SAMPLING AND BIOSTATISTICS Estimation of Hymenopteran Parasitism in Cereal by Using Molecular Markers

1 2 DOUGLAS B. JONES, KRISTOPHER L. GILES, YI CHEN, AND KEVIN A. SHUFRAN

Department of Entomology and Plant Pathology, Oklahoma State University, Stillwater, OK 74078Ð3033

J. Econ. Entomol. 98(1): 217Ð221 (2005) ABSTRACT Polymerase chain reaction (PCR) primers were designed and tested for identiÞcation of immature parasitoids in small grain cereal aphids and for estimation of parasitism rates. PCR technique was evaluated for 1) greenhouse-reared greenbugs, graminum (Rondani), parasitized by Cresson and 2) aphids collected from winter Þelds in Caddo County, Oklahoma. For greenhouse samples, parasitism frequencies for greenbugs examined by PCR at 0, 24, and 48 h after removal of L. testaceipes parasitoids were compared with parasitism frequencies as determined by greenbug dissection. PCR was unable to detect parasitism in greenbugs at 0 and 24 h postparasitism, but it was able to detect parasitoids 48 h after parasitoid removal at frequencies that were not signiÞcantly different from dissected samples. Field-collected samples were analyzed by rearing 25 aphids from each sample and by comparing parasitoid frequencies of mummies developed and PCR performed on another 50 aphids. samples included corn leaf aphids, (Fitch); bird cherry-oat aphids, Rhopalosiphum padi (L.); English grain aphids, Sitobion avenae (F.); and greenbugs. Mummies were isolated until adult emergence, whereupon each parasitoid was identiÞed to species (L. testaceipes was the only parasitoid species found). Parasitism detection frequencies for PCR also were not statistically different from parasitism frequencies of reared aphids. These results indicate that PCR is a useful tool for providing accurate estimates of parasitism rates and especially for identiÞcation of immature parasitoids to species.

KEY WORDS Lysiphlebus testaceipes, , PCR, parasitoid eggs

SMALL GRAIN APHIDS ARE attacked by several parasitoid difÞcult. Frequently, observable differences between species (Royer et al. 1998). Lysiphlebus testaceipes species consist of little more than a wing vein that is (Cresson), (: Aphidiidae) is arguably slightly longer in one species than another closely the most important parasitoid of small grain aphids related species, such as Aphidius colemani Viereck throughout the central and western wheat-growing versus L. testaceipes (Pike et al. 1997, Atanassova et al. regions of the United States (Jackson et al. 1970, 1998). Although it is relatively simple to Þnd aphid Walker et al. 1973, Archer et al. 1974, Summy et al. mummies because they differ in appearance from 1979, Rice and Wilde 1988, Giles et al. 2003). This their aphid host, detection of parasitoids in nonmum- oligophagous parasitoid frequently suppresses green- miÞed aphids is time-consuming, requiring rearing of bug, Schizaphis graminum (Rondani) (Homoptera: live specimens until the adult parasitoid emerges ), and other small grain aphid populations (Royer et al. 1998). Dissection of aphid hosts is useful below economic thresholds on crops such as wheat to conÞrm attack by parasitoids (Jones et al. 2003), but and grain in the southern Great Plains identiÞcation of the immature parasitoid to species is (Patrick and Boring 1990, Jones 2001, Giles et al. 2003). virtually impossible. A potential solution to each of Effective use of parasitoids for control of small grain these problems is the use of polymerase chain reaction aphids requires knowledge of species composition and (PCR). valid estimates of the rate of attack (Patrick and Bor- Polymerase chain reaction results in the rapid pro- ing 1990, Royer et al. 1998). IdentiÞcation of these duction of multiple copies of a speciÞc nucleotide aphid parasitoids by use of taxonomic keys is often sequence found in DNA (Mullis 1990). The PCR tech- nique makes it possible to analyze nucleotide se- Mention of trade names or commercial products in this article is quences in samples that contain amounts of DNA that solely for the purpose of providing speciÞc information and does not are either too small, or too degraded, to permit other imply recommendation or endorsement by the United States Depart- types of nucleic acid analysis (Glick and Pasternak ment of Agriculture. 1998). This feature of PCR has made possible the 1 Campus Box 2096, Alderson Broaddus College, Phillippi, WV 26416. development of experimental and diagnostic molec- 2 USDAÐARS, 1301 N. Western Rd., Stillwater, OK 74075. ular biology techniques, which were previously ex-

0022-0493/05/0217Ð0221$04.00/0 ᭧ 2005 Entomological Society of America 218 JOURNAL OF ECONOMIC ENTOMOLOGY Vol. 98, no. 1 tremely time-consuming or impossible to perform. For wheat aphid, Diuraphis noxia (Mordvilko); yellow example, Ward et al. (1990) successfully used PCR to sugarcane aphid, Sipha flava (Forbes); English grain diagnose an arboviral infection (Dugbe viral RNA) in aphid, Sitobion avenae (F.); and greenbug (Y.C., un- an vector. By using PCR, diagnoses of viral published data; GenBank accession nos. AY745772Ð infection were obtained within 48 h, whereas biolog- AY745781). Parasitoids consisted of Aphelinus asychis ical assays took at least8dtocomplete. Another Walker, Aphelinus albipodus Hayat & Fatima, Apheli- feature of PCR is that live specimens are not required nus varipes (Foerster), Aphelinus hordei (Kurdjumov), (Persing et al. 1990). Processing samples can be per- Aphytis melinus DeBach, (MÕIntosh), formed at the researcherÕs leisure because samples can Aphidius colemani, A. ervi Haliday, Aphelinus matricariae be kept for extended periods in the freezer (Glick and Haliday, and L. testaceipes (Chen et al. 2002; GenBank Pasternak 1998). accession nos. AF289130ÐAF289148). We used GCG We chose to use the 16S rDNA gene in our research. Wisconsin Package UNIX version 10 (Genetics Com- The 16S rDNA gene fragment is of interest because puter Group, Madison, WI) for alignment and analysis. little variation in the coding regions is found within The single-base detection technique (Kwok et al. 1990) species and could be used as a marker indicating the was used to design primers. presence or absence of parasitoids in a host. Addi- A forward and a reverse primer were designed on tionally, the 16S rDNA gene occurs as 100 or more the conserved part of the 16S rDNA gene to detect copies within each cell compared with single-copied the general presence of a parasitic wasp. These nuclear genes (Dover 1982, 1986). Finally, Chen et al. general primers were designated WaspF1 (5Ј-ACC (2002) already sequenced the 16S rDNA gene for all TG TTTAT CAAAA ACATG-3Ј) and WaspR (5Ј- of the common small grain aphid parasitoids found in CGAGG TCGCA ATCTT TTTTA-3Ј). A 465-bp PCR the southern Great Plains. product was produced by using of WaspF1 and Our objectives for this research were to 1) deter- WaspR primer pair. Development of these general mine accuracy of PCR primers designated LtepuF and primers was necessary because Þeld-collected aphids LtepuF for detection of L. testaceipes eggs and larvae could harbor any of the listed parasitoids. inside greenbugs at 0, 24, and 48 versus 72 h postovi- SpeciÞc primers were then developed on the con- position when dissection can provide reliable parasi- served part of the 16S rDNA gene as well, to detect the toid detection (Jones et al. 2003) in a controlled lab- presence of L. testaceipes DNA from any population, oratory experiment; and 2) further test the accuracy yet only amplify 16S rDNA from L. testaceipes (Chen of these primers for detection of L. testaceipes eggs and et al. 2002). These primers were designated LtepuF larvae inside aphids collected from winter wheat Þelds (5Ј-CAAAA ACATG TCTTA TTGAA-3Ј) and LtepuR in Oklahoma. (5Ј-CCCCA ATTAA ATATT AGTTT AA-3Ј). The LtepuF and LtepuR primers produced a 299-bp PCR product. Materials and Methods Laboratory Study. Approximately 500 second to Greenbug and Parasitoid Colonies. Biotype “E” third instars of greenbugs were placed on each of three greenbugs were obtained from colonies maintained pots of grain sorghum and allowed to settle for 4 h. at the USDAÐARS Plant Science and Water Con- Each sorghum plant was caged such that greenbugs servation Research Laboratory at Stillwater, OK, and were excluded from the soil, yet they were exposed to established on grain sorghum (ÔSG-925Õ) grown in a ample airßow. Twenty-Þve female L. testaceipes para- fritted clay and sphagnum moss mixture. sitoids from laboratory colonies were introduced into colonies and all sorghum plants were kept inside each cage and allowed to parasitize greenbugs for 4 h, double-walled Þne mesh cages within a greenhouse after which they were immediately removed by aspi- (Ϸ22Ð28ЊC). The double-walled cages prevented con- ration. Four groups of 100 greenbugs were removed tamination of colonies by feral greenbugs and parasi- randomly from each plant for analysis, at 0, 24, 48, and toids while permitting ample airßow (Jones et al. 72 h postparasitoid removal. The Þrst three groups of 2003). Fresh plants were rotated as needed into cages 100 greenbugs from each plant were each placed in housing colonies. 90% ethanol and frozen until PCR was performed. The L. testaceipes was isolated from specimens collected last group removed at 72 h was placed in 2% saline and at Perkins, OK, in fall 1999 (40 L. testaceipes adults dissected to detect presence of L. testaceipes larvae. isolated from Ϸ50 greenbug mummies). Pots of sor- Dissections were only performed on greenbugs re- ghum infested by greenbugs were placed in the col- moved at 72 h because parasitoid eggs are difÞcult to onies every 3Ð4 d to maintain a steady supply of detect, thus delaying dissections until after parasitoid parasitoids. Parasitoid colonies were maintained at eggs hatch greatly improved accuracy of the data 22 Ϯ 0.5ЊC and a photoperiod of 12:12 (L:D) in double- (Hofsvang and Hågvar 1978, van Steenis 1993). This walled Þne mesh cages kept in a growth chamber procedure was replicated three times. (Jones et al. 2003). Greenbugs destined for PCR analysis were placed Primer Design. Primers were designed using 16S individually in 1.5-ml microcentrifuge tubes. They rDNA sequences that represent commonly found ce- were then homogenized for 1 min with a pestle real aphids and their parasitoids. The aphids consisted mounted in a battery-powered drill in 80 ␮l of isola- of corn leaf aphid, Rhopalosiphum maidis (Fitch); bird tion buffer containing 0.1 M NaCl, 0.2 M sucrose, cherry-oat aphid, Rhopalosiphum padi (L.); Russian 0.1 M Tris-HCl (pH 9.1), 0.05 M EDTA, 1% SDS, and February 2005 JONES ET AL.: ESTIMATION OF PARASITISM IN CEREAL APHIDS 219

20 ␮g/ml RNAase A. The homogenate was vortexed brießy and incubated for 30 min in a 65ЊC water bath. The sample was then spun in a centrifuge for 3 min. The supernatant ßuid was removed and placed in a new tube. Five microliters of 3 M sodium acetate and 110 ␮l of ice cold 100% ethanol were then added. This solution was mixed well and then centrifuged for 30 min to collect the precipitated nucleic acid salts at the bottom of the tube. The salts were air-dried and then resuspended in 50 ␮l of distilled water. One microliter of the nucleic acid salt solution was placed in a new 0.5-␮l vial along with enzyme storage buffer A consisting of 50 mM Tris-HCl (pH 8.0 at 25ЊC), 100 mM NaCl, 0.1 mM EDTA, 1 mM dithio- threitol, 50% glycerol, and Triton X-100, along with

MgCl2, TaqDNA polymerase, NTPs (all supplied by Promega, Madison WI), primers LtepuF and LtepuR Fig. 1. Mean parasitism rates in laboratory prepared for L. testaceipes, and distilled water. Each vial was greenbug specimens as determined by PCR detection of placed in a PTC-100 thermocycler (MJ Research, parasitoid DNA in greenbugs at 0, 24, and 48 h after removal Watertown, MA). DNA was denatured for 3 min at of L. testaceipes females compared with parasitism rate de- Њ termined by dissection of greenbugs at 72 h after removal of 94 C, followed by 35 ampliÞcation cycles, with 30-s ϭ ϭ ϭ Њ Њ parasitoids (48 versus 72 h: t 0.18, df 6, P 0.86, SAS denaturing at 94 C, 30-s annealing at 55 C, and 1-min PROC MIXED). extension at 72ЊC. DNA was Þnally extended for 1 min at 72ЊC after ampliÞcation. After ampliÞcation, PCR products were separated on a 1.5% agarose electro- and those samples where the parasitism rate was mea- phoresis gel, stained with ethidium bromide, and pho- sured by PCR. tographed under UV light. Presence of parasitoid Voucher Specimens. Voucher specimens of aphid species have been deposited in the Department of DNA is conÞrmed by the migration of a band of DNA Entomology and Plant Pathology Museum at Okla- down its respective lane that matches the band in the homa State University, Stillwater, OK. Voucher spec- known DNA lane. Comparisons were made using imens of parasitoid species were reported previously PROC MIXED (SAS Institute 1999) between samples (Chen et al. 2002). where the parasitism rate was measured by dissection 0, 24, and 48 h postparasitism PCR. Field Study. Approximately 75Ð100 aphids were col- Results and Discussion lected from each of four wheat Þelds in Caddo County, We were unable to detect parasitism in greenbugs OK, during November 2001. For each group of aphids in the 0 and 24 h postparasitoid removal samples with collected, 25 randomly selected aphids were isolated PCR in the laboratory study (Fig. 1). However, we and reared on wheat plants for up to 7 d until mummies were able to detect L. testaceipes in 48 h postparasitoid ␮ developed. Each mummy was then isolated in a 1.5- l removal samples at frequencies not signiÞcantly dif- microcentrifuge tube, allowed to emerge, and identi- ferent from dissected samples (t ϭ 0.18, df ϭ 6, P ϭ Þed to species by taxonomic key (Pike et al. 1997). 0.86). In Þeld-collected samples consisting of corn leaf Another 50 aphids were randomly selected from each aphids, bird cherry-oat aphids, greenbugs, and English group of collected aphids and were frozen in 90% grain aphids, samples that tested positive for parasitoid ethanol to preserve until PCR ampliÞcation was per- presence (WaspF1 and WaspR primers) also tested formed. Because of the possibility of other parasitoid positive for L. testaceipes (LtepuF and LtepuR prim- species being present in the Þeld-collected samples, ers) in every instance. Previous research shows that we used the general parasitoid primers WaspF1 and this is not unusual during late fall and winter in the WaspR to determine the presence of any parasitoid southern Great Plains (Jones 2001, Giles et al. 2003). DNA. This PCR procedure used differed from the Samples reared to parasitoid emergence also were laboratory experiment by the primers used and the only parasitized by L. testaceipes. Statistical analysis thermocycler program in which DNA was denatured revealed that the percentage of hosts parasitized by for 3 min at 94ЊC, followed by 35 ampliÞcation cycles, L. testaceipes as detected by PCR was not statistically with 30-s denaturing at 94ЊC, 30-s annealing at 51ЊC, different from parasitism percentage as determined by and 1-min extension at 72ЊC. DNA was Þnally ex- rearing aphids (Fig. 2; F ϭ 1.85, df ϭ 3, P ϭ 0.27, PROC tended for 1 min at 72ЊC after ampliÞcation. If a para- MIXED). sitoid was detected in the sample by using general PCR techniques are able to detect and amplify DNA parasitoid primers, the sample was then reexamined fragments from very minute quantities (Johner et al. using LtepuF and LtepuR primers to discover whether 1999, Zhu and Greenstone 1999). Using PCR, they L. testaceipes was present. Comparisons were then were able to detect very small amounts of polydna- made using PROC MIXED between samples where virus DNA in the parasitized egg of Spodoptera litto- the parasitism rate was measured by rearing aphids ralis (Boisduval) (Lepidoptera: Noctuidae) shortly 220 JOURNAL OF ECONOMIC ENTOMOLOGY Vol. 98, no. 1

primary parasitoids and hyperparasitoids attacks ce- real grain aphids (Pike et al. 1997, Royer et al. 1998). As long as parasitoid DNA is present in the host, PCR should be able to detect its presence. At present, there is no deÞnitive method to determine which parasitoid would “win” the competition for the host aphid re- sources should there be more than one species of parasitoid present. Additionally, PCR cannot deter- mine whether the host aphid was parasitized multiple times by the same parasitoid species. Detection of parasitoid larvae within their hosts and identiÞcation of the parasitoid species by PCR will be very useful for researchers to examine aphid samples postmortem and derive useful data from them. Taxo- nomic keys are available to identify adult aphid para- Fig. 2. Mean parasitism rate determined by PCR detec- sitoids to species (Pike et al. 1997, Atanassova et al. tion of parasitoid DNA (using LtepuF and LtepuR primers) 1998). This necessitates live specimens and timely in samples of aphids collected in winter wheat Þelds com- inspection of those samples. Use of PCR could make pared with parasitism rate determined by key identiÞcation handling of aphid samples easier, eliminating the need of adult parasitoids that emerge from corresponding samples for host plant production and daily observations re- of aphids reared for7d(F ϭ 1.85, df ϭ 3, P ϭ 0.27, SAS PROC MIXED). quired for rearing procedures. Additionally PCR could be used to “resurrect” previously unusable samples where the aphid host died before the parasitoid could after oviposition by the parasitoid Chelonus inanitus develop into an adult. Archived specimens taken long (L.) (Hymenoptera: ). PCR should be able ago also could be reexamined to determine whether to amplify even one copy of the target gene. However, they were parasitized and, with the proper primers, it is more likely that multiple copies are needed to determine what species of parasitoid was responsible. provide a large enough titer of DNA for ampliÞcation (Glick and Pasternak 1998). Acknowledgments These results demonstrate that it is possible to re- liably use PCR to detect and identify the DNA of We thank Nathan Jones and Tim Johnson for technical parasitizing small grain aphids. However, it contributions toward this research project. We also thank does have its limitations. We were unable to detect Matt Greenstone, Richard Grantham, and Jack W. Dillwith parasitoid DNA at 0 and 24 h postparasitoid removal, for critically reviewing this manuscript. This work was ap- but we were very successful at 48 h postparasitoid proved for publication by the Director of the Oklahoma removal. These detection times differ somewhat from Agricultural Experiment Station and supported in part under projects OKL02334 and OKL02342. work done by Zhu and Greenstone (1999), who were able to detect cereal aphid parasitism by A. asychis by PCR. Additionally, Ratcliffe et al. (2002) were able to References Cited differentially detect larvae of four parasitoid species in Archer, T. L., R. H. Cate, R. D. Eikenbary, and K. J. Starks. the genus Muscidifurax (Hymenoptera: Pteromali- 1974. Parasitoids collected from greenbugs and corn leaf dae) within 24 h after they parasitized stable ßy, aphids in Oklahoma in 1972. Ann. Entomol. Soc. Am. 67: Stomoxys calcitrans (L.), pupae. 11Ð14. A possible reason for the lag time necessary for PCR Atanassova, P., C. P. Brookes, H. D. Loxdale, and W. Powell. to detect aphid parasitoids might be that the DNA of 1998. 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