Development and Evaluation of Methods To Detect Nucleopolyhedroviruses in Larvae of the Douglas-Fir Tussock Moth, Orgyia pseudotsugata (McDunnough)

Abstract

Various molecular methods are used to detect pathogenic microorganisms and viruses within their hosts, but these methods are rarely validated by direct comparison. Southern hybridization, enzyme-linked immunosorbent assay (ELISA), and a novel DNA extraction/PCR assay were used to detect Orgyia pseudotsugata multiple nucleopolyhedrovirus (OpMNPV) in Douglas-fir tussock moth larvae. PCR was more sensitive than Southern hybridization and ELISA at detecting semipurified virus. ELISA, however, was the most accurate method for detecting virus within larvae, given that Southern hybridization and PCR produced false-negative results (31% and 2.5%, respectively). ELISA may be preferable in some applications because virus infections can be quantified (r² = 0.995). These results may be applicable to both applied and academic research that seeks to accurately identify the incidence of viruses and microorganisms that regulate insect populations.

The collapse of many insect outbreak populations is correlated with infections by pathogenic microorganisms and viruses (2, 46). Baculoviruses, entomopathogens that play an important role in regulating many insect populations (33, 35), are prevalent after pest populations reach outbreak proportions and peak during the declining phase of the outbreak cycle (52). Pest management decisions may be guided by pathogen incidence, because increases are correlated with host declines (38, 49).

The Douglas-fir tussock moth, Orgyia pseudotsugata McDunnough (Lepidoptera: Lymantriidae) (DFTM), periodically reaches outbreak populations between the coastal and central mountain ranges in western North America (15, 32, 35, 50). Larvae feed on interior Douglas-fir trees, Pseudotsuga menziesii var. glauca Franco, and true fir species, Abies spp. Miller. DFTM fluctuate from outbreak populations that result in visible defoliation and economic losses to forestry to endemic populations below an economic threshold level (15, 30, 31).

DFTM populations are moderated by two native baculoviruses, Orgyia pseudotsugata multiple nucleopolyhedrovirus (OpMNPV) and O. pseudotsugata single NPV (OpSNPV) (19), which are prevalent during DFTM outbreak cycles and peak at the start of the host population decline (48). In the field, DFTM feed on foliage with other insects, including the western spruce budworm, Choristoneura occidentalis Freeman (38), and the rusty tussock moth, O. antiqua Linnaeus (18). Thus, DFTM larvae may be exposed to C. fumiferana multiple nucleopolyhedrovirus (CfMNPV) used to control C. occidentalis [37]), C. occidentalis granulovirus, or O. antiqua NPV (OaNPV) infections, although they are not known to reduce DFTM field populations.

An integrated pest management (IPM) program developed to suppress outbreak populations of the DFTM relies on monitoring the predicted density of DFTM larvae based on egg mass counts. The incidence of OpMNPV in larvae reared from the egg masses is also determined, and if an outbreak of DFTM is anticipated, OpMNPV is applied early in the outbreak cycle (36, 45, 48). The success of this IPM program is based on the accuracy of the monitoring methods, yet the current monitoring method, microscopic evaluation (38, 49), is neither specific, sensitive, nor accurate (21, 56). Detection methods should be both selective for, and sensitive to, DFTM-specific pathogens. They should be sensitive enough to detect early pathogenesis when viral copy number is low and should be amenable to processing large numbers of samples for accurate estimates of the pathogen incidence.

Pathogen detection should also be rapid to ensure rapid management decisions. DFTM cause the greatest foliage damage during the first year of significant defoliation (31). Severe outbreak populations may defoliate entire trees within a 2-month period (57), and OpMNPV requires 6 to 8 weeks to initiate epizootics and reduce the host population (38). Thus, management decisions must be made at the time of population outbreak to prevent economic damage. These requirements provide a strong rationale for rapid diagnosis.

PCR, Southern hybridization, and enzyme-linked immunosorbent assay (ELISA) have been used to identify baculoviruses. PCR can amplify femtogram to picogram quantities of baculovirus DNA (23) and may detect a single genome (3). Additionally, PCR can be specific by targeting unique genetic elements. Although improved DNA extraction methods have extended the application of PCR to field analyses (10, 13, 41, 51, 53, 55), PCR may not facilitate large sample sizes because of assay time and cost. In contrast, Southern hybridization is designed for large sample sizes. Hybridization is more sensitive than microscopic analysis (12, 21, 24, 56) but is less sensitive than PCR. ELISA is also specific and, like hybridization, is amenable for screening large numbers of samples. Furthermore, ELISA is rapid because detection can be made from crude extracts (34), but it is less sensitive than PCR.

In this study, we compared three candidate methods for DFTM pest management: PCR, Southern hybridization, and ELISA. We evaluated their sensitivity, specificity, accuracy, and efficacy for large sample sizes.

MATERIALS AND METHODS

Materials.

The following baculoviruses were extracted from insect cadavers: OpMNPV genotypic variants (including the variant used in the registered biocontrol agent, TM Biocontrol-1; Corporation CCIP, Charlesbourg, Quebec, Canada), OpSNPV genotypic variants, OaNPV, CfMNPV, Lambdina fiscellaria NPV (LfNPV), Lymantria dispar multiple nucleopolyhedrovirus (LdMNPV) variants, and Neodiprion abietis NPV (NeabNPV) (received from I. S. Otvos). Protein concentration was estimated by the Lowry method using an RC-DC protein assay purchased from Bio-Rad Corporation (Mississauga, Ontario, Canada).

A glassmilk-based DNA extraction kit was purchased from Fermentas (K0513; Burlington, Ontario, Canada). Proteinase K, DNase, deoxynucleoside triphosphates, and Taq polymerase were purchased from Invitrogen (Burlington, Ontario, Canada). Oligonucleotides were designed using OLIGO software (Molecular Biology Insights, Cascade, CO) and purchased from Alpha DNA (Montreal, Quebec, Canada). Gel purification (QIAquick Gel Extraction) and plasmid purification (QiaPrep spin minprep) kits were purchased from QIAGEN (Mississauga, Ontario, Canada). pBlueScript plasmid DNA was purchased from Stratagene (La Jolla, Calif.). A radioactive labeling kit (Bench Top Labeling) was purchased from Gibco (Burlington, Ontario, Canada).

Polyethylene glycol-purified monoclonal mouse anti-OpNPV antibodies were purchased from ImmunoPrecise Antibodies (Victoria, British Columbia, Canada). Horseradish peroxidase-labeled goat anti-mouse (Fc) secondary antibodies were purchased from Pierce (Rockford, Ill.). 5′3′-Tetramethylbenzidine substrate was purchased from BioFX (Owing Mills, Md.).

Hydrobond+ nitrocellulose membrane, Phosphor screens, and a Storm PhosphorImager were purchased from Amersham Biosciences (Pittsburgh, Pa.). Hybridization BioDot microfilter vacuum manifold and Costar 96-well flat-bottom EIA polystyrene plates were purchased from Bio-Rad Corporation (Mississauga, Ontario, Canada). Enzyme immunoassay (EIA) plates were read using a MicroTek plate reader purchased from Bio-Tek Instruments (Winooski, VT).

Extraction of NPV polyhedral inclusion bodies.

NPV polyhedral inclusion bodies (PIBs) were extracted from insect cadavers (29). Briefly, larvae were macerated with a sterile mortar and pestle in 500 μl/larva TEC buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA, 50 mM NaCl, 10 mM cysteine) supplemented with 0.1% sodium dodecyl sulfate (SDS). PIBs were separated from insect debris by centrifugation at 145 × g at 4°C for 4 min, and supernatant was aspirated. Supernatants were pelleted by centrifugation at 18,000 × g for 15 min at 4°C. PIB pellets were washed three additional times with 1 ml of TEC buffer each and pelleted as described and stored at −20°C. Protein content was estimated by Lowry assay. PIBs were counted using a hemocytometer under a 40× objective of a compound microscope to correlate protein concentration and PIB number.

Isolation and purification of baculovirus DNA (standard protocol).

OpMNPV DNA was isolated from infected DFTM cadavers (29) with the following modifications. Larvae were macerated by mortar and pestle as described above; however, TEC, 0.1% SDS buffer was supplemented with 0.6 μg/ml DNase I to degrade host DNA. Semipurified PIBs were resuspended in 0.1 M Na₂CO₃ and incubated for 20 min with gentle agitation, and released virions were isolated by centrifugation at 18,000 × g at 4°C for 30 min. Viral DNA was released by incubation at 55°C for 18 h in TES buffer (10 mM Tris-HCl, pH 7.6, 1 mM EDTA, 0.1% SDS) and 0.2 mg/ml proteinase K. DNA was then isolated from contaminating proteins by phenol-chloroform extraction and purified using glassmilk.

Isolation of baculovirus DNA (rapid DNA extraction protocol).

Virus DNA was extracted from OpMNPV PIBs by a novel alkaline/heat lysis protocol (modification of procedures in references 5 and 29). Virus PIBs (3.06 × 10⁷; isolated from cadavers and counted as described above) were lysed in 0.02 mM Na₂CO₃, pH 11.0 (final concentration), for 20 min at 25°C, with gentle mixing. The solution was neutralized with 0.16 M sodium acetate, pH 5.0 (final concentration). Enveloped virions were heat lysed for 10 min at 94°C to release virus DNA.

Sensitivity of PCR (rapid extraction protocol).

Virus DNA was confirmed with primers specific to a 750-bp region of the baculovirus polyhedrin (polh) gene (polhF, 5′-TCG ATT TAA TAC GCC GGG CCG-3′; polhR, 5′-TGC CAG ATT ACT CGT ACC GGC CG-3′). Serial dilutions of the lysed solution (306,000 to 8 PIBs/μl) were added to a PCR master mix (10 mM Tris HCl, pH 9.0, 60 mM KCl, 2.5 mM MgCl₂, 200 μM each deoxynucleoside triphosphate, 1 mM each primer, 0.045% Triton-X, 1.45% Tween 20, 1 mg/ml bovine serum albumin [BSA], 1 U Taq polymerase). PCRs were subjected to 94°C for 10 min; 30 cycles of 94°C for 30 s, 65°C for 45 s, and 72°C for 2 min; and 72°C for 7 min and held at 4°C. Amplified DNA were separated on 1% agarose-Tris-borate-EDTA gels (43).

Sensitivity of Southern hybridization.

A 750-bp polh amplicon was produced as described above with 10 ng of purified OpMNPV DNA (standard protocol). Amplified DNA was separated on a 0.6% Tris-acetate-EDTA gel (43), gel purified, cloned into pBlueScript KS II+, and confirmed by sequence analysis. The clone was labeled with [α-³²P]dCTP for hybridization reactions.

Southern hybridization. OpMNPV DNA was isolated from PIBs by alkaline and heat lysis as described above. Serial dilutions of the lysate (7.7 × 10⁵ to 6.0 × 10³ PIBs) were denatured and filtered to nitrocellulose membrane by vacuum (43). Unbound sites were blocked with 10 μg/ml herring sperm. Lysed virus samples were probed (43), and washed blots were exposed to a Phosphor screen for 4 h and then scanned.

Sensitivity of the ELISA method.

Extracted OpMNPV PIBs (serially diluted to 1.1 × 10⁶:77 PIBs/ml) were incubated in polystyrene EIA plates for 18 h at 4°C. Unbound sites were blocked with PBS-3% BSA for 30 min at 37°C. Mouse anti-NPV monoclonal antibodies were diluted optimally to 1/1,000 in PBS-0.1% Tween 20-0.05% BSA (PBSTB) and incubated for 1 h, with shaking at 37°C. Horseradish peroxidase-labeled goat anti-mouse immunoglobulin G antibodies (Fc portion) were diluted optimally (1/20,000) in PBSTB and incubated as described above. Plates were washed five times for 30 min with PBST after each antibody step and washed twice for 5 min after antigen and blocking incubations. Antibodies were visualized with 50 μl/well tetramethylbenzidine substrate and incubated for 1 h in the dark. A₆₅₀ values were compared to known concentrations of virus antigen. We defined positive samples as those with A₆₅₀ values that were 2.5 standard deviations greater than the mean absorbance value of uninfected larvae (n = 80) (39).

Specificity of detection methods.

Baculoviruses (10 ng DNA/reaction) were identified by amplification of a 750-bp region of the polh gene, as described above. We also tested putative OpNPV-specific primers designed to orf136 of OpMNPV (orf136F, 5′-TTT CTC TGG GCC TGC TGC TGG-3′; orf136R, 5′-GTT GGC CGG GAC TGT CGA CCT-3′ [1]). orf136 was amplified as described above but with annealing at 55°C. For identification by Southern hybridization, baculovirus DNA (100 ng) was denatured, vacuum filtered onto nitrocellulose, and probed as described above. Semipurified baculovirus PIBs (10 μg/ml) were incubated in polystyrene plates for identification by ELISA as described above.

Direct detection of OpMNPV infection in treated DFTM larvae.

DFTM larvae were reared from laboratory strain DFTM egg masses (Goose Lake) and decontaminated with a 2% bleach solution (Javex) at 25°C, 50% relative humidity, and a 16 h:8 h photoperiod until hatch. Hatched larvae were reared in a population maximum of 10 in 100- by 15-mm petri dishes containing artificial DFTM diet (54). Larvae were reared until second instar, transferred to new petri dishes, and then starved for 24 h before infection.

Viral infection method.

Fresh OpMNPV PIBs were produced as previously described (40). Diet plugs (20) were made from a DFTM diet, and each plug (4 ± 1 mg) was dosed with one 50% lethal dose OpMNPV (9.5 PIBs/second-instar larva [B. Kukan and I. S. Otvos, unpublished data]) in a total volume of 1 μl. Control larvae were dosed with 1 μl of distilled water administered to a diet plug. Larvae were reared individually in 24-well culture plates containing a single diet plug. Twelve control and 12 treated larvae that had consumed an entire diet plug within a 24-h period were randomly selected and frozen individually (day 1) for analysis. Selection was replicated to day 10 to a total of n = 120 for control and n = 120 for treatment. Larvae that died prior to selection were collected daily, frozen, and assayed to confirm infection.

Direct detection of baculovirus infection.

Larvae were macerated individually using a sterile mortar and pestle and an equal volume of TEC buffer to insect weight. Homogenates were diluted 1:10 in PBS, and 100 μl of each dilution was incubated in polystyrene EIA plates for 18 h at 4°C. The remainder of each sample was stored at −20°C for later use in PCRs, Southern hybridization reactions, or ELISA replicates. Baculovirus infection was determined by ELISA as described above. Tissue homogenates were subjected to alkaline and boiling lysis for hybridization and PCRs as described above (rapid extraction protocol). Baculovirus infection was determined by PCR amplification of a region of the polh gene as described above. Lysed tissue homogenates (100 μl/larva) were denatured, filtered onto nitrocellulose membrane, and probed by Southern hybridization as described above.

False-positive and false-negative results.

False-negative identification of virus infections were noted where there were concurrences of positive identification by only two methods. False-positive identification of virus infections was identified where more than one method did not identify infection. The criterion for false positives was refuted where virus infection was determined repeatedly (more than three times) by one method.

RESULTS

Sensitivity.

The rapid DNA extraction/PCR protocol was the most sensitive method for detecting OpMNPV from semipurified OpMNPV PIBs. The method was sensitive to a minimum of 7.8 PIBs (Fig. 1A). In contrast, Southern hybridization was sensitive to a minimum of 6,000 PIBs (Fig. 1B), and the ELISA method was sensitive to a minimum of approximately 850 PIBs (Fig. 1C).

FIG. 1.

Sensitivity of PCR, Southern hybridization, and ELISA to OpMNPV PIBs. (A) PCR amplification of the polh gene. Lanes 1 to 13, serial dilutions of OpMNPV viral DNA (extracted from 3.1 × 10⁵ to 7.65 PIBs); lane 14, negative control. L, 200 ng 1-kb ladder (New England Biolabs). (B) Samples probed with 750-bp OpMNPV polh gene. Row 1, lanes 1 to 8, lysate was serially diluted (7.65 × 10⁵ to 5.98 × 10³ PIBs); row 2, lanes 1 to 4, no DNA; row 2, lanes 5 to 8, insect DNA. (C) Lysate was serially diluted and identified by indirect ELISA using an OpNPV-specific antibody. Bars represent the standard deviation around the mean absorbance (n = 4). Uninfected DFTM larvae (n = 80) defined the background.

Specificity.

PCR amplification using polh primers was not specific to OpMNPV (Fig. 2A), because polh is highly conserved within nucleopolyhedroviruses (42, 58). In contrast, primers designed to orf136 of OpMNPV were specific to OpNPVs (Fig. 2B). The Southern hybridization method was most sensitive to baculoviruses with the greatest nucleotide sequence identity to the OpMNPV polh gene (Fig. 2C). The probe cross-reacted with CfMNPV (87% identical [9]), moderately cross-reacted with LdNPV and LffNPVs (76% identical [25, 27]), was minimally reactive to OpSNPV (76% identical [26]), and did not cross-react with NeabNPV (55% identical [11]). In contrast to PCR and Southern hybridization methods, the ELISA method was specific to O. pseudotsugata NPVs (Fig. 2D). The ELISA method did not cross-react with any other baculoviruses above the criteria for positive identification.

FIG. 2.

Comparative specificity of PCR, Southern hybridization, and ELISA. PCR amplification of the polh gene from various baculoviruses using (A) polh primers or (B) primers specific to OpMNPV's orf136. (A) L is 200 ng of a 100-bp ladder (Invitrogen). Lanes 1 to 6, OpMNPV variants; lane 7, CfMNPV; lanes 8 to 10, LdNPV variants; lane 11, LffNPV; lane 12, NeabNPV; lane 13, OpSNPV; lane 4, no DNA. (B) Lanes 1 to 3, OpMNPV variants; lane 4, OpSNPV; lane 5, CfMNPV; lanes 6 to 8, LdNPV variants; lane 9, LffNPV; lane 10, AcMNPV; lane 11, no DNA. Kb, 200 ng 1-kb ladder (New England Biolabs). (C) Specificity of Southern hybridization to baculovirus DNA. Lanes 1 to 9, OpMNPVs; lane 10, OpSNPV; lane 11, CfMNPV; lanes 12 to 14, LdNPVs; lane 15, LffNPV; and lane 16, NeabNPV. The ELISA method was specific to (D) OpMNPV and OpSNPV. Bars represent standard deviations (n = 8).

Identification of OpMNPV infection in DFTM larvae.

The initial ingested dose (9.5 OpMNPV PIBs/larva) was greater than the minimum detection level of PCR for semipurified PIBs (approximately 8 PIBs; Fig. 1A), but individual samples were divided equally for detection by PCR, ELISA, and Southern hybridization. This division effectively reduced the initial ingested virus to a level below the minimum detection level of all methods used in this study. Thus, the detection of OpMNPV in test larvae by all methods was indicative of infection and not initial ingestion of infective dose.

PCR amplification of the polh gene confirmed OpMNPV infection in 5% of survivors of 50% lethal doses of OpMNPV and 95% of DFTM larvae killed due to virus treatment (Table 1). OpMNPV infection was detected as early as day 3 postingestion in 12.5% of individuals tested (n = 8). In contrast, OpMNPV infections were detected as early as day 6 postingestion by Southern hybridization (Table 1). OpMNPV infections were detected in 2.5% of surviving insects and in 71% of larvae that were killed using Southern hybridization (Table 1). The ELISA method consistently detected OpMNPV infections as early as day 3 postingestion (Table 1). OpMNPV infections were identified in 7.5% of larvae that survived OpMNPV ingestion and in 95% of larvae that died between days 6 to 10 postingestion using the ELISA method (Table 1).

TABLE 1.

Direct detection of OpMNPV infections in DFTM larvae by PCR, Southern hybridization, and ELISA

Method	DFTM status	Incidencea of infection on postingestion day:										% Incidence	% False positiveb	% False negativec
		1	2	3	4	5	6	7	8	9	10
PCR	Survivor	0	0	(1)	0	0	2	0	2	0	0	5-6	Not observed	5
	Dead	NA	NA	NA	NA	NA	4/6	8/8	8/8	8/8	7/7	95	Not observed
	Controld	0	0	0	0	0	0	0	0	0	0	NA	NA	NA
Southern	Survivor	0	0	0	0	0	1	0	1	0	0	2.5	Not observed	31
hybridization	Dead	NA	NA	NA	NA	NA	0/6	5/8	7/8	8/8	7/7	73	Not observed
	Control	0	0	0	0	0	0	0	0	0	0	NA	NA
ELISA	Survivor	0	0	1	0	0	2	0	2	0	1	7.5	Not observed	Not observed
	Dead	NA	NA	NA	NA	NA	4/6	8/8	8/8	8/8	7/7	95	Not observed
	Control	0	0	0	0	0	0	0	0	0	0	NA	NA

^aSecond-instar larvae were fed 9.5 PIBs/larva. A total of 80 survivors (n = 8/day) and 38 dead larvae were tested. Scores in parentheses indicate that virus was detected in some, but not all, replicates.

^bFalse positives were defined as the detection of virus infection by one method but failure to detect virus by the other two methods. False positives were refuted when virus infection was detected more than three times by one method.

^cFalse negatives were defined as failure to detect virus by one method when the other two methods detected virus.

^dControl larvae were mock infected with distilled water. A total of 80 survivors (n = 8/day) and 8 dead larvae were tested for infection by all methods.

Quantification of virus infection by ELISA.

A significant linear relationship (r² = 0.995) was noted between absorbance (relative bound antibody) and baculovirus particles (Fig. 3A). Absorbance values were thus used to evaluate baculovirus infections. Median absorbance values of larvae that survived baculovirus inoculation were highly variable on a given day, reflecting that OpMNPV infections were infrequent in survivors (data not shown). In contrast, median absorbance values of larvae killed during experimentation increased linearly from days 6 to 8 postingestion (r² = 0.969), after which the quantity of infection was greater than the upper limit of the ELISA method (6) (Fig. 3B).

FIG. 3.

Sensitivity of the ELISA method. OpMNPV was detected by indirect ELISA using an OpNPV-specific monoclonal antibody. (A) Linear relationship between absorbance and OpMNPV PIBs. Bars represent the standard deviation around the mean absorbance (n = 4). The linear relationship between absorbance and OpMNPV PIBs was significant (r² = 0.99). (B) Box plot representation of the relationship between absorbance and lethal baculovirus infections over time. The median absorbance (▴) increased linearly between days 6 to 8 postingestion (r² = 0.969).

Frequency of false-positive and false-negative results.

ELISA and PCR shared equal sensitivity for detecting virus in infected larvae, even though the rapid DNA extraction/PCR method was significantly more sensitive to OpMNPV DNA from semipurified PIBs (Fig. 1; Table 1). In contrast, Southern hybridization was the least sensitive method and produced a 31% false-negative frequency (Table 1). There was no evidence of false-positive results for any of the methods tested (Table 1).

DISCUSSION

The goal of this study was to evaluate pathogen detection methods for the IPM program of the Douglas-fir tussock moth. Our results indicate that all three methods are appropriate for identifying OpMNPV infection in DFTM larvae, because all of the methods are more sensitive (21) than the current standard, virus counts using light microscopy (49). Thus, one may predict that use of any of the described methods will result in more accurate control decisions, i.e., accurate determination of when the registered virus product should be applied to suppress DFTM populations.

PCR has been used extensively to identify baculovirus infections because of the sensitivity of the method (3, 4, 7, 14, 16, 17). The rapid baculovirus DNA extraction protocol developed and described in this study was sensitive to a minimum of approximately 8 polyhedral inclusion bodies (PIBs), which was estimated to be the equivalent of 10 pg of extracted virus DNA (data not shown). A unique feature of our method is that we converted a standard 2-day extraction procedure into a 1-h extraction protocol. Reduced sensitivity and accuracy of this method may have been due to contaminants within the macerated insect tissues causing PCR inhibition or due to inefficient release of viral DNA from nucleocapsids.

Southern hybridization and ELISA are relatively sensitive methods for detecting baculovirus PIBs. We found that Southern hybridization was sensitive to a minimum of 6,000 PIBs, which is consistent (2,000 to 7,800) with previous reports (12, 21). The ELISA method was sensitive to a minimum of 850 PIBs, which is in the range (100 to 2,000) of previous reports (8, 22, 28, 39, 44, 47), even though we used a stringent criterion for positive identification.

Rapid analysis is essential for DFTM management, because larvae rapidly defoliate, yet the virus requires time to initiate epizootics. Assay time is reduced by directly detecting virus within larvae. PCR and Southern hybridization produced false-negative results, suggesting that the ELISA method was the most accurate technique for identifying virus infections.

The ELISA method can be used to quantify infection, unlike the other methods described in this paper. This feature may be applied to predicting the potential inoculum required for controlling future populations. Previously, it has been shown that the ELISA method can be used to quantify baculoviruses (6, 39). We found that there was a significant linear relationship between semipurified baculovirus PIBs (r² = 0.995) or macerated infected larvae (r² = 0.969) and absorbance values, suggesting that the ELISA method can be used to accurately quantify both virus PIBs and virus infections from tissue homogenates.

To integrate these tools into the IPM program for the DFTM, they must be compared to the current detection strategy, microscopic virus counts, during regular seasonal sampling. DFTM egg masses are collected, and 25 emergent larvae/egg mass are assayed for virus. If less than 15% of larvae are infected, supplementary virus is inoculated to initiate epizootics and prevent defoliation (49). Thus, the next logical step is to determine what level (incidence as determined by ELISA) corresponds to the need for biological control (less than 15% as determined by microscopic counts). We are currently determining the incidence of OpNPVs in larvae that have emerged from field-collected egg masses by the Stelzer method (49) and the indirect ELISA method described in this study.

The described methods may be effective and applicable to other IPM programs. To apply our results to other systems, however, one must first address whether or not species-specific identification is essential or beneficial. Once this is determined, primers can be designed and antibodies can be selected for the needs of the system. Finally, the new technology must be compared to the system's current technology to determine the value (incidence as determined by the new technology) that supports control measures.