CULTURE OF HOSTS FOR NATURAL
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The culture and colonization of natural enemies is fundamental to biological control work. The three principal reasons for culturing parasitoids, predators and pathogens are (1) for permanent field establishment, (2) periodic colonization0 and augmentation and (3) inundative releases.
For permanent establishment there are only relatively small numbers of a beneficial organism propagated for release at several dispersed sites. Successful organisms will persist in the new environment, spread and reduce the pest organism to a level, which is below the economic injury threshold in what has often been termed classical biological control. Once controlled, no further releases are required.
In periodic colonization and augmentation a beneficial organism is able perform well when the pest is seasonally present in damaging numbers, even though it is unable to persist in sizeable numbers the year round. Sailer (1976) gave an example of 3,000 Pediobius foveolatus Crawford, and eulophid parasitoid of the Mexican bean beetle, Epilachna varivestis Mulsant, being released in mid-spring. The parasitoid spread 595 km. by the end of October, with the near elimination of the host populations at locations in north central Florida. However, the parasitoid could not overwinter in the area and had to be recolonized annually. This procedure has been termed inoculative periodic colonization.
Similarly, the release of the tropical fish Tilapia zillii (Gervais) in irrigation canals in southeastern California for aquatic weed and mosquito habitat control is also usually a periodic requirement [Legner & Murray 1981 ] This fish species cannot always overwinter in canals when water temperatures drop below 10°C (Legner 1986b), or when competition with predator largemouth bass decimates its population.
In inundative releases, large liberations are made to effect short-term control of a pest. Inundative releases simulate pesticide treatments, and the agent simply reduces, rather than regulates, the pest population. Examples are the mass production and release of Lixophaga diatraeae (Townsend), a tachinid parasitoid of the sugarcane borer, Diatraea saccharalis (F.) (King et al. 1981). Mass releases are common for such organisms as the green lacewing, Chrysoperia carnea (Stephens), predaceous on soft-bodied insects; Spalangia and Muscidifurax pteromalid parasitoids of muscoid flies; and hydra against mosquitoes (Yu et al. 1974). However, the parasitoids most commonly released inundatively on a worldwide basis are egg parasitoids in the genus Trichogramma. Microbial pesticides, such as Bacillus thuringiensis Berliner, also come under this category. Such pesticides may also be used augmentively to control weeds. The fungus Colletotrichum gloeosporiodes f. spp. Aeschynomene (Penz) is more than 90% effective against northern jointvetch, Aeschynomene virginica (L.) B.S.P., a weed in American rice fields (van den Bosch et al. 1982).
Food employed in rearing the hosts of entomophagous organisms are, in decreasing order of difficulty, living plants, harvested plant parts, vegetables or fruit and prepared diets.
Living Plants.--The rearing of phytophagous insects on natural host plants requires purchases or farming, and are maintainable only at considerable cost of labor and space. Losses from plant diseases or pest arthropods are not unusual. The required holding time is important and related to host and entomophage life cycles. For example, the life cycle of the black scale, Saissetia oleae (Olivier), is about three months at 21°C on potted oleander. Since it must be nearly mature for acceptance by some parasitoids, which themselves may have a life cycle of three to 6 weeks, the oleander plants must be kept alive for several months after infestation with scale crawlers. Such maintenance may be complicated by diseases such as oleander knot or root rot, and by contaminating pests such as mealybugs.
Some plants used for insect production need only short durability, so that plant diseases are not usually a limiting factor. For example, certain parasitoids are raised on the pea aphid, Acyrthosiphon pisum (Harris) which in turn is raised on fava bean plants. These plants grow rapidly and are needed for only a short period after inoculation with host insects and parasitoids. Plant collapse in two weeks from aphid feeding and root rot does not interfere with parasitoid production.
The use of any practice to alleviate a problem should be thoroughly tested first for indirect effects. For example, the fungicide BenlateR is sometimes recommended to treat for certain plant fungal diseases. Because Benlate has a alight systemic action, aphids feeding on treated plants may consume sufficient quantities to kill their beneficial internal symbiotic microorganisms, which can cause their death. However, it is interesting to note that Benlate is recommended for suppressing certain protozoans that infect insectary-reared insects.
Forbes et al (1985) indicated that young, vigorously growing plants had to be used for raising aphids in order to achieve rapid growth and reproduction. They noted that rates of development, body size and fecundity can often be very different in reared versus wild aphids, and that these differences are partly due to variations between host plants in the field and in the laboratory. Furthermore, laboratory plants that are overcrowded have poor nutrition or are suffering from water stress, can stimulate alate production which may continue for several generations even after plant conditions have improved. Consequently, host plant quality affects parasitoid production by affecting the host insect.
Harvested Plant Parts.--Plant parts are sometimes used to feed insects, especially those that are voracious feeders on perennials. Potted perennials requiring lengthy developmental time might be destroyed in a few days by a pest, such as occurs with alfalfa consumption by the Egyptian alfalfa weevil, Hypera brunneipennis (Bohemon). The weevils consume so much food that it is necessary to feed them daily with cuttings taken from an alfalfa field and made into "bouquets" to retain foliage freshness.
Extended experimentation may be required to determine the type and condition of plant parts that are optimal for rearing pest insects. Willey (1985) found that dried dandelion green were preferred by the range grasshopper, Arphia conspersa Scudder, to dried Romaine or head lettuce or to assorted native grasses and alfalfa. Fresh dandelion leaves, however, were less favored. He noted that unprocessed dried leaves and buds of the dandelions could be stored frozen in polyethylene bags for later use.
Vegetables and Fruit.--Potatoes, citron melons and squash are commonly used to raise certain scale insects. Papacel & Smith (1985) reported that butternut pumpkins, Cucurbita moschata Duchesne, were the best substrate to grow oleander scale, Aspidiotus nerii Bouche. These in turn were used to mass produce the California red scale parasitoid, Aphytis lingnanensis Compere. A total quantity of 1.5 to 2 tons of pumpkins per week was required for annual production of 15-20 million parasitoids!
Rutabagas are used to grow cabbage maggots, Delia radicum (L.) which are hosts for the parasitic beetle Aleochara bilineata (Gyllenhal). Whistlecraft et al (1985b) provided at least one gram of rutabaga per cabbage maggot egg, in order to insure a uniform pupal size. Etzel (1985), rearing of the potato tuberworm, Phthorimaea operculella (Zeller), also found that one gram of substrate was sufficient for one individual. The tuberworms produced were processed as food for certain coccinellids and larvae of the common green lacewing.
Wight (1985) noted that insecticide residues could be troublesome with commercial produce. Because of such residues the outer leaves had to be stripped from lettuce purchased to feed the southern armyworm, Spodoptera eridania (Cramer).
The variety of produce is also important. The Russet potato is a mealy variety superior for tuberworm rearing, whereas White Rose with a smooth skin is best for raising California red scale, Aonidiella aurantii (Maskell).
Other significant problems associated with the use of vegetables and fruits are availability, durability and consistency. Citron melons are useful for rearing the brown soft scale, Coccus hesperidum L., but are not commercially available and must be specially grown. Commercial lots of other produce such as potatoes or rutabagas vary greatly in consistency and durability, sometimes rotting rapidly when removed from storage. Control of relative humidity during storage and use is important for reducing substrate deterioration. Decomposition not only ruins the food source, but may generate toxic gases. Such gases emitted by ripening grapefruit, e.g., are lethal to some parasitoid and host species in a confined space (Finney & Fisher 1964).
Chemical treatments might be useful to reduce deterioration of produce. In mass rearing the citrus mealybug, Planococcus citri (Risso), Krishnamoorthy & Singh (1987) treated ripe pumpkins, Cucurbita moschata with 1% Benlate and 5% formaldehyde solution.
Prepared Diets.--Singh (1985) reviewed 22 multiple-species rearing diets that together have been used to raise dozens of insect species. Prepared diets have been used to rear Lepidoptera and Diptera. Provided that they are nutritionally and physically adequate, diets provide the easiest and most consistent food source and eliminate most problems involved with host plants, plant parts, vegetables or fruit. However, adequate diets are more likely to be available for the least fastidious insects.
Omnivorous or polyphagous insects are obviously much easier to rear then are monophagous ones. Moore (1985) presented a systematic procedure and guidelines for choosing and modifying an artificial diet for a phytophagous arthropod. He discussed stimulants, repellents, nutrient requirements and microbial inhibitors, as well as physical and chemical adequacy, concentrations and proportions. Grisdale (1984) emphasized that consistently good artificial diets were produced with high quality fresh adequately mixed ingredients. However, both physical and chemical characteristics are important. Rearing success can often hinge on some critical step or technique in the physical presentation of a diet, as is true also with all aspects of insect production. Boller (1985) noted that cotton pads must only be coated with liquid larval diet on one side to provide a moisture gradient suitable for optimal development of certain fruit flies, and Bay & Legner (1963) had to feed blood mixture diets to chloropid eye gnats on dried prunes or filter paper.
Provision of food for adults of holometabolous insects is generally not as complicated as provision for larvae. Heather & Corcoran (1985) fed adults of the Queensland fruit fly, Dacus tryoni (Froggatt), sugar cubes, autolyzed brewers' yeast fraction and water. Hydrolysis of the yeast made the protein available for egg production. Tolman et al. (1985) fed adult onion maggots, Dellia antiqua (Meigen), with a dry diet consisting of 50% brewer's years, 33% yeast hydrolysate and 17% soybean flour. Bartlett & Wolf (1985) fed pink bollworm moths, Pectinophora gossypiella (Saunders), with 10% sugar water plus 0.2% methyl parasept (to retard microbial growth). Sometimes adult insect starvation simplifies production. Etzel (1985) held adult potato tuberworms without food or water and obtained adequate egg production.
Many of the considerations necessary in host culture apply as well to entomophage rearing, but separate treatment simplifies the often interacting factors.
The most prevalent and often most serious problem in the production of host arthropods is contamination by other arthropods, which may result in competition, disruption, parasitism, predation and or disease. Efforts to control undesired elements require costly labor, supplies, equipment and facilities. Some examples will indicate the range of contamination difficulties.
Phytophagous insects and mites frequently create problems in the production of hosts by competing for the substrate and interfering with a host-parasitoid system. Mealybugs, mites and aphids are frequent problems in rearing the black scale (Etzel & Legner 1999 ). Likewise, aphid infestations were troublesome on fava bean plants used to rear larvae of the red-banded leafroller, Argyrotaenia velutinana (Walker) (Glass & Roelofs 1985).
Mites have caused difficulties in laboratory cultures of Trogoderma beetles (Speirs 1985), Drosophila flies (Yoon 1985), the lesser peachtree borer, Synanthedon pictipes (Grote & Robinson, Reed & Tromley 1985b), the plum curculio, Conotrachelus nenuphar (Herbst) (Amis & Snot 1985), and the house fly, Musca domestica L. (Morgan 1985).
Papacek & Smith (1985) reported those ants, the citrus mealybug, and the scale-eating coccinellid Lindorus lophanthae (Blaisdell) were contaminants of insectary diaspid scale cultures used to rear an aphelinid parasitoid, Aphytis lingnanensis. Heather & Corcoran (1985) also had to cope with ants in a culture of the Queensland fruit fly, Dacus tryoni.
Wight (1985) found that phorid fly maggots were occasional problems in rearing the southern armyworm, and rapidly destroyed prepupae and pupae in open pupation pans. Gardiner (1985c) reported that the parasitoids, Cotesia (=Apanteles) glomerata (L.) and Pteromalus puparum L., are sometimes contaminants in laboratory cultures of the large white butterfly, Pieris brassicae L.
While it is common for parasitic insects to be impediments in insectary cultures, it is unusual to have other kinds of parasites. However, Gardiner (1985a) found that nematodes of the genus Mermis occasionally parasitize the desert locust, Schistocerca gregaria Forskal.
The degree of arthropod contamination depends on the generation time of the desired organism. Friese et al. (1987) found that when spider mites from a clean source colony were used to infest initially clean host plants, contamination by unwanted organisms was minimized since a spider mite generation is a short two weeks, and host plants can consequently be rapidly cycled. However, they also noted that greenhouse contamination by indigenous phytoseiid predators could be eliminated for up to three weeks without interfering with spider mites by treatment with an insecticide (carbaryl at 50% recommended dosage).
Microorganisms can cause severe contamination problems by being plant pathogens, saprophytic contaminants, saprophytic facultative insect pathogens, saprophytic true insect pathogens or obligatory true insect pathogens. Pathogens can readily destroy plants used to raise host insects. Saprophytic microorganisms compete with host insects for the same food, and destroy it. Fungi, bacteria and yeasts decompose plant parts, fruits and vegetables used as host food. Sikorowski (1984) noted that contaminating microbes growing on insect diets can biochemically change the nutritive value thereof, and may also produce harmful toxins. Shapiro (1984) concluded that fungi of the genus Aspergillus are the most common contaminants in insect cultures. These and other saprophytic fungi and bacteria are ubiquitous in nature and promptly appear in unsanitary conditions.
Saprophytic facultative pathogens include the bacterium Serratia marcescens (Bizzio), which can invade insects only through open wounds, which then causes acute disease. Saprophytic true insect pathogens, which are capable of direct invasion, are not common problems in insectaries. However, the bacterium Bacillus thuringiensis is occasionally troublesome. Stewart (1984) reported that it had interfered with mass production of the pink bollworm.
Obligatory true insect pathogens among the fungi, protozoa and viruses cause the most pervasive and difficult problems in host insect production. the fungus Nomuraea rileyi (Farlow) has been reported in a colony of the velvetbean caterpillar, Anticarsia gemmatalis Hübner; and Entomophthora spp. have been found attacking cultures of houseflies (Morgan 1985) and onion maggot adults (Tolman et al. 1985).
According to Goodwin (1984), protozoans (including Microsporidia) are the most important pathogens in insectaries, and many are not as host specific as originally thought. They can infect several closely related species and some may even infect insects in different orders or families. Protozoans are particularly troublesome because they typically cause chronic, debilitating diseases that are more difficult to detect and eliminate than are acute diseases. Protozoans of the microsporidian genus Nosema are very prevalent. They cause problems in mass production of the spruce budworm, Choristoneura fumiferana (Clemens) (Grisdale 1984), the western spruce budworm, Choristoneura occidentalis Freeman (Robertson 1985a), and the pink bollworm (Stewart 1984). Guthrie et al. (1985) in fact noted that it is very difficult to start a clean colony of the European corn borer, Ostrinia nubilalis (Hübner), because most field-collected larvae contain Nosema pyrausta (Paillot).
Mattesia is another bothersome genus. McLaughlin (1966) reported on efforts to eliminate Mattesia grandis McLaughlin from a colony of the boll weevil, Anthonomus grandis grandis Bohemon. In the entomophage insectary at the University of California at Albany, Mattesia dispora Naville causes a chronic disease in the Mediterranean flour moth, Anagasta kuehniella (Zeller). However, in the navel orangeworm, Amyelois transitella (Walker), also being reared at the Unversity's Lindcove Field Station, it causes an acute disease that destroys the colony. The navel orangeworm culture was used to rear the encyrtid parasitoid Pentalitomastix plethoricus Caltagirone and the bethylid Goniozus legneri Gordh for field release. Mattesia was the only major problem interfering with parasitoid rearing. Necessary measures to control the disease greatly restricted the level and ease of production. At Lindcove, California it was necessary to raise the rearing room temperature to 90°F to inactive the Mattesia.(Legner & Warkentin, unpublished data).
Three major groups of insect viruses can contaminate host insect cultures, making rearing very difficult. The diseases caused are typically acute, however, and consequently rather easily detected. Nuclear polyhedrosis viruses are the most prevalent. For example, such viruses have been reported in cultures of the Douglas-fir tussock moth, Orgyia pseudotsugata (McDunnough) (Robertson 1985b), the forest tent caterpillar, Malacosoma disstria Hübner (Grisdale 1985b), the Egyptian cotton leafworm, Spodoptera littoralis (Boisduval) (Navon 1985), the beet armyworm, Spodoptera exigua (Hübner) (Patana 1985b), and the cabbage looper, Trichoplusia ni (Hübner) (Guy et al. 1985). A cytoplasmic polyhedrosis virus caused severe effects in mass production of the pink bollworm (Stewart 1984), and Reed & Tromley (1985a) reported that a granulosis virus could interfere with rearing the codling moth, Laspeyresia pomonella (L.).
Although one microorganism may severely disrupt a rearing program, a group of them is intolerable. Stewart (1984) reported that the greatest difficulties in mass producing the pink bollworm were caused by the fungus Aspergillus niger van Tieghem, the protozoan Nosema sp., a cytoplasmic polyhedrosis virus, and the bacterium Bacillus thuringiensis. Another example of a complex of troublesome pathogens was reported by Henry (1985) who noted that colonies of grasshoppers, Melanoplus spp., can be contaminated with viruses, protozoa, bacteria and fungi.
Contamination problems and diseases must be prevented and eliminated. The practical mechanics of achieving these goals can be very difficult and costly. Consideration of source provides clues to control. Saprophytic contaminants cause disease indirectly by depriving insects of proper nutrition or environment. Such microorganisms are ubiquitous, and can increase rapidly in insectaries with poor sanitation or design. The source of obligate pathogens in an insectary has to be in or on insects introduced to initiate lab colonies, or on natural food used in rearing.
Shapiro (1984) recommended that in starting or adding to a colony, pathogen introduction could be decreased when insects were collected from less dense population areas; and Grisdale (1984, 1985b) suggested field collecting insects only from new infestation areas where disease is still at a low level. This advice is particularly useful for insects with widespread, high-incidence pathogens, such as the forest tent caterpillar attacked by a nuclear polyhedrosis virus (Grisdale 1985b), and the spruce budworm, widely infected by the microsporidian Nosema fumiferanae (Thomson) (Grisdale 1984).
Field-collected larval stages are generally the most seriously infected by pathogens. If possible it is best to collect another stage. Singh & Ashby (1985) noted that "... the egg is usually the best stage with which to start a colony since it is least likely to carry disease microorganisms." However, some viruses and protozoans are known to be transmitted on the surface of the egg, and some viruses can probably be transferred within the egg as well, as can certain protozoans. For example, when establishing a new colony of the forest tent caterpillar, Grisdale (1985b) surveyed field sites for the presence of the protozoan Nosema disstriae by microscopic examination of fully formed larvae removed from field collected eggs.
If eggs are difficult to field collect they may be obtained from field-collected adults. Leppla (1985) prevented fungus infection by Nomuraea rileyi in a colony of the velvetbean caterpillar by visual examination of field-collected adults and removal of dead ones, followed by surface sterilization of eggs laid in the laboratory.
Pathogens can also be accidentally introduced into an insectary colony on natural food. Patana (1985b) reported that colonies of the beet armyworm had frequently been lost to virus, attributed primarily to the use of natural food, cotton leaves in summer and Swiss chard in winter. After introducing prepared diet in 1965, Patana (1985b) reared the insect continuously without virus disease. Similarly Gardiner (1985a) used Brassica instead of field grass for rearing the desert locust, Schistocerca gregaria, because of the threat of introducing diseases and nematode parasites of local grasshoppers.
Contaminating microorganisms can likewise enter insectaries on ingredients for prepared diets. Shapiro (1984) found that more than 95% of the total bacteria recovered from various ingredients of gypsy moth diet occurred on the raw wheat germ. The pathogenic protozoan Mattesia dispora and the bacterium Bacillus thuringiensis may contaminate stored grain products used for insect diets, inasmuch as these microbes were originally isolated from stored grain insects.
Contaminating microorganisms may or may not be brought under control relatively easily, depending on the characteristics of the rearing programs, procedures and facilities. Fisher (1984) listed sources of contamination in an insectary and possible measures to control it. Grisdale (1984) found that rearing several species of insects in the same facility could result in serious microbial contamination, particularly if some species were reared on foliage and some on artificial diet. Even though he reared the eastern spruce budworm on artificial diet, balsam fir foliage was still used as an oviposition site, and was a principal source of fungal contamination. Stewart (1984) reported that cytoplasmic polyhedrosis virus caused severe continuing disease problems in a pink bollworm colony until he discovered that moth scales carried virus polyhedra on air currents from oviposition areas to larval rearing areas. Major changes where then instituted in procedures and facilities which virtually eliminated disease and highly increased insect production.
Microorganisms can be greatly reduced or eliminated by strict rigorous sanitation, proper rearing procedures and suitably designed insectaries. Controlling them requires recognition and monitoring. Specialists in large mass production facilities usually do this. However, all personnel should have some familiarity with microorganisms and sanitation procedures. Poinar & Thomas (1978) presented a useful manual on the diagnosis of insect pathogens, and Goodwin (1984) reviewed the recognition and diagnosis of diseases in insectaries and the effects of disease agents on insect biology. Shapiro (1984) discussed microorganismal contaminants and pathogens in insect rearing; Sikorowaski & Goodwin (1985) contaminant control and disease recognition in laboratory colonies; and Sikorowski (1984) occurrence, monitoring, prevention and control of microbial contamination in insectaries.
The first line of defense against contagious diseases in an insectary is exclusion by procedural, physical and chemical techniques, but initially and continuously. After laboratory introduction, insects are quarantined and reared individually for a few generations while they are monitored for disease presence (Goodwin 1984, Shapiro 1984). Diseased insects are destroyed by steam sterilization. Although initial individual rearing is highly laborious, it may guarantee a pathogen-free culture. When Grisdale (1984) added field collected eastern spruce budworms to an existing colony, the newly collected stock was reared in lab isolation for two generations, with only progeny from protozoan-free adults cultured. Forbes et al. (1985) likewise recommended that only progeny from field-collected aphids should be used to initiate laboratory colonies in order to reduce fungal disease.
In addition to quarantine for the elimination of pathogens, chemical surface disinfection of insect stages is often routinely used. This is particularly true with lepidopterous eggs, not only because obligate viruses and protozoans are frequently transmitted on these eggs, but because bacterial and fungal contaminants create problems on prepared diets typically used to rear lepidopterans.
Vail et al. (1968) and Sikorowski & Goodwin (1985) have recommended procedures for surface disinfecting insect eggs. Various techniques using sodium hypochlorite are most popular. Formalin is also used because it is a good viricide. Sodium hypochlorite concentrations and exposure times have to be adjusted to a particular insect species, depending on the susceptibility of its eggs to the action of the chemical. Guy et al. (1985) used a very weak solution of 0.02% for only five minutes to sterilize egg surfaces of the cabbage looper. A common solution contains 0.1%, which Reed & Tromley (1985b) used for five minutes to disinfect eggs of the lesser peachtree borer, whereas Robertson (1985b) employed it for 15 minutes twice with strong mechanical stirring to treat eggs of the Douglas-fir tussock moth, and Greenberg & George (1985) used it for 15 minutes with swirling to disinfect eggs of calliphorid flies.
Willey (1985) cautioned that although a solution of 0.25% sodium hypochlorite was used for 10 minutes to surface sterilize eggs of the range grasshopper, Arphia conspersa, it was used infrequently because treated eggs had a much lower hatching success than those incubated in situ. Similarly, L. Etzel (Etzel & Legner 1999 ) found that treatment of Mediterranean flour moth eggs for five minutes with 0.15% reduced hatchability by at least 50%, but was necessary to control disease caused by Mattesia dispora. Hatchability is best when eggs are not treated until nearly completely embryonated. Even then the eggs are extensively dechorionated so that they must be held on filter paper on a moist sponge in a petri dish to prevent desiccation.
In culturing Egyptian alfalfa weevil parasitoids, Etzel (pers. commun.) found that weevil eggs collected from alfalfa stems had to be treated with 1% sodium hypochlorite for one minute to retard saprophytic fungal growth if storage at 4°C followed. Finally, Grisdale (1985b) used full strength sodium hypochlorite (8%) for 1.5 minutes to disinfect egg masses of the forest tent caterpillar.
Although not as common, surface sterilization of eggs with formalin is also performed. Bartlett & Wolf (1985) used 9.5% formaldehyde for 30 minutes to disinfect pink bollworm eggs. Singh et al. (1985) noted that eggs of the light brown apple moth, Austrotortrix postvittana (Walker), have to be 4-5 days old before they can withstand surface disinfection with 5% formalin solution for 20 minutes, which prevents viral disease. Ashby et al. (1985) also cautioned that codling moth eggs should not be surface sterilized with 5% formalin until they are 48-6 days old. However, a satisfactory treatment for codling moth eggs is 0.15% sodium hypochlorite for 10 minutes.
Other chemicals are occasionally used to treat insect eggs. Speirs (1985) used 0.1% mercurous chloride in 70% ethanol plus 0.1 ml Triton X-100R /liter for three minutes to disinfect eggs of Trogoderma spp. Moore & Whisnant (1985) utilized 18% cupric sulfate (a fungicide) and a 0.3% solution of Mikro-QuatR (alkyl dimethylbenzylammonium chloride) to surface sterilize boll weevil eggs.
Insect larvae can also be chemically treated to prevent disease. The tachinid Lixophaga diatraeae was treated with 0.7% formalin for five minutes to control Serratia marcescens (King & Hartley 1985c); the European corn borer with a 0.01% phenylmercuric nitrate solution prior to diapause to control Nosema pyrausta (Guthrie et al. 1985); and the black cutworm, Agrotis ipsilon (Hufnagel), with a 1% solution of phenylmercuric nitrate before being placed in diet cups prior to parasitoid emergence (Cossentine & Lewis 1986).
It is not unusual for pupae to be surface disinfected to control contaminating microorganisms, where again sodium hypochlorite is the chemical of choice. Patana (1985b) treated pupae of the beet armyworm with a 0.03% solution for five minutes, and Guy et al. (1985) used 0.1% solution for 10 minutes for cabbage looper pupae.
Sodium hypochlorite is used to dissolve cocoon silk, as well as to disinfect the harvested larvae or pupae. Etzel (1985) used 1.3% sodium hypochlorite solution to dissolve cocoon silk and harvest larvae or pupae of the potato tuberworm from the layer of sand in which pupation occurred. Likewise, Grisdale (1985b) separated pupae of the forest tent caterpillar from their silken cocoons by exposure to a solution of 1:1 sodium hypochlorite (8%) in water, and Bartlett & Wolf (1985) utilized 3% sodium hypochlorite solution for 30 minutes to dissolve cocoon silk of the pink bollworm.
Other solutions used to surface disinfect pupae include 5% phenol for calliphorids (Greenberg & George 1985), and 0.2% mercuric chloride for the wood boring scolytid Xyleborus ferrugineus (F.) (Norris & Chu 1985).
In addition to the use of chemicals to sterilize insect eggs, larvae and pupae, ordinary disinfectants should be routinely used in normal sanitation. Sikorowski (1984) reviewed different antimicrobials available for cleaning and disinfection and noted in particular that wet-mopping floors after flooding with disinfectants is preferable to sweeping and dry-mopping. Stesart (1984) reported that disinfection and cleaning of equipment and facilities with bleach, quaternary ammonium and phenolic compounds and stabilized chlorine dioxide solutions were major factors in controlling microbial pathogens in mass production of the pink bollworm.
As with surface disinfection of insects, sodium hypochlorite is most commonly used for general sanitation. Concentrations range from ca. 0.026^ to 5.25%, but 1% is more common. The lower concentrations are often used to disinfect rearing containers. Baumhover (1985) employed a 0.026% solution to soak clean rearing containers for a minimum of four hours in culture of the tobacco hornworm, Manduca sexta (L.), and he mopped floors weekly with the same solution. Palmer (1985) used 0.05% sodium hypochlorite to soak water dishes and cheesecloths for 4-8 hours in rearing the chalcidid Brachymeria intermedia (Nees). Moore & Whisnant (1985) prevented microsporidian infection of the boll weevil by washing adult cages and emergence boxes with soap and 0.5% sodium hypochlorite. A 1% concentration is generally used for washing equipment and wiping down tables, etc. in the production of houseflies (Morgan 1985), and Melanoplus spp. grasshoppers (Henry 1985).
Some workers have used solutions of formaldehyde to spray walls, ceilings, cabinets and counters, or to fumigate rearing rooms or containers. These practices are to be discouraged since formaldehyde is a carcinogen.
Navon (1985) reported that treatment of rearing boxes overnight in 0.4% potassium hydroxide helped to prevent viral disease in rearing Spodoptera littoralis.
Insectary sanitation procedures have also included the use of commercial germicides, such as RoccalR (Reed & Tromley 1985a, Guthrie et al. 1985), Ves-pheneR (Riddiford 1985), and ZephiranR (O'Dell et al. 1985, Morgan 1985). Morgan (1985) employed 0.13% Zephiran as a surface disinfectant to kill the pathogenic fungus Entomophthora sp.
Physical means can likewise be employed in insectaries for sterilization or disinfection. Sterilization is most common for destroying unwanted laboratory organisms. However, steam deteriorates wooden cages. Legner (unpubl.) found that steam sterilization of pink bollworm cages was no longer required once smoking tobacco was banned from rearing rooms, after which host production increased several fold.
Heat has been used occasionally to directly treat insects for disease control. Etzel (1985) noted that treatment of potato tuberworm eggs in hot water at 48.3°C for 20 minutes, as described by Allen & Brunson (1947), is useful for controlling the protozoan Nosema. However, Etzel et al. (1981) reported that the same treatment performed on eggs of the weed-feeding chrysomelid Galeruca rufa Germar destroyed them within 10 minutes. Shapiro (1984) reviewed other examples of heat treatment that are helpful in disease control.
The physical design, structure and equipment of an insectary, especially as they relate to environmental control, are critical for the efficient production of healthy insects. In rearing gypsymoth larvae for parasitoid production, O'Dell et al. (1984) noted that in spite of egg disinfection and routine cleaning of work surfaces and equipment, there were still periodic severe problems with bacterial and fungal diseases, attributed to inadequate environmental control, other facility peculiarities and the stress of parasitization. Sikorowski & Goodwin (1985) remarked that proper facility design and traffic control aid significantly in controlling microbes. Dividing rearing facilities into a clean area for critical work and a conventional area for less critical work is advised. Of particular benefit is the use of high efficiency particulate air (HEPA) filters for clean rooms and laminar air flow work stations. Sikorowski (1984) believed one of the best methods for controlling microorganisms when working with insect diet preparation or infestation, or when performing other procedures where contamination was a threat, was to do the work in such a work station. He also recommended HEPA-type exhaust filters for vacuum cleaners.
Stewart (1984) virtually eliminated severe disease in mass producing the pink bollworm by making major procedural and facility changes, including centralization of egg disinfection and larval transfer, positive air pressurization of rooms for diet preparation and egg disinfection, and installation of HEPA filters for cleaning air in critical areas.
Careful control of temperature, humidity, moisture and light are also important for disease control. Finney et al. (1947) reported that bacterial diseases caused by facultative pathogens in potato tuberworm colonies are suppressed by preventing high humidities and by rearing temperatures of <30.6°C. Thus, environmental stress is a contributing factor in disease. Greany et al. (1977) documented another case of temperature caused stress, and subsequent insect disease. Rearing the Caribbean fruit fly, Anastrepha suspensa (Loew) and a braconid parasitoid Biosteres longicaudatus (Ashmead) above 30°C created stress that permitted the bacteria Serratia marcescens and Pseudomonas aeruginosa (Schroeter) to become pathogenic, causing high mortality of both insects. Lowering the rearing temperature controlled the diseases.
Gardiner (1985c) found that grossly overcrowding larvae of the large white butterfly, Pieris brassicae, accompanied by excessive humidity, contributed to occasional outbreaks of bacterial disease. He also noted that low humidities and avoidance of overcrowding are critical to preventing bacterial diseases in rearing the desert locust, Schistocerca gregaria (Gardiner 1985a). Henry (1985) likewise recommended controlling various grasshopper diseases by limiting relative humidity to 30-35%.
Moisture and stagnant air particularly favor fungal pathogen development. Ankersmit (1985) found that holding rearing containers of the summer fruit tortrix, Adoxophyes orana Fischer von Röslerstamm, at a constant temperature reduced chances for moisture condensation, correspondingly reducing microbial contamination. Patana (1985b) discovered in rearing the beet armyworm that mold contamination could be controlled on artificial diet by using rearing containers allowing slight diet drying. Likewise Roberson & Wright (1984) utilized porous polyethylene to seal polystyrene trays in mass producing the boll weevil, thus allowing air and moisture exchange in the rearing cavities. This, plus placing a sterile sand/corncob mixture on the diet to absorb moisture and force hatching larvae to feed, greatly reduced microbial contaminants. Proper ventilation was also recommended by Grisdale (1984) for control of fungal contamination. Even under conditions of very high humidity, which may be necessary for rearing some stages of some insects, fungal growth can be greatly reduced or controlled by providing constant clean air movement.
Other environmental factors can impact microbial contamination. Insect activity by itself can be significant. Whistlecraft et al. (1985a) remarked that a seedcorn maggot population, Hylemya platura (Meigen), large enough to actively feed on the available artificial diet would prevent mold development. Even light can be a factor. Heather & Corcoran (1985) found that a contaminant yeast would grow on a carrot based larval diet for the Queensland fruit fly unless light was excluded. How insect stages are handled is likewise important. Henry (1985) recommended leaving grasshopper eggs in situ in the oviposition substrate to protect hatching nymphs from lethal bacterial and fungal diseases.
The above procedural, physical and chemical means of controlling microbial contamination and insect diseases provide the best defenses. However, contamination and disease can still occur. Therefore, antimicrobial chemicals are sometimes used with insect food as a further control. Shapiro (1984) provided an excellent review of chemical antimicrobials as ingredients for prepared diets. Sikorowski (1984) and Goodwin (1984) reviewed different antimicrobial chemicals for diets, recommending against using antibiotics unless absolutely necessary because of the danger of selecting for resistant microbes.
Once diseases caused by obligate pathogens appear in a culture, it is usually best to destroy the culture, completely clean and sanitize the insectary and star a new colony. However, if the culture is too valuable to discard, then isolation, quarantine and rigorous sanitary procedures can be used to try to recover healthy specimens.
Contamination in production of beneficial organisms does not occur only from parasitoids, predators, pathogens and interspecific competitors. The desired organism can also contaminate if it appears spatially or temporally where unwanted. Plants being grown for host insect production might be destroyed by contamination by that species before being suitable for purposeful infestation. Similarly a source colony of host insects could be decimated if contaminated by the entomophage. In mass producing pteromalids for filth fly control, one species may contaminate the culture of another. In such cases continuous manual elimination of contaminants is required if spatial separation of cultures in impractical (Legner unpub.)
Intraspecific competition or cannibalism can also be troublesome, especially with host insect production. In detailing the history of Heliothis spp. rearing, Raulston & King (1984) noted that a major problem was cannibalism. Consequently the reared larvae must be separated. One method was to use compartmented disposable plastic trays covered with Mylar film, as pioneered by Ignoffo & Boening (1970), and later automated (Sparks & Harrell 1976). Another type of compartmenting was described by Hartley et al. (1982). However, Patana (1985a) developed a different technique for separating larvae of these species. He placed 75 Heliothis larvae in a plastic box with a layer of diet covered by a layer of dried diet flakes. The dried flakes separated the larvae and greatly reduced cannibalism. Such rearing units will yield 65% pupae for corn earworm or 85% for the tobacco budworm. Hippelates eye gnat larvae undergo severe competition and stunting if crowded in the rearing medium (Legner 1966 ).
Obviously in mass production it is highly desirable to develop a system for rearing cannibalistic insects together. This is in spite of the fact that a major advantage of individual rearing is facilitation of disease control. Brinton et al (1969) reared another cannibalistic species "gregariously by using a sawdust based diet for codling moth larvae. Not only did the sawdust tend to separate the larvae, but the diet was more economical than if agar based.
It is sometimes possible to avert cannibalism by seeking a naturally noncannibalistic race. This was accomplished with the planarian mosquito predator Dugesia dorotocephala (Woodworth), which is normally cannibalistic (Legner & Tsai 1978).
Not all cannibalistic insects need to be kept physically separated. Grisdale (1985a) found that although the hemlock looper, Lambdina fiscellaria fiscellaria (Guenée) is cannibalistic, providing an acceptable artificial diet allowed gregarious development. In fact 10-20 larvae could be reared on diet in small 22-ml cups until the third instar, at which time four larvae were transferred to each new cup to complete development.
Some insects are gregarious in nature, making rearing relatively easy. Grisdale (1985b) found that the first three instars of the forest tent caterpillar seemed to develop better when crowded on artificial diet. Nasonia vitripennis Walker and Muscidifurax raptorellus Kogan & Legner, pteromalids for filth fly control, are also mass produced gregariously. In fact, the latter species exists in nature as several races demonstrating both solitary and gregarious development (Legner 1987c, 1988c), suggesting that similar racial types might exist for other species. [ Please refer also to Related Research ]
The genetic composition desired in a laboratory culture depends on its purpose. Either genetic uniformity or variability may be preferred. A high homozygosity or genetic uniformity is desirable in a culture used for insecticide testing to provide a relatively stable standard for treatment comparisons (Wheeler 1984). The same is true for insect colonies used to assay pathogens for microbial control. However, a high genetic variability is desired in entomophages produced for biological control as discussed in a previous section.
With respect to host provision for entomophage rearing, primary production goals are ease, rapidity and quality maintenance. However, host strain effects on parasitoid production are also important. For example, ODell et al. (1984) reported significantly different puparial weights of two groups of the tachinid Blepharipa pratensis Meigen when the parasitoid was reared on two different gypsy moth strains. The host strain differences were related to their field densities and geographic sources.
Geographic strain differences can also be important to ease of rearing. Diapause in the life cycle is a particularly aggravating production problem, and so it is advantageous to obtain nondiapausing field strains. With the plum curculio, which has a northern strain with diapause and a southern one without, Amis & Snow (1985) chose the southern one for culture. Bartlett & Wolf (1985) noted that the pink bollworm probably has a facultative diapause since no diapause is known for the insect in latitudes between 10°N and 10°S, such as in southern India. In California pink bollworm diapausing strains are interspersed with nondiapausing in different seasons [Legner 1979c ], whereas diapausing naval orangeworm occurs at such a low frequency as to go largely undetected (Legner 1983). Henry (1985) reported that the migratory grasshopper, Melanoplus sanguinipes (F.), widely distributed in North America, has diapausing strains. Throughout most of the range it is univoltine, with an obligatory egg diapause. In southern areas, however, there may be two or three generations a year, and the egg stage may simply enter an extended quiescent stage during the winter. Grasshoppers collected from a southern area would thus be best for initiating a laboratory culture.
Even if a nondiapausing field strain does not exist, it may be possible to develop such a strain by selection over a number of generations. For example, Jackson (1985) noted that although the wild strain of the western corn rootworm, Diabrotica virgifera LeConte, has a diapause in the egg stage, a laboratory nondiapause strain also exists.
Development of a nondiapausing insect strain illustrates planned genetic adaptation of a species to the laboratory. Whether planned or unplanned, some degree of such adaptation typically occurs before a species becomes easily reared. The problem is to balance the need for laboratory adaptation against the possible need to retain genetic diversity or heterozygosity, and certainly to prevent genetic deterioration of the stock. Gardiner (1985c) noted that the large white butterfly, Pieris brassicae, is relatively easy to rear, but only after it has become adapted to the lab. In this case the basic problem of adaptation is that adults have to be fed by hand for several generations until they will feed at artificial flowers. Heather & Corcoran (1985) used ripe, fresh and whole fruit for rearing the Queensland fruit fly for the first couple of generations in the laboratory until the population could be increased, and adaptation to a prepared diet could be initiated. In starting a colony of the Mediterranean fruit fly, Ceratitis capitata (Wiedemann), Boller (1985) recommended rearing field-collected specimens at low densities during the early colony establishment period since high adult fly mortality occurs due to irritation and unnatural densities in lab cages. This can result in unwanted selection of laboratory ecotypes.
Once a species is adapted to laboratory culture, maintenance of genetic vigor depends on the culture's genetic plasticity , the number of deleterious genes in the population and the number of parent individuals and their degree of mixing for each generation. Some insect cultures have been maintained satisfactorily for years, whereas some have to be replenished from field stock annually. Wight (1985) reported that the southern armyworm had been reared continuously since 1938, giving remarkably consistent responses in pesticide testing, the consequence of genetic homogeneity developed during long-term culturing. Guthrie et al. (1985) noted that the European corn borer had been reared on artificial diet for 200 generations over 19 years with no genetic deterioration in terms of fecundity, fertility and pupal weight. However, after about 14 generations there was a loss of adaptiveness to corn plants. Similarly, Baumhover (1985) continuously reared a laboratory colony of the tobacco hornworm for 170 generations (18 years) with no apparent genetic deterioration. Field tests of sterilized laboratory reared male moths showed nearly complete competitiveness with native males.
Most laboratory colonies cannot be kept indefinitely without replacement or replenishment with newly collected stock. Reed & Tromley (1985) recommended renewing a laboratory colony of the codling moth after 20 to 30 generations on artificial diet. Leppla (1985) maintained genetic variability of a laboratory colony of the velvetbean caterpillar by annually mixing the eggs from about 50 wild type and 50 lab females.
Many species deteriorate genetically in culture. Belloncik et al. (1985) found that the white cutworm, Euxoa scandens (Riley) and the darksided cutworm, Euxoa messoria (Harris) genetically deteriorated after only four laboratory generations (ca. one year): there was a loss of vigor and fertility, and the appearance of adult malformations. Jones (1985) discovered that annual recolonization with wild stock was necessary to maintain vigorous laboratory colonies of the southern green stink bug, Nezara viridula (L.). Inbreeding depression was minimized by starting five laboratory families from each of five field collected females and then mating progeny to those from different families in a planned pattern.
Various workers have recommended planned mixing in a colony to reduce inbreeding depression. O'Dell et al. (1985) advised the mating of males from one gypsy moth egg mass with females from another egg mass. In maintaining a culture of the beet armyworm for over 18 years, Patana (1985b) believed that continual mixing of larvae from different groups of parents provided a limited random mixing of genetic material that prevented the effects of absolute inbreeding. Young et al. (1976) studied genetic changes in a corn earworm colony and developed a crossing procedure to reduce inbreeding, thereby improving mating, fecundity and fertility. Hoffman et al. (1984) described a system using genetic selection to improve the characteristics of an already existing colony of the cabbage looper. The colony was divided into 26 subcolonies, set up on consecutive days, with the eggs for each sub colony obtained from the parent colony on different days to try to maintain genetic diversity. Performance was monitored by rating fecundity, hatch percentages, number of larvae reaching the fourth instar, pupation and emergence with set rearing regimes at certain fixed time periods. Subcolonies not reaching expected performance levels in two consecutive generations for hatch, larval development, pupation and emergence were discarded. Hoffman et al. (1984) were able to increase mean colony fecundity by 30% within three generations with subcolony selection. The fractional colonization scheme also enables better control of insect diseases since contaminated subcolonies can be immediately discarded.
The genetic vigor of laboratory colonies can be determined by standard quality control tests such as size, fecundity, fertility and longevity (Legner 1988b ). Sophisticated technical tests have also been used (Brown 1984, Bush et al. 1978, Goodenough et al. 1978). Robertson (1985a) recommended using starch gel electrophoresis to monitor genetic quality of laboratory colonies of spruce budworms in the genus Choristoneura. On the basis of her testing, she suggested that wild stock collected in the same area as the founder group should be introduced into the colony at two to three year intervals to prevent excessive homozygosity.
The actual laboratory production of insects, involving factors already discussed, is obviously dependent upon environmental conditions. Combinations of light, temperature and humidity and their sequences, are particularly critical in managing development of insects that undergo facultative or obligatory diapause. Obligatory diapauses especially cause severe production problems, but both facultative and obligatory diapauses can be advantageously used to enable long term insect storage. For example, the darksided cutworm overwinters in the egg stage, which can be kept in storage at least one year at 4°C (Belloncik et al. 1985).
Generally, light and temperature are the most important physical factors in initiating and terminating diapause. To illustrate, the environmental regime for diapause prevention in colonies of the cabbage moth, Mamestra brassicae L., is 20°C, 60% RH, and a photophase of 18 hrs, for rearing the larvae, after which the pupae are nondiapausing (Gardiner 1985b). Diapause can be initiated by rearing the larvae with a 9-hr photophase. Gardiner (1985b) also noted that prevention of diapause in lab colonies of the cabbage moth had been difficult for many early workers, and that larval food quality and insect strain had been two factors involved.
Moisture can also be a factor regulating diapause. According to Henry (1985) a subspecies of Melanoplus differentialis (Thomas) (s.s. nigricans), occurs in the Central Valley of California and apparently undergoes a winter obligatory diapause, which may be more conditioned by moisture than by temperature. Density is an occasional diapause factor as well, as Speirs (1985) noted that overcrowding in Trogoderma cultures might increase the rate of diapause.
Facultative hibernal diapause can usually be prevented in host insects by using long light with temperatures >20°C, depending upon the species. Such an environment mimics the natural summer when insects with a facultative hibernal diapause usually continue to reproduce. Daily photophases used to prevent diapause typically range from 16 hrs for the onion maggot (Tolman et al. 1985) and spruce budworms (Robertson 1985a), to 18 hrs for the codling moth (Ashby et al. 1985), and the large white butterfly, Pieris brassicae (Gardiner 1985c), to continuous light for the tobacco hornworm (Baumhover 1985) and the European corn borer (Guthrie et al. 1985).
Some insects, such as the Egyptian alfalfa weevil, have an aestival diapause and are active in nature in the spring. New generation adults aestivate until fall. Under laboratory conditions of 21°C and a daily photophase of 8 hours, at least some individuals of each generation will forego aestivation and produce eggs.
Chilling insects for several weeks to several months, whether facultative or obligatory, typically breaks diapause. Egg diapause has been broken in the grasshopper genus Melanoplus by exposure to 10°C for 3-12 months (Henry 1985); in the Douglas-fir tussock moth by conditioning at 5-10°C for 4-6 months (Robertson 1985b); and in the hemlock looper by storage at 2°C for 3-9 months (Grisdale 1985a). Examples of chilling requirements to terminate diapause in larvae include 1°C for 18-35 weeks for eastern spruce budworms (Grisdale 1984), and 5"2°C for 2-6 months for the red oak borer, Enaphalodes fufulus (Haldeman) (Galford 1985).
Pupal hibernal diapause may be terminated similarly. Tolman et al. (1985) were able to break diapause in the onion maggot by chilling at 1"0.5°C for 2-12 months. The same procedure works well for the cabbage maggot, except chilling must be a minimum of four months (Whistlecraft et al. 1985b). Bolle (1985) noted that pupae of the European cherry fruit fly, Rhagoletis cerasi (L.), required refrigeration at 4°C for 3-5 months to break an obligatory diapause. There is, however, a time limit beyond which insects cannot be safely refrigerated.
The length of diapause conditioning of the egg stage can affect the sex ratio of emerging gypsy moth adults. After a short chilling period of 120 days, the sex ratio of the first 25% of hatching larvae will be male biased: after a long chilling period of 180 days it will be female biased (O'Dell et al. 1985).
Different host insect stages and different species vary in developmental environmental requirements. Some examples indicate the range of variations and similarities. Phytophagous insect eggs frequently require moisture or high humidity to prevent desiccation, and providing just the right amount of moisture to maintain the eggs is critical. Singh et al. (1985) held eggs of the light brown apple moth in airtight containers to maintain egg turgidity. However, the container had to be checked frequently to remove condensed moisture in order to prevent fungus contamination.
Another way to control fungus contamination while providing moisture to eggs was developed by Clair et al. (1987). They cut elm leaf beetle, Xanthogaleruca luteola (Müller), and clusters from elm leaves and placed them on cloth and filter paper in a plastic petri dish. This combination was kept moist by a wick of dental cotton extending through a hole in the petri dish to a water reservoir. The eggs were then exposed to air circulation, preventing stagnant air which is conducive to fungal growth. This type of system is useful for maintaining a variety of eggs.
Eggs treated with sodium hypochlorite need to be held on moist cloth and filter paper to prevent desiccation. However, this can usually be done in closed containers since the egg treatment also reduces fungal contamination.
Varying conditions in temperature and relative humidity are commonly used, with only periodic conditions for lighting. For example, Navon (1985) reared Spodoptera littoralis with a photoperiod of 16 hr, 24°C and RH of 50-70%. Sometimes these workers used completely aperiodic environmental conditions (i.e., constant temperature, RH and light) for the rearing. Insects reared in this manner include the southern armyworm (Wight 1985), the lesser peachtree borer (Reed & Tromley 1985b), the European corn borer (Guthrie et al. 1985), and the tobacco hornworm (Baumhover 1985).
Fluctuating environmental rearing conditions retain and promote insect vigor. Greenberg & George (1985) cited Kamal (1958) who reported that fluctuating temperature and humidity increased the longevity of several laboratory reared calliphorid and sarcophagid species, as did a larger cage size.
The optimum rearing temperature must be experimentally determined for each insect and strain. Orthopterans frequently require high rearing temperatures, although some need cool conditions. McFarlane (1985) found that crickets do best at temperatures between 28°C and 35°C. When reared at 20°C the mean weight of the emergent adults was greater than at higher temperatures, but they would not reproduce. However, the range grasshopper, Arphia conspersa, requires much lower laboratory rearing temperatures than some other species. Willey (1985) raised the various stages at 22°C and variable RH, with a photoperiod of 12 hr, at which a generation could be completed in an average of 6 months. Temperatures above 30°C resulted in lower hatch and weak grasshopper.
A few insects change forms (morphotypes) depending on the rearing conditions. Forbes et al. (1985) reported that aphids would reproduce parthenogenetically in the laboratory at 20"1°C with a photophase of 16 hr. The production of sexual forms necessitated a maximum photophase of 8-12 hours with a temperature of 15°C or less. Medrano & Heinrichs (1985), however, found that production of the two distinct morphotypes of the brown planthopper, Nilaparvata lugens (Stal), was governed by nymphal density and food availability. They noted that a short winged form developed with low nymphal density and abundant food, whereas a long winged form developed under opposite conditions.
Humidity, moisture and substrate are often critical for insect pupation. Baumhover (1985) noted that pupation requirements of the tobacco hornworm are precise. Humidity must be controlled near 85% as higher or lower values will prevent adult ecdysis. A dehumidifier may be necessary to remove air moisture, since each prepupa loses 4 ml of water by the time of ecdysis. Further, prepupae require complete darkness to make them inactive and must be held individually in flat cells to allow proper pupation. Pupae must be well hardened before harvesting, as teneral individuals are easily injured.
Pupation substrates for various insects include materials such as sand for the potato tuberworm (Etzel 1985), and Hippelates eye gnats (Legner & Bay 1964, 1965), sawdust for the Queensland fruit fly (Heather & Corcoran 1985), a sawdust / ground corn cob mixture for the lesser peachtree borer (Reed & Tromley 1985b) and vermiculite for Spodoptera littoralis (Navon 1985). The pupation medium can be quite critical, as it is in rearing the southern armyworm. Wight (1985) noted that vermiculite no larger than 6-mm mesh must be used for this insect, and with the proper moisture content (400 ml water in 1200 ml vermiculite). If the medium is too wet, there is a high pupal mortality, and if too dry, dead pupae or defective moths result.
Lighting conditions seem to be of particular importance to adult insects. The photoperiod under which immature insects are reared can even have a pronounced effect on the subsequent adults. For example, McFarlane (1985) found a dramatic photoperiodic effect on the house cricket, Acheta domesticus (L.), with adults surviving up to twice as long with a 14- rather than with a 12-hr nymphal rearing photophase.
Adults of many insects mate and oviposit best if they are provided with natural light through laboratory windows. Such insects include the Queensland fruit fly, Dacus tryoni (Heather & Corcoran 1985), the saltmarsh caterpillar, Estigmene acrea (Drury) (Vaile & Cowan 1985) and the light brown apple moth (Singh et al. 1985).
Lighting conditions required for different species vary greatly. Robertson (1985a) reported that spruce budworm adults mated most successfully in the day within 24-hr after emergence, and optimum oviposition also occurred in the dark at 23-26°C. She also noted that the best laboratory conditions for oviposition by the Douglas-fir tussock moth were complete darkness and 23-26°C (Robertson 1985b). However, hemlock loopers will not mate well in continuous light, and therefore require a light/dark cycle (Grisdale 1985a).
Sometimes adults mate and oviposit best if they are provided with a weal light during the scotophase. Guy et al. (1985) held cabbage looper moths under a photophase of 14-hr, with a 0.25-watt night-light during the scotophase. Leppla & Turner (1975) earlier had shown that maximum fecundity of the cabbage looper can be achieved with long intensity night illumination. Gardiner (1985b) likewise made use of long intensity light by utilizing a 7.5-watt bulb during the scotophase at a distance of 3-6 ft for mating and oviposition of the cabbage moth, Marnestra brassicae. A 60-100-watt bulb was used at the same location during the 12-hr photophase.
Guthrie et al. (1985) employed a slight asynchrony in light and temperature phases to provide for mating oviposition by European corn borer moths, with two more daily hours of higher temperature than of light. A room temperature of 27°C was maintained for 16-hr with 18-20°C prevailing for 8 hours. The lights were on for 14 hours starting one hour after initiation of 27°C.
Temperature can be critical by itself, though, without interacting with the photoperiod. Tolman et al (1985) showed that survival and fecundity of the onion maggot were substantially greater at 20°C than at 15°C, 25°C or 30°C.
Humidity must also be considered in providing optimum mating and oviposition conditions. Leppla (1985 reported that the velvet bean caterpillar mated and oviposited best with a relative humidity in excess of 80% and with a source of liquid food. Wight (1985) held the oviposition cage for southern armyworm moths over a pan of water, and covered the cage with black cloth to encourage oviposition. The humidity in the cage had to be in excess of 50% to obtain good mating, oviposition and egg hatch.
Mating can sometimes be quite difficult to achieve in the laboratory and may involve a variety of factors. Although Reed & Tromley (1985b) reared immatures of the lesser peachtree borer under aperiodic conditions, the adults were held under a 16-hr photophase for mating and oviposition. It was noted that proper environment was important to achieve mating, outdoor conditions being simulated whenever possible. Indoor conditions required adequate lighting and ventilation (to avoid pheromone accumulation). Moths were observed for mating and pairs in copula were removed, after which the females were allowed to oviposit.
Density may affect optimum mating and oviposition. Laboratory mating and oviposition of the large white butterfly, Pieris brassicae, requires a relatively large cage (100 x 90 x 75 cm) in which 200 adults are placed (Gardiner 1985c). Tobacco hornworm moths also require a large cage (137 x 121 x 125 cm) for just 50 pairs (Baumhover 1985). Low light conditions are also necessary (15 watt light for 12 hr and rheostat-reduced 7-1/2 watt light for 12 hr). On the other hand, a high density is not detrimental to mating and oviposition of the spruce budworm. Grisdale (1984) reported that up to 300 pairs of these moths could be crowded into a screened cage (35 x 35 x 25 cm).
The Age of adult insects is a further factor that must be considered for mating and oviposition. O'Dell et al. (1985) noted that gypsy moth females, Lymantria dispar (L.), would not mate once they began to lay eggs. Codling moth adults held for more than five days before mating have considerably reduced fecundity (Singh & Ashby 1985). Similarly, Grisdale (1985b) recommended mating female moths of the forest tent caterpillar as soon after eclosion as possible for optimum results.
Even members of the same insect family can vary dramatically in the ease of laboratory mating and oviposition. This is certainly true of the mosquito family Culicidae. Friend & Tanner (1985) reported that Culiseta inornata (Williston) males often initiate mating before females have completely emerged without special flight cages. Munstermann & Wasmuth (1985a) noted that Aedes aegypti (L.) also mates easily in confined spaces. However, these workers had to use beheaded, impaled males of the eastern tree hole mosquito, Aedes triseriatus (Say), in a forced copulation technique (Munstermann & Wasmuth 1985b). They noted that the Walton strain of A. triseriatus will mate satisfactorily in a cubical cage of at least 60 cm3. Bailey & Seawright (1984) reviewed a system useful to achieve rapid laboratory colonization of Anopheles albimanus Wiedemann, a vector of malaria. It was discovered that field collected females individually placed in 5-dram vials would lay more than 100 times the number of eggs of an equal number (500) of females placed together in a single cubical cage of 61 cm3. The degree of clustering of ovipositing adults and the amount of space provided can affect fertility as well as oviposition and must be considered in a production program.
Frequently insects will have a preoviposition period between emergence and oviposition during which they feed and develop their eggs. For example, adult cabbage maggots have a preoviposition period of 6-7 days at 19"1°C (Whistlecraft et al. 1985b), and adult onion maggots have a similar period, but at 22"1°C (Tolman et al. 1985).
After insects have been provided with appropriate mating conditions, they must be stimulated to oviposit. Insects that lay eggs in crevasses may often be easily induced to oviposit on crinkled wax paper or on cloth. A good oviposition substrate for the hemlock looper is a six-layer thick cheesecloth (Grisdale 1985a). Reed & Tromley (1985b) found that the lesser peachtree borer is also rather easily motivated to oviposit if moist cotton balls are provided. Other insects can be far more fastidious in their ovipositional requirements and ingenious systems must be devised. Heather & Corcoran (1985) used hollowed-out half apples as an ovipositional substrate for the Queensland fruit fly to enable easy egg collection. Boller (1985) devised a clever dome made of ceresin wax which served as an oviposition substrate for the European cherry fruit fly, Rhagoletis cerasi. Baumhover (1985) described artificial leaves composed of outdoor carpeting sandwiched between layers of polypropylene, which served as substitutes for tobacco leaves and were sprayed daily with a tobacco leaf extract to stimulate oviposition by tobacco hornworm moths.
Host Culture (Behavior)
Knowledge of insect behavior is obviously crucial to a successful rearing program. This is especially important in order to obtain an optimum ovipositional situation. Blenk et al. (1985) discovered that six or seven reared noctuid species would oviposit on the underside of a paper towel on top of an oviposition cage. The black cutworm, on the other hand, would only oviposit on paper toweling in the bottom of the cage. Guthrie et al. (1985) observed that European corn borer moths would oviposit only on smooth laboratory surfaces.
Chemical ovipositional stimuli may also be necessary. Adult seedcorn maggots oviposit in response to moist soil, decaying vegetation, germinating seeds and metabolites produced by seed borne microorganisms (Whistlecraft et al. 1985a). In some cases insect-produced chemicals may deter oviposition. Boller (1985) found that the European cherry fruit fly and the Mediterranean fruit fly produced oviposition deterring pheromones that lowered egg deposition in artificial devices. Consequently, the devices required frequent washing.
Insect responses to stimuli can simplify rearing. For example, the positive phototaxis of scale crawlers makes them easily collected. Papacek & Smith (1985) lighted a rearing room for oleander scale two hours before work began so that scale crawlers would accumulate on top of butternut pumpkins. Positive phototaxis is a common attribute of many insects and can be similarly used in their collection.
In some cases the combining of behavioral characteristics may be disadvantageous. Grisdale (1985a) reported that first instar hemlock loopers are strongly photopositive and active, but also cannibalistic. Since the larvae drink readily they can be sprayed lightly with distilled water and held in the dark at 18°C to reduce cannibalism.
Light can also affect insect emergence. Willey (1985) noted that range grasshopper nymphs, Arphia conspersa, hatch from eggs daily about 5-8 hr after start of the light cycle, so daily collection can be timed accordingly. Similarly Boller (1985) observed that the European cherry fruit fly and the Mediterranean fruit fly both emerge mostly during the morning and oviposit in the afternoon.
Host Culture (Techniques)
Many and varied techniques have been developed for arthropod rearing. Some have already been mentioned, other examples follow:
Stockpiling refrigerated hosts is advantageous. Effects of refrigerated host material on parasitoid and predator production must be resolved on a case-by-case basis. Legner (1979b) found that house fly pupae could be successfully refrigerated only at 10°C for <21 days before being used to raise three pteromalid parasitoids. Parasitoids given nonrefrigerated pupae produced significantly more female progeny, however. As the progeny did not differ significantly in biomass, the decreased reproductive potential when refrigerated hosts are used may not be readily apparent.
Precise storage temperatures can often be very critical in rearing insects. Ankersmit (1985) reported that newly laid eggs of the summer fruit tortrix, Adoxophytes orans, are killed when held at 5°C. However, embryonated eggs can be held at 5°C, but not for more than four days. There is also a critical temperature above which A. orana eggs hatch. Eggs held at 13°C will not hatch, but at 15°C there is about 70% hatchability.
The insect stage put in cold storage is also important. Adults of Arphia spp. range grasshoppers cannot be stored in the cold or without food for more than one day without losing vigor. However, eggs in diapause left in situ in soil can be stored at 2°C for 1-2 years if the soil remains moist, and eggs not in diapause may be stored for several months at 10-17°C (Willey 1985).
Glass & Roelofs (1985) reported that newly hatched red-banded leafrollers could be stored at least 7 days at 5-7°C with 100% RH. However, leafroller pupae can be stored for six months at 5°C by inducing diapause through larval exposure to an 11-hr photophase.
Differential cold storage of the sexes can be used to synchronize emergence, since males of most insects develop more rapidly than females and emerge first. To achieve synchronized emergence of male and female hemlock lookers, Grisdale (1985a) sexed freshly formed pupae and stored the males initially at a temperature 4°C lower than that for the females.
To maximize and quantify insect production, suitable methods of determining insect numbers are necessary. One way is to estimate by weight. Baumhover (1985) weighed tobacco hornworm eggs to ascertain their numbers (400 eggs weight 0.534g). He cautioned to weigh only fresh eggs because they lose 20% of their initial weight by hatching time. Similarly Moor & Whisnant (1985) estimated numbers of reared boll weevil adults by first weighing a sample of 10 and then the total collection.
Egg number determination is useful for adjusting available food per individual to maximize production and food usage. For example, Guy et al. (1985) reared cabbage looper larvae gregariously on artificial diet, with diet amount per individual adjusted by placing an appropriate number of eggs in each container. Eggs were applied to squares of paper toweling with a medicine dropper with 50-60 eggs per spot. Squares with dried egg spots were glued to each container lid with casein glue.
Various means have been developed to separate insects from a substrate or from each other. Rahalkar et al (1985) separated eggs of the red palm weevil, Rhyncophorus ferrugineus Oliver, from shredded sugar cane by placing this ovipositional substrate into a 30% aqueous solution of glycerol. After the sugarcane shreds sink, the floating eggs are collected with a strainer. Similarly Martel et al. (1975) developed a method of extracting eggs of the carrot weevil, Listronotus oregonensis, from carrot pieces.
Morgan (1985) separated viable from nonviable house fly eggs by placing them in water where viable eggs sink. The technique also works to separate pupae from larval medium since the pupae float. Tolman et al. (1985) designed a simple flotation device to separate pupae of the onion maggot from the cut onion and sand larval substrate. However, the pupae must be at least 48 hours old before they will float on water. Greenberg & George (1985) separated blowfly eggs by using 1% sodium sulfite to dissolve the adhesive holding them together.
Anesthetizing insects makes handling easier. Carbon dioxide is used when the brevity of the effect is not a hindrance. Longer activity is provided by a combination of ethyl ether and carbon dioxide (Etzel 1985). Munstermann & Wasmuth (1985b) made a device utilizing nitrogen gas saturated with water vapor to anesthetize adult eastern tree hold mosquitoes. Nettles (1987) used nitrogen anesthesia to enable sexing of the adult tachinid Eucelatoria bryani Sabrosky.
A general problem in producing lepidopterans is the accumulation of moth scales, which can be highly allergenic to humans. The scales are commonly removed from the rearing environment with air filtration. However, another method was used by Baumhover (1985) in rearing the tobacco hornworm. He noted that 85% RH in mating and oviposition cages prevented most scale pollution if the moths remained inactive.
Methods of containing arthropods in rearing units are varied. Sleeve cages of assorted sizes with wooden frames, organdy cloth sides, glass topes and cloth sleeves to enable manipulation of cage contents are commonly used. Some arthropods, particularly mites, are often reared in open units. Margolies (1987) used a mixture of 4ml clove oil in 100 g lanolin applied to the edge of a petri dish to stop tetranychid mites escapes. Physical handling of insects can be critical to their rearing. Some insects are particularly fragile in at least one or more stages. Boller (1985) observed that young fruit fly pupae have to be handled very gently to prevent ruptures of the fly muscles.
Host Culture (Quality)
The required quality of cultured insects depends on their intended use. Waage et al. (1985) noted that species, size and stage are factors affecting the quality of a host for parasitoids and predators.
Three aspects of quality are standards, assessment and control. A survey of the literature revealed that the most commonly used quality criterion was fertility. The fertility test was used in producing Heteroptera (Jones 1985), Coleoptera (Jackson 1985, Rahalkar et al. 1985), and Lepidoptera (Robertson 1985a, 1985b; Bartlett & Wold 1985, O'Dell et al. 1985, Guy et al. 1985, Navon 1985, Reed & Tromley 1985). King & Hartley (1985a) used this criterion in raising the sugarcane borer as a host for the tachinid Lixophaga diatraeae, but also considered fecundity, larval and adult survival and percent adult emergence. These quality criteria are in common use, although mortality is often employed instead of survival. Another typical criterion was size, either in dimensions or weight.
Patana (1985b) believed that continued reproduction and survival are the most significant indicators of insect quality in long term laboratory cultures of at least one to five years. Leppla et al. (1984) commented that even the best programs for mass producing cabbage loopers can depend only on quantity, since quality control is difficult.
Exercise 28.1--In culturing hosts, what principal biological characteristics does a researcher strive to maintain? Give a few procedural examples of how such traits might be maintained?
Exercise 28.2--What operational procedure must be routinely and rigorously followed to guarantee healthy cultures of hosts?
Exercise 28.3--Name an arthropod behavioral trait that facilitates the removal of hosts from culture media.
Exercise 28.4--How would you practically counteract the trend toward homozygosity in cultures of entomophages?