Technical Bulletin 218 - Protozoan Diseases Microspora Nosema heliothidis Signs and Symptoms Life Cycle Penetration Infection Effect of N. heliothidis on Helicoverpa zea Predators and Parasites Geographic Distribution Rearing Nosema-free Heliothines and Selection of Feral Insects for Laboratory Colony Vairimorpha necatrix Infection Spore Production Microbial Control Production of Microsporidia Spores Laboratory Insects Tissue Culture Other Protozoa Class Neogregarinida   Mattesia grandis   Ophryocystis sp References The word "Protozoa" means first animal, primitive animal. The protozoa are small, unicellular microorganisms. Protozoa are eukaryonts (organisms with a nucleus containing chromosomes encircled by a nuclear membrane), with numerous organelles whose functions are similar to the more complex, multicellular organs of higher animals. The structure of a nucleus with a single set of chromosomes with one genome is called haploid; those with a double complement are called diploid; and those possessing several genomes, polyploid. One cell can contain one or more nuclei. Protozoa reproduce both by sexual and asexual means, but some groups lack the ability for sexual or asexual reproduction. In some species, autogamy (self fertilization) takes place. Pasteur (1870) contributed to the establishment of insect pathology as a science through his classical study of the protozoan infection, pebrine, of the silkworm, Bombyx mori. The majority of entomogenous protozoa-insect associations produce chronic, nonlethal infections of which a common feature is a reduction in the host reproductive output (Hurd, 1993). Most infections of protozoa exhibit nonspecific signs and symptoms of disease such as sluggishness, irregular growth, loss of appetite, malformed larvae, pupae, or adults, or adults with reduced vigor, fecundity, and longevity. Protozoa offer little potential as short-term, quick-acting microbial insecticides, but they are being considered as candidates for long-term application or introduction programs (Canning, 1982). Entomogenous protozoa are commonly transmitted on the surface of the egg (transovum) or within the egg (transovarian) of their host. In the 1980 report of the Committee on Systematics and Evolution of the Society of Protozoologists, the protozoa are treated as a subkingdom, and seven phyla are recognized, five of which contain entomogenous species. Most members of protozoa found so far in heliothine larvae in Mississippi belong to the Phylum Microspora, Class Microsporea, Order Microsporida, and Genus Nosema. For more information on taxonomy of entomogenous protozoa, see the comprehensive treatment of this group of microbes by Brooks (1988) and Tanada and Kaya (1993). Microspora Members of this group are commonly called microsporidia. Of the entomogenous protozoa, the members of microsporidia appear to have the best potential for use in insect control, although they generally induce mostly chronic infections and are difficult to mass produce (McLaughlin, 1971). However, current use and improvement of artificial diets, improvement in mass-rearing technologies, and an availability of trained personnel have made production of required numbers of Heliothis virescens or Heli coverpa (=Heliothis) zea larvae for production of N. heliothidis spores for large-scale experiments feasible. The microsporidia are known for their resistant spores, which survive well in the environment. Nosema heliothidis Figure 1. Spores of N. heliothidis from H. zea (Courtesy of Dr. Wayne M. Brooks.) Nosema heliothidis was first described by Lutz and Splendore (1904) in Brazil from H. zea and was later found associated with both H. zea and H. virescens by Kramer (1959). Kramer also redescribed the life cycle of this protozoan. Lipa (1968) reported that N. heliothidis attacks the midgut epithelium, trachea, and gonads of H. zea. Brooks (1968) presented data to show that N. heliothidis is transmitted transovarially in H. zea and that both the male and female can transmit the protozoan to the next generation. For example, under laboratory conditions, eggs produced by diseased H. zea (240 examined) were 100% infected with N. heliothidis. Lipa (1968) reported that under laboratory conditions a high percentage of diseased larvae failed to pupate or produced abnormal pupae. Results of preliminary field tests conducted by Sikorowski (unpublished) in 1/16-acre cages showed a significant decrease in the number of larvae produced by adult H. zea infected with N. heliothidis. In Mississippi, we have isolated N. heliothidis from both H. zea and H. virescens field collected larvae. In general, we found N. heliothidis more frequently in H. zea than in H. virescens larvae. Our studies (in the laboratory environment) showed that the average longevity of 20 pairs of healthy H. zea moths was 23.1 days (24.1 for male and 22.1 for female) and that for 20 pairs of diseased moths was 14.4 days (15.7 for male and 13.2 for female). Thompson and Sikorowski (1979) reported that infected H. zea larvae accumulated fatty acids more rapidly than healthy larvae in the first 10 days, but then they decreased rapidly until death. Experimental infections have been obtained in H. zea and H. virescens with several other species of microsporidia (Brooks, 1988). Signs and Symptoms Infected larvae of H. zea and H. virescens differ little, if any, from healthy larvae. Diseased larvae are not as active and lose their appetite. Infected larvae may fail to pupate, and some pupae and adults may be deformed. Thus, proper diagnosis (the final decision) requires microscopic examination of susceptible tissues such as fat bodies, midgut epithelium, etc. Life Cycle The Nosema life cycle can be defined by two events: 1)mergony, the vegetative phase, and 2)sporogony, the production of spores. During merogony, the microsporidium multiplies rapidly by primary fission, plasmotomy (fragmentation of a multinucleate cell or multiple fusion), or multiple budding. From binucleate meronts, sporonts are formed. Nuclear division within the sporont gives rise to a tetranucleate sporont, which divides to produce two sporoblasts (prespore stage), each of which develops into a spore. Spores are ovoid, with one pole more pointed than the other. The fresh spores of N. heliothids measured in water were 3.3 to 6.0 µm wide (Lipa, 1968). Penetration Nosema heliothidis can enter the host by three ways: 1)per os, 2)ovarial (transovum and transovarial), and 3)cuticular. Entrance by the oral and cuticular means results in horizontal transmission. Oral entrances include feeding of the heliothine larvae on spore-contaminated food or feeding on dead or moribund insects. Entrance by the ovarial portal (vertical transmission) is accomplished either by contamination of the surface of the host eggs and infecting emerging larvae during eating of the contaminated egg chorion (shell) or by an ovarian transmission in which N. heliothidis meronts, sporonts, and/or spores are integrated into the egg or embryo within the female's reproductive tract. Offspring from such females are also infected (Brooks, 1988). Helicoverpa zea-infected females may produce 100% infected offspring (Sikorowski, unpublished). Entrance by the cuticular portal is accomplished by inoculation of spores by the ovipositor of a N. heliothisdis-contaminated parasitoid ( Brooks, 1973). Infection The infective stage of N. heliothidis is the spore. After ingestion by heliothine larvae, spores of N. heliothidis release a sporoplasm by injection through an everted polar tube. For detailed discussion on polar filament function, extrusion, and extrusion apparatus see Brooks (1988) and Tanada and Kaya (1993). Only a short discussion of this topic is given in this publication. Microsporidian spores are among the smallest and most complicated of eukaryotic cells. The extrusion apparatus places the sporoplasm into a host cell. The sporoplasm is the infective unit of microsporidia. Once ingested, the spore is stimulated to evert its polar tube, which then serves as an injection needle through which the sporoplasm is injected. Under pressure, the rigid tube penetrates the peritrophic membrane and epithelial gut cells and deposits the sporoplasm directly into susceptible host cells. Afterwards, the sporoplasm eventually undergoes merogony and sporogony to produce new spores (Brooks, 1988). Effect of N. heliothidis on H. zea Predators and Parasites Nosema infected H. zea were fed to mymphs of Nabis roseipennis (84 insects used), N. capsiformis (72 insects used), N. sordidus (27 insects used), and N. alternatus (3 insects used), and Chrysopa sp. larvae (100 insects used). At maturity, the nymphs were crushed in drops of water on glass slides and then examined with a phase interference compound microscope to determine the presence or absence of Nosema spores. However, none of the nymphs were infected with Nosema heliothidis. (Sikorowski, unpublished), although Brooks and Cranford (1972) documented the susceptibility of the braconid Campoletis sonorensis to N. heliothidis. Geographic Distribution Nosema heliothidis occurs regularly in corn field populations of H. zea throughout much of the eastern United States, including Mississippi. During a 1979 North Carolina survey, H. zea adults were collected in light traps at weekly intervals from mid-May to late October. Prevalence of the disease ranged from 10 to 80% infection in these adults and averaged about 30% (Brooks and Cranford, 1978). Larvae collected from the same location during 1971 were 84.9% infected (n=106) when collected from corn, 62.3% (n=61) when collected from tobacco, and 52.8% (n=53) when collected from soybeans. Our studies showed that almost 100% of eggs produced by infected females are infected (internally) or contaminated (externally) with N. heliothidis . Thus, neonate larvae are infected either before they emerge from the eggs or by eating Nosema-contaminated chorion during emergence. Thus, offspring of infected females of earworm can aid in spread of Nosema throughout the earworm population in a particular ecosystem. Although N. heliothidis produces chronic, frequently nonlethal infections, it is likely to have significant effects on the dynamics of H. zea (Gaugler and Brooks, 1975). Rearing Nosema-free Heliothines and Selection of Feral Insects for Laboratory Colony Nosema heliothidis is a frequent problem in insectary colonies of H. zea, H. virescens, and Spodoptera exigua in Mississippi. This is caused by the incorporation of field-collected insects into laboratory colonies preceding quarantine and examination for such pathogens as protozoa, viruses, fungi, and bacteria. Nosema heliothidis-infected heliothine rearing populations usually crash after several months when eggs from heavily infected heliothine breeding colonies are used. In our laboratory, we rear the first generation of field-collected adults in individual 32-ml plastic diet cups with flash-sterilized wheat-germ-soy flour diet (Shaver and Raulston, 1971). Upon pupation, all pupae are disinfected in 0.5% sodium hypochlorite (Sikorowski and Goodwin, 1985) and placed individually in empty diet cups. The meconium, the substance excreted by adults soon after emergence from pupae, is examined microscopically using 450 to 950X magnification to determine the presence or absence of N. heliothidis spores. This is a very simple method, and meconium smears may be examined unstained (in water) with a phase-contrast microscope or stained (using a stain such as Giemsa) with a simple light microscope. Two experienced technicians, one preparing smears and the other examining the smears with a compound microscope (at 450X), can check up to 50 smears per hour. Nosema-free adults are generally quarantined for another generation before they are incorporated into the breeding colony. Vairimorpha necatrix Figure 2. Vairimorpha necatrix infected H. virescens. H. virescens larva infected with V. necatrix showing white fat body visible through the integument. Light microscope photographs of sections of fat body showing spores and a total loss of cell differentiation. Light microscope of V. necatrix spores (Courtesy of Dr. Wayne M. Brooks)) Vairimorpha necatrix, a microsporidium with two distinctly different spore forms, was first reported from Pseudaletia unipuncta larvae from Hawaii (Tanada and Chang, 1962). One form has spore sizes (length = L and width = W) that are highly dependent upon temperature (for example at 32 °C L=2.4 to 5.2 µm and W=2.2 µm, but at 15 °C L=4.2 to 6.9 µm and W=2.4 µm). The other spores are smaller and less variable (ranging from 2.82 to 3.98 µm in length by 1 .74 to 2.40 µm in width). This protozoan is infective for more than 20 noctuids (Maddox, 1966), including H. zea and H. virescens. All known hosts are species of Lepidoptera, and many are major pests of agricultural crops (Brooks, 1988). Infection The insect fat bodies are the primary infection site. In advanced stages of disease, the fat body cells enlarge many times, giving the fat tissue a lobated appearance. The lobated fat tissues are filled with microsporidian spores. The abnormally large, white fat body is usually obvious through the integument. The degree of infection depends on spore dosage, temperature, and larval age. Feeding of the insect is usually normal at first, decreasing to very little for the last few days before death. Vairimorpha is transmitted to the next generation both on and in the eggs of infected adults. Vairimorpha necatrix causes two different types of diseases resulting in mortality in its hosts, death that results from gut damage followed by bacterial septicemia, and death that results from microsporidiosis after ingestion of even a light spore dose (Maddox, 1966). A low dosage of V. necatrix results in a chronic infection of mainly the fat bodies and some muscular tissues, whereas high dosages produce an acute infection of mainly midgut tissues (Chu and Jaques, 1979). In the advanced stage of infection, the abnormally large, white fat body is usually conspicuous, and a dorsal swelling may appear on the last two or three abdominal segments (Pilley, 1976). Spore Production Vairimorpha necatrix develops only in living cells. Many species of noctuid larvae, in particular H. zea or T. ni, are suitable hosts for spore production. Spore production per infected larva is about 2 x 1010 spores/g of host larva. Mass production and storage of V. necatrix spores were evaluated by Fuxa and Brooks (1978). Microbial Control Considerable progress has been made in recent years to evaluate the potential usefulness of protozoa as microbial control agents. The present status of protozoa as biological control agents can be summarized as follows: production, quality control, safety to vertebrates and beneficial insects, and large acreage field tests are all features that need further study before protozoa in general and Nosema and Vairimorpha in particular can be made more available for use as part of a new generation of microbial insecticides. However, both groups of protozoa have many features of a good potential biocontrol agent. Maddox et al. (1981) reported that V. necatrix has many such characteristics: for example, a high degree of virulence and a wide host range. Spore production in its hosts (usually H. zea or H. virescens) is straight-forward, inexpensive, and safe to man and the environment. Nosema heliothidis, within a few days of larval infection, becomes a continuous source of fresh spores excreted in the feces for other larvae. Thus, infected adults and larvae can serve as a source and as disseminators of spores until death. Field tests with V. necatrix showed that high infection rates in pest populations can be obtained with appropriate spore formulations. However, the use of this species as a short-term microbial control agent on a large scale would not be practical. Production of Microsporidian Spores Microsporidia are obligate pathogens and must be produced in laboratory-reared insects or in cell-tissue culture. Laboratory Insects Generally, neonatal or early stage larvae are exposed to a low dose of spores placed on artificial diet or on a suitable food substrate. Several days later, depending on the rearing temperature, stage of insects, and diet, mature spores are harvested from insects by various methods as described by Brooks (1988). The habitual host(s) or permissive hosts, such as H. zea or H. virescens larvae, that can support production of spores of various microsporidia are frequently used for spore production. Tissue Culture Some species of microsporidia have been grown in tissue culture, but the high cost of culture media, the low yields in spores, and the imperfect production techniques for mass production still limit the usefulness of this method at this present time (Brooks, 1988). In Mississippi, we have been able to produce N. heliothidis and V. necatrix spores in H. zea larvae for large-scale field tests. Other Protozoa Class neogregarinida The gregarines of the order Neogregarindia are known as neogregarines. A number of entomopathogenic neogregarines produce lethal infections in important insect pests in the orders Diptera, Coleoptera, and Hemiptera. Mattesia Grandis. Mattesia grandis, a neogregarine of the cotton boll weevil (Anthonomus grandis), was experimentally transmitted to H. zea and H. virescens by Ignoffo and Garcia (1965). The symptomology of this disease in the boll weevil is associated with a progressive destruction of the fat bodies and the reduction in the oviposition of eggs (McLaughlin, 1965). Ophryocystis Sp. We observed the presence of this neogregarine in heliothine pupae that originated from an insectary-reared population (received in our laboratory for diagnosis). Large amber-colored spores were occasionally found in the dark areas under translucent heliothine pupal cases. The spores can be seen on the scales of the moths under a dissecting microscope. Or, if the moths are allowed to emerge in clean plastic cups, the spores can be seen easily with a hand lens on the sides of the cups. McLaughlin and Myers (1970) described for the first time O. elektroschirrha from naturally occurring populations of the monarch butterfly (Danaus plexippus) and the Florida queen butterfly D. glippus berenice. They reported that O. elektroscirrha infected the hypodermal tissue, that it remains in micronuclear schizogony until after pupation of the host, and then completes morphogenesis in the tissue that becomes the scales of the adult butterfly. The adult thus carries the spores externally. References Brooks, W. M. 1968. Transovarian transmission of Nosema heliothidis in the corn earworm, Heliothis zea. J. Invertebr. Pathol. 11: 510-512. Brooks, W. M. 1973. Protozoa: Host-parasite-pathogen interrelationships. Misc. Publ. Entomol. Soc. Am. 9: 105-111. Brooks, W. M. 1988. Entomogenous protozoa, pp. 1-149. In C. M. Ignoffo and N. B. Mandava (eds.) Handbook of natural pesticides, Vol V, microbial pesticides, part A. CRC Press, Inc., Boca Raton, FL. Brooks, W. M., and J. D. Cranford. 1972. Microsporidoses of the hymenopterous parasites, Campoletis sonorensis and Cardiochils nigriceps, larval parasites of heliothis species. J. Invertebr. Pathol. 20: 77-94. Brooks, W. M., and J. D. Cranford. 1978. Host-parasite relationships of Nosema heliothidis Lutz and splendor. Misc. Publ. Entomol. Soc. Am. 11: 51. Canning, E. U. 1982. An evaluation of protozoal characteristics in relation to biological control of pests. Parasitology 84: 119-149. Chu, W. H., and R. P. Jaques. 1979. Pathologie d'une microsporidiose de l'arpenteuse du chou, Trichoplusia ni [Lep.: Noctuidae], par Vairimorpha necatrix. Entomophaga 24: 229-235. Fuxa, J. R., and W. M. Brooks. 1979. Mass production and storage of Vairimorpha necatrix (Protozoa: Microsporida). J. Invertebr. Pathol. 33:86-94. Gaugler, R.R., and W. M. Brooks. 1975. Sublethal effects of infection by Nosema heliothidis in the corn earworm, Heliothis zea J. Invertebr. Pathol. 26: 57-63. Hurd, H. 1993. Reproductive disturbances induced by parasites and pathogens of insects, pp. 87-93. In N. E. Beckage, S. N. Thompson, and B. A. Federici (eds.) Parasites and pathogens of insects, Vol. 1 parasites. Academic Press, Inc., San Diego, CA. Ignoffo, C. M., and C. Garcia. 1965. Infection of the cabbage looper, bollworm, tobacco budworm, and pink bollworm with spores of Mattesia grandis McLaughlin collected from boll weevils. J. Invertebr. Pathol. 7: 260-262. Kramer, J. P. 1959. Observations of the seasonal incidence of microsporidosis in European corn borer populations in Illinois. Entomophaga 4: 37-42. Lipa, J. J. 1968. Some observations on Nosema heliothidis Luts et Splendore, a microsporidian parasite of Heliothis zea (Boddie) (Lepidoptera, Noctuidae). Acta Protozool. Pol. 4: 237-278. Lutz A., and A. Splendore. 1904. Uber perbrine and verwandte Mikorsporidien. Nachtrag zur ersten Mitteilung. Zentr. Bakt. (I) Orig. 36: 645-650. Maddox, J. V. 1966 Studies on a Microsporidosis of the armyworm, Pseudaletia unipuncta (Haworth). Ph.D. thesis, University of Illinois, Urbana, IL. Maddox, J. V., W. M. Brooks, and J. R. Fuxa. 1981. Vairimorpha necatrix a pathogen of agricultural pests: potential for pest control, pp. 587-594. In H. D. Burges (ed.) Microbial control of pests and plant diseases 1970-1980. Academic Press, New York, NY. McLaughlin, R. E. 1965. Some relationships between the boll weevil, Anthonomus grandis Boheman, and Mattesia grandis McLaughlin (Protozoa: Neogregarinida). J. Invertebr. Pathol. 7: 464-473. McLaughlin, R. E., and J. Myers. 1970. Ophryocystis elektroscirrha sp. N., a neogregarine pathogen of the monarch butterfly Danaus plexippus (L.) and the Florida queen butterfly D. glippus berenice Cramer. J. Protozool. 17: 300-305 . McLaughlin, R. E. 1971. Use of protozoans for microbial control of insects, pp. 151-172. In H. d. Burges and N. W. Hussey (eds.) Microbial control of insects and mites. Academic Press, London, UK. Pasteur, L. 1870. Etudes sur la maladie des vers a soie, Tome I and II. Gauthier-villars, Paris, France. Pilley, B.M. 1976. A new genus, Vairimorpha (Protozoa: Microsporida) for Nosema necatrix Kramer 1965: pathogenicity and life cycle in Spodoptera exempta (Lepidoptera: Nocuidae). J. Invertebr. Pathol. 28: 177-183. Shaver, T. N., and J. R. Raulston. 1971. A soybean wheat-germ diet for rearing of the tobacco budworm. Ann. Entomol. Soc. Am. 64: 1077-1079. Sikorowski, P.P., and R. H. Goodwin. 1985. Contaminant control and disease recognition in laboratory colonies, pp. 85-105. In P. Singh and R.F. Moore (eds.), Handbook of insect rearing, vol. 1. Elsevier, Amsterdam, Netherlands. Tanada, Y., and G. Y. Chang. 1962. An epizootic resulting from a microsporidian and two virus infections in the armyworm, Pseudaletia unipuncta (Haworth). J. Invertebr. Pathol. 4: 128-131. Tanada, Y., and H. K. Kaya. 1993. Insect pathology. Academic Press, New York, NY. Thompson, A.C. and P.P. Sikorowski. 1979. Effects of Nosema heliothidis on fatty and amino acids in larvae and pupae of the bollworm Heliothis zea. Comp. Biochem. Physiol. 63A: 325-328.