Viral Diseases

  1. General Features
  2. Taxonomy
  3. Baculoviridae - Occluded
    1. Taxonomy
    2. Infection
    3. Gross Pathology
      1. Larvae (Early Instars)
      2. Larvae (5- to 6-day-old)
      3. Pupae
      4. Adults
    4. Light Microscopy
    5. Transmission
    6. Host Range and Geographical Distribution
    7. Microbial Control
  4. Baculoviridae - Nonoccluded
    1. Infection
    2. Host Range and Geographical Distribution
    3. Microbial Control and Elimination from Microplitis croceipes and Heliothis virescens Colonies
  5. Poxviridae
  6. Ascoviridae
    1. Gross Pathology
    2. Cytopathology
    3. Host Range and Geographical Distribution
    4. Microbial Control
  7. Gonad-specific Virus (GSV)
    1. General Pathology
    2. Detection
    3. Effects on Oviposition
    4. Hosts and Geographical Distribution
    5. Microbial Control and Elimination from Helicoverpa zea and H. virescens Colonies
  8. Reoviridae - Cytoplasmic Polyhedrosis Viruses (CPVs)
    1. Taxonomy
    2. Histopathology
    3. Infection
    4. Transmission
    5. Host Range and Geographical Distribution
    6. Microbial Control
    7. Rearing CPV-free Heliothines
  9. Iridoviridae
    1. Gross Pathology
    2. Inapparent Infection
    3. Host Range and Geographical Distribution
    4. Microbial Control
  10. References

General features

Viruses are infectious units (obligate parasites) comprised of either DNA or RNA enclosed in a protective coat. Their nucleic acid contains information necessary for their replication in a susceptible host cell. They contain no energy-producing enzyme system, no functional ribosomes or other cellular organells; these are supplied by the cell in which they replicate. The cell may also supply some of the enzymes necessary for viral replication (Hull et al., 1990). The sixth report of the International Committee on Taxonomy of Viruses (Murphy et al., 1995) placed all viruses in one order, 71 families, 9 subfamilies, 164 genera, more than 3,600 virus species, and hundreds of unclassified viruses.


Insect pathogenic viruses have been placed in 12 viral families but many remain unclassified. Viruses in three families, Baculoviridae, Poxviridae, and Reoviridae, produce occlusion bodies in which virions are occluded randomly. Many insect viruses are assigned to families of vertebrate viruses, but some are in families with viruses specific for insects and related invertebrates (e.g., Baculoviridae, Polydnaviridae, and Ascoviridae). The tobacco budworm, Heliothis virescens, and the cotton bollworm, Helicoverpa (=Heliothis) zea (Lepidoptera: Noctuidae), have been reported to be susceptible to several viruses from the families Baculoviridae, Poxviridae, Reoviridae, Ascoviridae, Iridoviridae, and Gonad-specific virus.

Baculoviridae - Occluded

Fig. 1 H. virescens nuclear polyhedrosis virus (NPV).
Infected grossly disintegrated larval body.
Fat body cells showing enlarged nuclei packed with polyhedra (arrows)
Epidermal nuclei of infected cells showing enlarged nuclei.
TEM micrograph of cross section of a polyhedron showing virons randomly scattered through it.)


The family Baculoviridae is composed of two genera, Nucleopolyhedrovirus (NPV) and Granulovirus (GV) (Murphy et al., 1995). Baculoviruses are characterized by occlusion of virions in a paracrystalline protein matrix. Nuclear polyhedrosis viruses have polyhedral occlusion bodies (OBs) composed of polyhedrin, and granulosis viruses have OBs made up of a protein called granulin.

More than 520 NPVs have been identified in insects (Martignoni and Iwai, 1986). The nucleocapsid is similar in all members of Baculoviridae and has a DNA core surrounded by a protein capsid. The virions measure from 40-60 x 200-400 nm and are encased by an outer lipoprotein envelope. The nucleocapsids are packaged as one (S) or multiple (M) within a single viral envelope. Murphy et al. (1995) lists H. zea virus as SNPV.


The most common mode of entry of NPVs into the host is by ingestion of virus during feeding by an insect. Polyhedra are dissolved by the alkaline gut juices, and the liberated virions penetrate the gut epithelium and multiply in the epidermis, tracheal matrix, fat bodies, and blood cells of lepidopterous larvae. In hymenopterous larvae, the virus proliferates in the midgut epithelium. The disease is usually fatal. Other less common ports of entry and infection include transovarial and transo vum passage, through the spiracles, and by parasitism.

Gross Patholo

(H. virescens and H. zea)

Larvae (Early Instars)
Early instar larvae may die within 1 to 2 days post infection (Ignoffo, 1966).

Larvae (5- to 6-day-old)
Signs and symptoms of disease caused by NPVs are not apparent for several days (room temperature 5 to 6 days). Larvae exhibit a loss of appetite. Color change may be evident (day 6). Integument becomes opaque and assumes a shiny and swollen appearance (day 6). The internal tissues are in a state of disintegration (day 6). The larvae are difficult to remove from their habitats without breaking the skin and liberating the liquefied contents (day 7). Time from infection until death for 5-day-old larvae averages 6 to 8 days.

There is no external change observed in pupae during the greater part of the incubation period. But, at the end of the disease cycle, the skin is very easily ruptured on handling, and the pupal body becomes liquefied. Sometimes black markings are observed on the body surface of the pupae about the time of death.

Even though death usually occurs in the larval stage, larvae infected in the late instars may survive to the adult stage. Such infections are followed by breakdown of the skin and subsequent liberation of body contents into the environment.

Light Microscopy

A puncture of the cuticle of an infected larva reveals a "broken-down" hemolymph, which is filled with polyhedra. Smears prepared from diseased larvae show the presence of large numbers of polyhedra that can be readily seen with a light microscope.


Transmission is horizontal when the pathogen is transferred from individual to individual but not directly from parent to offspring. Transmission is vertical when there is direct transfer of the pathogen from parent insects to their progeny (Andreadis, 1987). Nuclear polyhedrosis viruses and GVs are transmitted both horizontally and vertically.

Host Range and Geographical Distribution

The majority of insects susceptible to NPV are in Lepidoptera, but some species are in Hymenoptera, Coleoptera, Trichoptera, and Diptera. Many of the NPVs appear to be genus- or family-specific.

Microbial Control

Federici (1993) and Tanada and Kaya (1993) reported that NPVs and GVs will continue to gather the most interest for use as either conventional or recombinant viral insecticides because they are known "from" numerous important insect pests, safe to non-target species including man, easy to produce, easy to store, can be applied with existing technology, and are relatively easy to manipulate. In the United States, several NPV-based microbial insecticides have been registered by commerical companies and governmental agencies. According to Tanada and Kaya (1993), Heliothis NPV was registered in the United States for control of cotton bollworm and tobacco budworm in 1975. A new NPV formulation (Gemstar®; based on H. zea NP V for control of H. virescens and H. zea) was introduced in 1996. However, even a new generation of microbial insecticides are not used on a large scale or are not commercially obtainable. Probably the major reasons for the lack of interest in viral pesticides are limited market size, questionable efficacy, high dose and the long period requried to kill late-instar larvae, uncertain profitability for the producers, and growers' general fear of using biocides (and viruses in particular). With th e increasing emphasis on genetically engineered viral insecticides, a new generation of NPVs that are safe to nontarget organisms, have a wide host range, and kill targeted pests fast will, in our opinion, be commercially available in the near future.

Baculoviridae - Nonoccluded

Figure 2. Electron micrographs of Microplitis croceipes nonoccluded baculovirus (MC-NOBV) in M. croceipes and H. virescens. (Last two electron micrographs courtsey of A.A.A. Gothama.)
Parts of three hypertrophic infected nuclei of M. croceipes fat body cells.
Enlarged virions.
Mc-NOBV virions in hypertrophic nuclei of infected fat body cells of H. virescens.
MC-NOBV in intercellular spaces of H. virescens fat body cells

Recently, Murphy et al. (1995), transferred the subfamily Nudibaculovirinae to the unclassified group of viruses. The subfamily consists of viruses that are not occluded (in any type of occlusion body) at any stage of their replication cycles. The virions measure approximately 75 x 270 nm. The nonoccluded baculovirus of Microplitis croceipes (Mc-NOBV) was the first virus reported to be pathogenic to a hymenopteran endoparasitoid (Hamm et al., 1988). Gothama et al., 1997 (in press) have recently found that Mc-NOBV also replicates in H. virescens.

Chittihunsa and Sikorowski (1995) reported high concentrations of Mc-NOBV in most tissues of infected wasps, including the alimentary canal, fat body, circulatory system, respiratory system, and integument. This is in agreement with Hamm et al. (1992), who reported that the principal sites of infection in M. croceipes wasps are the fat body and midgut. In H. virescens the virus was present in the midgut, tracheal matrix, fat body, and hemolymph (Gothama et al., in press).


Infection of H. virescens, as well as in other hosts of Mc-NOBV, occurs per os (by mouth) and during parasitization. No noticeable signs nor symptoms were observed during the early stages of the H. virescens larval infection. In the prepupal stage, some larvae fail to pupate and some pupae fail to eclose to adults. In general, infected larvae weigh less than healthy larvae (Gothama et al., in press).

Histopathological studies, bioassays and enzyme-linked immunosborent assay (ELISA) showed that feces from Mc-NOBV infected H. virescens and M. croceipes contained virulent virus, capable of producing infections. Thus, virus-contaminated foliage may provide inoculum for H. virescens larvae feeding on it.

During parasitization of infected H. virescens larvae, healthy M. croceipes females may contaminate their ovipositors with Mc-NOBV. These females may in turn infect many H. virescens larvae during oviposition by direct insertion of a virus-contaminated ovipositor into the body of a healthy H. virescens larva. Infected males may also infect females during mating (veneral transmission).

Host Range and Geographical Distribution

Presently, M. croceipes is the primary host while H. virescens, Cotesia marginiventris, and Cardiochiles nigriceps are secondary hosts of this virus (Gothama et al., in press). Not much is known of the distribution of this virus, though it is probably coincident with the hosts.

Microbial Control and Elimination from Endoparasitoids and H. virescens Colonies

Microplitis croceipes is a larval endoparasitoid that attacks two important agricultural pests, H. virescens and H. zea. Release of diseased endoparasitoids, particularly on a mass scale, could have negative effects on the feral populations of the endoparasitoid. Also, virus-infected H. virescens would magnify the spread of Mc-NOBV throughout the parasitoid population. Thus, this virus could have damaging effects on mass rearing and field releases of M. crociepes as a biological control agent of H. virescens.

Elimination of Mc-NOBV from a laboratory colony of M. croceipes and H. virescens is difficult for two reasons: 1) Mc-NOBV can be transmitted transovarially (within the egg), and 2) Mc-NOBV can be present in H. virescens larvae without any apparent signs and symptoms. Three methods are available to suppress or eliminate the disease from M. croceipes and H. virescens colonies:

Adult selection method. (M. croceipes). Use for breeding colony wasps that live for 7-days or longer (virus-infected wasps usually live fewer than 7 days) (Hamm et al., 1988). The life span of healthy M. croceipes adults is about 2 weeks or longer. This method would eliminate most of the virus-infected adults.

Transmission electron microscope (TEM) method.
(M. croceipes and its hosts). Newly-emerged adults from apparently normal colonies are paired for mating and oviposition. About 20 healthy H. virescens larvae (5-day-old) are exposed per day to each pair of wasps until the death of the female. Then, each parent pair is examined by negative-stain TEM for the presence of Mc-NOBV. Only progeny from virus-free parents are used for establishment of virus-free M. croceipes colonies. To check for virus infection, the internal tissues including the fat bodies and midguts of wasps or heliothine larvae are extracted, macerated, and suspended in a 0.9% sodium chloride solution. A drop of this suspension is placed on a formvar and carbon-coated 200 mesh copper grid for 5 minutes. The liquid is gently removed with a filter paper wick and the grid stained immediately with 2% phosphotungstic acid for 1 minute and examined with TEM. This method is expensive but in combination with the first or third method would help select and maintain virus-free colonies.

Enzyme-linked immunosorbent assay (ELISA) method. (M. croceipes and its hosts). A double antibody sandwich ELISA is performed to detect Mc-NOBV infection in M. croceipes and other possible host insects. Antiserum is prepared in rabbits, and the resulting immunoglobulin (IgG) is purified. The IgG is conjugated with alkaline-phosphatase as described by Clark and Bar-Joseph (1984). The assay is performed using a modified method of Clark and Adams (1977) and Oien and Ragsdale (1992), and the results are verified using negative-stain TEM (Gothama et al., in press). This method is applicable for routine examinations in large rearing facilities of the parasitoid and its host colonies for the presence of Mc-NOBV.


Poxviridae is a large family containing members infecting vertebrates and invertebrates. Those infecting vertebrates cause diseases such as smallpox, cowpox, fowlpox, etc. Insect poxviruses are placed in the subfamily Entomopoxvirinae with three probable genera (Murphy et al., 1995):

Enomopoxvirus A - This genus includes poxviruses of Coleoptera.
Entomopoxvirus B - This genus includes poxviruses of Lepidoptera and Orthoptera.
Entomopoxvirus C - This genus includes poxviruses of Diptera.

Milner and Beaton (1987) reported infection in H. zea by Entomopoxvirus (in Goodwin et al., 1991). The virus size is 340 x 270 nm, the shape is elipsoid, and the affected tissue is the fat body. The results of surveys by Smith et al. (1976) and Sikorowski (unpublished) of the pathogens of H. virescens and H. zea in Mississippi have not detected the virus in either species.

Entomopoxviruses closely resemble the pox viruses of vertebrate animals; for this reason, this group of viruses is not considered as a good candidate for insect control, even though there is no record of any member of the group¹s causing disease in vertebrates. With increasing knowledge and decreasing fear of this group of viruses, we may use them as control agents of agricultural pests in the near future.


Figure 3. Ascovirus virions. Negatively stained mature virions illustrating the reticulate surface. (Photos courtesy of Dr. John J. Hamm.)
Ascovirus viriions isolated from H. virescens.
Ascovirus virion isolated from H. zea

Viruses belonging to this family are currently known only from larvae of species in the lepidopteran family Noctuidae, in which they cause a chronic and fatal disease (Federici et al., 1991). Enveloped virions are approximately 130 x 400 nm.

Gross Pathology

Laboratory infected heliothine larvae turn an opaque yellowish-white and have difficulty in molting. Field-collected larvae differ little, if any, in appearance from healthy larvae (Federici et al., 1991). Larvae typically survive for 2 to 5 weeks after infection but feed only intermittently, gain little weight, and eventually die.


Infection begins with hypertrophy of the nucleus of infected cells. Hypertrophy of the nucleus continues, leading to cellular hypertrophy. After fragmentation of the nuclear membrane, sheets of cytoplasmic membranes assemble throughout the cell and coalesce, partitioning the cell into a cluster of vesicles (virion-containing vesicles are formed by the cleavage of host cells). Once vesicle formation is complete, the vesicles dissociate from each other and accumulate within the tissue in which they are formed. As the disease progresses, the basement membrane of infected tissues is disrupted, and vesicles are released into the hemolymph (Federici et al., 1991).

Host Range and Geographical Distribution

The first ascoviruses were discovered by Adams et al. (1979) in the cotton bollworm, H. zea, and the clover cutworm, Scotogramma trifolii. Since then, ascoviruses have been isolated from larvae of H. virescens (Carner and Hudson, 1983) and other noctuid species (Federici et al., 1991). Ascovirus isolates from Heliothis spp. have been transmitted per os into larvae of several other species of noctuids (Federici, 1983).

Microbial Control

Ascoviruses, because of their poor infectivity per os, possess virtually no potential for use as viral control agents (Federici, 1993).

Gonad-specific Virus (GSV)

Figure 4. H. zea infected with gonad-specific virus (GSV). (Photos courtesy of Dr. John J. Hamm.)
Abdomen of GSV-infected female moth showing waxy plug.
Section of waxy plug from GSV-infected female showing vesicles containing rod-shaped GSV particles
Intranuclear virions of GSV showing longitudinal and cross sections.

Atrophy of the ovarian and testicular organs in the culture of adults of H. zea was reported for the first time by Herzog and Phillips (1982). The abnormal trait occurred through five succesive generations of the culture and ranged in prevalence from 63 to 78%. Raina and Adams (1995) first reported that the disease was caused by the Gonad-specific virus (GSV).

General Pathology

The syndrome characteristic of this disease is primarily manifested in adults examined by dissection. Infected females have grossly deformed common and lateral oviducts. The common oviduct is full of a white mass of occlusion bodies. The female moths are without functional reproductive systems. They have no ovaries, bursa copulatrix, accessary glands, or spermatheca. In the GSV-infected males, seminal vesicles, vasa deferentia, duplexes, and accessory glands are absent (Raina and Adams, 1995; Hamm et al., 1996). Nucleocapsids measure 382 x 77 nm (Raina and Adams, 1995).

The virus produces atypical occlusion bodies, which contain large numbers of virions. The occlusion bodies differ from a typical NPV polyhedra in that they have a granular matrix rather than a typical polyhedrin protein matrix, and they have a host-derived membrane rather than a virus-derived polyheron membrane. Raina and Adams (1995) also found that the virus replicates in nuclei of cells in the distal part of the oviduct. Gonad-specific virus does not cause larval mortality. The virus was observed in asymptomatic adult females.


Infected males showed no external indication of the agonadal conditon. However, most agonadal females could be identified externally by a white, waxy plug protruding from the vulva (Hamm et al., 1996).

Effects on Oviposition

As is apparent from the publications of Herzog and Phillips (1982), Raina and Adams (1995), and Hamm et al. (1996), fecundity of infected females is greatly reduced or nonexistent because of the degradation or elimination of reproductive organs. However, lightly infected females can be asymptomatic.

Hosts and Geographical Distribution

Presently, the virus has been reported only as GSV of H. zea adults. However, it may be carried asymptomatically in H. virescens (Raina, personal communication). The host range and the geographical distribution of this virus have not been investigated.

Microbial Control and Elimination from H. zea and H. virescens Colonies

The suitability of GSV as a biological control agent has not yet been investigated. The prevalance and range of this virus in the feral heliothine is also unknown at this time.

In insectaries, a number of H. zea adults that appear normal are frequently infected inapparently with GSV, and the presence of the virus is usually evidenced by a decline in egg production and general vigor of the colony. Microscopic examination of the adult reproductive organs is required for final diagnosis.

Perhaps the most efficient method to eliminate GSV from the heliothine colonies is to destroy the infected colony, sanitize the laboratory with sodium hypochlorite (0.5%) or paint the laboratory furniture and walls with latex or other paint, and sanitize equipment with sodium hypochlorite; then introduce into the laboratory a disease-free colony. This is the only method that eliminates GSV from the colony.

An ELISA may be used to distinguish partially infected colonies of healthy and diseased adults. In this method, surface-disinfected pupae are placed individually into plastic cups, and the meconium from each newly emerged adult is analyzed for the presence of the virus. This is a quick, inexpensive method for detection of the virus in heliothine colonies and may be a method of choice for a large lepidopterous, insect-producing insectary; however, some weak positives may escape detection by this method.

Control of GSV in H. zea colonies can be accomplished by using eggs only from the first or second night of oviposition. Hamm et al. (1996) reported that the incidence of agonadal progeny in a colony increased with the day of oviposition from 5. 4% for day 1 and 17% for day 2 to 62.8% for day 3. Thus the presence of GSV in the H. zea colony may be reduced by using eggs only from the first or second night of oviposition. This method, if used routinely, may eliminate most, but not all, of the infected insects.

Polyhedrosis Viruses (CPVs)

Figure 5. Midgut epithelium of H. virescens larvae infected with cytoplasmic polyhedrosis virus (CPV).
Infected epithelium showing polyhedra (arrowheads, microvilli (mv), and degenerated microvilli (dmv) immediately prior to rupture of the cell (arro w).

Figure 6. H. virescens infected with cytoplasmic polyhedrosis virus (CPV)
High magnification of polyhedra, scanning electron micrograph, arrows indicate polyhedra, bar = 2 µm.
Thin section of polyhedra showing virus particles (arrows) and virogenic stroma (vs), transmission electron micrograph.
Virus particles of CPV released from polyhedra by treatment with dilute alkali and negatively stained with phosphotungstic acid showing spikes (arrows, bar = 50 nm.

Polyhedroses, virus diseases of insects characterized by the formation of polyhedral inclusions in tissues of infected insects, have been known for many years and include the bulk of viral diseases (Smith, 1963). Later, these were separated into two separate groups of viruses causing different disease symptoms. The existence of a separate type of polyhedrosis virus, the virions of which are near-spherical instead of rod-shaped, was reported for the first time by Smith and Wycoff (1950) in the larvae of Arctia caja and A. villica. This may be considered the first record of a cytoplasmic polyhedrosis virus (CPV). The CPV from H. zea was reported by Smith and Rivers (1956).

Cytoplasmic polyhedrosis virus was noticed first by Ishimori in 1934, when he observed polyhedra in the cytoplasm of midgut cells of a diseased silkworm larva (Aruga, 1971). Since then, cytoplasmic polyhedrosis has been recognized as one of the most important diseases of the silkworm, inflicting substantial economic losses to the sericulture industry in Japan (Aruga, 1971). Cytoplasmic polyhedrosis viruses have a very wide host range, affecting mainly lepidopterous insects. Martignoni and Iwai (1981) listed 221 known insect hosts of CPVs. Studies of H. virescens showed that CPV-infected larvae weighed 5% less than the healthy ones (Simmons & Sikorowski, 1973). Infected larvae required 4 days longer to reach the pupal stage than healthy larvae, and diseased pupae weighed 60 mg less than healthy pupae. Diseased moths lived 9 days less than the healthy moths, and the production of eggs was reduced by 68% in diseased females.


According to the International Committee on Taxonomy of Viruses (ICTV), CPVs are classified under the family Reoviridae and genus Cypovirus (Francki et al. 1991). The CPVs of insects are the only group of the family Reoviridae characterized by the presence of polyhedral inclusion bodies.


Cytoplasmic polyhedrosis viruses are distinguished by the production of crystal-like inclusion bodies, or polyhedra, normally located in the cytoplasm of virus-infected cells in the midgut epithelium. A few to several hundred virus particles may be occluded in the inclusion body.

Polyhedra can range widely in diameter from a fraction of a micron to several microns. They are characteristic of the terminal stage of infection, are highly stable, and serve as the vehicle for transmission of infectious virus from one susceptible host to another.

The histopathology of Heliothis CPV is based on work of Bong and Sikorowski (1991 and 1991a). Small polyhedral inclusion bodies (PIBs) are present in columnar cells of the midgut 1 or 2 days after treatment with CPV. Virions are partially or completely occluded in a polyhedral matrix to form PIBs at the periphery of virogenic stroma. Polyhedral inclusion bodies are dodecahedral in shape with hexagonal outlines in cross-section. Virions measure from 47 to 52 nm in diameter. The microvilli o f infected columnar cells are not affected until immediately prior to rupture of the cell. At this time, the microvilli are partially or almost completely absent from infected cells. Infected columnar cells may rupture to release PIBs into the gut lumen 2 or 3 days after infection. Polyhedral inclusion bodies are also released into the lumen by extrusion of heavily infected columnar cells. Mitochondria and endoplasmic reticulum deteriorate as infection progresses, and in many cells with advanced infection, the nucleus is obscured or appears absent. Polyhedral inclusion bodies are found in goblet cells 5 days after infection. Infected goblet cells degenerate to such an extent that only a few of the original microvilli-like cytoplasmic projections and cell organelles remain.


Heliothis CPV infections produce a chronic disease, which causes structural damage to the midgut epithelium. The virus infection causes cessation of feeding and reduction of the larval size. As a result, CPV-infected larvae produce smaller pupae and adults and require a longer time to complete the larval stage.

Heliothis CPV may be acquired either per os or as a result of injury, e.g. through parasitism, with the larval stages being the most susceptible to infection. Oral infection is the most common mode of acquisition. Virus infection of larvae follows the ingestion of food contaminated with CPV polyhedra. The polyhedra are dissolved in the alkaline gut juices of the larva. The free virions then infect susceptible cells. At the final stage of infection, the cells rupture and release polyhedra into the gut lumen.


In H. virescens, polyhedra are found mainly in the epithelial cells of the midgut. In the advanced stage of disease development, polyhedra are liberated from the damaged cells into the gut and excreted with the feces (Sikorowski et al., 1973; Bong and Sikorowski, 1991). Feces from diseased larvae contain virulent virus, which is produced throughout the larval stage. Adults contaminate the environment with CPV present in their meconium and feces. Thus, almost all stages of CPV-infected insects may contaminate their environment with the virulent virus.

Sikorowski et al. (1973) reported that a major means of transmission of CPV to the next generation is surface contamination of eggs. Larvae are infected during emergence from the egg when they consume a portion of the CPV-contaminated chorion (egg shell).

Host Range and Geographical Distribution

Cytoplasmic polyhedrosis viruses appear to have a very wide host range, but the assessment is complicated by the possible activation of occult (masked) CPVs. Thus, the possibility of the activation of an occult CPV cannot be ruled out (Smith, 1976).

Cytoplasmic polyhedrosis viruses are among the most prevalent of insect pathogens and have been isolated from 173 species of Lepidoptera, 11 Hymenoptera, 32 Diptera, 2 Neuroptera, 2 Coleoptera (Martignoni and Iwai, 1981), and a freshwater crustacean, Simocephalus expinosus (Federici and Hazard, 1975). Ignoffo and Adams (1966) successfully transmitted the pink bollworm CPV to the cabbage looper, Trichoplusia ni; bollworm, H. zea; and tobacco budworm, H. virescens; but not to the southern armyworm, Prodenia eridania; European corn borer, Ostrinia nubilalis; lobster roach, Nauphoeta cinerea; American cockroach, Periplaneta americana; greater wax moth, Galleria mellonella; and the housefly, Musca domestica.

Sikorowski and Lawrence (unpublished data) fed chicks (starting age 1- and 7-day-old) with CPV-infected H. virescens larvae for 14 days. Histological examination of various tissues of the chicks and bioassay of blood with 5-day-old H. virescens larvae were both virus-free. Temperatures at 35 °C and above inhibit replication of the virus (Tanada and Chang, 1968; Kobayashi and Kawase, 1980), even in insect hosts. The early stages of silkworm CPV infection, including RNA synthes is, are limited at these temperatures (Kobayashi and Kawase, 1980). Ignoffo (1968) reported that the use of CPVs against lepidopterous larvae does not appear to harm plants or beneficial arthropods (parasites and predators).

Sunlight is one of the major factors affecting the stability of viruses. Persistence of Heliothis CPV in field conditions was evaluated by Ali and Sikorowski (1986). On cotton foliage, 8 hours of exposure to sunlight at 28 - 32 °C and 400 langley (a unit for measuring solar radiation equal to one calorie per cm2) reduced CPV infectivity by 93%.

Microbial Control

Viral disease of insects provides a large reservoir of potential control agents. Cytoplasmic polyhedrosis viruses are among the most prevalent of insect viruses for which potential as biological control agents has not yet been evaluated in the United States. Insect control by CPV was reviewed by Katagiri (1981).

Rearing CPV-free Heliothines

Laboratory colonies of Heliothis spp. are frequently contaminated with CPV without any apparent evidence of the disease (Sikorowski et al., 1971b). This may occur when a few field-collected diseased insects are introduced into an established healthy colony. Eventually, more insects in the colony become infected and the moths may show reduced reproduction rates.

Sikorowski et al. (1971a) described a simple method designed for detection of CPV in H. virescens larvae. Summary of method: The larvae are starved for 12-24 hours, killed in 100% ethyl alcohol, removed from the alcohol, and allowed to air dry. Using CPV-free scissors, each larva is divided into three parts. Only that portion of the larva between the last pair of thoracic legs and the first pair of prolegs is used for smear preparation. The midgut is removed from this portion of the larv a. Smears are prepared with the midgut. A good thin smear has a single layer of polyhedra. Smears are air-dried for 1-2 hours at room temperature. The following staining solution and procedure was developed by Sikorowski et al. (1971a):

Staining Solution

a. Buffalo Black NBR (Allied Chemical)

0.1 g

b. 100% Methyl Alcohol

50.0 ml

c. Distilled Water

20.0 ml

d. Glacial Acetic Acid

30.0 ml

The staining solution should be freshly prepared and filtered before use.


Staining Procedure
a. Place the smear on a slide warmer heated to 40 °C and cover it with staining solution for 5 minutes.
   Caution: Do not allow the staining solution to dry on the slide during this period.
b. Remove the slide from the warmer, stand it on end, and allow it to drain and air dry.
c. Dip it or wash gently in tap water for 5 seconds.
d. Dry and examine without cover glass, using oil immersion objective (1,000X).

A method for selection of healthy individuals from a partially infected colony has also been developed (Sikorowski et al., 1971a): The meconium (the substance excreted by lepidopterous insects soon after emergence from pupa) of CPV-infected adults contains a large number of polyhedra mixed with urate crystals. The previously described method may be used to differentiate polyhedra from urate crystals in meconium because the urate crystals dissolve in the staining solution (procedure given by Sikorowski et al., 1971a). Pupae selected for this test are placed separately in 20-ml clear plastic cups that are closed with paper lids. Meconium discharged by newly emerged moths is collected from the cup wall with a moist flat toothpick, smeared on a clear glass slide, and stained by the method given in Staining Procedure. Both of these methods stained polyhedra navy blue, while the background was light blue.

Rearing CPV-free H. virescens is essential for production of vigorous insects for laboratory and field studies.


Figure 7. H. zea infected with iridescent virus (HIV).
H. zea larva infected with HIV showing iridescent lavender-blue coloration of entire larva.
Small iridescent patches frequently are the only signs of infection.
Transmission electron micrograph of decomposed infected fat body.
High magnification of HIV virions, bar = 120 nm.

Iridescent viruses (IVs) are double-stranded DNA viruses (virions measure 120 to 140 nm in diameter) that are pathogenic to a wide variety of insects and are found in the cytoplasm of infected cells. Most members of this family infect invertebrates, but some are found in vertebrates (Hall, 1985; Williams, 1996). There are five genera in this family with two of them infecting insects: Iridovirus and Chloriridovirus. The remaining three genera occur in vertebrates.

The first insect iridescent virus was reported in the crane fly, Tipula paludosa (TIV) (Xeros, 1954). Subsequently, another IV was detected in a lepidopteran (the rice stem borer, Chilo suppressalis ) by Fukaya and Nasu (1966). A Heliothis iridescent virus (HIV) was isolated from an insectary stock of H. armigera by Carey et al. (1978) and from field collected H. zea by Stadelbacher et al. (1978).

Iridescent viruses produce systemic infections, with the highest concentrations in the epidermis and the fat bodies. The loss of cell differentiation and cellular organization was found in the fat bodies of HIV-infected H. zea (Thompson and Sikorowski, 1981). In many insects, the virus is thought to be transmitted through cannibalism as shown in IVs of T. paludosa (Carter, 1973), Scapteriscus acletus (Boucias et al., 1987), and Aedes taeniorhynchus (Linley and Nielson, 1968). Oral transmission also has been shown in H. zea (Sikorowski and Tyson, 1984).

In many previous studies of oral infection with iridescent viruses, the criterion for infection was the appearance of iridescent coloration. This criterion was used in studies with Galleria mellonella (Yule and Lee, 1973), A. taeniorhynchus (Linley and Nielson, 1968), H. zea (Sikorowski and Tyson, 1984), Aedes sollicitans (Becnel and Fukuda, 1989) and Simulium vittatum (Erlandson and Mason, 1990). In these studies, only infections with virus at high enough concentrations to produce iridescence coloration were detected. It is likely that infections with lower concentrations of IV may have escaped detection, since they would not show iridescent coloration. Houston (1991) found asymptomatic HIV infections in pupae and adults of H. zea.

Gross Pathology

Sikorowski and Tyson (1984) reported that H. zea larvae infected with HIV developed iridescent lavender-blue, blue, and blue-green coloration. In per os transmission, the first signs of infection were small iridescent patches in the prolegs, clypeus, labrum, and intersegmental membranes. In many instances, only a few such patches were found per larva. Seldom was the entire body iridescent colored, even after prolonged incubation. In older infections, the iridescent color was generally more vivid, and frequently there were also small, extremely brilliant islands, usually less than 1 mm2. Electron microscope examination of fat bodies, silk glands, and body-wall muscles of larvae infected per os or by injection confirmed the presence of numerous intracytoplasmic virions that frequently occurred in paracrystalline arrays. No virus particles were seen in the nuclei.

Infection of larvae by hemocoelic injection. Inoculation of IV into the hemocoel was uniformly successful and produced almost 100% infection.

Per os infection of neonate larvae.

Larvae from IV-contaminated eggs (more than 400 larvae examined) produced 29% infection. None of the controls (more than 200 larvae examined) were infected. Larvae with a patent infection failed to reach the adult stage. Only a few reached the pupal stage, and almost all of the HIV-infected pupae were deformed.

Per os infection of 5- and 9-day-old larvae. Both groups of larvae fed HIV-contaminated foliage had a high percent of infection. Although the 5-day-old larvae appeared to be more susceptible to virus infection, the 9-day-old larvae fed HIV only 4 days before they would normally pupate showed up to 30% infection.

Longevity experiment. About 90% of the controls (217 used) pupated in 12 to 14 days. From the HIV-infected larvae (225 used), only a few deformed puape were produced; more than 95% died in the larval stage. A few infected insects remained in the larval stage for up to 89 days, when the experiment was terminated.

Inapparent Infection

Houston (1991) reported that a number of H. zea pupae and adults that appeared normal were found to be inapparently infected with HIV. She found that there were no significant differences between the percentage of insects with inapparent infections from the different ages of larvae and dilutions of viral inoculum. However, in many instances, the percent of insects with inapparent infection was higher than or equal to those with apparent infection. Low titers of the virus allow more hidden infectio ns to occur because the infections are at low enough levels not to overwhelm the insect and cause death.

Host Range and Geographical Distribution

Smith et al. (1961) listed 7 species of Diptera, 11 species of Lepidoptera, and 3 species of Coleoptera experimentally infected with TIV. Helicoverpa IV was isolated from H. armigera by Carey et al. (1978) and H. zea by Stadebacher et al. (1978).

Microbial Control

Studies by Houston (1991) showed that HIV may have a far higher infectivity than previously assumed, because challenging heliothines with HIV inocula lead to few patent infections yet frequent covert infections. Thus several factors such as infectivity, host range, fecundity of infected females, and longevity of infected adults should be reevaluated before HIV applicability as a biocontrol agent can be determined.


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