Steinernema glaseriSurface Coat Protein Suppresses the Immune Response ofPopillia japonica(Coleoptera: Scarabaeidae) Larvae

Biological Control 14, 45–50 (1999) Article ID bcon.1998.0672, available online at http://www.idealibrary.com on

Steinernema glaseri Surface Coat Protein Suppresses the Immune Response of Popillia japonica (Coleoptera: Scarabaeidae) Larvae Yi Wang and Randy Gaugler Department of Entomology, Rutgers University, New Brunswick, New Jersey 08903-0231 Received April 21, 1998; accepted October 12, 1998

1990; Klein and Georgis, 1992; Selvan et al., 1994; Wang et al., 1995). The entomopathogenic nematode, Steinernema glaseri (Steiner), is naturally associated with scarab larvae (Glaser and Fox, 1930), and was originally isolated from P. japonica (Steiner, 1929). Defense mechanisms in P. japonica larvae have developed against nematode infection. The larvae can detect nematodes on its cuticle and initiate evasive and aggressive behaviors (Gaugler et al., 1994). Aggressive behaviors are effective in removing and even killing nematodes. Scarab larvae also have morphological defenses against nematode entry such as a thick peritrophic membrane and sieve plates over the spiracles that serve as barriers of nematode infection via the gut and cuticle, respectively (Forschler and Gardner, 1991). Chemical defenses within the gut are effective against other nematode species such as Heterorhabditis bacteriophora (Poinar) (Wang et al., 1995). One of the most important insect defense mechanisms against metazoan parasites is encapsulation. An envelope is formed by deposition of multiple layers of hemocytes and/or a melanin coat that encloses and kills the intruder. A counter mechanism for parasites to cope with the host immune system is suppression. Host humoral or cellular immune responses may be suppressed by viruses, venom, or other anti-immune factors associated with parasites (Go¨tz et al., 1981; Brown, 1986; Yokoo et al., 1992; Webb and Luckhart, 1994). Entomopathogenic nematodes release symbiotic bacteria in the host hemocoel to suppress the host immune system (Dunphy and Thurston, 1990). In S. glaseri, the virulence of its symbiotic bacterium, Xenorhabdus poinarii (Akhurst), to P. japonica larvae has been demonstrated by injection (Yeh and Alm, 1992; Wang unpublished data). However, the bacteria are released 4–6 h after nematode entry into the host hemolymph (Wang et al., 1995). To assure bacterial release, S. glaseri escapes or defeats encapsulation (Wang et al., 1994; 1995). The mechanism involved in this immunoprotection is unclear. Nematode surface coat proteins can be secreted from

The host immune response is a key obstacle to entomopathogenic nematodes in making the transition from the free-living state to parasitism. The entomopathogenic nematode Steinernema glaseri has evolved mechanisms to evade immune encapsulation in larvae of the Japanese beetle, Popillia japonica. Host intrahemocoelic injection tests show that live axenic nematodes of S. glaseri not only avoid host melanotic encapsulation but also protect dead nematodes injected after the live ones. This result indicates that the nematodes release anti-immune factor(s). We extracted the nematode surface coat proteins and found that at least one protein (SCP3a) from the S. glaseri surface coat can suppress the host immune system. This suppression protects unrelated nematode species from encapsulation and latex beads from phagocytosis. We conclude that S. glaseri uses an antiimmune protein to defeat the host immune system, thereby protecting itself from encapsulation. Presumably its symbiotic bacteria are similarly protected from phagocytosis. r 1999 Academic Press Key Words: entomopathogenic nematode; immune suppression; melanotic encapsulation; Japanese beetle; Steinernema glaseri; surface coat protein.

INTRODUCTION

Entomopathogenic nematodes from the families Steinernematidae and Heterorhabditidae are receiving considerable attention from biological control workers (Kaya and Gaugler, 1993). The nematode infective juveniles live in the soil where they search for insect hosts. After entering the host hemocoel via natural openings (gut, spiracles), the nematodes release symbiotic bacteria that kill their hosts within 48 h. The Japanese beetle, Popillia japonica (Newman), is an important turfgrass pest (Ahmad et al., 1983; Tashiro, 1987) and a major control target for entomopathogenic nematodes. Numerous field trials and laboratory tests have been performed with nematodes against scarab larvae (Villani and Wright, 1988; Klein, 45

1049-9644/99 $30.00 Copyright r 1999 by Academic Press All rights of reproduction in any form reserved.

46

WANG AND GAUGLER

the excretory pores, amphids, and phasmids (Premachandran et al., 1988; Politz and Philipp, 1992). In free-living nematodes, the surface coat plays a role in lubrication and protection against abrasion, dehydration, and predation (Blaxer et al., 1992). In animalparasitic nematodes, the surface coat has evolved functions such as interaction with host immune defenses (Cookson et al., 1992). We hypothesize that the S. glaseri surface coat protein(s) suppresses the host immune response at the early stage of infection against P. japonica. MATERIALS AND METHODS

Culture. Infective juveniles of S. glaseri (NC strain) and H. bacteriophora (HP88 strain) were reared in last-instar Galleria mellonella (L.) at 25°C (Dutky et al., 1964). Nematodes were harvested from White traps and used after 5–10 days storage in distilled water at room temperature. Axenic S. glaseri were cultured on beef liver extract containing streptomycin (100 µg/ml) and kanamycin (20 µg/ml) following the sterilization process described by Go¨tz et al. (1981). Axenic infective juveniles were harvested from the medium and rinsed in sterile water three times before experiments. Injection. To determine if S. glaseri releases antiimmune factor(s), we injected five axenic nematodes into the hemocoel of each P. japonica larva at the base of the first leg. At 0, 1, 2, 4, 8, and 16 h post-injection, we injected 10 additional freeze-killed axenic nematodes into the hemolymph of the same larva. Eight hours after the freeze-killed nematode injections, we dissected the larva and counted the encapsulated and melanized nematodes. Ten scarab larvae were used for each time interval. Crude surface coat extraction. To extract the nematode surface coat proteins, we used the procedure described by Page et al. (1992) with modifications. Briefly, 15 g of infective juveniles was desheathed using 0.5% sodium hypochlorite at room temperature for 15 min. The nematodes then were rinsed by centrifugation in distilled water three times, and stored in 2 liters of distilled water with bubbled air. The nematodes were allowed 2 days to redevelop their surface coat after desheathment. After three distilled water rinses, the nematodes were put in 35, 50, or 100% ethanol for 30 min. The resulting extracts were lyophilized and stored at ⫺20°C for later use. To test the activity of crude extract of nematode surface coat in suppressing the host immune system, we dissolved the nematode surface coat extracts in distilled water, and adjusted the protein concentration to 1 ng/µl using the technique developed by Bradford (1976). Twenty surface-sterilized (0.1% methylbenzethonium chloride for 5 min) infective juveniles of H. bacteriophora were injected into each larval hemocoel

with 5 µl (1 ng/µl) of each extract solution (from 35, 50, and 100% ethanol) following Wang et al. (1994). Twenty larvae were used for each extract. Ten larvae were dissected 8 h post-injection and examined for encapsulated nematodes. Ten larvae were dissected 72 h postinjection to determine nematode survival. In the control group, 5 µl (1 ng/µl) of bovine serum albumin (SIGMA) was used. Separation and assays of proteins from 35% ethanol extract. The most effective crude extract from the above test, the 35% ethanol extract, was run on an 8% nondenaturing polyacrylamide gel. The protein bands were cut separately after Coomassie brilliant blue R250 staining. The proteins were collected by electric elutor and dialyzed in distilled water at 4°C overnight. Proteins were quantified using the Bio-Rad technique (Bradford, 1976). We conducted two activity assays for the surface coat proteins (SCP): encapsulation protection and hemocyte cell count reduction. In the encapsulation protection test, the same procedure was used as described above, except instead of using crude extracts, we used 5 µl of each purified protein (1 ng/µl). In the hemocyte reduction test, we withdrew hemolymph into an anticoagulation saline (0.1 M dextrose, 0.03 M sodium citrate dihydrate, 0.026 M citric acid, 0.01 M disodium ethylenediamine tetracetic acid, and LPS-free 3% NaCl) developed by Gupta et al. (1991). The hemocytes were rinsed in the saline three times by centrifugation (1000g, 5 min, 4°C) before being suspended at a concentration of 80,000 ⫾ 3,500 hemocytes/µl. Five microliters of each SCP (1 ng/µl) solution was added to 400 µl of the hemocyte solution in each test tube. The hemocytes were counted 4 h post-SCP inoculation (room temperature) with a hemocytometer. Each protein isolate was tested three times. Activity tests of SCP3a. From the above assays, we found that the surface coat protein 3a (SCP3a) was most effective in protection against encapsulation and hemocyte lysis. We conducted three experiments to test SCP3a in suppression of immune responses. (1) To test the protein’s effect on phagocytosis, fluorescent-conjugated latex beads (1 M) with 5 µl of SCP3a (1 ng/µl) were injected into the hemocoel of each P. japonica larva. Hemolymph was withdrawn into the anticoagulant saline at 4, 5, 6, and 7 h post-injection by cutting forelegs. Phagocytosed and free latex beads were counted under a fluorescent microscope, and the percentage of phagocytosed beads was calculated. Six larvae were used for each time interval. In the control group, water instead of SCP3a solution was used. The symbiotic bacterium, X. poinarii, was not used in this experiment because we wanted to eliminate any effect of the bacterial surface components. (2) To test SCP3a ability to lyse hemocytes in vivo, we conducted total hemocyte counts at 0.5, 1, 2, 4, 8, and 20 h post-

HOST IMMUNE SUPPRESSION BY S. glaseri

injection of SCP3a. The withdrawn hemolymph was diluted (1:100) into anti-coagulant saline before counting. Six larvae were used for each time interval. Distilled water was used for control injections. (3) In insects, phenoloxidase is an important factor for nonself recognition and melanin formation. We used the procedure described by Dunphy and Webster (1991) to test the SCP3a inhibition of the phenoloxidase activity and its conversion from prophenoloxidase. Briefly, hemolymph was withdrawn into distilled water (45:200 µl) by cutting the forelegs of P. japonica larvae at 2 and 4 h post-injection of 5 µl SCP3a (1 ng/µl). The hemolymph was pipetted vigorously for 5 min, and debris spun down (10,000g, 5 min, at 4°C). The supernatant was incubated with 20 µl Laminarine (1 mg/ml, SIGMA) for 1 h at room temperature to activate phenoloxidase. Phenoloxidase was determined spectrophotometrically by adding 500 µl L-dihydroxyphenylalamine (1.6 mg/ ml) and measuring melanization at 490 nm. To test for SCP3a inhibition of phenoloxidase activity, hemolymph was withdrawn from healthy larvae and pipetted as described above. The supernatant was incubated with 5 µl SCP3a for 2 h after Laminarine activation. Phenoloxidase activity was then measured. Statistical analyses. Distribution(s) of percentage data was normalized using the arcsine square-root transformation prior to analysis. Analysis of variance and Tukey’s multiple range test (␣ ⫽ 0.05) were used for mean comparisons. Data in the text are presented as mean ⫾ standard error of the mean.

47

FIG. 2. (A) Encapsulation of Heterorhabditis bacteriophora 8 h post-injection with surface coat extract from Steinernema glaseri. (B) Survival of H. bacteriophora 72 h post-injection with surface coat extract from S. glaseri. Extract I, 35% ethanol extract; extract II, 50% ethanol extract; extract III, 100% ethanol extract.

RESULTS

Live axenic nematodes of S. glaseri protect freezekilled nematodes from encapsulation by suppressing the host immune system (Fig. 1). Virtually all freezekilled nematodes coinjected (0 h) with live nematodes were encapsulated. However, encapsulation of freezekilled nematodes dropped sharply (30.3 ⫾ 6.2%) if the

FIG. 1. Encapsulation of freeze-killed Steinernema glaseri injected at different times after axenic nematode injection into Popillia japonica larvae.

nematodes were injected 4 h after axenic nematode injection. No axenic nematodes were encapsulated. This indicates that initially, nematodes can impair the host immune response only locally. Whole system suppression requires 4 to 8 h. The crude extract of nematode surface coat from 35% ethanol protected H. bacteriophora from host melanotic encapsulation (Fig. 2A) and thereby significantly enhanced nematode survival in the host hemocoel (Fig. 2B). In the control group, 97.0 ⫾ 2.5% of injected nematodes were encapsulated, whereas only 15.4 ⫾ 3.1% were encapsulated when co-injected with the extract from 35% ethanol. The survival of nematodes co-injected with the 35% ethanol extract increased by four times compared with controls (60.3 ⫾ 5.1% vs 16.1 ⫾ 2.3%). Clearly, enhanced survival can be attributed to the suppression of the host immune system by S. glaseri surface extract. The surface coat proteins were separated on nondenaturing PAGE, and five protein bands were detected (Fig. 3). When the hemocytic lysing activity of each protein was tested in vitro by exposure to larval hemocytes, two proteins (SCP3a or SCP3b) significantly reduced the hemocytes by 16.8–18.2% (Fig. 4A). Sur-

48

WANG AND GAUGLER

FIG. 3. Surface coat proteins in 35% ethanol extract separated on 8% nondenaturing polyacrylmade gel (lane A), and the protein extracted from homogenized nematodes with 35% ethanol (lane B).

face coat protein SCP3a also significantly reduced melanotic encapsulation of H. bacteriophora (Fig. 4B). In the control group, 95.2 ⫾ 4.1% of injected nematodes were encapsulated, whereas only 18.5 ⫾ 5.2% were encapsulated if the H. bacteriophora nematodes were co-injected with SCP3a. No significant reduction of encapsulation was found for any other protein bands, indicating that the SCP3a is most responsible for suppressing the host immune system.

FIG. 5. Latex beads phagocytosed in Popillia japonica larvae at different times post-injection with 5 µl of SCP3a.

SCP3a reduced hemocyte phagocytosis to latex beads in vivo (Fig. 5). In controls, 55.2 ⫾ 9.6% latex beads were phagocytized by hemocytes in the host hemocoel. However, phagocytosis was reduced by 10 times (5.3 ⫾ 3.5%) if the beads were co-injected with SCP3a. The injection of SCP3a caused hemocyte reduction compared with bovine protein (Fig. 6). In both treatment and control groups, a decline of hemocytes was observed 30 min post-injection. In controls, hemocyte numbers recovered to normal by 4 to 8 h post-injection. In SCP3a injections, however, complete recovery was not observed until 20 h post-injection. At each time interval, SCP3a injection caused a 20 to 30% hemocyte reduction compared with controls. Neither the conversion of phenoloxidase from prophenoloxidase nor the activity of phenoloxidase was inhibited by SCP3a (data not shown). DISCUSSION

The nematode–bacteria complex of S. glaseri uses anti-immune proteins to defeat the host immune re-

FIG. 4. (A) Hemocyte reduction following 4-h exposure to different surface coat proteins isolated from 35% ethanol extract. (B) Encapsulation of 20 Heterorhabditis bacteriophora 8 h post-injection with different surface coat proteins. *Indicates significantly different from controls (Tukey’s multiple range test, P ⬍ 0.05).

FIG. 6. Total hemocyte count (number of hemocytes per microliter hemolymph) of Popillia japonica larvae at different times postinjection with 5 µl of SCP3a.

49

HOST IMMUNE SUPPRESSION BY S. glaseri

sponse in P. japonica larvae. This process is an important component maintaining the mutual relationship between the parasitic nematode and its symbiotic bacterium X. poinarii. To kill the host, the nematode first must release its symbiont. To assure bacterial release, the nematode has to cope with the host immune encapsulation. This immune suppression may also provide a safe environment for the bacteria. In return, the bacterium kills the host and provides essential nutrients. Without its nematode partner, the symbiotic bacterium cannot be delivered into host hemocoel. The nematodes may also protect the bacteria from phagocytosis or host humoral factors. Similar protection of symbiotic bacteria X. nematophilus (Poinar and Thomas) by the nematode Steinernema carpocapsae (Weiser) has been observed (Go¨tz et al., 1981). Encapsulation provides insects with defense against many invading microorganisms. However, it is ineffective against certain species which have evolved strategies to avoid or inactivate the host immune system. One strategy is to secrete anti-immune factors into the host hemocoel. The action varies according to different parasites or pathogens. Destruction of hemocytes impairs the host immune system and causes hemocytopenia. This action has been found in bacterial pathogens (Ratcliffe and Walters, 1983; Dunphy and Webster, 1991) as well as some parasitoid wasps (Rizki and Rizki, 1984). Interaction with immune active molecules (i.e., cecropins and opsonins) reduces host ability to attack the surface of parasites or pathogens (Bloom, 1979; Go¨tz et al., 1981; Jarosz and Stefaniak, 1993). Inhibition of phenoloxidase activity can suppress insect immune encapsulation (Stoltz and Cook, 1983; Yokoo et al., 1992) since phenoloxidase activity plays an important role in foreign body recognition and melanin formation in many insect species (Gupta, 1991; Gillespie et al., 1997). The action mode of SCP3a has not yet been investigated except insofar as it has not been found to impact phenoloxidase activity. Note that the SCP3a concentration we used for our experiments could be higher than the amount which is naturally secreted. However, freshly secreted protein in natural conditions should be more active than extracted protein (Burman, 1982). Because of the complexity of the insect immune system and the diversity of mechanisms involved, parasite–host interaction patterns may vary according to different nematode or host species. The immune suppression mediated by S. glaseri may only apply to P. japonica larvae or ecologically related insect species. Injection of S. glaseri infective juveniles into the hemocoel of the cricket Acheta domesticus (L.) induces strong nematode encapsulation and low host mortality (Wang et al., 1994), indicating that A. domesticus possesses a different immune system, one which is not affected by this nematode’s surface coat protein.

In summary, our data show that the surface coat protein SCP3a from S. glaseri protects nematodes from melanotic encapsulation, protects latex beads from phagocytosis, and destroys host hemocytes. We conclude that this protein plays an important role, at least at the early infection stage, in the survival and establishment of the nematode and its symbiotic bacteria. Studies of the protein’s structure and its action mode in the host will elucidate a molecular understanding of interaction mechanisms between parasite and host. ACKNOWLEDGMENTS Support from the Rutgers Center for Turfgrass Science is gratefully acknowledged. This is New Jersey Agricultural Experiment Station Publication No. D-08256-07-98 and was supported by state funds and Regional Research Funds.

REFERENCES Ahmad, S., Streu, H. T., and Vasvary, L. M. 1983. The Japanese beetle: A major pest of turfgrass. Am. Lawn Appl. 4, 2–10. Blaxer, M. L., Page, A. P., Rudin, W., and Maizels, R. M. 1992. Nematode surface coats: Actively evading immunity. Parasitol. Today 8, 243–247. Bloom, B. R. 1979. Games parasites play: How parasites evade immune surveillance. Nature 279, 21–26. Bradford, M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of proteindye binding. Anal. Biochem. 72, 248–254. Brown, F. 1986. The classification and nomenclature of viruses: Summary of results of meetings of the International Committee on Taxonomy of Viruses in Sendai, September, 1984. Intervirology 25, 141. Burman, M. 1982. Neoaplectana carpocapsae: Toxin production by axenic insect parasitic nematodes. Nematologica 28, 62–70. Cookson, E., Blaxter, M. L., and Selkirk, M. E. 1992. Identification of the major soluble cuticular glycoprotein of lymphatic filarial nematode parasites (gp29) as a secretory homolog of glutathione peroxidase. Proc. Natl. Acad. Sci. USA 89, 5837–5841. Dunphy, G. B., and Thurston, G. 1990. Insect immunity. In ‘‘Entomopathogenic Nematodes in Biological Control’’ (R. Gaugler and H. K. Kaya, Eds.), pp. 301–323. CRC Press, Boca Raton, FL. Dunphy, G. B., and Webster, J. 1991. Antihemocytic surface components of Xenorhabdus nematophilus var. dutki and their modification by serum of nonimmune larvae of Galleria mellonella. J. Invertebr. Pathol. 58, 40–51. Dutky, S. R., Thompson, J. V., and Cantwell, G. E. 1964. A technique for the mass propagation of the DD-136 nematode. J. Insect Pathol. 6, 417–422. Forschler, B. T., and Gardner, W. A. 1991. Parasitism of Phyllophaga hirticula (Coleoptera: Scarabaeidae) by Heterorhabditis heliothidis and Steinernema carpocapsae. J. Invertebr. Pathol. 58, 386–407. Gaugler, R., Wang, Y., and Campbell, J. 1994. Aggressive and evasive behaviors in Popillia japonica (Coleoptera: Scarabaeidae) larvae: Defense against entomopathogenic nematode attack. J. Invertebr. Pathol. 64, 193–199. Gillespie, J. P., Kanost, M. R., and Trenczek, T. 1997. Biological mediators of insect immunity. Annu. Rev. Entomol. 42, 611–643. Glaser, R. W., and Fox, H. 1930. A nematode parasite of the Japanese beetle (Popillia japonica Newm.). Science 71, 16–17.

50

WANG AND GAUGLER

Go¨tz, P., Boman, A., and Boman, H. G. 1981. Interactions between insect immunity and an insect-pathogenic nematode with symbiotic bacteria. Proc. R. Soc. Lond. B 212, 333–350. Gupta, A. P. 1991. Insect immunocytes and other hemocytes: Roles in cellular and humoral immunity. In ‘‘Immunology of Insects and Other Arthropods’’ (A. P. Gupta, Ed.), pp. 19–118. CRC Press, Boca Raton, FL. Gupta, A. P., Orenberg, S. D., Das, Y. T., and Chattopadhyay, S. K. 1991. Lectin-binding receptors, Na⫹, K⫹-ATPase, and acetylcholinesterase on immunocyte plasmamembrane of Limulus polyphenmus. Exp. Cell Res. 194, 83–89. Jarosz, J., and Stefaniak, M. 1993. Entomocidal proteases released by rhabditoid nematodes depress inducible cell-free antibacterial immunity in insects. Entomonematologia 2, 1–11. Kaya, H. K., and Gaugler, R. 1993. Entomopathogenic nematodes. Annu. Rev. Entomol. 38, 181–206. Klein, M. G. 1990. Efficacy against soil-inhabiting insect pests. In ‘‘Entomopathogenic Nematodes in Biological Control’’ (R. Gaugler and H. K. Kaya, Eds.), pp. 195–214. CRC Press, Boca Raton, FL. Klein, M. G., and Geogis, R. 1992. Persistence of control of Japanese beetle (Coleoptera: Scarabaeidae) larvae with steinernematid and heterorhabditid nematodes. J. Econ. Entomol. 85, 727–730. Page, A. P., Rudin, W., Fluri, E., Blaxter, M. L., and Maizels, R. M. 1992. Toxocara canis: A labile antigenic surface coat overlying the epicuticle of infective larvae. Exp. Parasitol. 75, 72–86. Politz, S. M., and Philipp, M. 1992. Caenorhabditis elegans as a model for parasitic nematodes: A focus on the cuticle. Parasitol. Today 8, 6–12. Premachandran, D., Von Mende, N., Hussey, R. S., and McClure, M. A. 1988. A method for staining nematode secretions and structures. J. Nematol. 20, 78–78. Ratcliffe, N. A., and Walters, J. B. 1983. Studies on the in vitro cellular reactions of insects: Clearance of pathogenic and nonpathogenic bacteria in Galleria mellonella. J. Insect Physiol. 29, 407–415. Rizki, R. M., and Rizki, T. M. 1984. Selective destruction of a host

blood cell type by a parasitoid wasp. Proc. Natl. Acad. Sci. USA 81, 6154–6158. Selvan, M. S., Grewal, P. S., Gaugler, R., and Tomalak, M. 1994. Evaluation of steinernematid nematodes against Popillia japonica (Coleoptera: Scarabaeidae) larvae: Species, strain, and rinse after application. J. Econ. Entomol. 87, 605–609. Steiner, G. 1929. Neoaplectana glaseri, n. g., n. sp. (Oxyuridae), a new nemic parasite of the Japanese beetle (Popillia japonica Newm.). J. Wash. Acad. Sci. 19, 346–440. Stoltz, D. B., and Cook, D. I. 1983. Inhibition of host phenoloxidase activity by parasitoid hymenoptera. Experimentia 39, 1022–1024. Tashiro, H. 1987. Turfgrass insects of the United States and Canada. Cornell Univ. Press, Ithaca, NY. Villani, M. G., and Wright, R. J. 1988. Entomogenous nematodes as biological control agents of European chafer and Japanese beetle (Coleoptera: Scarabaeidae) larvae infecting turfgrass. J. Econ. Entomol. 81, 484–487. Wang, Y., Campbell, J. F., and Gaugler, R. 1995. Infection of entomopathogenic nematodes Steinernema glaseri and Heterorhabditis bacteriophora against Popillia japonica (Coleoptera: Scarabaeidae) larvae. J. Invertebr. Pathol. 66, 178–184. Wang, Y., Gaugler, R., and Cui, L. 1994. Variations in immune response of Popillia japonica and Acheta domesticus to Heterorhabditis bacteriophora and Steinernema species. J. Nematol. 26, 11–18. Webb, B., and Luckhart, S. 1994. Evidence for an early immunosuppressive role for related Campoletis sonorensis venom and ovarian proteins in Heliothis virescens. Archiv. Insect Biochem. Physiol. 26, 147–163. Yel, T., and Alm, S. R. 1992. Effects of entomopathogenic nematode species, rate, soil temperature, and bacteria on control of Japanese beetle (Coleoptera: Scarabaeidae) larvae in the laboratory. J. Econ. Entomol. 85, 2144–2148. Yokoo, S., Tojo, S., and Ishibashi, N. 1992. Suppression of the prophenoloxidase cascade in the larval hemolymph of the turnip moth, Agrotis segetum by an entomopathogenic nematode, Steinernema carpocapsae and its symbiotic bacterium. J. Insect Physiol. 38, 915–924.