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CULTURE OF HOSTS FOR NATURAL ENEMY
PRODUCTION
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Purpose 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 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 which 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 can 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 that 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. This is usually done by specialists in large mass production
facilities. 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 undiseased 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 subcolony 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. Diapause, whether facultative or obligatory, is typically
broken by chilling insects for several weeks to
several months. 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), 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 |