Differential oviposition timing by seasonality and oviposition timing of parasitoids of the invasive roseau cane scale (Nipponaclerda biwakoensis) as revealed by multiplex PCR assays

Differential oviposition timing by seasonality and oviposition timing of parasitoids of the invasive roseau cane scale (Nipponaclerda biwakoensis) as revealed by multiplex PCR assays Violet Sparks1, Ilgoo Kang2, Morgan Carrere1, Hannah Broadley3, Rodrigo Diaz1 1 Department of Entomology, 404 Life Sciences, Louisiana State University, Baton Rouge, LA 70803 USA 2 Department of Entomology, Kyungpook National University, 2559 Gyeongsang-daero, Sangju, Gyeongsangbuk-do 37224 Republic of Korea 3Forest Pest Methods Laboratory, USDA APHIS PPQ S&T, 1398 West Truck Road, Buzzards Bay, MA 02542 USA Corresponding author: Violet Sparks ([email protected]) Academic editor: Petr Janšta|Received 18 February 2025|Accepted 1 December 2025|Published 17 December 2025 https://zoobank.org/69831A5D-574D-48ED-ACF2-215F68C278BF Citation: Sparks V, Kang I, Carrere M, Broadley H, Diaz R (2025) Differential oviposition timing by seasonality and oviposition timing of parasitoids of the invasive roseau cane scale (Nipponaclerda biwakoensis) as revealed by multiplex PCR assays. Journal of Hymenoptera Research 98: 1133–1152. https://doi.org/10.3897/jhr.98.150721 Abstract Roseau cane scale, Nipponaclerda biwakoensis (Hemiptera: Aclerdidae) is an invasive species negatively impacting Phragmites australis (Cyperales: Poaceae) stands within the Mississippi River Delta. Previous research generated a PCR multiplex capable of identifying the three species of parasitoid wasps (Hymenoptera: Encyrtidae) that attack the scale in its invasive range. The appearance of a fourth species, Aprostocetus sp. (Hymenoptera: Eulophidae), within Louisiana in 2022 necessitated updating the multiplex with primers specific to the new species. The newly updated multiplex can identify the presence of all four parasitoid species within their host. DNA was extracted from the whole bodies of scales collected at different life stages and three different dates during the growing season and PCR performed with the new multiplex. Our findings show each species within the complex oviposits within a specific window of nymphal development which might help explain how successful multiparasitism frequently occurs in this system. The frequency of multiparasitism and relative abundance of each parasitoid species was consistent across the three sampling dates, except Astymachus lasallei (Hymenoptera: Encyrtidae), which was found in higher abundance during the end of the growing season. Keywords Competition, genetic identification, multiparasitism, oviposition, Phragmites, roseau cane scale, temporal niche partitioning JHR 98: 1133–1152 (2025) doi: 10.3897/jhr.98.150721 https://jhr.pensoft.net Copyright: This is an open access article distributed under the terms of the CC0 Public Domain Dedication. RESEARCH ARTICLE Violet Sparks et al. / Journal of Hymenoptera Research 98: 1133–1152 (2025) 1134 Introduction Roseau cane scale, Nipponaclerda biwakoensis (Kuwana) (Hemiptera: Aclerdidae) (hereafter referred as RCS), is invasive to the Gulf Coast, United States where it feeds exclusively on Phragmites australis (Cav.) Trin. Ex Steud. It is native to east Asia in the countries of China, Japan, South Korea, and Taiwan and has been collected as far as Beijing, China in the north and Hong Kong in the south (Cortez et al. 2022; Schneider et al. 2022). Scales aggregate under the leaf sheathes of host plants where they feed on the sap from vascular tissues. Mature females are approximately 5 mm long, brown, and give birth (ovoviparity) between 300 and 600 1st instar nymphs (crawlers) at the end of their lives (Kaneko 1995; Lopez 2022). Crawlers are the most mobile of the life stages, complete with six legs, eyes, and antennae (McConnell 1953) (Fig. 1A) and disperse from under the mother scale in search of appropriate feeding locations along the stem. Once they begin feeding, the immature scale molts, losing its appendages and eyes, and begins producing wax around the cuticular margin (McConell 1953; Kaneko 1995). Males of the species are less than 2 mm, apterous, and have simple legs Figure 1. Roseau cane scale development stages. A 1st instar (crawler) B 2nd instar C 3rd instar D 4th instar E mature female. Scale bars: 0.5 mm (A, B); 1.0 mm (C, D, E). Oviposition timing by RCS parasitoids 1135 they use to move towards nearby females with which they mate (McConell 1953). The life cycle takes between one and two months to complete with three to six overlapping generations each growing season, with faster development times and more generations associated with warmer climates like Taiwan (Kaneko 1995, 2004; Cortez et al. 2022). RCS was first reported outside its native range on Phragmites stands in the Lower Mississippi River Delta, Plaquemines Parish, Louisiana, United States in 2017 (hereafter referred to as MRD). Widespread die-back of Phragmites stands in the MRD (see Suppl. material 1) were associated with RCS populations as high as 2,000 scales per stem, suggesting that these outbreaks could be a major plant stressor (Knight et al. 2018). Given that Phragmites is the dominant vegetation within the MRD, this dieback raised concerns about potential ecological harm and wetland loss (Ramsey and Rangoonwala 2017; Cronin et al. 2020; Knight et al. 2020). Management of RCS within the MRD faces several challenges that make typical pest management tools ineffective. Insecticides are a not feasible tool, as the scales are difficult to spray due to their location underneath the leaf sheathes, and their use could endanger non-target wildlife inhabiting Phragmites stands. Prescribed burns, as used in cultivated Phragmites in China (Brix et al. 2014), risks damaging the numerous oil pipelines and pumping stations present in the MRD (Suppl. material 2). This leaves biological control of RCS as the most suitable option to achieve management. Initial investigations into RCS parasitism found three species present in Louisiana: Boucekiella depressa Hoffer, Neastymachus japonicus Tachikawa, and Astymachus lasallei Noyes and Higashiura (Hymenoptera: Encyrtidae) (Fig. 2A, B, D) (Knight et al. 2018; Cortez et al. 2022; Pruett et al. 2025). These three are found throughout the RCS native range and are thought to have arrived in Louisiana alongside their host (Kaneko 1995; Knight 2018; Cortez et al. 2022; Pruett et al. 2025). Species within the complex are noted as being gregarious (multiple individuals develop from the same host) endoparasitoids that consume all but the exocuticle of their host over the course of their development, with adults exiting from adult female mummies (Kaneko 1995). Astymachus lasallei and N. japonicus are known only to attack RCS while B. depressa has been collected from multiple hosts (Kaneko 1995; Herting 1997; Noyes and Higashiura 2020). In the fall of 2022 collections of parasitized RCS in Louisiana reared a species of Aprostocetus Westwood (Hymenoptera: Eulophidae) (Fig. 2C) that was morphologically similar to those found in the native range (Kaneko 1995, 2004, 2005a, b; Cortez et al. 2022) and was previously unreported in the region. Within a year of its discovery in Louisiana, Aprostocetus sp. was found in nearly every Phragmites patch infested by RCS in the adventive range (V. Sparks unpublished data). Previous work with RCS parasitoids has relied on the rearing of adult wasps from isolated hosts to make species level identification (Kaneko 1995, 2004, 2005a, b; Knight et al. 2018; Cortez et al. 2022). The larvae of these parasitoids are morphologically indistinct, and it cannot be guaranteed that all larvae within the host are of the same species due to the frequency of multiparasitism. Rearing adult wasps is not a perfect method either as it takes time for parasitoids to emerge, only parasitoids in the later stages of development may survive having their host removed from the stem, and infor- Violet Sparks et al. / Journal of Hymenoptera Research 98: 1133–1152 (2025) 1136 mation is lost for those that fail to complete development (Agusti et al. 2005). Polymerase chain reaction (PCR)-based molecular identification techniques are frequently used in parasitoid research due to their ability to accurately identify a species regardless of life Figure 2. Dorsal (left) and lateral (right) view of A Boucekiella depressa B Neastymachus japonicus C Aprostocetus sp. D Astymachus lasallei. Scale bars: 1.0 mm. Oviposition timing by RCS parasitoids 1137 stage and can differentiate between morphologically similar cryptic species (Smith et al. 2006, 2008; Gariepy et al. 2007). Diagnostic PCR-based assays are able to detect even small amounts of parasitoid DNA within a host, including eggs and early instars, because they can be designed to bind and amplify to specific DNA sequences (Tilman et al. 2000; Greenstone 2006). Combining multiple species-specific primers into a viable multiplex allows for single-step identification of parasitoids within a host in systems where multiple parasitoid species may be expected, and in the case of RCS, multiple species present within the same host (Traugott et al. 2006; Pruett et al. 2025). One of the outstanding aspects of the RCS parasitoid complex is the frequency with which successful multiparasitism, two or more species completing development from the same host, occurs. All combination pairs of each species and as many as three species have been observed emerging from the same host (Kaneko 1995; Cortez et al. 2022; Pruett et al. 2025; V. Sparks pers. obs.). Successful multiparasitism is supposed to be a rare phenomenon in Hymenoptera (Miller 1982). The larvae of solitary parasitoids typically come equipped with large mandibles and are highly mobile, allowing them to kill and consume other competitors within the host (Harvey et al. 2009). In typical intrinsic competition scenarios, there is an inferior and a superior competitor that is more likely to win the competition, but the inferior competitor can reclaim the advantage by ovipositing and developing ahead of the superior species (Hood et al. 2021). However, gregarious parasitoids, like the RCS parasitoids, lack these weapons and instead cohabitate with the other larvae of the same species (superparasitism) inside the host, whom are most likely also their siblings (Godwin and Odell 1984) or with the larvae of other parasitoid species in cases of multiparasitism. The first descriptions of the RCS parasitoid complex by Kaneko (1995) claim oviposition by all species occurs within immature females in their latest stage of development; however, it was not stated how this was concluded (such as first appearance of parasitoid larvae, dissection for eggs, or direct observation of oviposition behavior). The simultaneous oviposition by multiple species, if true, could explain why multiparasitism frequently occurs within the system as no one species has a head start on development over the others (Kaneko 1995). It is necessary in a biological control program to identify when the pest is vulnerable to attack by its natural enemies and so additional research on multiparasitism and oviposition timing in the RCS parasitoid community in the adventive range is needed. As Abell et al. (2020) indicates, releases performed at inopportune times may see a loss of agent effectiveness simply because populations of the target pest are not currently at a life stage the natural enemy can utilize. Previous work by Pruett et al. (2025) successfully produced species-specific PCR primers that were then combined into a multiplex for the molecular identification of B. depressa, N. japonicus, and A. lasallei DNA within RCS. With the arrival of Aprostocetus sp. to Louisiana, the multiplex needs to be updated to allow for the identification of all RCS parasitoids currently within the adventive range. The objectives of this study were to (1) develop species-specific PCR primers for Aprostocetus sp., (2) incorporate the newly developed primers into the already existing encyrtid multiplex, and (3) determine the host life stage window each species parasitizes their host by performing PCR with the multiplex on different RCS life stages. Violet Sparks et al. / Journal of Hymenoptera Research 98: 1133–1152 (2025) 1138 Methods Collection of scales and parasitoid wasps RCS were collected from field sites within the MRD, Plaquemines Parish, Louisiana, USA in October 2022 (Fall), May 2023 (Spring), and August 2023 (Summer). Roseau cane stems were haphazardly selected, cut at the soil or water line, and transported back to Louisiana State University, Baton Rouge, Louisiana where leaf sheathes were removed to expose the scales underneath. Living scales were collected and preserved in ethanol at 4 °C for later DNA extraction. Collected scales were measured across their longest point using 10 mm microscope rulers under a National DC3-420T digital microscope (National Optical, Schertz, TX, USA) and sorted into 1st instar larvae, 2nd through 4th instar larvae, and mature males and females (Table 1, Fig. 1). Size ranges for the 2nd through 4th instars were arbitrarily created based on a general trend of observing three size classes of immobile larvae (small, medium, and large). Visual indications (eggs, larvae, pupae present) of current parasitism were noted for each scale specimen. Parasitized mature scales with fully developed larvae were placed into gel capsules and adult wasps reared for morphological identification instead of storing in ethanol. Reared adult wasps were used for positive control DNA and for development of species-specific primers. Images of scale life stages and parasitoids were taken using the Visionary Digital BK Plus imaging system (Dun, Inc.) equipped with a Canon® EOS 5DS DSLR and were stacked using Zerene Stacker™ v. 1.04 (Zerene Systems LLC.). Adobe Photoshop® CS 6 was used to edit images. DNA extraction Genomic DNA was extracted from immature and adult scales as well as parasitoid wasps using the DNeasy Blood and Tissue Kit (QIAGEN, Hilden, Germany). The full body was ground using Fisherbrand™ Rnase-Free Disposable Pellet Pestles (Fisher Scientific, Carlsbad, CA, USA) in 180 µL of ATL buffer and 20 µL of proteinase K solution (Qiagen, Hilden, Germany). Samples were then incubated overnight at 56 °C. The remaining steps were followed according to the manufacturer’s protocol, except the final elution step was performed twice using 100 µL AE buffer each time for a total of 200 µL final DNA product. Table 1. Size and physical descriptions used to categorize life stages of the roseau cane scale, Nipponaclerda biwakoensis. Development is split into a mobile 1st instar nymph (crawler), three immobile nymphal instars (2nd-4th), and mature males and female. Instar Length (mm) Physical Characters 1st < 0.5 Legs present, no wax secretions 2nd 0.5-1.5 Loss of legs and antennae, wax secretion 3rd 1.5-3.0 Wax secretion buildup 4th > 3.0 Lack of melanization Mature (female) > 3.0 Complete melanization Mature (male) < 2.0 Pink, legs present, cylindrical body Oviposition timing by RCS parasitoids 1139 Aprostocetus sp. primer development and incorporation into multiplex PCR Previous research with RCS parasitoids in Louisiana designed species-specific PCR primers for Boucekiella depressa, Neastymachus japonicus, and Astymachus lasallei that produce 155 bp, 221 bp, and 495 bp bands, respectively (Pruett et al. 2025). We designed species-specific forward primers that produce 375 and 303 bp DNA sequences of Aprostocetus sp. when amplified with the encyrtid reverse primer previously developed by Pruett et al. 2025 (Table 2). To find appropriate forward primer regions, we downloaded 28S DNA sequences of B. depressa (656 bp, accession number: MN705818), N. japonicus (593 bp, accession number: MN705819), and A. lasallei (620 bp, accession number: MN705815) originally submitted to GenBank by K. Pruett. 28S sequences from Aprostocetus sp. were acquired from PCR using 28S-D2-3665F and 28S-D2-4068R universal primers (Hancock et al. 1988; Campbell et al. 1994) with DNA extracted from whole specimens. PCR products were then submitted to the LSU genomics facility (Baton Rouge, LA) and cleaned up using a primer depletion clean-up method developed by the facility. Purified products were bi-directionally sequenced on the 3130xl Genetic Analyzers (Applied Biosystems) using the BigDye Terminator v3.1 chemistry (Applied Biosystems). The obtained DNA sequences were edited and assembled using Geneious Prime 2021.2 (https://www.geneious.com) and compared with previously deposited parasitoid DNA sequences in the GenBank database of the National Center for Biotechnology Information (NCBI) using the Basic Local Alignment Search Tool (BLAST). Sequences of 28S gene region of the four RCS parasitoids were aligned using the Multiple Sequence Alignment Viewer 1.23.0 (https://www.ncbi.nlm.nih.gov/projects/ msaviewer/) and compared. We found two potential regions within the 28S gene sequence that could be used for designing forward species-specific primers for Aprostocetus sp. producing 375 and 303 bp DNA sequences when conducting PCR with the encyrtid reverse primer. Using the Oligo Analysis Tool provided on the website of Eurofins (https://eurofinsgenomics.com/en/resources/tools/oligo-analysis/), the properties of two potential forward primers were calculated, and the cross-dimer formation for each pair was evaluated. Analysis results indicated both forward primers could be used effectively in PCR. Table 2. Primer information including gene region, name, and sequence for all PCR primers discussed in this project. Gene Primer name and direction Sequence (5’ → 3’) Source 28S BD_F (F) CAAGCCAGGCAAAAAATCG Pruett et al. 2025 28S NJ_F (F) GACGAAACCGGTCCAACC Pruett et al. 2025 28S AL_F (F) GACCAAAGACCGGAGCCA Pruett et al. 2025 28S Encyrtid_28S_R (R) CACGAGACCGATAGCGAACA Pruett et al. 2025 28S 28S-D2-3665F (F) AGAGAGAGTTCAAGAGTACGTG Hancock et al. 1988 28S 28S-D2-4068R (R) TTGGTCCGTGTTTCAAGACGGG Campbell et al. 1994 28S Aprostocetus_F1 (F) TTGCGGCCCATTACCGTGA Newly developed 28S Aprostocetus_F2 (F) GCCCGGAGGCGCAAAC Newly developed Violet Sparks et al. / Journal of Hymenoptera Research 98: 1133–1152 (2025) 1140 The two newly designed primers were tested in singleplex with the encyrtid reverse primer and when incorporated into the encyrtid multiplex PCR using wasp DNA extracted both directly from wasps and from wasp immatures present in RCS samples. Singleplex PCR amplification and gradient PCR PCR amplification was run to confirm whether each species-specific primer amplifies DNA of their respective target organism when used with the Encyrtid_28S_R primer. Each 25 µl PCR amplification reaction volume contained 12.5 µl of DreamTaq Green PCR Master Mix (2X) (Thermo Scientific), 1 µl of DNA template, 9.5 µl of ddH2O, and 1 µl of the respective forward and reverse primer at 10 µM. PCR conditions were 94 °C for 3 minutes; 35 cycles of 94 °C for 30 seconds, either 52 °C for Aprostocetus_F1 or 57 °C for Aprostocetus_F2 and all other forward primers for 30 seconds and 72 °C for 1 minute; and a final extension at 72 °C for 5 minutes. To find out the optimal PCR annealing temperature of each primer set, we conducted gradient PCR using a PTC-200 thermal cycler (MJ Research, Massachusetts, USA). The gradient PCR condition was 94 °C for 3 minutes; 35 cycles of 94 °C for 30 seconds, 45–64 °C for 30 seconds and 72 °C for 1 minute; and a final extension at 72 °C for 7 minutes. Bands from all five forward primers were visible at 58 °C and so that temperature was selected as the annealing temperature for latter multiplex PCR. Multiplex PCR Multiplex PCR was performed using a T100 thermal cycler (Bio-Rad, Hercules, USA) with a total volume of 25 µl per reaction, containing 12.5 µl of DreamTaq Green PCR Master Mix (2X) (Thermo Scientific), 2.5 µl of primer mix, 1 µl of DNA template, and 8–9 µl of ddH2O. Optimization of primer concentrations was performed through trial and error testing of the multiplex with stock DNA extracted from adult parasitoids. The primer concentrations found to be most optimal for amplification of DNA products from all four species were: each forward primer for B. depressa, N. japonicus, and A. lasallei 0.096 µM and for Aprostocetus sp. was 0.064 µM, and the final concentration of the reverse primer was 0.35 µM (i.e., 50 µl of primer mix contains 1.2 µl of 100 µM forward primers for B. depressa, N. japonicus, and A. lasallei, 0.8 µl of 100 µM forward primer for Aprostocetus sp., 4.4 µl of the reverse primer, and 41.2 µl of ddH2O). PCR cycling conditions comprised of an initial denaturation at 94 °C for 3 minutes; followed by 40 cycles of denaturing at 94 °C for 30 seconds, annealing at 58 °C for 30 seconds, and extension at 72 °C for 50 seconds, and the final extension at 72 °C for 5 minutes. Serial dilutions of DNA extracted from specimens of each RCS parasitoid species were prepared and used in multiplex PCR assay to determine primer sensitivity to lowest DNA concentration. The stock concentration of each parasitoid sample was: 2.89 ng/µl for B. depressa, 6.0 ng/µl for N. japonicus, 4.5 ng/µl for Aprostocetus sp., and 4.8 ng/µl for A. lasallei. Stock solution was added or diluted so each 25 µl reaction contained 10, 5, 1, 0.5, 0.1, 0.05, 0.01, or 0 ng (negative control) of parasitoid DNA. Results are viewable in Suppl. material 3. Oviposition timing by RCS parasitoids 1141 Gel electrophoresis PCR products were visualized on 2.0% TAE agarose gels, made with TopVision Agarose Tablets (Thermo Fisher Scientific, Waltham, MA, USA), and stained with 1X SYBR™ Safe DNA Gel Stain (Invitrogen, Carlsbad, CA, USA) to confirm the success of PCR amplification. Initial multiplex PCR test using actual RCS samples The first test of the multiplex PCR with field-collected samples was conducted using DNA extracted from possibly parasitized RCS collected from the MRD in October 2022. Depending on the size of the immature and adult scales, we included multiple individuals within a sample, increasing the potential amount of parasitoid DNA within and making successful detection more likely. The sample set included 20 1st instar nymphs, 10 2nd instar nymphs, five 2nd instar nymphs, five 3rd instar nymphs, three 3rd instar nymphs, a single 3rd instar nymph, three 4th instar nymphs, two 4th instar nymphs, a single 4th instar nymph, three adult female scales, two adult female scales, a single adult female scale, and three male scales. Additionally, we included five positive controls, which consisted of DNA from B. depressa, N. japonicus, Aprostocetus sp., A. lasallei, all four wasps, and a negative control. Singleplex PCR was then performed on the same samples to confirm the results from the multiplex. Life stage and seasonality parasitism frequency Multiplex PCR was conducted using DNA extracted from individual scales collected at different times of the year to identify whether the frequency of parasitism by each species was variable by life stage and season. 100 1st instar nymphs, 20 2nd instar nymphs, 35 3rd instar nymphs, 35 4th instar nymphs, and 15 adult females were used from the October 2022 collection date. Only the 1st instar nymphs were pooled into the same sample, due to their small size, and they were never used in later dates due to the failure to detect any parasitoid DNA from them. 20 2nd instar nymphs, 30 3rd instar nymphs, 26 4th instar nymphs, and 14 adult females were used from the May 2023 collection date and 19 2nd instar nymphs, 16 3rd instar nymphs, 22 4th instar nymphs, and 20 adult females were used for August 2023. Statistical analysis The Pearson chi-square test was performed using JMP Pro 18.0.1 (SAS Institute, Cary, NC) to check whether observed species and multiparasitism frequencies from the PCR multiplex varied by seasonality. We tested the null hypothesis that the observed species frequencies and multiparasitism frequency were the same for all three sampling dates versus the alternative hypothesis that the frequencies were different for at least one sampling date. Violet Sparks et al. / Journal of Hymenoptera Research 98: 1133–1152 (2025) 1148 Agusti N, Bourguet D, Spataro T, Delos M, Eychenne N, Folcher L, Arditi R (2005) Detection, identification and geographical distribution of European corn borer larval parasitoids using molecular markers. Molecular Ecology 14(10): 3267–3274. https://doi.org/10.1111/ j.1365-294X.2005.02650.x Brix H, Ye S, Laws EA, Sun D, Li G, Ding X, Yuan H, Zhao G, Wang J, Pei S (2014) Largescale management of common reed, Phragmites australis, for paper production: A case study from the Liaohe Delta, China. Ecological Engineering 73: 760–769. https://doi. org/10.1016/j.ecoleng.2014.09.099 Campbell BC, Steffen‐Campbell JD, Werren JH (1994) Phylogeny of the Nasonia species complex (Hymenoptera: Pteromalidae) inferred from an internal transcribed spacer (ITS2) and 28S rDNA sequences. Insect molecular biology 2(4): 225–237. https://doi. org/10.1111/j.1365-2583.1994.tb00142.x Cortez Jr AO, Chu CL, Broadley HJ, Lo YS, Chen YC, Gates MW, Meyerson LA, Hoelmer KA, Gould JR, Hwang SY (2022) Exploratory surveys in Taiwan of the roseau cane scale Nipponaclerda biwakoensis Kuwana (Hemiptera: Aclerdidae) and its associated parasitoids. Journal of Applied Entomology 146(5): 596–606. https://doi.org/10.1111/jen.12982 Cronin JT, Johnston J, Diaz R. (2020) Multiple Potential Stressors and Dieback of Phragmites australis in the Mississippi River Delta, USA: Implications for Restoration. Wetlands. https://doi.org/10.1007/s13157-020-01356-8 Oligo Analysis Tool (2023) Eurofins Oligo Analysis Tool. https://eurofinsgenomics.com/en/ resources/tools/oligo-analysis/ [Accessed on 15.05.2023] Gariepy TD, Kuhlmann U, Gillott C, Erlandson M (2007) Parasitoids, predators and PCR: the use of diagnostic molecular markers in biological control of Arthropods. Journal of Applied Entomology 131(4): 225–40. https://doi.org/10.1111/j.1439-0418.2007.01145.x Godwin PA, Odell TM (1984) Laboratory study of competition between Blepharipa pratensis and Parasetigena silvestris (Diptera: Tachinidae) in Lymantria dispar (Lepidoptera: Lymantriidae). Environmental entomology 13(4): 1059–1063. https://doi.org/10.1093/ ee/13.4.1059 Greenstone MH (2006) Molecular methods for assessing insect parasitism. Bulletin of entomological research 96(1): 1–3. https://doi.org/10.1079/BER2005402 Hancock JM, Tautz D, Dover GA (1988) Evolution of the secondary structures and compensatory mutations of the ribosomal RNAs of Drosophila melanogaster. Molecular Biology and Evolution 5(4): 393–414. https://doi.org/10.1093/oxfordjournals.molbev.a040501 Harvey JA (2005) Factors affecting the evolution of development strategies in parasitoid wasps: the importance of functional constraints and incorporating complexity. Entomologia experimentalis et applicata 117(1): 1–13. https://doi.org/10.1111/j.1570-7458.2005.00348.x Harvey JA, Gols R, Strand MR (2009) Intrinsic competition and its effects on the survival and development of three species of endoparasitoid wasps. Entomologia Experimentalis et Applicata 130(3): 238–248. https://doi.org/10.1111/j.1570-7458.2008.00816.x Harvey JA, Poelman EH, Tanaka T (2013) Intrinsic inter-and intraspecific competition in parasitoid wasps. Annual review of entomology 58(1): 333–351. https://doi.org/10.1146/ annurev-ento-120811-153622 Oviposition timing by RCS parasitoids 1149 Herting B (1997) Section A. Host or Prey/Enemy. In: Hymenoptera. A catalogue of parasites and predators of terrestrial arthropods (Vol. 4). Commonwealth Agricultural Bureaux, Institute of Biological Control, 206 pp. Hood GR, Blankinship D, Doellman MM, Feder JL (2021) Temporal resource partitioning mitigates interspecific competition and promotes coexistence among insect parasites. Biological Reviews 96(5): 1969–1988. https://doi.org/10.1111/brv.12735 JMP Pro 18.0.1 (1989–2023) JMP Pro®, Version 18.0.1. SAS Institute Inc., Cary, NC, 1989– 2023. Kaneko S (1995) Frequent successful multiparasitism by five parasitoids attacking the scale insect Nipponaclerda biwakoensis. Population Ecology 37(2): 225–228. https://doi. org/10.1007/BF02515824 Kaneko S (2004) Within‐plant vertical distributions of the scale insect Nipponaclerda biwakoensis and its five parasitoids that exhibit frequent successful multiparasitism on the common reed. Entomological Science 7(4): 331–339. https://doi.org/10.1111/j.14798298.2004.00081.x Kaneko S (2005a) Seasonal population changes of five parasitoids attacking the scale insect Nipponaclerda biwakoensis on the common reed, with special reference to predation by wintering birds. Entomological Science 8(4): 323–329. https://doi.org/10.1111/j.14798298.2005.00131.x Kaneko S (2005b) Abundances of five parasitoids attacking the scale insect Nipponaclerda biwakoensis on morphologically changed reed shoots due to damage by a stem-boring caterpillar. Ecological research 20: 555–561. https://doi.org/10.1007/s11284-0050068-3 Klomp H, Teerink BJ (1978) The elimination of supernumerary larvae of the gregarious egg-parasitoid Trichogramma embryophagum [Hym.: Trichogrammatidae] in eggs of the host Ephestia kuehniella [Lep.: Pyralidae]. Entomophaga 23(2): 153–159. https://doi. org/10.1007/BF02371721 Knight IA, Wilson BE, Gill M, Aviles L, Cronin JT, Nyman JA, Schneider SA, Diaz R (2018) Invasion of Nipponaclerda biwakoensis (Hemiptera: Aclerdidae) and Phragmites australis dieback in southern Louisiana, USA. Biological Invasions 20: 2739–2744. https://doi. org/10.1007/s10530-018-1749-5 Knight IA, Cronin JT, Gill M, Nyman JA, Wilson BE, Diaz R (2020) Investigating Plant Phenotype, Salinity, and Infestation by the Roseau Cane Scale as Factors in the Die-Back of Phragmites australis in the Mississippi River Delta, USA. Wetlands 40: 1327–1337. https:// doi.org/10.1007/s13157-020-01307-3 Lopez LAA (2022) Ecology of the Roseau Cane Scale (Nipponaclerda biwakoensis, Hemiptera: Aclerdidae) in Coastal Louisiana. Master’s Thesis, Louisiana State University, Louisiana. https://doi.org/10.31390/gradschool_theses.5202 McConnell HS (1953) A Classification of the Coccid Family Aclerdidae (Coccoidea, Homoptera). Bulletin Maryland Agriculture Experiment Station A75: 1–121. Miller JC (1982) Life history of insect parasitoids involved in successful multiparasitism. Oecologia 54: 8–9. https://doi.org/10.1007/BF00541100 Violet Sparks et al. / Journal of Hymenoptera Research 98: 1133–1152 (2025) 1150 Noyes JS (2019) Universal Chalcidoidea Database. http://www.nhm.ac.uk/chalcidoids [Accessed on 07.08.2024] Noyes JS and Higashiura, Y (2020) The species of Astymachus Howard (Hymenoptera: Encyrtidae), potentially important parasitoids of Aclerdidae (Hemiptera: Coccoidea) associated with grasses (Poaceae), with descriptions of three new species. Journal of Natural History 54(9–12): 665–679. https://doi.org/10.1080/00222933.2020.1747654 Pruett KN, Knight IA, Kang I, Sparks VL, Diaz R (2025) Species-specific PCR primers provide insights into spatial distributions and interspecific interactions of roseau cane scale parasitoids in Louisiana. Biocontrol Science and Technology. https://doi.org/10.1080/0958315 7.2025.2563728 Ramsey III EW, Rangoonwala A (2017) Mapping the change of Phragmites australis live biomass in the lower Mississippi River Delta marshes (No. 2017-1098). US Geological Survey. https://doi.org/10.3133/ofr20171098 Schneider SA, Broadley HJ, Andersen JC, Elkinton JS, Hwang SY, Liu C, Noriyuki S, Park JS, Dao HT, Lewis ML, Gould JR (2022) An invasive population of Roseau Cane Scale in the Mississippi River Delta, USA originated from northeastern China. Biological Invasions 24: 2735–2755. https://doi.org/10.1007/s10530-022-02809-3 Smith MA, Woodley NE, Janzen DH, Hallwachs W, Hebert PD (2006) DNA barcodes reveal cryptic host-specificity within the presumed polyphagous members of a genus of parasitoid flies (Diptera: Tachinidae). Proceedings of the National Academy of Sciences, 103(10): 3657–3662. https://doi.org/10.1073/pnas.0511318103 Smith MA, Rodriguez JJ, Whitfield JB, Deans AR, Janzen DH, Hallwachs W, Hebert PD (2008) Extreme diversity of tropical parasitoid wasps exposed by iterative integration of natural history, DNA barcoding, morphology, and collections. Proceedings of the National Academy of Sciences 105(34): 12359–12364. https://doi.org/10.1073/ pnas.0805319105 Tilmon KJ, Danforth BN, Day WH, Hoffmann MP (2000) Determining parasitoid species composition in a host population: a molecular approach. Annals of the Entomological Society of America 93(3): 640–647. https://doi.org/10.1603/0013-8746(2000)093[0640:DP SCIA]2.0.CO;2 Traugott M, Zangerl P, Juen A, Schallhart N, Pfiffner L (2006) Detecting key parasitoids of lepidopteran pests by multiplex PCR. Biological Control 39(1): 39–46. https://doi. org/10.1016/j.biocontrol.2006.03.001 Wylie HG (1971) Observations on intraspecific larval competition in three hymenopterous parasites of fly puparia. The Canadian Entomologist 103(1): 137–142. https://doi. org/10.4039/Ent103137-1 Xia BC, Zhao YQ, Shen BY (1993) The ecological control of pests and diseases of reed in Shenyang beach. Journal of Plant Resources and Environment 2: 31–36. Zhou QS, Xi YQ, Yu F, Zhang X, Li XJ, Liu CL, Niu ZQ, Zhu CD, Qiao GX, Zhang YZ (2014) Application of DNA barcoding to the identification of Hymenoptera parasitoids from the soybean aphid (Aphis glycines) in China. Insect Science 21(3): 363–373. https:// doi.org/10.1111/1744-7917.12095 Oviposition timing by RCS parasitoids 1151 Supplementary material 1 Phragmites australis die-back in the Mississippi River Delta Authors: Violet Sparks, Ilgoo Kang, Morgan Carrere, Hannah Broadley, Rodrigo Diaz Data type: jpg Explanation note: Die-back of Phragmites australis surrounded by healthy stands within the Mississippi River Delta, Louisiana, USA. Copyright notice: This dataset is made available under the Open Database License (http://opendatacommons.org/licenses/odbl/1.0/). The Open Database License (ODbL) is a license agreement intended to allow users to freely share, modify, and use this Dataset while maintaining this same freedom for others, provided that the original source and author(s) are credited. Link: https://doi.org/10.3897/jhr.98.150721.suppl1 Supplementary material 2 Oil and natural gas infrastructure is dispersed throughout the Mississippi River Delta Authors: Violet Sparks, Morgan Carrere Data type: jpg Explanation note: An oil pumping station is surrounded by Phragmites australis within the Mississippi River Delta, Louisiana, USA. Copyright notice: This dataset is made available under the Open Database License (http://opendatacommons.org/licenses/odbl/1.0/). The Open Database License (ODbL) is a license agreement intended to allow users to freely share, modify, and use this Dataset while maintaining this same freedom for others, provided that the original source and author(s) are credited. Link: https://doi.org/10.3897/jhr.98.150721.suppl2 Violet Sparks et al. / Journal of Hymenoptera Research 98: 1133–1152 (2025) 1152 Supplementary material 3 Serial dilutions of stock parasitoid DNA for testing primer sensitivity Authors: Violet Sparks, Ilgoo Kang, Morgan Carrere, Hannah Broadley, Rodrigo Diaz Data type: png Explanation note: Results of primer specificity testing. Serial dilutions were performed so each 25 µl reaction contained 10, 5, 1, 0.5, 0.1, 0.05, 0.01, or 0 ng (negative control) of stock parasitoid DNA (lanes 1–8 from greatest to least DNA concentration). Copyright notice: This dataset is made available under the Open Database License (http://opendatacommons.org/licenses/odbl/1.0/). The Open Database License (ODbL) is a license agreement intended to allow users to freely share, modify, and use this Dataset while maintaining this same freedom for others, provided that the original source and author(s) are credited. Link: https://doi.org/10.3897/jhr.98.150721.suppl3