Entomopathogenic nematodes for control of Phyllophaga georgiana (Coleoptera: Scarabaeidae) in cranberries

Biocontrol Science and Technology, Vol. 18, No. 1, 2008, 2131 Entomopathogenic nematodes for control of Phyllophaga georgiana (Coleoptera: Scarabaeidae) in cranberries Albrecht M. Koppenhöfer a *, Cesar R. Rodriguez-Saona b, Sridhar Polavarapu b$ and Robert J. Holdcraft b a Department of Entomology, Rutgers University, New Brunswick, NJ, USA; b Blueberry and Cranberry Research and Extension Center, Rutgers University, Chatsworth, NJ, USA (Received 5 September 2007; final version received 2 October 2007) A series of laboratory and greenhouse experiments evaluated the entomopathogenic nematodes Steinernema scarabaei Stock & Koppenhöfer, Heterorhabditis bacteriophora Poinar, and H. zealandica Poinar for control of second- and thirdinstar cranberry white grub, Phyllophaga georgiana Horn (Coleoptera: Scarabaeidae), in cranberries. Steinernema scarabaei was the most effective species with 76100% control at a rate of 2.510 9 IJ/ha in the greenhouse experiments. H. zealandica and especially H. bacteriophora were generally less effective and required rates of 510 9 IJ/ha for acceptable control. Larval stage had no effect on H. zealandica and H. bacteriophora performance, whereas S. scarabaei was more effective against third instars than second instars in the laboratory but not in the greenhouse experiments. Steinernema scarabaei, should it become commercially available, could be an effective alternative to chemical insecticides for P. georgiana management. Keywords: Phyllophaga georgiana; entomopathogenic nematodes; Steinernema; Heterorhabditis Introduction The larvae of several species of white grubs (Coleoptera: Scarabaeidae) are pests of cranberries, Vaccinium macrocarpon Ait., in the USA (Averill and Silvia 1998; Cowles, Polavarapu, Williams, Thies and Ehlers 2005). The larvae of white grubs feed in the soil on underground roots, runners and stems. This causes stunting and spindling of plants which in severe cases is followed by the death of sporadic patches of cranberries. The resulting bare patches may be colonised by weeds, making reestablishment of plants difficult and expensive. White grub species that cause damage in cranberries include the cranberry root grub, Lichnanthe vulpina Hentz, the grape Anomala, Anomala lucicola Fab., the oriental beetle, Anomala ( Exomala) orientalis Waterhouse, Hoplia modesta (Haldeman), Hoplia equina (LeConte), and the cranberry white grubs, Phyllophaga anxia LeConte and Phyllophaga (Subgenus Phytalus) georgiana Horn. In New Jersey, P. georgiana appears to be the most common white grub pest of cranberries (C.R. Rodriguez-Saona, R.J. Holdcraft, personal observations). Corresponding author. Email: koppenhofer@aesop.rutgers.edu $ Deceased 7 May 2004. ISSN 0958-3157 print/issn 1360-0478 online # 2008 Taylor & Francis DOI: 10.1080/09583150701721705 http://www.informaworld.com

22 A.M. Koppenhöfer et al. The life cycle of P. georgiana has not been studied in detail. Based on our limited observations we believe that the species has a 2-year life cycle with a new generation starting every year. Adults fly in June and early July and lay eggs among the roots of host plants. The eggs hatch in about 3 weeks and first instars occur during July and August. Second instars are found from late August onward, overwinter, and resume feeding in spring. Third instars start to appear in mid-june and will feed voraciously into September. This is the stage believed to cause the most severe damage, typically from July through September. Third instars collected in September will typically purge their intestines within a few weeks and will become much less active. Thus, the larvae appear to enter a prepupal stage in early fall, in which the second winter is spent, followed by pupation in the following spring around MayJune. The USA produces about 77% of the world cranberry crop with 15,245 ha harvested in 2006 representing a value of $251 million (NJASS 2006). Cranberries are a major component of the southern New Jersey economy (annual value $17 million in 2006) and are grown on approximately 1840 ha, mainly in the ecologically sensitive pine barrens (NJASS 2006). Cranberries are grown over naturally acidic peat bogs in beds that have been drained, cleared, leveled, and covered with sand before planting. Presently, the only effective control option for white grubs in cranberries is the neonicotinoid systemic insecticide imidacloprid. However, imidacloprid is expensive, particularly because it is most effective when applied preventively against the first-instar grubs. There is a dire need for alternative control options, particularly control options that are effective as curative treatments against the later larval stages. Entomopathogenic nematodes (Heterorhabditidae and Steinernematidae) are used for the biological control of insect pests (Grewal, Ehlers and Shapiro-Ilan 2005a), primarily in an inundative approach and against soil insects. These nematodes may offer an environmentally safe and IPM compatible option for curative white grub control (Grewal, Koppenhöfer and Choo 2005b). Very little research has been conducted on the control of white grubs in cranberries using entomopathogenic nematodes. In several field experiments the nematodes Steinernema carpocapsae (Weiser), S. feltiae (Filipjev), S. glaseri (Steiner), and Heterorhabditis bacteriophora Poinar generally provided little grub suppression (Cowles et al. 2005). In a laboratory experiment in 30-mL cups Koppenhöfer, Fuzy, Crocker, Gelernter and Polavarapu (2004) tested three nematode species against third instars of Phyllophaga crinita Burmeister, Phyllophaga congrua LeConte, and P. georgiana. S. glaseri was ineffective against all three Phyllophaga species, H. bacteriophora showed some promise against P. georgiana but was ineffective against the other species, but Steinernema scarabaei Stock & Koppenhöfer was highly virulent against all three species. However, the substrate used in this laboratory study was a sandy loam. Koppenhöfer and Fuzy (2006) showed that the infectivities of S. glaseri, H. bacteriophora, H. zealandica Poinar, and S. scarabaei were lower in acidic sand (in that study originating from blueberry fields) than in less acidic typical agricultural soils (sandy loam to clay loam; ph range 5.97.0). More recent research in the turfgrass system has demonstrated the high virulence and efficacy of recently isolated strains of H. bacteriophora (GPS11 strain) and H. zealandica (X1 strain) and of the recently isolated species S. scarabaei against several white grub pests (Grewal PS, Grewal SK, Malik and Klein 2002; Grewal PS, Power, Grewal SK, Suggars and Haupricht 2004; Grewal et al. 2005b; Koppenhöfer

Biocontrol Science and Technology 23 and Fuzy 2003a). In particular, S. scarabaei has shown very high virulence and field efficacy against several species of the Melolonthinae, the scarab subfamily to which Phyllophaga spp. belong (Cappaert and Koppenhöfer 2003; Koppenhöfer and Fuzy 2003b; Koppenhöfer et al. 2004; Koppenhöfer, Grewal and Fuzy 2006). In addition, S. scarabaei has also shown excellent efficacy against A. orientalis in blueberries in acidic sands similar to those in which cranberries are grown (Polavarapu, Koppenhöfer, Barry, Holdcraft and Fuzy 2007). The objective of this study was therefore to determine the potential of the entomopathogenic nematode S. scarabaei, H. bacteriophora (GPS11) strain, and H. zealandica (X1 strain) for control of P. georgiana in cranberries. Since larval stage can affect nematode efficacy against some white grub species (Koppenhöfer and Fuzy 2004) we also studied the effect of P. georgiana larval stage on nematode performance as this information would help predict the best timing of field applications. Material and methods General methods Second- and third-instar P. georgiana were collected from commercial cranberry bogs located in Burlington Co., NJ, in May/June and August/September. None of the sites had been treated with insecticides during the previous year. The larvae were kept individually in the cells of 24-well plates in acidic cranberry bog sand at 158C for short-term storage (B1 week) and at 108C for long-term storage (16 weeks). The larvae were warmed up at room temperature (21258C) for 24 h before use in experiments. Heterorhabditis bacteriophora (GPS11 strain) and H. zealandica (X1 strain) were cultured in late instar greater wax moth larvae, Galleria mellonella (L.) (Lepidoptera: Pyralidae) (Kaya and Stock 1997). Steinernema scarabaei (AMK001 strain) was cultured in P. japonica and A. orientalis larvae because its production in wax moth larvae was unreliable. It should be noted that the use of a different host for rearing may have affected to a limited degree the relative virulence of the three nematode species (see Discussion). The infective juvenile stage nematodes (IJs) emerging from infected larvae were harvested from emergence traps, i.e. modified White traps, over 7 days and stored in water at 108C for 730 days (Kaya and Stock 1997). The soils used in the experiments were typical cranberry acidic peat bog sands (ph 4.05.0; Hancock 1995). The soil had been carefully air-dried on flats in the greenhouse (21 238C) for at least 3 weeks before use to eliminate native entomopathogenic nematodes. Baiting of soil samples with wax moth larvae never revealed native nematodes, and no infections by entomopathogenic nematodes or fungi observed in the untreated larvae in any experiment. A series of laboratory and greenhouse experiments were conducted to compare nematode species efficacy and determine optimal release rates. Laboratory experiments Laboratory experiments were conducted at room temperature (22258C) in 30-mL plastic cups (10 cm 2 ) filled with 25 g of moist acidic cranberry bog sand (final soil water potential after addition of treatments 10 kpa10%, w/w, soil moisture) with cranberry roots added as food for the larvae. Second and third instars were

24 A.M. Koppenhöfer et al. collected in late August/early September. Individual larvae that had been held at room temperature for 24 h were released into the cups. Larvae that did not enter into the soil within 2 h were replaced and the cups were treated 1 day later. Treatments were applied in 0.5 ml of water; controls received water only. Larval mortality was assessed at 7 and 14 days after treatment (DAT). In the first laboratory experiment, second instars and third instars were exposed to 100 or 400 S. scarabaei. There were four replicates with 10 cups each per treatments. In the second experiment, second and third instars were exposed to 400 S. scarabaei, H. bacteriophora,orh. zealandica. In addition, second instars were also exposed to 100 or 200 S. scarabaei, H. bacteriophora, or H. zealandica. There were four replicates with five cups each per treatment. Greenhouse experiments Greenhouse experiments were conducted in 2-L pots (200 cm 2 at soil surface, 8-cm soil depth) planted with cranberry vines in acidic cranberry bog sand. Average temperature was 21238C and photoperiod was 14:10 light to dark. Phyllophoga georgiana larvae were added 34 days before treatment application. Larvae that did not burrow into the soil within 2 h were replaced. Treatments were applied in 100 ml water. Control pots were treated with water only. The pots were watered every 34 days until evaluation. Treatments were evaluated destructively at 21 DAT by carefully going through the soil and noting the number of live, dead, and nematode-infected larvae. For the first experiment (2002), larvae were collected in late September and the experiment conducted in October. Each pot received either five second instars or five third instars. Second instars were exposed to S. scarabaei (2.510 9 IJ/ha) and third instars were exposed to S. scarabaei (1.2510 9,2.510 9,or510 9 IJ/ha) or H. bacteriophora (510 9 IJ/ha). For the second experiment (2004), larvae were collected in late August and the experiment conducted in September. Pots received either eight second instars or six third instars. Second and third instars were exposed to S. scarabaei, H. bacteriophora, orh. zealandica (each at 510 9 IJ/ha). Third instars were additionally exposed to S. scarabaei at 2.510 9 IJ/ha. For the third experiment (2005), larvae were collected in mid- to late-may and the experiment conducted in June. To prevent the second instars from molting into third instars during the experiment, they were kept at 108C for 24 weeks between collection and the start of the experiment. Pots received eight second instars or six third instars. Second instars were exposed to S. scarabaei (0.6310 9 IJ/ha). Third instars were exposed to S. scarabaei (0.3110 9, 0.6310 9, 1.2510 9, or 2.510 9 IJ/ha), H. bacteriophora (2.510 9 or 510 9 IJ/ha), or H. zealandica (2.510 9 or 510 9 IJ/ha). For the fourth experiment (2007), larvae were collected in mid- to late-may (mostly second instars) and in June (third instars) and the experiment was conducted in July. Larvae were kept at 108C for 46 weeks between collection and the start of the experiment to prevent the second instars from molting into third instars during the experiment. Pots received either eight second instars or six third instars. Second and third instars were exposed to S. scarabaei (0.3110 9, 0.6310 9, 1.2510 9,or 2.510 9 IJ/ha), H. bacteriophora (2.510 9 ) IJ/ha, or H. zealandica (2.510 9 IJ/ha).

Biocontrol Science and Technology 25 Data analysis Before analyses, data were normalised where necessary by square root (number of surviving larvae) or arcsine square root (control-corrected percent mortality data) transformation. To determine if treatments caused significant mortality the numbers of surviving larvae were subjected to analysis of variance (ANOVA), by larval stage if different numbers of larvae were released per pot in the greenhouse experiments (Experiments 24), and means separated using Tukey s test. To compare treatment effects between larval stages the data were corrected for control mortality (Abbott 1925) and analysed using ANOVA and Tukey s test. Differences among means were considered significant at P B0.05. Means 9 SE are shown. Results Laboratory experiments In the first experiment, there was a significant interaction between larval stage and S. scarabaei rate whether data were analysed as number of larvae surviving or as control-corrected percentage mortality. Therefore, data were analysed with larval stage-rate combination as treatments. Both S. scarabaei rates had significantly lower larval survival than the controls in both second and third instar at 7 and 14 DAT (F]50.67; df5, 18; P B0.0001). Control-corrected mortality was significantly higher in third instars exposed to 400 S. scarabaei than in any other treatment at 7 and 14 DAT (F]22.24; df3, 12; PB0.0001) (Figure 1). The lower S. scarabaei rate caused higher mortality in third instars than in second instars at 14 DAT but not at 7 DAT. The higher S. scarabaei rate caused higher mortality in third instars than in second instars at 7 and 14 DAT. In the second experiment, larval survival at 7 DAT was significantly lower than in the controls at 400 S. scarabaei for second and third instars and 400 H. zealandica for third instars (F10.89; df13, 42; PB0.0001). At 14 DAT, larval survival was significantly lower than in the controls at 400 IJs for all three nematode species in both instars and at 200 S. scarabaei against second instars (F9.31; df13, 42; PB 0.0001). At 7 DAT, control-corrected mortality was higher than in all other treatments for the 400 S. scarabaei rate against third instars (Figure 1). At 14 DAT, third instar mortality by 400 S. scarabaei was higher than in all other treatments except the 400 S. scarabaei rate against second instars (F]8.29; df11, 36; P B0.0001). Against third instars, S. scarabaei caused at least 2 higher mortality than the other nematodes species at 7 and 14 DAT. However, against second instars there was no significant difference among nematode species at any rate. If only data for second instars were included in the analysis, there was a significant effect at 7 and 14 DAT of nematode species (F]4.00; df2, 27; PB0.05) and nematode rate (F]7.00; df 2, 27; P B0.005) but no interaction between species and rate. At 7 and 14 DAT, S. scarabaei caused significantly higher mortality than H. bacteriophora with H. zealandica not different from the other species, and 400 IJs caused significantly higher mortality than 100 IJs with 200 IJs not differing from the other rates.

26 A.M. Koppenhöfer et al. Figure 1. Effect of treatment with the entomopathogenic nematodes Steinernema scarabaei (Ss), Heterorhabditis bacteriophora (Hb) and H. zealandica (Hz) on control-corrected mortality of second-instar and third-instar Phyllophaga georgiana in 30-mL cups with acidic sand and cranberry roots. Numbers in labels indicate number of infective juvenile nematodes applied in treatment. Lower (upper) case letters indicate significant differences in controlcorrected mortality at 7 (14) days after treatment (DAT) (PB0.05). An * indicates that treatment caused significant mortality compared to untreated cups in a separate analysis (PB 0.05). Greenhouse experiments In the first experiment, the only treatment causing a significant reduction in larval survival was S. scarabaei (2.510 9 IJ/ha) against second instars (F27.42; df6, 63; P B0.001) which also caused significantly higher control-corrected mortality (79%) than all other treatments (F 19.97; df4, 45; P B0.001). However, the majority of the third instars recovered at evaluation had purged their intestines and showed very little activity. In the second experiment, second-instar survival was significantly lower in the H. bacteriophora treatment than in the control and significantly lower in the

Biocontrol Science and Technology 27 H. zealandica and S. scarabaei treatments than in the H. bacteriophora treatment (all treatments 510 9 IJ/ha) (F30.93; df3, 16; PB0.001). Third instar survival was significantly lower than in the control in the H. bacteriophora and H. zealandica treatments (both 510 9 IJ/ha) and significantly lower in both S. scarabaei treatments (2.510 9 and 510 9 IJ/ha) than in all other treatments. Controlcorrected mortality (both instars analysed together) was higher in all S. scarabaei treatments than in the other treatments except H. zealandica against second instars (F 12.10; df6, 40; P B0.001) (Figure 2). There was no significant difference between second and third instar for any nematode treatment. In the third experiment, second-instar survival was significantly lower in the S. scarabaei treatment (0.6310 9 IJ/ha) than in the control (F15.78; df1, 6; PB 0.01). Third-instar survival was significantly lower than in the control in all S. scarabaei treatments (0.3110 9 IJ to 2.510 9 IJ/ha) and in the higher rate of H. zealandica (5 10 9 IJ/ha) (F11.74; df8, 63; PB0.001). Control-corrected mortality (both instars analysed together) was higher at the highest S. scarabaei rate than both H. bacteriophora rates and the lower H. zealandica rate (F7.38; df8, 59; PB0.001) (Figure 2). There was no significant dose effect for S. scarabaei (5376% mortality). Against third instars at the 2.510 9 IJ/ha rate, S. scarabaei caused 2.9 higher mortality than H. zealandica and 4.8 higher mortality than H. bacteriophora. Instar had no significant effect on mortality caused by S. scarabaei (at 0.6310 9 IJ/ha). In the fourth experiment, larval survival for both second and third instars was significantly lower than in the control in the S. scarabaei treatments but not in the H. bacteriophora and H. zealandica treatments (F]7.58; df6, 49; PB0.001). Controlcorrected mortality (both instars analysed together) was significantly affected by treatment (F10.33; df5, 84; PB0.001) but not by instar, and there was no significant interaction between treatment and instar. The highest S. scarabaei rate caused significantly higher mortality than the second lowest rate of S. scarabaei and than H. bacteriophora and H. zealandica (Figure 2). The lowest S. scarabaei rate caused significantly higher mortality than H. bacteriophora and H. zealandica. At the 2.510 9 IJ/ha rate, S. scarabaei caused 2.3 higher second instar mortality and 2.63.7 higher third instar mortality than H. zealandica and H. bacteriophora. Discussion The entomopathogenic nematode S. scarabaei has excellent potential for the control of P. georgiana in cranberries as it provided 76100% control at a rate of 2.510 9 IJ/ha in greenhouse experiments. Steinernema scarabaei was more effective against third instars than second instars in laboratory but not in greenhouse experiments. Heterorhabditis zealandica and especially H. bacteriophora were less effective than S. scarabaei and required rates of 510 9 IJ/ha to achieve 5589% and 2463% control, respectively. Larval stage had no effect on H. zealandica and H. bacteriophora performance. In the first greenhouse experiment, neither S. scarabaei nor H. bacteriophora caused significant mortality of third-instar P. georgiana. However, most of the third instars had purged their intestines during the experiment and had become inactive compared to normal feeding third instars. Based on our limited life cycle observations for P. georgiana we suspect that these larvae were preparing to spend the winter in this inactive stage to turn into pupae in spring. In the Japanese beetle, Popillia japonica Newman, and A. orientalis susceptibility to H. bacteriophora and

28 A.M. Koppenhöfer et al. Figure 2. Effect of treatment with the entomopathogenic nematodes Steinernema scarabaei (Ss), Heterorhabditis bacteriophora (Hb), and H. zealandica (Hz) on control-corrected mortality of second-instar and third-instar Phyllophaga georgiana in 2-L pots with acidic sand and cranberry vines at 21 days after treatment. Graphs represent three greenhouse experiments conducted in 2004, 2005, and 2007. Numbers in labels indicate nematode application rates ( 10 9 infective juveniles/ha). To facilitate comparisons among experiments, same treatments are placed at the same position on the x-axis of each experiment. Letters indicate significant differences in control-corrected mortality (PB0.05). An * indicates that treatment caused significant mortality compared to untreated pots in a separate analysis (PB0.05). especially S. scarabaei drops dramatically from the feeding third instar over the purged third instar to the prepupal and pupal stage (Koppenhöfer and Fuzy 2004). We suspect that purged third-instar P. georgiana are also much less nematodesusceptible than feeding third instars.

Biocontrol Science and Technology 29 Observations in two other studies suggest that S. scarabaei, H. bacteriophora, and H. zealandica may be less effective in acidic soils than in more ph-neutral soils. Koppenhöfer and Fuzy (2006) observed reduced efficacy in acidic sand (from blueberry fields) of these species under laboratory condition but no significant reduction in 1-L pots with grass (Koppenhöfer and Fuzy 2006). Polavarapu et al. (2007) observed that higher S. scarabaei rates were required to control A. orientalis in blueberries (acidic sand) than generally in turfgrass (loamy more ph neutral soils) (e.g. Koppenhöfer and Fuzy 2003a; Koppenhöfer et al. 2006) but this difference may be more due to the deeper root-system and with that the deeper distribution of the larvae in blueberries compared to turfgrass. Our observations on P. georgiana susceptibility in the laboratory were similar to those made by Koppenhöfer et al. (2004) despite the use of acidic sand as a substrate rather than sandy loam. Based on our laboratory and greenhouse observations, the best timing for S. scarabaei applications should be in May/June when individuals from the P. georgiana generation initiated 1 year earlier are second and third instars and after soil temperatures have risen above 158C. However, at this time P. georgiana from the generation initiated 2 years earlier will be prepupae, pupae, and adults, the nematode-susceptibility of which still needs to be determined. From late-june into late-august soil temperature will often be too high for good S. scarabaei performance (Koppenhöfer and Fuzy 2003c). Application from late-august to mid-september when soil temperatures are more favorable again and P. georgiana are first/second instars (generation initiated 1 year earlier) and third instars (generation initiated 2 years earlier) should also be effective. However, at this time white grub feeding damage to the cranberries may be imminent or may already have occurred. Optimal timing for H. zealandica and H. bacteriophora may require somewhat higher soil temperatures than for S. scarabaei due to the lower efficacy of these species. In addition, H. bacteriophora was generally ineffective at soil temperatures below 208C against P. japonica in turfgrass (Georgis and Gaugler 1991). The temperature range for infectivity under laboratory conditions (constant temperature, small arenas) is 12.5308C for S. scarabaei (using A. orientalis as a host) (Koppenhöfer and Fuzy 2003c) vs. 15328C for H. bacteriophora (using wax moth larvae as a host) (Grewal, Selvan and Gaugler 1994). Thus, compared to S. scarabaei optimal timing for H. bacteriophora application may be 12 weeks later in spring and 12 weeks earlier in late summer. The temperature range of H. zealandica has not been studied in detail. Observations in turfgrass suggest that S. scarabaei may be even more effective over longer periods of time. Thus, when applied at the low rates of 0.10.2510 9 IJ/ ha in September S. scarabaei provided 5077% A. orientalis control at 31 DAT, 86 100% by the following spring, and 7677% 13 months after application (i.e. in the next larval generation) (A.M. Koppenhöfer, unpublished data). Laboratory data suggest that the low ph of typical blueberry soil has no negative effect on S. scarabaei persistence (Koppenhöfer and Fuzy 2006). Steinernema scarabaei may therefore also have good potential for long-term white grub suppression in cranberries. In fact, the overlap of two biannual generations in P. georgiana will provide a more constant supply of susceptible hosts in cranberries compared to turfgrass where the annual life cycle of most white grub species results in an absence of susceptible hosts in June and July. This is critical for the perpetuation of nematode populations because S. scarabaei appears to be rather specialised to scarab larvae as

30 A.M. Koppenhöfer et al. hosts (Koppenhöfer and Fuzy 2003c). Thus, in cranberries the effect of S. scarabaei after an application in late-may could accumulate throughout the growing season and result in very high levels of P. georgiana suppression. However, S. scarabaei survival through the winter may be poor due to the flooding of the cranberry bogs from December through April and the resulting oxygen deficiency in the soil. As indicated in the material and methods sections, the use of a different host species for rearing may affect the relative virulence S. scarabaei vs. H. bacteriophora and H. zealandica. However, S. scarabaei- and H. bacteriophora-virulence to thirdinstar A. orientalis did not differ when produced for one rearing cycle in wax moth larvae or in scarab larvae (Koppenhöfer and Fuzy 2003a). Steinernema scarabaeivirulence to third-instar A. orientalis was also not significantly affected by three consecutive rearing rounds in A. orientalis vs. G. mellonella (A.M. Koppenhöfer, unpublished data). While it is possible that more rearing rounds in different hosts would affect the relative virulence, S. glaseri-virulence to third-instar P. japonica was not affected after 12 rearing rounds in different hosts (G. mellonella vs. P. japonica) (Stuart and Gaugler 1996). Our observations will need to be confirmed under field conditions. However, at least S. scarabaei has excellent potential as a biological control agent for P. georgiana and probably also other cranberry white grubs. Unfortunately, thus far it has not been possible to mass-produce S. scarabaei effectively. If H. zealandica or H. bacteriophora can provide similar control levels under field condition as observed in our greenhouse studies, these species could offer a feasible albeit expensive control option for P. georgiana. Acknowledgements We thank Eugene Fuzy and Matthew Resnik for laboratory assistance and Dan Rice, Mark Cappuccio, Alyssa DeStefano, and Dan Goldbacher for greenhouse and field assistance. This research was funded in part by the Cranberry Institute, a USDA-CSREES Special grant to the P.E. Marucci Center for Blueberry and Cranberry Research ad Extension Rutgers University (No. 2006-06058), and a grant from EPA Region 2 to CRS. References Abbott, W.S. (1925), A method for computing the effectiveness of an insecticide, Journal of Economic Entomology, 18, 265267. Averill A.L., and Sylvia M.M. (1998), Cranberry insects of the northeast. East Wareham, MA, USA: UMass Cranberry Experimental Station. Cappaert, D.C., and Koppenhöfer, A.M. (2003), Steinernema scarabaei, an entomopathogenic nematode for control of the European chafer, Biological Control, 28, 379386. Cowles, R.S., Polavarapu, S., Williams, R.N., Thies, A., and Ehlers, R.-U. (2005), Soft fruit applications, in Nematodes as biocontrol agents, eds. P.S. Grewal, R.-U. Ehlers and D.I. Shapiro-Ilan, Wallingford, UK: CABI Publishing, pp. 231254. Georgis, R., and Gaugler, R. (1991), Predictability in biological control using entomopathogenic nematodes, Journal of Economic Entomology, 84, 713720. Grewal, P.S., Selvan, S., and Gaugler, R. (1994), Thermal adaptation of entomopathogenic nematodes: niche breadth for infection, establishment, and reproduction, Journal of Thermal Biology, 19, 245253.

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