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Introduction 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). Host
Food 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 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 ] Genetic
Considerations 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. Physical
Environment 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? REFERENCES: Please refer to <bc-30.ref.htm> [Additional references may be found at
MELVYL Library ] |