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Insect Molecular Genetics and Biotechnology

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Research Group: Molecular Genetics of Insects and Biotechnology

Research Staff

Luc Swevers, Research Director

Vassiliki Labropoulou, Senior Researcher

Kostas Iatrou, Emeritus Scientist

Panagiota Tsitoura, Postdoctoral Fellow

Aristeidis Zografidis, Postodoctoral Fellow

Nadia Sdralia, Graduate Student

Konstantinos Ioannidis, Graduate Student

Anna Kolliopoulou, Graduate Student

Dimitra Stefanou, Technical Specialist

Christos Meristoudis, Research Technician

Dimitris Kopanelis, Research Technician (retired)

Research Interests

1. Regulatory mechanisms controlling insect physiological functions.

(i)   Oogenesis in lepidopteran insects: a model differentiation program induced by ecdysteroid hormones.

(ii)    Mechanisms of immunosuppression in lepidopteran insects following parasitization by hymenopteran endoparasitoids: the role of interactions between proteins of hymenopteran endosymbiotic viruses and proteins of the hemocytes of the lepidopteran hosts.

(iii)  Mechanisms controlling olfactory function in the malaria mosquito vector Anopheles gambiae.

(iv)  Analysis of small RNA (miRNA, siRNA) pathways in lepidopteran insects.

(v)    Analysis of the antiviral immune response against RNA virus infection in lepidopteran insects: small RNAs and “cytokines”.

2. Molecular biology and genetic manipulation of insect viruses.

(i)     Viruses that express proteins toxic for the insect hosts.

(ii)   Genetically modified baculoviruses as vectors for insect genetic transformation.

(iii)  Genetically modified baculoviruses as vectors for gene therapy and cellular immunization applications.

(iv)  Genetically modified RNA viruses for delivery of RNAi triggers.

3. Functional genomics.

(i)     Systems for production of proteins of economic importance in lepidopteran insect and mammalian cell lines.

(ii)    High throughput screening systems for detection of bioactive substances (activators and inhibitors of pharmacological targets) in chemical libraries and collections of natural products (plants and microorganisms).

4. Insect pest management.

(i)     Cell-based assays for molting-accelerating compounds (ecdysone agonists): development, high-throughput screening and validation in larvicidal assays.

(ii)   Functional expression and characterization of detoxification enzymes of insecticides.

(iii)  Screening assays for identification of synergists/stabilizers of insecticides in natural plant extracts.

(iv)  Biotechnological methods for production of protein-based insecticides: delivery vehicles for toxins and dsRNA triggers.

(v)  Development of RNAi as tool for assessment of mechanism of insecticide resistance in insect larvae.

Participation in research projects

1) Bioinsecticides from Insect Parasitoids (BIP). 2001-2004. EU/FP5-QL Basic Research (RTD). Coordinator: C. Malva.

2) Baculovirus Artificial Chromosomes (BVACs) and technologies for gene therapy and continuous high-level expression of therapeutic proteins in insect production systems. 2004-2007. GSRT (Greece) – Research Consortia (EPAN). Coordinator: Κ. Iatrou.

3) Methods for overexpression and production of recombinant proteins from cloned genes and development of technologies and their application for high-throughput screening of new pharmacological effectors in plant extracts. 2004-2005. GSRT (Greece) - Praxi – Fase Α. Coordinator: Κ. Ιatrou.

4) Endocrine and paracrine control mechanisms of the gonadal ecdysteroid response in insects. 2004-2007. FWO – Vlaanderen G.0469.04 (Belgium). Coordinator: J. Vanden Broeck.

5) The olfactory odorant binding proteins of the malaria mosquito Anopheles gambiae as targets for vector control. 2004-2006. Bilateral S & T Cooperation (Greece-USA). Coordinator: Κ. Ιatrou.

6) Regulatory mechanisms that govern oogenesis in Lepidopteran insects: identification and functional characterization of factors that play a key role during successive stages of oogenesis in the silkworm Bombyx mori. 2006-2008. GSRT (Greece) – Basic Research (PENED). Coordinator: L. Swevers.

7) Hormonal disruptors for environmentally-safe control of the Mediterranean climbing cutworm Spodoptera littoralis. 2006-2008. Bilateral S & T Cooperation (Greece-Spain). Coordinator: L.  Swevers.

8) Insect cell-based high-throughput screening systems for the identification of compounds with ecdysteroid mimetic and order-specific insecticide activities in synthetic libraries. 2006-2008. Bilateral S & T Cooperation (Greece-Japan). Coordinator: L.  Swevers.

9) Key mechanisms of systemic RNA interference (RNAi) in insects. 2009-2012. FWO – Vlaanderen F 6/12 (Belgium). Coordinator: G. Smagghe.

10) ENAROMaTIC - European Network for Advanced Research on Olfaction for Malaria Transmitting Insect Control. 2008-2011. European Union (EC FP7-HEALTH-2007-B- Research Consortia). Coordinator: K. Iatrou.

11) Genomic and functional approach to understand insecticide resistance in insects and mites and development of applications for their control. 2012-2015. Ministry of education, Lifelong Learning and Religious Affairs (Greece), Operational Program Education & Lifelong Learning “Thalis”. Coordinator: J. Vontas.

12) Virus-induced mechanisms regulating RNAi in insects. 2013-2016. FWO – Vlaanderen G028013N (Belgium). Coordinator: G. Smagghe.

13) New Approaches for Insect Transformation. 2013-2014. Bilateral S & T Cooperation (Greece-Slovakia). Coordinator: L. Swevers.

Lab equipment

Insect cell culture: incubators, BIOWAVE bioreactor, laminar flow, inverted microscope, inverted fluorescence microscope, microcentrifuges with cooling, osmometer.

Insect culture incubator and maintenance room (for silkmoth).

Protein production: affinity chromatography, antibody purification, HPLC.

Biochemistry and molecular biology: DNA, RNA and protein electrophoresis, microcentrifuges, electroporation apparatuses, sonicator, microphotospectrometer (Nanodrop).

Screening systems for detection of bioactive molecules: fluorescence/absorbance plate reader (Galaxy), fluorescence/luminescence/absorption plate reader (Tecan).

Collaborations

Dr. J. Vontas, FORTH - Institute of Molecular Biology & Biotechnology, Heraklion, Crete.

Dr. A. Kourti, Department of Biotechnology, Agricultural University of Athens, Athens, Greece.

Dr. R. Matsas, Laboratory of Cellular and Molecular Neurobiology, Hellenic Pasteur Institute, Athens, Greece.

Dr. K. Kalantidis, FORTH - Institute of Molecular Biology & Biotechnology, Heraklion, Crete.

Dr. M. Konstantopoulou, Chemical Ecology and Natural Products, NCSR Demokritos, Athens, Greece.

Dr. G. Smagghe, Faculty of Bioscience Engineering, Ghent University, Belgium.

Dr. J. Vanden Broeck, Animal Physiology and Neurobiology, University of Leuven, Belgium.

Dr. Y. Nakagawa, Graduate School of Agriculture, Kyoto University, Japan.

Dr. D. Zitnan, Institute of Zoology, Slovak Academy of Sciences, Bratislava, Slovakia.

Dr. D. Martin, Institute of Evolutionary Biology, Barcelona, Spain.

Dr. J. Sun, College of Animal Science, South China Agricultural University, Guangzhou, People's Republic of China.

Recent Progress:

Regulatory mechanisms controlling insect physiological functions

The RNAi response in the silkmoth, Bombyx mori

RNA interference (RNAi) has recently been developed as a potent reverse genetics technique to analyse gene function with possible application in insect pest control (3,13).

In the silkmoth, B. mori (Lepidoptera), no potent RNAi response is induced following injection or feeding of dsRNA (1). This observation prompted us to evaluate factors that could contribute to the (lack of) RNAi efficiency in the silkmoth, such as:

Expression pattern of basic intracellular RNAi factors

Expression studies suggested that the absence of R2D2 expression, an essential co-factor of Dicer-2 and Ago-2, may play a role in the refractoriness of the systemic RNAi response in Bombyx (2). However, functional studies indicate that the intracellular RNAi machinery can work efficiently in the absence of R2D2 in silkmoth-derived Bm5 cells (5).

Expression of dsRNA-degrading enzymes

It was demonstrated that a non-specific DNA/RNA nuclease (“dsRNase”) has a broad expression in many different tissues and is capable both to degrade dsRNA intracellularly and to interfere with dsRNA-mediated gene silencing (4).

DsRNA as (non-specific) “pathogen-activated molecular pattern” (PAMP)

It was observed that injection of dsRNA into the hemolymph induces the expression of genes of the RNAi machinery (Dicer-2, Ago-2) and dsRNase in the midgut, while the expression of the innate immune Toll9-1 receptor was inhibited (6). Ectopic expression of Toll9-1 receptor in Bm5 cells was observed to modulate the response against the PAMPs dsRNA and lipopolysaccharide (LPS) with respect to the expression of the RNAi machinery and innate immunity genes (12).

Persistent RNA virus infection

It is hypothesized that persistent virus infection can severely affect the function of the RNAi machinery according to several different molecular mechanisms (8). Since the Daizo strain of Bombyx was found to be persistently infected with cytoplasmic polyhedrosis virus (CPV), characterized with a segmented dsRNA genome (Cypovirus, Reoviridae), it was decided to investigate whether the persistent infection could affect the immune response (including RNAi) against pathogenic infection of the same virus. Analysis by next-generation sequencing reveals a unique response to dsRNA virus infection in the silkmoth, with no overlap with the classical innate immune pathways triggered by bacteria or fungi (15). More specifically, transcriptome analysis reveals a complex response to pathogenic BmCPV infection that involves differential expression of genes belonging to categories such as physical barrier, immune response, proteolytic/metabolic enzymes, heat-shock proteins, hormonal signaling and uncharacterized proteins (15). Analysis of virus-derived small RNAs indicates a clear activation of the RNAi response against BmCPV infection, both in persistently and pathogenically infected larvae (15). The induction of the RNAi response, as indicated by the amounts of observed viral small RNAs, could be correlated with the severity of the viral infection (persistent versus pathogenic). Interestingly, earlier persistent infection did not seem to influence significantly the subsequent response to pathogenic infection (comparison with data from literature).

 

Overview of RNAi research in insects.

Publications:

1. Terenius, O., Papanicolaou, A., Garbutt, J.S., Eleftherianos, I., Huvenne, H., Sriramana, K., Albrechtsen, M., An, C., Aymeric, J.-L., Barthel, A., Bebas, P., Bitra, K., Bravo, A., Chevalier, F., Collinge, D.P., Crava, C.M., de Maagd, R.A., Duvic, B., Erlandson, M., Faye, I., Felföldi, G., Fujiwara, H., Futahashi, R., Gandhe, A.S., Gatehouse, H.S., Gatehouse, L.N., Giebultowicz, J., Gómez, I., Grimmelikhuijzen, C.J., Groot, A.T., Hauser, F., Heckel, D.G., Hegedus, D.D., Hrycaj, S., Huang, L., Hull, J., Iatrou, K., Iga, M., Kanost, M.R., Kotwica, J., Li, C., Li, J., Liu, J., Lundmark, M., Matsumoto, S., Meyering-Vos, M., Millichap, P.J., Monteiro, A., Mrinal, N., Niimi, T., Nowara, D., Ohnishi, A., Oostra, V., Ozaki, K., Papakonstantinou, M., Popadic, A., Rajam, M.V., Saenko, S., Simpson, R.M., Soberón, M., Strand, M.R., Tomita, S., Toprak, U., Wang, P., Wee, C.W., Whyard, S., Zhang, W., Nagaraju, J., ffrench-Constant, R.H., Herrero, S., Gordon, K., Swevers, L., and Smagghe, G. (2011). RNA interference in Lepidoptera: an overview of successful and unsuccessful studies and implications for experimental design. J. Insect Physiol. 57, 231-245.

2. Swevers, L., Liu, J., Huvenne, H., and Smagghe, G. (2011). Search for limiting factors in the RNAi pathway in silkmoth tissues and Bm5 cells: the RNA-binding proteins R2D2 and Translin. PLoS ONE 6:e20250.

3. Swevers, L. and Smagghe, G. (2012). Use of RNAi for control of insect crop pests. In:” Arthropod-Plant Interactions, Novel Insights and Approaches for IPM”, Progress in Biological Control, Volume 14. G. Smagghe & I. Diaz (Eds.), pp 177-197. Springer-Verlag, Dordrecht.

4. Liu, J., Swevers, L., Iatrou, K., Huvenne, H., and Smagghe, G. (2012). Bombyx mori DNA/RNA non-specific nuclease isoforms: expression in insect culture cells, subcellular localization and functional assays. J. Insect Physiol. 58, 1166-1176.

5. Kolliopoulou, A., and Swevers, L. (2013). Functional analysis of the RNAi response in ovary-derived silkmoth Bm5 cells. Insect Biochem. Mol. Biol. 42, 654-663.

6. Liu, J., Smagghe, G., and Swevers, L. (2013). Transcriptional response of BmToll9-1 and RNAi machinery genes to exogenous dsRNA in the midgut of Bombyx mori. J. Insect Physiol. 59, 646-654.

7. Kontogiannatos, D., Swevers, L., Maenaka, K., Park, E.Y., Iatrou, K., and Kourti, A. (2013). Functional Characterization of a Juvenile Hormone Esterase Related Gene in the Moth Sesamia nonagrioides through RNA Interference. PLoS ONE 8, e73834.

8. Swevers, L., Vanden Broeck, J., and Smagghe, G. (2013). The possible impact of persistent virus infection on the function of the RNAi machinery in insects: a hypothesis. Frontiers in Physiology 4, Article 319.

9. Swevers, L., Huvenne, H., Menschaert, G., Kontogiannatos, D., Kourti, A., Pauchet, Y., ffrench-Constant, R., and Smagghe, G. (2013). Colorado potato beetle (Coleoptera) gut transcriptome analysis: expression of RNA interference-related genes. Insect Mol. Biol. 22, 668-684.

10. Christiaens, O., Swevers, L., and Smagghe, G. (2014). DsRNA degradation in the pea aphid (Acyrthosiphon pisum) associated with lack of response in RNAi feeding and injection assay. Peptides 53 (2014) 307–314.

11. Swevers, L., Kolliopoulou, A., Li, Z., Daskalaki, M., Verret, F., Kalantidis, K., Smagghe, G., and Sun, J. (2014). Transfection of BmCPV genomic dsRNA in silkmoth-derived Bm5 cells: Stability and interactions with the core RNAi machinery. J. Insect Physiol. 64 (2014) 21–29.

12. Liu, J., Kolliopoulou, A., Smagghe, G., and Swevers, L. (2014). Modulation of the transcriptional response of innate immune and RNAi genes upon exposure to dsRNA and LPS in silkmoth-derived Bm5 cells overexpressing BmToll9-1 receptor. J. Insect Physiol. 66 (2014) 10–19.

13. Smagghe, G., and Swevers, L. (2014). Editorial overview: Pests and resistance – RNAi research in insects. Current Opinion in Insect Science 6, iv-v.

14. Kolliopoulou, A., and Swevers, L. (2014). Recent progress in RNAi research in Lepidoptera: intracellular machinery, antiviral immune response and prospects for insect pest control. Current Opinion in Insect Science 6, 28-34.

15) Kolliopoulou, A., Van Nieuwerburgh, F., Stravopodis, D.J., Deforce, D., Swevers, L., and Smagghe, G. (2015). Transcriptome analysis of Bombyx larval midgut during persistent and pathogenic cytoplasmic polyhedrosis virus infection. Submitted for publication.

Bio-insecticides for crop protection – wasp parasitism

Parasitism of lepidopteran insects by parasitoid hymenopterans

For the parasitization of Manduca sexta larvae by the endoparasitoid Cotesia congregata together with its endosymbiotic virus, Cotesia congregata Bracovirus (CcBV), the expression of viral proteins  is necessary for successful wasp parasitization. Bracoviruses (PDV), is a group deriving from Nudiviruses and are baculovirus-related type of viruses.

Innate immune responses caused by an endoparasitoid insect virus.

To study the mechanisms of immunosuppression in lepidopteran insects following parasitization by hymenopteran endoparasitoids, we have expressed CcBV proteins and investigated their interactions with host’s (Manduca sexta) haemocyte proteins. In fact, CcV1 protein has been shown to interfere with host cellular and humoral immune responses (1,2,3,4).

Differential inhibition of the Imd and Toll pathways in lepidopteran cells by CcBV ankyrin-repeat proteins.

One of the largest bracovirus (CcBV) protein families is the ankyrin-repeat protein family which is homologous to mammalian IκBa. This family includes nine members Ank1-9 and is thought to interfere with the host’s induced innate immune responses. Introducing transcriptional assays we found that CcBV Anks exhibit differential inhibition on the Bombyx mori, Rel/NFκB (Relish1-d2 and RelB), transcriptional activities. Subcellular localization of Anks suggests functional and spatial differences between the various members of this family (5).

 

  1. Lapointe R, Wilson R, Vilaplana L, O'Reilly DR, Falabella P, Douris V, Bernier-Cardou M, Pennacchio F, Iatrou K, Malva C, Olszewski JA. Expression of a Toxoneuron nigriceps polydnavirus-encoded protein causes apoptosis-like programmed cell death in lepidopteran insect cells. J Gen Virol. 2005, 86:963-71.
  2. Espagne E, Douris V, Lalmanach G, Provost B, Cattolico L, Lesobre J, Kurata S, Iatrou K, Drezen JM, Huguet E. A virus essential for insect host-parasite interactions encodes cystatins.J Virol. 2005, 79: 9765-76.
  3. Douris V, Swevers, L, Labropoulou V, Andronopoulou E, Georgoussi Z. and Iatrou  K. Stably transformed insect cell lines: tools for expression of secreted and membrane-anchored proteins and high throughput screening platforms for drug and insecticide discovery. Adv. in Virus Res., 2006, 68: 113-156.
  4. Labropoulou V, Douris V, Stefanou D, Magrioti C, Swevers L, and Iatrou K. Endoparasitoid wasp bracovirus-mediated inhibition of hemolin function and lepidopteran host immunosuppression. Cell. Microbiol. 2008, 10: 2118-28.
  5. Magkrioti C, Iatrou K. and Labropoulou V.  Differential inhibition of BmRelish1-dependent transcription in lepidopteran cells by bracovirus ankyrin-repeat proteins. Insect Biochem Mol Biol 2011, 12: 993-1002.

Molecular biology and genetic manipulation of insect viruses

Genetically modified baculoviruses as vectors for insect transformation and mammalian cell transduction

Baculoviruses, a group of insect viruses with large DNA genome, have several properties that make them very suitable for development as gene transduction vectors, such as large genome size, capability to enter cells by a non-specific mechanism and lack of toxicity. Efforts have been made to engineer baculoviruses as gene transduction vectors for both mammalian and insect cells.

Transduction vectors targeting mammalian cells

Baculovirus vectors were engineered with mammalian expression cassettes that could drive GFP and therapeutic protein expression in cell lines and primary Schwann cells (1,4). More specifically, baculovirus-mediated transduction of the L1 adhesion molecule could elicit physiological effects in Schwann cells in ex vivo assays, with potential therapeutic benefits (4). Besides vectors based on a wild-type baculovirus genome, alternative vectors with a deletion of the essential ie-1 gene were also developed which showed reduced endogenous viral gene expression in the transduced mammalian cells (3), which is considered an enhanced safety feature.

Transduction vectors for insect transformation

A similar strategy resulted in the construction of baculovirus vectors with insect expression cassettes. Baculovirus-based transduction vectors were used to induce RNAi in the lepidopteran Sesamia nonagrioides through the expression of RNA hairpin cassettes (5). More recently, research has focused on the generation of baculovirus vectors with deletions in essential genes, such as lef8, which encodes a subunit of the viral RNA polymerase (6), and ie1, which encodes the master regulator of the viral infection cycle. Because such viral particles (produced in engineered rescue lines) are deficient in viral amplification, they are less prone to cause toxic effects that could confound experimental results. Finally, baculovirus vectors that have incorporated the PiggyBac transposition system were generated (based on both wild-type and deficient baculovirus genomes) with as aim the development of a new easy applicable method of insect transformation. Hybrid PiggyBac-baculovirus vectors will be tested on the model lepidopteran Bombyx mori.

Transduction of Schwann cells by baculovirus vectors (ex vivo).

Publications:

1. Iatrou, K., and Swevers, L. (2005). Transformed lepidopteran cells expressing a protein of the silkmoth fat body display enhanced susceptibility to baculovirus infection and produce high titers of budded virus in serum-free media. J. Biotech. 120, 237-250.

2. Kenoutis, C., Efrose, R. C., Swevers, L., Lavdas, A.A., Gaitanou, M., Matsas, R., and Iatrou, K. (2006). Baculovirus-mediated gene delivery into Mammalian cells does not alter their transcriptional and differentiating potential but is accompanied by early viral gene expression. J Virol. 80, 4135-4146.

3. Efrose, R., Swevers, L., and Iatrou, K. (2010). Baculoviruses deficient in ie1 gene function abrogate viral gene expression in transduced mammalian cells. Virology 406, 293-301.

4. Lavdas, A.A., Efrose, R., Douris, V., Gaitanou, M., Papastefanaki, F., Swevers, L., Thomaidou, D., Iatrou, K., and Matsas, R. (2010). Soluble forms of the cell adhesion molecule L1 produced by insect and baculovirus-transduced mammalian cells enhance Schwann cell motility. J. Neurochem. 115, 1137-1149.

5. Kontogiannatos, D., Swevers, L., Maenaka, K., Park, E.Y., Iatrou, K., and Kourti, A. (2013). Functional Characterization of a Juvenile Hormone Esterase Related Gene in the Moth Sesamia nonagrioides through RNA Interference. PLoS ONE 8, e73834.

6. Ioannidis, K., Swevers, L., and Iatrou, K. (2015). The lef8 gene of BmNPV: effects of deletion and implications for gene transduction applications. Submitted for publication.

Insect Pest Management

Molting accelerating compounds (MACs) or ecdysone agonists

MACs act as insecticides by inducing a premature, lethal moult in the target insects. Because MACs can induce the moult at much lower concentrations than the natural hormone, 20-hydroxy-ecdysone (20E), and are not cleared efficiently from the insect body, the insect remains trapped in the moulting process and dies from desiccation and starvation.

A cell-based screening system for MACs was developed based on the activation of an ecdysone-responsive reporter gene (1) and was used to test a collection of > 200 dibenzoyl hydrazine compounds (2). The activity of the most active compounds in the screen was validated in toxicity tests on larvae of the cotton leafworm, Spodoptera littoralis (3).

Cell-based detection systems for ecdysone agonists or MACs were developed for lepidopteran (3), dipteran (4), coleopteran (5,6) and crustacean species (7,8). Recently, potent activity of dibenzoyl hydrazine compounds was observed in toxicity assays against malaria mosquito (Anopheles gambiae) larvae (9). Ecdysteroid screening systems were also employed to assess the endocrine disruption activity of the pollutant bisphenol A against arthropods (11).

General schedule for screening of ecdysone agonists or antagonists using transformed or transfected cell lines.

Publications:

1. Swevers, L., Kravariti, L., Ciolfi, S., Xenou-Kokoletsi, M., Ragoussis, N., Smagghe, G., Nakagawa, Y., Mazomenos, B., and Iatrou., K. (2004). A cell-based high-throughput screening system for detecting ecdysteroid agonists and antagonists in plant extracts and libraries of synthetic compounds. FASEB J. 18, 134-136. FASEB J. 10.1096/fj.03-0627fje.

2. Wheelock, C.E., Nakagawa, Y., Harada, T., Oikawa, N., Akamatsu, M., Smagghe, G., Stefanou, D., Iatrou, K., and Swevers, L. (2006). High-throughput screening of ecdysone agonists using a reporter gene assay followed by 3-D QSAR analysis of the molting hormonal activity. Bioorganic & Medicinal Chemistry 14, 1143-1159.

3. Soin, T., De Geyter, E., Mosallanejad, H., Iga, M., Martín, D., Ozaki, S., Kitsuda, S., Harada, T., Miyagawa, H., Stefanou, D., Kotzia, G., Efrose, R., Labropoulou, V., Geelen, D., Iatrou, K., Nakagawa, Y., Janssen, C.R., Smagghe, G., and Swevers, L. (2010a). Assessment of species specificity of molting accelerating compounds in Lepidoptera: comparison of activity between Bombyx mori and Spodoptera littoralis by in vitro reporter and in vivo toxicity assays. Pest Management Science 66, 526–535.

4. Soin, T., Swevers, L., Kotzia, G., Iatrou, K., Janssen, C.R., Rougé, P., Harada, T., Nakagawa, Y., and Smagghe G (2010b). Comparison of the activity of non-steroidal ecdysone agonists between dipteran and lepidopteran insects, using cell-based EcR reporter assays. Pest Manag. Sci. 66, 1215-1229.

5. Soin, T., Masatoshi, I., Swevers, L., Rougé, P., Janssen, C.R., and Smagghe, G. (2009). Towards Coleoptera-specific high-throughput screening systems for compounds with ecdysone activity: development of EcR reporter assays using weevil (Anthonomus grandis)-derived cell lines and in silico analysis of ligand binding to A. grandis ligand-binding pocket. Insect Biochem. Mol. Biol. 39, 523-534.

6. Ogura, T., Nakagawa, Y., Swevers, L., Smagghe, G., and Miyagawa, H. (2012). Quantitative evaluation of the molting hormone activity in coleopteran cells established from the Colorado potato beetle, Leptinotarsa decemlineata. Pesticide Biochemistry and Physiology 104, 1–8.

7. Verhaegen, Y., Parmentier, K., Swevers, L., Rougé, P., Soin, T., De Coen, W., Cooreman, K., and Smagghe, G. (2010). The brown shrimp (Crangon crangon L.) ecdysteroid receptor complex: Cloning, structural modeling of the ligand-binding domain and functional expression in an EcR-deficient Drosophila cell line. Gen. Comp. Endocr. 168, 415-423.

8. De Wilde, R., Swevers, L., Soin, T., Christiaens, O., Rougé, P., Cooreman, K., Janssen, C.R., and Smagghe, G. (2013). Cloning and functional analysis of the ecdysteroid receptor complex in the opossum shrimp Neomysis integer (Leach, 1814). Aquat. Toxicol. 130-131, 31-40.

9. Morou, E., Lirakis, M., Pavlidi, N., Zotti, M., Nakagawa, Y., Smagghe, G., Vontas, J., and Swevers, L. (2013). A new dibenzoylhydrazine with insecticidal activity against Anopheles mosquito larvae. Pest Manag. Sci. 69, 827–833.

10. Zotti, M.J., De Geyter, E., Swevers, L., Braz, A.S., Scott, L.P., Rougé, P., Coll, J., Grutzmacher, A.D., Lenardão, E.J., and Smagghe, G. (2013). A cell-based reporter assay for screening for EcR agonist/antagonist activity of natural ecdysteroids in Lepidoptera (Bm5) and Diptera (S2) cell cultures, followed by modeling of ecdysteroid-EcR interactions and normal mode analysis. Pestic Biochem Physiol. 107, 309-320.

11. Kontogiannatos, D., Swevers, L., Zakasis, G., and Kourti, A. (2015). The molecular and physiological impact of bisphenol A in Sesamia nonagrioides (Lepidoptera: Noctuidae). Ecotoxicology (In Press).