Cactoblastis cactorum 

Life > Eukaryotes > Opisthokonta > Metazoa (animals) > Bilateria > Ecdysozoa > Panarthropoda > Tritocerebra > Phylum: Arthopoda > Mandibulata > Atelocerata > Panhexapoda > Hexapoda > Insecta (insects) > Dicondyla > Pterygota > Metapterygota > Neoptera > Eumetabola > Holometabola > Panorpida > Amphiesmenoptera > Lepidoptera (moths and butterflies) > Glossata > Coelolepida > Myoglossata > Neolepidoptera > Heteroneura > Ditrysia > Apoditrysia > Obtectomera > Pyraloidea > Family: Pyralidae > Subfamily: Phycitinae > Genus: Cactoblastis

Figure 1a. Cactoblastis cactorum female laying eggstick. [photo H. Roberson ©]	Figure 1b (top). Cactoblastis cactorum first instar larvae penetrating cladode surface, at base of spine (note base of eggstick at top right).Figure 1c (bottom): First instar larvae embedded in gum. Gum is a hazard for them when they first penetrate the cladode surface. [photos H. Roberson ©]
Figure 2a (top). Cactoblastis cactorum 3rd or 4th instar larvae in search of a new cactus cladode to feed in, having eaten out the previous one. Note the silken path they have constructed for their journey. [photo H. Roberson ©] Figure 2b (bottom). Cactoblastis cactorium cocoon with soil particles adhering to it (left), cocoon without soil (middle), and pupa that has been removed from the cocoon (right). [photo H. Roberson ©]	Figure 2c. Cactoblastis cactorum final instar larvae on Opuntia aurantiaca (Jointed cactus), either seeking out a new cladode to feed in or about to find a place in the leaf litter to pupate. The cladode in the centre is pale-coloured because most of the tissue inside has been eaten by the larvae. [photo H. Roberson ©]

Distribution and habitat

Native to northern Argentina, Uruguay, Paraguay and southern Brazil. In addition, it has been introduced to the following regions for the biological control of Opuntia cactus weeds (Zimmermann et al. 2004):

• Australia (introduced 1926, from Argentina)

• New Caledonia (introduced 1933, from Australia)

• South Africa (introduced 1933, from Australia)

• Mauritius (introduced 1950, from South Africa)

• Hawaii (introduced 1950, from Australia)

• Nevis in the Leeward Islands, Caribbean (introduced 1957, from South Africa)

• Antigua in the Leeward Islands, Caribbean (introduced 1960, from Nevis)

• Kenya (introduced 1966, from Antigua; establishment unconfirmed)

• Cayman Islands, Caribbean (introduced 1970, from Nevis and Antigua)

• St Helena, mid-South Antlantic (introduced 1971, from Nevis and Antigua)

• Ascension Island, mid-South Atlantic (introduced 1973, from St Helena)

• Pakistan (introduced 1994, from Australia; establishment unconfirmed)

Cactoblastis cactorum has also become established by accident in the USA (first recorded from Florida in 1989) either through migration from the Carribean islands and/or importation on ornamental cacti. This is a serious issue as it is attacking the indigenous cacti in the region (Zimmermann et al. 2000).

Life cycle

Adults

Cactoblasis cactorum adults emerge from pupae in the evening, usually within two hours after dusk. Calling and mating takes place mainly in the period of about an hour and a quarter before sunrise (Hight et al. 2003 - USA study). Females adopt a calling posture with the abdomen protruding upwards through the wings, at an angle of 45 degrees and for those that ultimately are successful in mating, the calling period only lasts for about five minutes before a male arrives and mates. Unsuccessful females call for a lot longer (about 40 minutes). Calling involves releasing a sex pheromone that attracts a male that is downwind from the female. The copulation period lasts for an average of 32 minutes (Hight et al. 2003). 

During the day, adults usually remain inactive and sit motionless in the vegetation in the vicinity of their host plants. Very rarely, if ever, do they shelter on the host plant itself (I searched for eggsticks daily on hundreds of plants in the field over summer and winter egg laying periods without once seeing adults on the host plants). Oviposition (egg laying), like emergence, usually occurs within two hours after dusk (Dodd 1940 p. 122 - Australia) although oviposition can continue until about midnight (Pettey 1948 p. 42).

Adults do not feed and therefore have to rely on energy stores accumulated in the larval stage. Hence, they have a short life, surviving for about five or six days although at lower temperatures (18ºC) they can survive for about 12 days (Legaspi and Legaspi 2007; Legaspi et al. 2009).

The dispersal capability of adults C. cactorum has become an important issue in developing a strategy for controlling its spread in the USA (Sarvary et al. 2008). Myers et al. (1981) found that among a group of plants, those that had C. cactorum moths placed on them before evening commenced, were more likely to have eggsticks laid on them than those which did not. Similarly, plants attacked in the previous generation were more likely to have eggsticks laid on them in the following generation (Dodd 1940 p. 121; Myers et al. 1981; Robertson 1985a, 1987), suggesting that moths emerging from pupae in the vicinity laid at least some of their eggs near the emergence site (although one can't discount the possibility that these plants were inherently more attractive to ovipositing females).

Although there is abundant evidence of C. cactorum dispersing very little, there are also a number of records of long distance dispersal by C. cactorum females. Dodd (1940 p. 121) remarks "many instances on record indicate that individuals have flown as far as fifteen miles to oviposit". In the Leeward Islands, C. cactorum appears to have dispersed naturally from some of the islands to others. For instance, it colonised the island of St Kitts from Nevis four miles away (Garcia Tuduri et al. 1971). Similarly, in the Hawaiian Island chain, C. cactorum has apparently dispersed naturally through these islands over a period of about seven years (C.J. Davis in Garcia Tuduri et al. 1971).

On the basis of the above evidence for short and long distance dispersal in C. cactorum, and as proposed by Osmond and Monro (1981), it is likely that the dispersal behaviour of C. cactorum females is age-related, with females laying their first eggstick(s) near the emergence site and then dispersing to lay the rest of their egg complement further afield. Osmond and Monro (1981) suggest that oviposition of the eggs near the emergence site by C. cactorum "probably acts as an insurance against flying into regions devoid of prickly-pear and thus dying without finding a host". Another possible reason for oviposition prior to dispersal is to reduce the energy cost of flight. In C. cactorum, a full egg complement constitutes about 46% of female mass (Robertson 1985a) and thus the tendency may be to reduce wing-loading by laying part of the egg complement before dispersal. This type of behaviour is commonly recorded in moths that emerge gravid from the pupa (Johnson 1969 pp. 181-182; Baker 1978). Puzzlingly, Sarvary et al. (2008) in their study of flight performance of C. cactorum considered unmated versus mated males and females but do not appear to have considered the effect of the reduction of egg load on flight performance.

Eggs and egg laying

The most distinctive feature of C. cactorum biology is the clumping of the eggs in eggsticks (Figure 1a). The number of eggs per eggstick typically ranges from 4 to 111 eggs, averaging about 56 eggs (Robertson 1985a). Dodd (1940) and Pettey (1948) recorded averages higher than this, ranging from 68 to 96 but I suspect there was a bias to the larger eggsticks. They each independently recorded a maximum number of 150 eggs per eggstick.

The laying of an eggstick (Figure 1a) involves first depositing an amber-coloured substance near the tip of the selected cactus spine which is used to glue the first egg to the spine. The positioning of eggs on top of one another is guided by setae that surround the ovipositor. Each egg is pressed against the last egg in the eggstick. According to Pettey (1948 p. 41), eggs are laid at an average rate of one every 16 seconds (fastest: 1 every 10 seconds; slowest: 1 every 24 seconds) so that the average 56 egg eggstick takes about 9-22 minutes to lay.

Under near optimal temperature conditions, fecundity (number of eggs laid by each female) varies from 54 to 281 eggs (Robertson 1985a). Each female usually lays 3 to 4 eggsticks (range 2-7). Fecundity is detrimentally affected by low temperatures (Robertson 1989), such that, in the temperature range from 8 - 20ºC  there is a linear relationship between the mean number of eggs laid per female and the mean minimum temperature at the time of egg laying. Under cold conditions females often fail to lay all the eggs inside them.

In the Eastern Cape, South Africa, there are typically two generations a year: At a study site on Thursford Farm, 21 km NW of  Grahamstown (Robertson 1985a), the egg laying period in the summer generation lasted from the beginning of October through to mid-December and by the end of January they had all hatched. Winter generation eggs were laid from the beginning of March to early May. Because of the cooling temperatures, late laid winter eggs took a long time to hatch so it was only by mid-August that they had all hatched. Egg laying in other parts of the Eastern Cape can start as early as September in the summer generation and February in the winter generation.

In the Thursford study, the egg development period lasted an average of 51 days in the summer generation (range: 41-64) and 75 days in the winter generation (range 58-114). Pettey (1948 p. 64) recorded much shorter average development periods at other localities in the Eastern Cape (e.g. 33 days in summer generation and 50 days in winter generation at Graaff-Reinet), and Dodd (1940 p. 117) recorded even shorter development periods in Australia (28-30 days in summer generation and 23-38 days in winter generation in Southern Queensland). Temperature has a direct effect on egg development: development halts altogether at about 12ºC and reaches its maximum rate at about 30ºC (Robertson 1985a, Appendix 4; Legaspi and Legaspi 2007). By 35ºC, the egg development rate has dropped off slightly. Under constant near optimal temperature conditions in the laboratory (30ºC), the minimum egg development time was about 20 days (Legaspi and Legaspi 2007) although Dodd (1940) recorded a minimum period of 18 days.

Larvae

The larvae that hatch from an eggstick penetrate the plant together (Figure 1b). The first larvae that emerge move down to the base of the spine where they surround themselves with a web of silk spun between the spine and the cladode surface, perhaps as a way of protecting them from predators such as ants. When the number of larvae reaches a certain undetermined threshold level, the larvae begin penetrating, usually adjacent to the spine on which the eggstick was laid. As few as nine larvae are able to successfully penetrate a cladode. To penetrate, the larvae chew away the cuticle at one spot on the cladode surface and deposit it in a ring around the penetration site (Figure 1b). If penetration is prevented by a tough cuticle or because of gum exudation (Figure 1c), the surviving larvae usually attempt to penetrate elsewhere on the plant.

After successful penetration, the larvae feed gregariously on the parenchymous tissue inside the cladode. Fibres are not eaten. Larvae can tunnel from cladode to cladode thus avoiding having to move outside and start penetrating the plant elsewhere. However, larvae sometimes do have to vacate the cladode when:

• the cladode becomes detached from the plant and contains no more suitable tissue for consumption;

• they destroy the entire plant in which they are feeding;

• they are unable or unwilling to bore into adjacent cladodes, particularly when the adjacent cladode is woody;

• the internal cladode temperature is too high. Under the latter circumstances, larvae, when outside the cladode, sometimes suspend themselves on a web of silk that is spun in the shade beneath a cladode.

When locating a new penetration site, larvae spin a silken path (Figure 2a), which presumably maintains colony cohesion as well as enabling the larvae to retrace their route if necessary. After about the third instar, larvae in some colonies split up into smaller groups, especially when the cladodes are small.

Larvae usually pass through six instars (each instar is separated by a moult of the old cuticle) although there can be as many as eight (Robertson 1985a). As larvae get bigger, they develop vivid orange and black banding, which is believed to act as a warning to predators that they are distasteful (such colouring is termed aposematic colouration).

Regarding the phenology of larvae (i.e. the times of the year when they were present), at the Thursford site in the Eastern Cape, the summer generation larvae were present from near the beginning of December through to mid-March although there were a few colonies that skipped a winter generation and passed right through winter, only pupating in August. Winter generation larvae were present from the beginning of April through to the beginning of November.

In the Thursford study, the larval development period  ranged from 46 to 53 days in summer (excluding the overwintering larvae) and 90 to 141 days in winter. The larval development period of the overwintering summer generation larvae was 200-210 days. Pettey (1948) at Uitenhage and Graaff-Reinet in the Eastern Cape, recorded larval development periods ranging from 47-82 days in summer and 99 to 164 days in winter.

Pupae

When larvae reach maturity they move to the ground where they spin a cocoon in a secluded place, typically among the leaf litter or in loose soil. Within the cocoon the larva moults its skin to reveal the pupal form beneath. Within the pupa, major redevelopment happens with genes for adult characters such as wings being turned on and genes for larval characters being turned off. 

In the study by Pettey (1948 pp. 61, 65), the pupal development period ranged from 20 to 32 days in the summer generation and 56 to 86 days in the winter generation.

Ecological interactions in southern Africa

Larval host plants

Unless otherwise indicated, information is from Zimmermann et al. (2000) and Zimmermann et al. (2004). The comments about impact are from a biological control perspective, in terms of how effective the larvae are in destroying the plant.

• Opuntia aurantiaca (Jointed cactus). Moderate to significant impact, mainly on large plants.
• Nopalea cochenillifera (= Opuntia cochenellifera)
• Opuntia engelmannii. Moderate impact
• Opuntia ficus-indica (Prickly pear). Ineffective in killing large woody plants but highly effective in destroying small plants.
• Opuntia humifusa. Significant impact.
• Opuntia monacantha. Has a moderate to significant impact.
• Opuntia pusilla. Cultivated in South Africa. Recorded as being attacked by Cactoblastis cactorum in Florida, USA.
• Opuntia robusta. Has a significant impact, especially damaging to small plants. Cactoblastis cactorum is a pest of the cultivated spineless varieties of this species.
• Opuntia salmiana. Moderate impact, mainly on larger plants.
• Opuntia spinulifera
• Opuntia stricta. This is the species that Cactoblastis cactorum decimated in Australia but in South Africa, it has been less effective.
• Opuntia tomentosa. Cultivated in southern Africa. Has an insignificant impact.
• Cylindropuntia imbricata.  It does not thrive on this plant

Predators

Figure 3a. Crematogaster liengmei ants eating eggs in eggstick of Cactoblastis cactorum. [photo H. Roberson ©]	Figure 3b (top). Unidentified mite extracting the contents of an egg in an eggstick of Cactoblastis cactorum. [photo H. Roberson ©] Figure 3c (bottom). Nysius bugs (Lygaeidae) eating eggs in eggstick of Cactoblastis cactorum. [photo H. Roberson ©]

Eggs are eaten by:

• ants (Hymenoptera: Formicidae). Egg predation by ants varied from 55-77% % at a site 22 km NW of Grahamstown (Robertson 1985a, 1985b, 1988; Robertson and Hoffmann 1989). Ant taxa attacking eggs included:
  • Crematogaster liengmei
  • Pheidole sp.
  • Tetramorium erectum
  • Monomorium albopilosum
  • Monomorium sp. (monomorium-group)
  • Camponotus niveosetosus

  Pettey (1948 p. 78) recorded the ant Technomyrmex albipes feeding on C. cactorum eggs, although this ant is almost certainly Technomyrmex pallipes as, until a recent revision of the genus, T. albipes was confused with T. pallipes in southern Africa.

  Ants are attracted on to Opuntia ficus-indica, Opuntia aurantiaca, and probably other opuntias, by extrafloral nectaries situated at the base of areoles (from where the spines eminate).

• Nysius (Hemiptera: Lygaeidae). Of minor importance in comparison to ants (Robertson 1985a; Robertson and Hoffmann 1989).
• mites (Arachnida: Acari). Of minor importance in comparison to ants (Robertson 1985a; Robertson and Hoffmann 1989)..

Larvae are eaten by:

• ants (Hymenoptera: Formicidae).
  • Anoplolepis steingroeveri (Robertson 1985a; Robertson and Hoffmann 1989).
  • Pheidole sp. (Robertson 1985a; Robertson and Hoffmann 1989).

Pupae are eaten by:

• ants (Hymenoptera: Formicidae)
  • Dorylus helvolus (a mainly subterranean driver ant). Robertson (xxx) recorded xx% mortality of pupae by these ants.

Parasitoids

Eggs are parasitised by:

• Trichogrammatoidea sp. (Hymenoptera: Trichogrammatidae). An incidental parasitoid of C. cactorum larvae (Robertson 1985a; Robertson and Hoffmann 1989; Pettey 1948 recorded Trichogrammatoidea lutea).

Larvae are parasitised by:

• Pseudoperichaeta sp. (Diptera: Tachinidae). An incidental parasitoid of C. cactorum larvae (Robertson 1985a; Robertson and Hoffmann 1989).

Pupae are parasitised by:

• Hymenoptera > Chalcididae
  • Brachymera sp. (Pettey 1948)
  • Invreia sp. (Robertson 1985a; Robertson and Hoffmann 1989)
  • Euchalcidia sp. (Robertson 1985a; Robertson and Hoffmann 1989)