wood, 1968a). The difference is the three tubercles on the thoracic ridge for O. gnu, instead of two in O. rhinoceros. The larvae of Xylotrupes gideon are more hairy than those of O. rhinoceros. The head capsule at each larval stage is also smaller (Wood, 1968a). Aegus chelifer larvae are relatively hairless, relatively more translucent, with an orange head and a longitudinal anal slit (Wood, 1968a). Coelodera diardi larvae appear to be very similar to the second-instar O. rhinoceros larvae but are less curved. When placed on a slippery surface they swiftly move away on their backs. See Description for keys to larvae and adults, also Bedford (1976a) for a description of the Chalcosoma atlas larva from Malaysia, and Bedford (1979) for the larva of O. monoceros from the Seychelles.
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.
A method for trapping adults has been developed and tested by Hoyt (1963) and Bedford (1973). Hoyt (1963) designed a simple, cheap trap, consisting of a piece of coconut trunk, the cap with a beetle-size hole drilled through the centre, and resting on a tin can a tin can placed right below it leaving no space between them. The whole trap is set at a height of 1.8 m from the ground. There was no chemical attractant used in this trap: the decaying trunk served as the attractant. When a small quantity of the synthetic chemical attractant ethyl dihydrochrysanthemumate (chrislure) was applied to the coconut cap of the Hoyt trap, catch was increased (Bedford, 1973) compared to dispensing the lure from a more expensive metal vane-type trap (Barber et al., 1971). Chrislure was subsequently superseded by ethyl chrysanthemumate (rhinolure) (Maddison et al., 1973). Hoyt's trap was used by Bedford (1975) to monitor population trends at a density of 23 traps per 8 hectares. To control beetle infestation, the density of the traps should be increased at the borders of a known source of infestation rather than inside the field (Young, 1972). The use of light traps for controlling populations has been found to be ineffective: Wood (1968a) indicated that beetles do not often enter traps although they are attracted to the light source. However, light traps may be useful for monitoring purposes.
A male-produced aggregation pheromone, ethyl 4-methyloctanoate (E4-MO) was discovered (Hallett et al., 1995; Morin et al., 1996). It has been synthesised and is available commercially (for details of synthesis and types of traps available, see Bedford (2013a)). It is reported to be 10 times more attractive than ethyl chrysanthemumate. The pheromone is stored in a small, heat-sealed, polymer membrane bag and placed between interlocking metal vanes mounted on a plastic bucket. The beetles attracted by the pheromone are trapped inside the bucket. It is very useful as a monitoring tool, and as an economical control method particularly in young oil palm replant areas when placed at one trap per 2 ha (Norman and Basri, 2004; Oeschlager, 2007; Bedford, 2014).
Sanitation within and surrounding the plantations, especially destruction of the potential or existing breeding sites of this pest, provides an important basis for its control (Wood, 1968a; Turner, 1973). Manure heaps and pits may have to be covered or alternatively turned regularly for the removal of the grubs (Catley, 1969). The establishment of a good, fast-growing ground cover crop provides a vegetative barrier that hampers the movement of the adult beetle looking for suitable breeding sites or young oil palms in replant areas (Wood, 1968b). This restricts the damage in oil palm to low levels (Wood et al., 1973). Removal of the adults from the point of attack in young palms by using a hooked piece of wire (winkling) can be considered a common mechanical control technique to reduce the number of adults in an infested area. This practice is often costly, labour intensive and needs to be conducted regularly, provided that sufficient labour is available. However, some additional damage may be inflicted to the young palms during the search for the adults, making the practice not entirely satisfactory.
Early attempts at biological control of O. rhinoceros concentrated on the introduction of predators and scoliid parasitoids of other Oryctes species mainly from Africa. None of those that became established was able to provide satisfactory control. However, biological control effort concentrated on Oryctes rhinoceros nudivirus (OrNV) after its discovery in Malaysia in 1965 (Huger, 1966) and its successful introduction into Western Samoa in 1967 (Swan, 1974; Waterhouse and Norris, 1987). Endemic natural enemies of O. rhinoceros offer a cheap and long-term control of the pest, leading to a reduction in the use of chemical insecticides. OrNV and the pathogenic fungus Metarhizium anisopliae have been utilized further for field control of this pest in several countries (George and Kurian, 1971; Latch and Falloon, 1976; Zelazny, 1979b; Bedford, 1986; Darwis, 1990). For OrNV, the adult beetles are dipped in a suspension of ground, infected grubs. They are then allowed to crawl about for 24 hours through sterilized sawdust mixed with the above suspension. They are then released back into the plantation to infect other adults and larvae in the breeding sites (Bedford, 1976d). OrNV suspension may also be applied directly to the mouthparts of adults to infect them for release (Ramle et al., 2005). A supply of beetles for infecting and release may be obtained from a mass-rearing facility. The fungus Metarhizium anisopliae var. major may be produced commercially or in bulk by various methods, for release by suitable means into breeding sites, particularly into chipped decaying oil palm trunk material in oil palm replant areas (Sivapragasam and Tey, 1995; Tey and Ho, 1995; Ramle et al., 1999, 2006, 2007, 2009, 2011; Ramle and Kamarudin, 2013).
Most of the chemicals applied are targeted to control the adult stages attacking the spear of the palm. The point of application is therefore at the base of the leaf sheath (Sadakathulla and Ramachandran, 1990). Granular formulations of the insecticide gamma-BHC were effective as this facilitates applying them in the frond axils and thereby lowering costs compared to spraying liquid formulations (Ho and Toh, 1982). Eleven types of insecticides were evaluated for the control of the adults in the nursery and immature stages of the oil palm (Chung et al., 1991). On immature oil palm, the most effective treatment in reducing the number of broken spears and spear dieback, was lambda cyhalothrin. Cypermethrin was effective in reducing the number of holes on spears and fronds. Systemic insecticides have been used in trunk injections in tall coconut palms in Malaysia, but their effectiveness is uncertain. Insect-repellent naphthalene applied (as moth balls) fortnightly to the frond axils provided 95% control of the pest (Singh, 1987). Juvenile hormone mimics have been tested on pupae of O. rhinoceros and indicated that methoprene was effective in causing death of the developing adult at the pupal stage (Dhondt et al., 1976). The use of long residual insecticides for drenching the breeding sites (i.e. coconut stumps) has been found effective to suppress the larval stages up to 7 months (Ho and Toh, 1982). Insecticidal treatment at the bottom soil of manure pits may also be useful to suppress the larval stages (Visalakshi et al., 1988).
Field Monitoring/Economic Threshold Levels
On oil palm, the economic threshold for O. rhinoceros still needs to be investigated and developed. Wood (1968a) developed four categories of damage to young oil palms which may be used to determine the severity of infestation in a particular area.
Category 1: Slight damage - limited signs of damage to fronds with little or no damage to spears.
Category 2: Medium damage - numerous damaged fronds but with new healthy spears.
Category 3: Severe damage - many fronds damaged and only short, distorted spears emerging. Spear rot does not affect all spears.
Category 4: Dead palms - fronds severely damaged, all spears rotting and can be lifted out easily with no live spears to emerge.Some palms of this category may recover.
These categories may change between the observation periods, thus indicating the beetles' activity. If a census of 1 row in every 10 is conducted and more than 10% incidence of damage (those found in the categories mentioned) is found, this indicates high beetle activity and control measures should be conducted immediately (Wood, 1968a). The number of beetles found affecting young palms per hectare can also be considered as threshold levels for initiating control measures. IRHO (1991) recommended the detection of 3-5 adult beetles per hectare of oil palms for the first 2 years of the crop, to initiate control measures. However, the level can be increased to 15-20 adults per hectare once the palms have grown for more than 2 years (IRHO, 1991). Winkling frequency also depended on the numbers found per hectare. Monthly winkling is suggested for 3-5 adults, fortnightly for 5-10 adults and weekly for more than 10 adults per hectare (IRHO, 1991).
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19/05/15 Review by:
Geoff Bedford, Department of Biological Sciences, Macquarie University, NSW 2109, Australia
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