# Response of a lepidopteran pest to xenobiotics: exploring transduction pathways by a transcriptomic approach

Maeva Giraudo

► To cite this version:

Maeva Giraudo. Response of a lepidopteran pest to xenobiotics: exploring transduction pathways by a transcriptomic approach. Life Sciences [q-bio]. Université Côte D'Azur, 2010. English. NNT: .tel-04447503

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Submitted on 8 Feb 2024

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UNIVERSITE DE NICE-SOPHIA ANTIPOLIS – UFR SCIENCES
Ecole Doctorale des Sciences de la Vie et de la Santé

THESE

Présentée pour obtenir le titre de

DOCTEUR en SCIENCES

De l'Université de Nice-Sophia Antipolis

Spécialité: Aspects Moléculaires et Cellulaires de la Biologie

Par

Maeva GIRAUDO

Réponse d'un lépidoptère ravageur polyphage aux xénobiotiques: exploration des mécanismes de transduction par une approche transcriptomique

Travail réalisé sous la co-direction du Dr René Feyereisen et du Dr Gaëlle Le Goff
Équipe Evolution et Spécificité des Interactions Multitrophiques
UMR 1301 INRA-CNRS-Université de Nice Sophia Antipolis

Soutenue publiquement en anglais le 22 juin 2010, devant un jury composé de:

Pr Richard FFRENCH-CONSTANT

Dr Jean-Marc PASCUSSI

Dr Gaëlle LE GOFF

Dr David PAURON

Dr René FEYEREISEN

University of Exeter, UK

INSERM, Montpellier

INRA, Sophia Antipolis

INRA, Sophia Antipolis

INRA, Sophia Antipolis

Rapporteur

Rapporteur

Examinatrice

Président

Directeur de thèse
<figure><img src="image_1.png" /></figure>
**Abstract**

Herbivorous insects have developed mechanisms to metabolically inactivate some of the potentially toxic xenobiotics that they ingest, such as plant allelochemicals and insecticides. These defence mechanisms involve mostly a cytochrome P450-based metabolic detoxification. A large portion of them is inducible by xenobiotics, and for vertebrates including humans, the nuclear receptors (or « xenosensors ») that control P450s induction are increasingly understood.

We have shown in *Drosophila* that only a small subset of P450s are inducible by xenobiotics and that the specific *Drosophila* CYP6A2 showed pattern of induction similar to that of mammalian CYP2 genes that are induced by PXR/CAR nuclear receptors. We have then extended our study to gene expression in *Spodoptera frugiperda* (Lepidoptera, Noctuidae) in response to xenobiotics. Transcriptional analysis with an oligonucleotide microarray of 10K genes revealed that the xenobiotic response in the midgut of this insect and in the derived Sf9 cells involves the regulation of many detoxification genes. The patterns of differentially transcribed genes were specific to each chemical tested, suggesting multiple, and partially overlapping transduction mechanisms. qRT-PCR analysis showed that the plant allelochemicals were the strongest inducers of a small number of P450s *in vivo*, when insecticides induced P450 expression manly *in vitro*. Specific focus on the effects of two hormone mimics, methoxyfenozide and methoprene, in Sf9 cells showed that both of these compounds induced cell cycle arrest. In order to identify the link between ingestion of the toxin and P450 gene induction, we have cloned SfHR96, a nuclear receptor ortholog of the vertebrate xenosensors CAR and PXR. We found that this receptor is not inducible by xenobiotics and is expressed constitutively in all tissues and developmental stages. Strategies to demonstrate the function of SfHR96 by inactivation in vitro by RNAi and in vivo with dominant negative mutants in transgenic silkworms have been initiated. We hypothesize that SfHR96 dimerizes with ultraspiracle (USP), the insect ortholog of vertebrate RXR.
<figure><img src="image_2.png" /></figure>
Remerciements

Les remerciements c'est un peu l'occasion de faire « ma vie mon œuvre » et surtout d'avoir une pensée pour tous ces gens à côté, autour ou plus loin qui ont d'une manière ou d'une autre participé au déroulement et à l'accomplissement de ce travail. Parce qu'une thèse ça s'écrit tout seul, mais ça se gère à plusieurs. Et vous avez été nombreux!

My first and foremost huge thank you will go to Big Chief aka Mr P450 aka Dr. René Feyereisen, the one and only. I do not know any word that could be powerful enough to express all my gratitude. You are by far the best PhD supervisor that anyone can dream of and I feel like the luckiest PhD student to have had the opportunity to work with you for almost 4 years. Even though it tends to be annoying to have a boss who knows everything about everything about science, now how am I supposed to find a new boss half as good as you are? Thanks a lot for trusting me, for your weird and crazy sense of humour, your kindness, your modesty and your infinite scientific knowledge. You make P450 rock!

J'aimerais maintenant adresser mes remerciements à la plus géniale des co-directrices de thèse, celle sans qui rien de tout cela n'aurait été possible, celle qui a su être plus qu'une chef, une amie, que je respecte profondément pour ses qualités humaines et scientifiques, Dr. Gaëlle Le Goff. Je n'aurais pas pu imaginer de meilleures conditions de thèse si je ne t'avais pas eue à mes côtés durant ces 4 années. Merci de m'avoir fait confiance, d'être une source de motivation et d'inspiration quotidienne, d'avoir mangé des trucs bizarres avec moi en Chine, de m'avoir soutenue quand j'étais au pays des sushis, merci pour tout. Sincèrement. Si j'arrive un jour à devenir ne serait-ce que la moitié de ce que tu es, j'aurai le sentiment d'avoir réussi ma vie. Et j'envie le ou la prochain(e) étudiant(e) qui aura la chance de t'avoir comme chef parce que c'est un privilège qui donnerait presque envie de refaire une thèse. (j'ai dit presque)

I would like to deeply thank Pr. Richard Ffrench-Constant, Professor at the University of Exeter for accepting to judge this work and be part of the jury on the big day. It is a real honour to have such a brilliant scientist to judge my work, I really appreciate it. Besides, I would like to thank you for being such a nice and passionate person and for the good times in Italy.

Mes remerciements s'adressent également au Dr. Jean-Marc Pascussi, Directeur de Recherches à l'INSERM de Montpellier, pour m'avoir fait l'honneur de juger ce travail. Je vous remercie sincèrement pour votre regard critique sur mon manuscrit et mon travail de thèse et pour m'avoir fait partager vos précieux conseils au tout début de l'aventure lors du comité de thèse.

J'aimerais remercier très chaleureusement le Pr. David Pauron, Directeur de Recherches à l'INRA de Sophia-Antipolis, pour avoir accepté d'être le président de mon jury. Merci pour ton esprit critique toujours juste et honnête et sache que je suis très honorée que tu aies accepté. Te côtoyer pendant ces 4 années a rendu mes pauses café et mon quotidien au labo plus qu'agréable.
Un merci particulier aux Dr. Christine Risso et Alain Devaux pour avoir été les moteurs de
mes choix par votre passion et votre dévotion, pour votre humilité, pour m'avoir permis
d'être là et pour être mes maitres Jedi qui ont su me transmettre la passion pour l'écotox
aquatique. J'espère avoir la chance un jour de pouvoir re-travailler avec vous.

Je remercie également David Rouquié pour m'avoir accompagnée de ses conseils avisés depuis
le début de l'aventure en faisant partie de mon comité de thèse.

Un grand arigato à l'équipe du Dr. Shinoda de Tsukuba au Japon pour m'avoir accueillie pendant 3 mois, pour m'avoir fait bénéficier de leurs compétences et de leurs connaissances scientifiques et humaines et m'avoir permis de découvrir un peu plus du quotidien à la japonaise. Merci d'avoir rendu ce séjour inoubliable et d'avoir mis un peu de zénitude dans mes valises.

Je voudrais remercier du fond du cœur la dreamteam GFI, l'élite de l'équipe de choc, ceux qui ont été mon quotidien pendant tout ce temps et qui m'ont apporté tout leur soutien. Je pense particulièrement à Pascaline, Fred, Alex, Josiane et à tous les membres de l'équipe qui ont réussi à me supporter au quotidien : Hideki, Thierry, Trang, Isabelle, Leslie, Olivia et Nathalie.

Je voudrais à présent remercier les gens d'à côté, ceux qui ont été un soutien moral, physique et parfois logistique au quotidien et sans qui je n'aurais jamais réussi à atteindre la ligne d'arrivée.

Je pense bien sûr à Alice Baudot, ma louloute, mon amie, ma copine. Celle qui a essuyé toutes mes galères tout en ayant les siennes, celle qui a eu les mots qu'il fallait au moment où il fallait, celle qui m'a permis de survivre dans un quotidien japonais pas toujours facile, celle qui m'a redonné confiance en moi, tout le temps, sans relâche, celle qui est sans conteste la plus chouette rencontre de ces 4 dernières années. Merci pour ton canapé. Merci pour tout. Je n'y serais jamais arrivé sans toi. Il me faudrait 4 pages de remerciements rien que pour toi ! J'en profite pour remercier celui qui partage ta vie, Marc Ciosi, qui du haut de son plus qu'1m90 a su gérer les conditions de crises, a toujours eu la main tendue avec son cœur dessus et décroche haut la main l'oscar du mec qui m'a le plus fait rire, même quand il s'entête à me contredire. Merci à vous deux, vous avez été mon plus fort soutien et mes meilleurs copains.

Je voudrais remercier de tout mon cœur ma choup'choup' blonde, celle qui m'a accompagnée en dehors comme au boulot, Melle Laetitia Paganelli. Partager ton quotidien a été un réel bonheur pendant ces quelques années. Merci d'avoir fait le chemin avec moi, d'avoir bu des bières avec moi, d'avoir la classe américaine, d'avoir été la meilleure infirmière pour le zoziaux à la patte cassée, de m'avoir soutenue et d'avoir été une amie hors compète ! J'aimerais tellement pouvoir t'apporter la moitié du soutien que tu m'as donné quand viendra ton tour.
Un immense merci à ma Gégé nationale (c'est toi la bas dans le noir ?), pour tout ce que tu es et tout ce que tu m'as apporté et ta contribution nationale à l'appellation d'origine contrôlée Sussabi'. Pour tous tes conseils si judicieux, ta bonne humeur communicative et sans égal, pour être celle sur qui on peut compter, celle qui fait rigoler, pour m'avoir souvent sorti la tête de l'eau, pour être une amie hors du commun et m'avoir aidée au quotidien, même exilée à la capitale.

Un gigantesque merci à tous les potos du labo, ceux qui m'ont donné une réelle bouffée d'oxygène pendant les pauses déjeuner et en dehors, et ont rendu mon quotidien au labo beaucoup plus funkyl: Anto, Nico Hardcore, Dr Waffle, Maëlle, Marc, Pierpoljak-Didier, Eric. Je vous kiffe. Heureusement qu'il y a encore des gens comme vous à l'INRA, sans quoi ça serait vachement moins drôle.

Ludo, toi, jeune renard des sables, je te remercie infiniment pour... tout ! Et bien plus encore.

Je tiens à remercier particulièrement mon Gérard trop loin pour m'avoir accompagnée tout au long de la rédaction, ton soutien sans faille, ton OLO réconfortant. Maintenant c'est mon tour de te soutenir, futur grand avocat de la poutre (le barreau c'est surfait).

Merci à toi, Laurent T., pour ton arrivée imprévue et plus qu'appréciable dans ma vie, et avoir ainsi été une source de soutien et de motivation inestimable dans ces ultimes semaines de rédaction. You rock my world, t'as vu ?

Merci à tous les collègues de boulot passés ou encore là qui ont rendu mon quotidien tellement plus agréable, merci à Claude et ses pauses Bounty, Manuella, David, Lolo, Benoit, Laury, Etienne, Alex, Sophie, Petite SanSan, Emilie, Anaïs, Nico P. et Cédric. Un grand merci à Seb, mon Brandon adoré, pour ces quelques mois de colocation de bureau je sais que tu ne seras pas là le jour J mais que le cœur y est.

J'adresse mes plus sincères remerciements à mes meilleurs amis, ceux qui m'ont toujours soutenue et qui ont toujours cru en moi. Merci infiniment à Peggy, Cyril, Alice (et Mayeul !), Baptiste, Gaëlle (et Ambre !), Mary, Olive, Kévou et Fred. Merci d'être fiers de moi, de votre sincère amitié et d'être encore là, après toutes ces années. Merci à Eléonore, Ninie, Marion, Vanessa, Ophélie. A huge thank you to Ross for proof-reading my introduction. Un merci particulier à ma Chacha sur son îlot un peu trop loin, merci à toi et ta bordille d'être là malgré la distance. Vous manquez terriblement. Un merci en passant à Ben Mazuet, docteur à ses heures et chanteur pour les barmitzvah, ainsi qu'à tous ceux qui constituent la BO de ma thèse pour avoir accompagné mes oreilles pendant ces 4 années.

Un grand merci à Macbook Pro, aux expériences qui marchent, aux cellules pas contaminées, aux PCR niquels, aux clonages qui réussissent, aux séquençages pas foireux, et... aux insectes, mais bon pas trop non plus, hein, faut quand même pas abuser.
Je remercie sincèrement ceux qui espèrent que je vais « avoir mes partiels », qui me souhaitent de « bonnes vacances d'été » et de « trouver un cabinet bien placé », ceux qui me demandent « à quoi ça sert » ce que je fais, qui pensent que je vais être « étudiante toute ma vie » et qui m'ont souhaité bon courage pour écrirema thèse, parce que pour eux aussi « le mémoire de stage c'était dur, t'as vu ». Merci à vous, du fond du cœur, pour me faire apprécier d'autant plus ce que j'ai réalisé.

Je voudrais enfin remercier ma famille de survivants : ma maman, Agnès, Sandra et les enfants pour être ma raison de vivre et ma principale source de motivation au quotidien.

Un grand merci à vous tous pour votre soutien et m'avoir supportée pendant ces 4 années.

<figure><img src="image_3.png" /></figure>

Bon Maeva,
les insectes c'est
bren gentil,
mais quand est-ce
que tu nous inventes
la chantilly sans calories?

Un sincère quoi ?

gros bisous

<figure><img src="image_4.png" /></figure>
A mon Papa

Au club des survivants
<figure><img src="image_5.png" /></figure>
Table of contents

<table><tr><td>INTRODUCTION</td><td>1</td></tr><tr><td>1. Navigating the chemical space through plant-insect interactions</td><td>3</td></tr><tr><td>2. Cytochrome P450s</td><td>7</td></tr><tr><td>2.1 History</td><td>7</td></tr><tr><td>2.2 Nomenclature</td><td>7</td></tr><tr><td>2.3 Catalytic mechanisms</td><td>10</td></tr><tr><td>2.4 Structure</td><td>12</td></tr><tr><td>2.5 Functions</td><td>14</td></tr><tr><td>2.6 Regulation</td><td>15</td></tr><tr><td>3. P450s in insects</td><td>17</td></tr><tr><td>3.1 Overview</td><td>17</td></tr><tr><td>3.2 Substrate diversity</td><td>19</td></tr><tr><td>3.3 Functions of insect P450s</td><td>19</td></tr><tr><td>3.3.1 Metabolism of endogenous substrates</td><td>20</td></tr><tr><td>3.3.2 Metabolisms of xenobiotics</td><td>21</td></tr><tr><td>3.3.2.1 Plant allelochemicals</td><td>21</td></tr><tr><td>3.3.2.2 insect P450s role in plant-insect interactions</td><td>22</td></tr><tr><td>3.3.2.3 Insecticide resistance</td><td>24</td></tr><tr><td>3.4 Regulation of insect P450s</td><td>25</td></tr><tr><td>4. Nuclear Hormone Receptors</td><td>27</td></tr><tr><td>4.1 General properties</td><td>27</td></tr><tr><td>4.2 Structure</td><td>27</td></tr><tr><td>4.2.1 The A/B domain</td><td>28</td></tr><tr><td>4.2.2 The DBD domain</td><td>29</td></tr><tr><td>4.2.3 The D region</td><td>30</td></tr><tr><td>4.2.4 The LBD domain</td><td>30</td></tr><tr><td>4.2.5 The F region</td><td>31</td></tr><tr><td>4.3 Nomenclature</td><td>32</td></tr><tr><td>4.4 Evolution</td><td>34</td></tr><tr><td>4.5 NR binding and response elements</td><td>35</td></tr><tr><td>4.6 Nuclear receptors and P450s regulation: example of CAR and PXR</td><td>37</td></tr><tr><td>5. Nuclear receptors in insects</td><td>41</td></tr><tr><td>5.1. The Ecdysone (20E) receptor: EcR/USP</td><td>43</td></tr><tr><td>5.2 Insect nuclear receptor and xenobiotic metabolism: DHR96</td><td>46</td></tr></table>
<table>
  <tbody>
    <tr>
      <td><strong>OBJECTIVES</strong></td>
      <td>51</td>
    </tr>
    <tr>
      <td><strong>CHAPTER 1</strong></td>
      <td>57</td>
    </tr>
    <tr>
      <td>Article: Regulation of cytochrome P450 in Drosophila: genomic insights.</td>
      <td>57</td>
    </tr>
    <tr>
      <td><strong>CHAPTER 2</strong></td>
      <td>69</td>
    </tr>
    <tr>
      <td>Manuscript: Xenobiotic gene induction response in <em>Spodoptera frugiperda</em> midgut and Sf9 cells.</td>
      <td>69</td>
    </tr>
    <tr>
      <td><strong>CHAPTER 3</strong></td>
      <td>109</td>
    </tr>
    <tr>
      <td>Manuscript: Effects of hormone agonists on the Sf9 cells, cellular proliferation and cell cycle arrest.</td>
      <td>109</td>
    </tr>
    <tr>
      <td><strong>CHAPTER 4</strong></td>
      <td>145</td>
    </tr>
    <tr>
      <td>Manuscript: Molecular cloning and expression of nuclear receptors in <em>Spodoptera frugiperda</em> midgut and Sf9 cells.</td>
      <td>145</td>
    </tr>
    <tr>
      <td>1. RNAi impairment of SfHR96.</td>
      <td>181</td>
    </tr>
    <tr>
      <td>1.1 in vivo experiments</td>
      <td>181</td>
    </tr>
    <tr>
      <td>1.2 in vitro experiments</td>
      <td>183</td>
    </tr>
    <tr>
      <td>2. Bombyx mori transgenic construction.</td>
      <td>186</td>
    </tr>
    <tr>
      <td><strong>GENERAL CONCLUSION</strong></td>
      <td>191</td>
    </tr>
    <tr>
      <td><strong>REFERENCES</strong></td>
      <td>199</td>
    </tr>
  </tbody>
</table>
List of illustrations

List of tables

**Table 1** Updated number of P450 sequences identified as of August 2009.

**Table 2** Number of CYP genes found in each insect P450 clade.

**Table 3** Different sub-families of human nuclear receptors.

**Table 4** Chemical properties of the xenobiotic compounds used as inducers in *S. frugiperda* larvae and Sf9 cells.

**Table 5** Levels of P450 expression measured by qRT-PCR analysis in *S. frugiperda* larvae injected with dsRNA targeted against SfHR96 and exposed to xanthotoxin. Larvae injected with dsPAL from *A. thaliana* served as control.

List of figures

**Figure 1** Major mechanisms in plant allelochemicals-insect interactions and resistance.

**Figure 2** The detoxification process.

**Figure 3** P450 nomenclature.

**Figure 4** Single family tree of 107 P450 sequences.

**Figure 5** Catalytic cycle of cytochrome P450s.

**Figure 6** An example of a P450 system in the endoplasmic reticulum.

**Figure 7** Monooxygenation reactions catalysed by P450s.

**Figure 8** 3D structure of P450BMP protein.

**Figure 9** Examples of involvement of P450s in biological processes.

**Figure 10** Schematic representation of activation mechanisms of AhR.

**Figure 11** The four clades of insect P450 genes and relationships with major P450 families from other organisms.

**Figure 12** Hormonal regulation of growth and development in insects.

**Figure 13** Scheme of 20-hydroxyecdysone (20E) biosynthesis in Drosophila.

**Figure 14** Graphical representation of the nuclear receptor structure.
**Figure 15** Comparison of the crystal structures of the apo-retinoid X receptor-α (RXRα) ligand-binding domain (LBD) with the holo-RXRα LBD complexed with 9-cis retinoic acid).

**Figure 16** The different types of receptor binding to their response elements.

**Figure 17** PXR and CAR ligands.

**Figure 18** Activation of the mammalian xenosensors PXR and CAR.

**Figure 19** Classification of identified insect nuclear receptors.

**Figure 20** Chemical structure of the 20-hydroxyecdysone.

**Figure 21** Chemical structures of naturally occurring JHs.

**Figure 22** Phylogenetic tree representing human and drosophila nuclear receptors.

**Figure 23** The combinatorial problem in insect detoxification mechanism.

**Figure 24** Chemical structures of the 11 xenobiotic compounds used in *S. frugiperda* induction study.

**Figure 26** Preliminary results of gene expression levels of SfHR96 (A), CYP9A30 (B) and CYP9A31 (C) in Sf9 cells transfected with 3 concentrations (10, 25 and 50 nM) of siRNA targeted against SfHR96 (siHR96) and untransfected cells (ctrl) exposed to 50µM deltamethrin (delta), 250µM indole (Ind) or 25µM methoprene (Mtp).

**Figure 27** The GAL4/UAS system principle

**Figure 28** Restriction map of the plasmid DNA used as a vector for generating transgenic *B. mori*.
# INTRODUCTION
# 1. Navigating the chemical space through plant-insect interactions

The efficient identification of small molecules that modulate protein function *in vitro* and *in vivo* is at the heart of chemical biology and drug discovery (Koch et al., 2005). All these “biologically relevant” molecules represent an infinitely small fraction of what is called the “chemical space”. Identifying that chemical space – which encompasses all possible small organic molecules (Dobson, 2004) – would mean to navigate through an estimated number of 10⁶⁰ molecules that populate that space, a number that could as well be infinite and limited only by the chemist’s imagination (Lipinski and Hopkins, 2004). Given this, a thorough examination of all chemical space is practically impossible but can rather be focused on the biologically relevant part of this space for which limits could be defined by the specific binding of small molecules to biological molecules such as proteins, DNA or RNA (Lipinski and Hopkins, 2004).

Those biologically relevant molecules comprise all synthetic man-made compounds such as drugs, pesticides and industrial by-products as well as the plethora of natural compounds that are synthesised and used by living organisms. If we must look at the living matter of earth to identify these natural compounds producers, there is no doubt that green plants are a most voluminous and prolific sector (Zhu-Salzman et al., 2005). Plants synthesize a broad range of secondary metabolites that are essential for their growth and development but also for the defence against their natural enemies. These so-called phytotoxins or plant allelochemicals are very well documented by the immense diversity of plant secondary metabolism with some 200 000 compounds well known to pharmacologists (Hartmann, 2004) that account for a large part of the chemical space. About 25% of the drugs prescribed worldwide come from the plants (Rates, 2001). Recent developments in the field of constitutive plant allelochemicals have been extensively reviewed by (Wittstock and Gershenzon, 2002).

The plant metabolome therefore represents a well-documented part of the chemical space that we can navigate through in order to identify biologically relevant molecules. Such a task can be achieved by looking at other biological molecules that will interact with the plant metabolome, and thus intrinsically to the living organisms that harbour these molecules. When looking at plants, there is no doubt that herbivorous insects represent the organisms
that are the major interacting species. As Strong stated (Strong, 1988) "The overwhelming majority of all species interactions occur between herbivorous insects and plants, simply because these two groups comprise about half of the macroscopic species on Earth".

Plant and insects have co-existed on land for as long as 350 million years and have therefore developed a series of close relationships, some of them mutually beneficial like pollination, but the most common interaction however involves insect predation on plants and the defence of plants against the attack of herbivorous insects (Gatehouse, 2002). The process of co-evolution between plants and insects is widely believed to have generated much of the Earth's biological diversity (Rausher, 2001). Ehrlich and Raven in their famous paper "butterflies and plants: a study in co-evolution" have suggested back in 1964 that insects feeding on plants has been a determining factor in increasing species diversity in both herbivores and host plants. According to their co-evolution theory, evolution of plant chemical defences against their natural enemies is closely followed by biochemical adaptation in insect herbivores, and that newly evolved counter-resistance mechanism result in adaptive radiation of herbivore lineages (Wheat et al., 2007). Therefore, herbivorous insects impose natural selection that cause the evolution of a new plant resistance character and because most resistance characters reduce the survival of herbivorous insects, their evolution generates characters that circumvent the newly evolved plant resistance (Rausher, 2001).

The success of phytophagous insects as herbivores results therefore from their ability to successfully counteract the defensive strategies of their plant hosts (Gatehouse, 2002). They have developed for that matter multiple mechanisms to overcome plant chemical defences as explained in figure 1.
<figure><img src="image_6.png" /></figure>

**Figure 1:** Major mechanisms in plant allelochemicals-insect interactions (blue arrows) and resistance (red arrows). As a first step, insects can avoid (1) contact with the plant allelochemicals (green circle), mainly by behavioural strategies. After ingestion of the allelochemical, insects have the ability to excrete (2), sequester (3) or metabolize (4) the compound before excretion. As allelochemicals reaches its molecular target, a last mechanism involves the mutation of that target so that allelochemicals effects are reduced or eliminated. (from Despres et al., 2007)

Among these mechanisms, the metabolism of plant allelochemicals (number 4 in figure 1) is one of the major weapons that insects have evolved in their co-evolutionary arms race with plants. This detoxification process allows insects to metabolically inactivate the plant toxin that they ingest and rely on two main successive steps, as is shown in figure 2 below. First, in the phase I, or the biotransformation phase, a variety of enzymes acts to introduce reactive and polar groups into their substrates. One of the most common modifications is hydroxylation catalysed by the cytochrome P450-dependent mixed-function oxidase system. These enzyme complexes act to incorporate an atom of oxygen into nonactivated hydrocarbons, which can result in either the introduction of hydroxyl groups or N-, O- and S-dealkylation of substrates (Guengerich, 2001).
<figure><img src="image_7.png" /><figcaption>**Figure 2:** The detoxification process. Main enzymes involved in the phase I and phase II steps are indicated.</figcaption></figure>

The substrate can then be readily excreted or goes to the second step of the detoxification pathway: the Phase II or the conjugation phase. In this phase, activated xenobiotic metabolites are conjugated with charged species such as glutathione (GSH), sulfate, glycine, or glucuronic acid. These reactions are catalysed by a large group of broad-specificity transferases, which in combination can metabolize almost any hydrophobic compound that contains nucleophilic or electrophilic groups (Jakoby and Ziegler, 1990). The most important ones of this group are the glutathione S-transferases (GSTs). The addition of large anionic groups (such as GSH) detoxifies reactive electrophiles and produces more polar metabolites that cannot diffuse across membranes, and may, therefore, be actively transported. Conjugated forms of xenobiotics can then be recognized by specific membrane-associated transporters such as ABC transporters in phase III detoxification, resulting in their vacuolar sequestration or release into the apoplasmic space via exocytosis.

Insect detoxification mechanisms rely therefore on the overexpression of specific class of enzymes, including the cytochrome P450s (Despres et al., 2007; Li et al., 2007) which will be discussed in the following chapter, after an overview of general properties and functions of these enzymes in vertebrates.
## 2. Cytochrome P450s

### 2.1 History

Cytochromes P450s are probably the most common and well-characterised detoxification enzymes that constitute one of the largest gene family with representatives in virtually all living organisms from vertebrates to insects, except in the bacteria *Escherichia coli* (Werck-Reichhart and Feyereisen, 2000).

Klinbergerg (Klingenberg, 1958) and Garfinkel (Garfinkel, 1958) were the first to report the presence of a specific pigment in rat and pig hepatocytes. This pigment showed an absorption peak at 450nm and has been later characterised by Omura and Sato (Omura and Sato, 1964) as a heme-containing protein. These proteins were rapidly identified as proteins involved in the steroid metabolism (Estabrook et al., 1963). It was then established that hepatic microsomal oxidation of xenobiotics and the increase of metabolic activity in animals treated with drugs or xenobiotics, could be the result of cytochrome P450 activity and inducibility (Remmer and Merker, 1963).

### 2.2 Nomenclature

As increasing numbers of P450s were isolated and named by different groups, there became more and more confusion in the literature regarding the relatedness of these isoforms. To date, more than 10 000 sequences coding for P450s have been reported (D. R. Nelson, P450 Gene Superfamily Nomenclature Committee, University of Tennessee Health Science Center, Memphis, http://drnelson.uthsc.edu/CytochromeP450.html, august 2009). This huge number of sequences identified as P450 forced the international scientific community to adopt a standardised nomenclature system: a new nomenclature was proposed in 1987 (Nebert et al., 1987) and, although subject to revisions (Nebert and Nelson, 1991; Nebert et al., 1989; Nelson et al., 1993), it remains the preferred system (Nelson et al., 1996). This nomenclature is based on percentage of protein sequence identities: two P450s that are more than 40% identical in their sequence belong to the same family, and to the same sub-family when that percentage exceeds 55%. Sequences are
named as follow: the CYP prefix (CYtochrome P450) is followed by an Arabic numeral that designate the family (e.g. CYP1), a capital letter indicating the sub-family (e.g. CYP1A) and finally an Arabic numeral if there is more than one gene in a single sub-family (e.g. CYP1A1, CYP1A2...) (figure 3).

<figure><img src="image_8.png" /><figcaption>**Figure 3:** P450 nomenclature (from Feyereisen, 2005)</figcaption></figure>

From the latest update in august 2009, there are currently 977 families of P450 identified and 2519 sub-families (see table 1).

<table><caption>**Table 1:** updated number of P450 sequences identified as of August 2009</caption><thead><tr><th></th><th></th><th>CYP families</th><th>CYP sub-families</th></tr></thead><tbody><tr><td>Animals</td><td>3282</td><td>120</td><td>500</td></tr><tr><td>Insects</td><td>1675 (part of the animal total)</td><td>59</td><td>338</td></tr><tr><td>Animals (not insects)</td><td>1607</td><td>69</td><td>169</td></tr><tr><td>Plants</td><td>4266</td><td>126</td><td>464</td></tr><tr><td>Fungi</td><td>2570</td><td>459</td><td>1011</td></tr><tr><td>Protists</td><td>247</td><td>62</td><td>119</td></tr><tr><td>Bacteria</td><td>905</td><td>196</td><td>409</td></tr><tr><td>Archaea</td><td>22</td><td>12</td><td>14</td></tr><tr><td>Viruses</td><td>2</td><td>2</td><td>2</td></tr><tr><td><strong>Total</strong></td><td>11294</td><td>977</td><td>2519</td></tr></tbody></table>

As number of P450 families identified extended, another level of classification has been proposed based on grouping together larger families as “clans” as seen in figure 4. In this figure we can see an alignment of 107 P450 sequences, assembled by merging existing alignments. The plants have four main clans, the largest has been called the group A plant
P450s. The fungi have four clans and a few non-affiliated sequences. Animal's clans are named for a representative family as in the 2, 3, 4, and 7 clans, a unique location as in the mitochondrial clan, or for a specific organism as in the *Caenorhabditis elegans* clan. The clan number 3 has the mammalian 3 and 5 families, the insect 6 and 9 families, as well as the C. *elegans* 13 and 25 families (Nelson, 1999).

<figure><img src="image_9.png" /><figcaption>**Figure 4:** Single family tree of 107 P450 sequences (from http://drnelson.uthsc.edu/P450trees.html as of September 1998)</figcaption></figure>
## 2.3 Catalytic mechanisms

Cytochrome P450s catalyse a large range of chemical reactions many of which are monooxygenation reactions, whereby they catalyse the transfer of one atom of molecular oxygen to a substrate while reducing the other to water (Feyereisen, 2005). The overall reaction of P450 monooxygenase-mediated metabolism can be expressed as follows:

$$RH + O_2 + NADPH + H^+ \rightarrow ROH + H_2O + NADP^+$$

Where RH is the substrate. The different steps are represented in figure 5.

<figure><img src="image_10.png" /><figcaption>**Figure 5:** Catalytic cycle of cytochrome P450s (from Werck-Reichhart and Feyereisen, 2000).</figcaption></figure>

In the initial step (1), the heme protein in its oxidised ($Fe^{3+}$) form binds a given substrate. This fixation facilitates the formation of the P450-reductase complex and transfer of a single electron from the redox partner (2). The iron reduced to $Fe^{2+}$, then binds oxygen (3). A second electron reduction step occurs (4) before the complex reactins in which molecular oxygen is split and an atom of oxygen is inserted into the substrate, the other one being
reduced to water. The product is then released and the P450 goes back to its initial oxidised
form.

The monooxygenases reaction requires the intervention of a redox partner that is most of
the time a flavoprotein but that can differ according to the substrate, the organism or the
localisation. The majority of P450s in eukaryotes are located in the endoplasmic reticulum
and require the flavoprotein NADPH cytochrome P450 reductase for reducing equivalents
(figure 6). In addition to P450 and reductase, cytochrome b5 is sometimes needed,
depending upon the substrate and/or the P450 involved. Cytochrome b5 can be important in
donating the second electron from NADH to P450 or by allosterically regulating substrate
binding to P450 (Murataliev et al., 2008). Mitochondrial P450s require ferredoxin and an
NADPH ferredoxin reductase, and are therefore somewhat similar to the soluble P450
system found in bacteria.

<figure><img src="image_11.png" /><figcaption>**Figure 6:** an example of a P450 system in the endoplasmic reticulum. (From Despres et al., 2007)</figcaption></figure>

P450s can catalyse a large number of chemical reactions that have been ably reviewed in
Mansuy (1998) and are not only based on the oxygen atom transfer. Indeed, P450 also show
activity as oxidases, reductases, desaturases, isomerases, etc.... Figure 7 summarizes some of
the main monooxygenation reactions.
<figure><img src="image_12.png" /><figcaption>**Figure 7:** Monooxygenation reactions catalysed by P450s (from Mansuy, 1998)</figcaption></figure>

## 2.4 Structure

P450s are made of an apoprotein of about 43 to 60 kD, from which one of the cysteine residue is coordinated with the heme iron atom. Three-dimensional structures of P450s are difficult to obtain in eukaryotes, as these proteins are associated with the membrane of the endoplasmic reticulum or the inner mitochondrial membrane. However P450s are soluble in bacteria and allowed the determination of the first 3D structure back in 1985, P450cam (CYP101) from *Pseudomonas putida* (Poulos et al., 1986). It took several years before the structure of another P450 became available, that of P450BM-3 (CYP102), from *Bacillus megaterium* (Ravichandran et al., 1993). Since then, different P450s structure have been identified such as CYP120A1 from the cyanobacteria *Synechocystis* sp.(Kuhnel et al., 2008), the human CYP3A4 (Scott and Halpert, 2005) and CYP2C5 (Johnson et al., 2002). From the P450BM-3D structure, Graham and Peterson (1999) made a model of the secondary and tertiary structures of P450s as it can be seen in figure 8 (Graham and Peterson, 1999). Basically P450s are formed of two main domains: one rich in α-helix and the other one in β-sheets. The heme is located in the center of the α-domain, as part of the L helix that
contains the conserved cysteine involved in iron binding to the heme. Substrate fixation
occurs through a hydrophobic canal formed by the β-sheets rich domain.

<figure><img src="image_13.png" /><figcaption>**Figure 8:** 3D structure of P450BMP protein. The secondary structure of P450BMP is shown from the "distal" face with helices in green coils and strands of α-sheets in blue. The random coil is shown in orange. Those elements that can be readily identified from this perspective are labelled. The N- and C-termini are labelled with N, and C, respectively. (from Graham and Peterson, 1999)</figcaption></figure>

From the peptidic sequences alignment, we can point out that P450s are proteins of about
500 amino acids that can be identified by the following characteristic sequence with 4
conserved residues: FXXGXXXCXG. These 4 residues are located in the carboxyterminal
region the heme binding part, the cysteine allowing the thiol-ligand binding to the heme.
The N-terminal region of microsomal P450 is rich in hydrophobic AA and allows the fixation
of the protein to the membrane.
## 2.5 Functions

Virtually any moderately to highly hydrophobic organic chemical is likely, perhaps certain, to be a substrate for one or another CYP (Stegeman and Livingstone, 1998). This large variety of potential substrates and the numerous different reactions that P450 can catalyse underline the essential and polyvalent role of P450s in the organism. Figure 9 represents how diverse is the array of biological processes into which P450s may participate.

<figure><img src="image_14.png" /><figcaption>**Figure 9:** examples of involvement of P450s in biological processes. (from Stegeman and Livingstone, 1998)</figcaption></figure>

Basically, P450s physiological functions can be divided into two main roles: first they are also able to synthesise, activate or inactivate many endogenous compounds, some of which are toxic and some of which (e.g. steroids) are important regulatory molecules. For example, several P450s have a role in conversion of sterols to hormones, vitamin D and bile acids (Nebert and Russell, 2002). Second, they are known to be involved in the metabolism of exogenous chemicals and xenobiotics such as drugs, pollutants or natural products. From the 57 identified human P450s, at least 15 belonging essentially to the 1, 2 and 3 P450 families are involved in the detoxification of xenobiotics, including drugs (Gueguen et al., 2006). Indeed, 90% of drug metabolism is achieved by CYP activity with more than 2000 substrates
identified (Rendic, 2002). To cite a few examples, expression of the CYP1 family (including CYP1A1, 1A2 and 1B1) is induced by polycyclic aromatic hydrocarbons (PAH), such as those found in industrial incineration products, cigarette smoke, and charcoal-grilled food (Nebert and Russell, 2002; Shimada and Fujii-Kuriyama, 2004) and members of the CYP2B family are induced by the barbiturate Phenobarbital (PB) (Waxman and Azaroff, 1992) which has long been an archetypal model inducer of P450s. In addition, ethanol increases levels of CYP2E1 and peroxysome proliferators are able to induce the expression of CYP4A (Handschin and Meyer, 2003).

## 2.6 Regulation

Since P450s play key roles in biosynthetic and catabolic pathways of a variety of compounds, their expression must be highly regulated. Some P450s are expressed only in some tissues and specific cells within this tissue. Similarly, the expression pattern of a number of P450s is different in developmental stages and in females and males (Handschin and Meyer, 2003).

In most cases, induction of CYPs occurs by a process involving de novo RNA and protein synthesis. An alternative mechanism of CYP induction involves compounds that stabilise translation or inhibit the protein degradation pathway. For example, ethanol can induce CYP2E1 by such mechanisms leading to enhanced formation of reactive acetaminophen metabolites and enhanced hepatotoxicity (Woodcroft and Novak, 1998).

The induction of many CYPs occurs nonetheless by a similar mechanism, where ligand activation of key receptor transcription factors leads to increased transcription (Tompkins and Wallace, 2007). Such transcription factors include nuclear receptors that will be discussed later in this manuscript, as well as other receptors such as the Aryl Hydrocarbon Receptor (AhR), which will serve as an example to illustrate P450 regulation.

The molecular mechanisms governing the inducible expression of CYPs have been successfully elucidated for the CYP1 family (Fujii-Kuriyama and Mimura, 2005). Polyaromatic hydrocarbons such as 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) and halogenated aromatic
hydrocarbon (HAHs) such as Benzo(a)pyrene induce CYP1A and CYP1B through binding to the aryl-hydrocarbon receptor (AhR) (Honkakoski and Negishi, 2000; Nebert et al., 2004). The induction of CYP1A1 in particular is one AhR-dependent response that has been consistently observed in most vertebrate species, and it has been used as the model system to define the mechanism by which the AhR regulates gene expression (Denison and Nagy, 2003). The typical model of AhR action is presented in Figure 10. A ligand such as TCDD binds to AhR, which then associates with its dimerisation partner, AhR nuclear translocator (Arnt), and this ligand-bound complex then translocaes into the nucleus. AhR and Arnt belong to a family of transcription factors that contain basic-helix-loop-helix (bHLH) and Per-Arnt-Sim (PAS) domains. The AhR–Arnt heterodimer binds to xenobiotic response elements (XRE) such as “TNGCGTG” in the proximal and distal promoter regions of the CYP1A gene and induces its expression (Tompkins and Wallace, 2007; Whitlock, 1999).

<figure><img src="image_15.png" /><figcaption>**Figure 10:** Schematic representation of activation mechanisms of AhR.</figcaption></figure>

Transduction mechanisms of P450 induction are therefore well established in mammals. But how is the situation in insects? Insect P450s functions and regulation seem to be overall very similar to mammalian mechanisms and will be discussed in the following chapter.
## 3. P450s in insects

### 3.1 Overview

The first evidence of total P450 in insects was reported in 1967 by Ray (Ray, 1967) and the first insect P450 (CYP6A1) was cloned in 1989 by screening a cDNA expression library prepared from phenobarbital treated house flies with an antiserum raised against partially purified P450 (Feyereisen et al., 1989). Since then an increasing amount of studies have been carried out on insect P450s.

As shown in table 1, to date 59 CYP P450 families and 338 sub-families have been identified in insects, represented by 1675 sequences (http://drnelson.uthsc.edu/cytochromeP450.html). Given the tremendous diversity of insect species and the ever increasing number of fully sequenced insect genomes, there is no doubt that this number will exponentially increase in the next few years. For example, there are to date 90 fully identified sequences in the silkworm *Bombyx mori* genome, 85 active P450 genes in the fruit fly *Drosophila melanogaster* (Tijet et al., 2001), and about 143 sequences have been found in the genome of the red flour beetle *Tribolium castaneum* (Richards et al., 2008). The analysis of these available sequences revealed that insect P450s fall into 4 clades as shown in figure 11 and the summary of the number of P450 genes and P450 families found in each of these clades is presented in Table 2.

<table><caption>**Table 2:** Number of CYP genes found in each insect P450 clade (from Feyereisen, 2006, updated with insect P450 sequences as of March 2010)</caption><thead><tr><th>species</th><th>CYP2</th><th>mito</th><th>CYP3</th><th>CYP4</th><th>approx. total</th></tr></thead><tbody><tr><td><i>Drosophila melanogaster</i></td><td>7</td><td>11</td><td>36</td><td>32</td><td>86</td></tr><tr><td><i>Anopheles gambiae</i></td><td>10</td><td>9</td><td>40</td><td>46</td><td>105</td></tr><tr><td><i>Aedes aegypti</i></td><td>12</td><td>9</td><td>82</td><td>57</td><td>160</td></tr><tr><td><i>Bombyx mori</i></td><td>7</td><td>12</td><td>29</td><td>33</td><td>87</td></tr><tr><td><i>Apis mellifera</i></td><td>8</td><td>6</td><td>28</td><td>4</td><td>46</td></tr><tr><td><i>Nasonia vitripennis</i></td><td>7</td><td>7</td><td>48</td><td>30</td><td>92</td></tr><tr><td><i>Tribolium castaneum</i></td><td>8</td><td>9</td><td>72</td><td>45</td><td>134</td></tr><tr><td><i>Pediculus humanus</i></td><td>8</td><td>7</td><td>12</td><td>9</td><td>36</td></tr><tr><td><i>Acyrthosiphon pisum</i></td><td>10</td><td>8</td><td>23</td><td>23</td><td>64</td></tr></tbody></table>
<figure><img src="image_16.png" /><figcaption>**Figure 11:** the four clades of insect P450 genes and relationships with major P450 families from other organisms. The color code is: black, insects; red, vertebrates; brown, fungi; green, plants; blue, bacteria. Triangles represent families with large numbers of genes. Plant A-type P450 genes are found only in plants. CYP55 and CYP102 are P450foxy and P450BM3, fatty acid hydroxylases fused with a P450 reductase domain. CYP51 is the sterol 14a-demethylase, the only P450 ortholog with CYP74 (not shown) found in different phyla. (From Feyereisen, 2006)</figcaption></figure>
## 3.2 Substrate diversity

Functional identification of P450s can only be achieved through the biochemical characterization of its substrate selectivity. As the membrane-bound localisation of P450 make it difficult to study protein activity of individual P450 isolated from insect tissues, the expression of insect P450 cDNA in heterologous system has become a standard way of characterizing insect P450 proteins (Feyereisen, 2005). Insect P450s have therefore been expressed into different systems such as *Escherichia coli*, Baculovirus transfected lepidopteran cells, yeasts and Drosophila S2 cells. These techniques have allowed the identification of a large array of substrates that insect P450 can metabolize from endogenous compounds such as hormones to exogenous substrates such as plant allelochemicals and insecticides. For example CYP6A1 from the housefly expressed in *Escherichia coli* has been shown to metabolize the insecticide diazinon (Sabourault et al., 2001), some members of the CYP6AS family from the honeybee *Apis mellifera* expressed in baculovirus system were able to metabolize quercetin (Mao et al., 2009) or transfected S2 cells with CYP314A1 were reported to metabolize the insect hormone, ecdysone (Petryk et al., 2003).

## 3.3 Functions of insect P450s

From this ever increasing number of studies of functional characterization of insect P450s, we can divide their functional roles into two main categories: the metabolism of endogenous compounds, including signal molecules involved in growth, development and feeding, and the protection against xenobiotics, including resistance to pesticides and tolerance to plant toxins. Both of these aspects have been reviewed in Feyereisen (1999), Feyereisen (2005) and Schuler (1996).
### 3.3.1 Metabolism of endogenous substrates

Regarding the first aspect, it is now well known that P450s are involved in biosynthetic pathways of ecdysteroids and juvenile hormones, which are at the center stage of insect growth, development, and reproduction (Tijet et al., 2001) as it is depicted in figure 12 below. For example, it was found that Drosophila Halloween genes namely Phantom (Phm; CYP306A1), Disembodied (Dib; CYP302A1) and Shadow (Sad; CYP315A1) are involved into the biosynthesis pathway that transforms cholesterol into ecdysone. Shade (Shd; CYP314A1) catalyses the final hydroxylation step that transforms ecdysone into 20E (20-hydroxyecdysone) the principal insect molting hormone (Rewitz et al., 2006) (figure 13).

Another example of the role of insect P450s in the biosynthetic pathways of endogenous molecules is reported in the cockroach *Diploptera punctata*, where CYP4C7 and CYP15A1 have been shown to be involved in the biosynthesis and the regulation of the juvenile hormone III (JHIII) (Helvig et al., 2004; Sutherland et al., 1998).

<figure><img src="image_17.png" /><figcaption>**Figure 12:** hormonal regulation of growth and development in insects. Metamorphosis of insect larvae into adults is under the control of two main hormones: the juvenile hormone (JH) and the moulting hormone, 20-hydroxyecdysone (2OE). (from Dhadialla et al., 2005)</figcaption></figure>
<figure><img src="image_18.png" /></figure>

**Figure 13:** Scheme of 20-hydroxyecdysone (20E) biosynthesis in Drosophila. P450s involved in the last four hydroxylation steps leading to the formation of the moulting hormone 20-hydroxyecdysone are indicated. (from Gilbert and Warren, 2005)

### 3.3.2 Metabolisms of xenobiotics

#### 3.3.2.1 Plant allelochemicals

The metabolism of plant toxins by insects P450 has been extensively studied back in the BC era (Before Cloning, ©Feyereisen) as the measurement of the enzyme activity of the different reactions catalyzed by P450s, known as Mixed Function Oxidases (MFO) (reviewed in Brattsten, 1979), as well as the P450 protein content of insect microsomes (Yu, 1985) and NADPH reductase-dependent activities (Yu, 1986).

Based on the measurement of these MFOs, many plant compounds have been shown to induce P450-based oxidases activities as well as microsomal P450 content. For example plant terpenoids in *Spodoptera eridania* (Brattsten et al., 1984), *Peridroma saucia* (Yu et al., 1979) and *Spodoptera frugiperda* (Gunderson et al., 1986; Yu, 1982);
glucosinolates/glucosinolate derivatives in *S. frugiperda* (Yu, 1983), flavonoids in *Heliothis zea* (Neal, 1987) and *S. frugiperda* (Wheeler et al., 1993) and nicotine in *Manduca sexta* (Snyder et al., 1993).

These indirect methods are a good first step in the characterization of P450-based plant
allelochemicals metabolism in insects, however they do not give qualitative and quantitative
information on which type of P450 is involved and at which extent. As it has been discussed
in 3.2, the identification of individual P450 sequences has allowed the expression of their
cDNAs in heterologous systems in order to investigate substrate selectivity but has also led
to the correlation of mRNA induction levels with the previously identified protein levels and
enzyme activities. For example in *Manduca sexta*, the plant allelochemical 2-tridecanone is
an inducer of CYP4M1 and CYP4M3 (Snyder et al., 1995) and some P450s from the CYP9A
family are induced by indole 3-carbinol, xanthotoxin and 2-tridecanone (Stevens et al., 2000)
and in *Helicoverpa armigera*, the plant terpene gossypol is a potent inducer of CYP9A12 and
CYP9A17 (Zhou et al., 2009).

3.3.2.2 insect P450s role in plant-insect interactions

As it was discussed in the first chapter of this manuscript, plants and insects arms
race has led both parts to evolve sophisticated mechanisms to defend against each other,
and insect P450s play an important role in this co-evolutionary process. In order to
understand this, it is important to introduce the definition of host-plant specialization.

Herbivorous insects are classified into two groups: polyphagous species (generalists) which feed on a wide range of plant species, potentially encountering an array of toxic substances, and oligophagous species (specialists) feed on one or a small number of plant species and, thus, encounter a limited range of allelochemicals (Schuler, 1996). The majority of herbivorous insects belong to the second category and encounter large amounts of predictable chemistries, and have therefore characteristically high P450-based metabolism towards such chemistries (Stevens et al., 2000).
Probably the best story to illustrate this host plant specialization is the case of Papilio species detoxification of plant furanocoumarins through metabolism by P450s from the CYP6B family.

Furanocoumarins are plant allelochemicals found as two types in different plant species: the linear furanocoumarins (e.g. xanthotoxin and bergapten) and the angular type (e.g. angelicin). These substances are toxic to non-adapted herbivores (Berenbaum, 1990) by a photoreactivation mechanism which leads to the formation of DNA adducts. However, despite the toxicity of these substances, the black swallowtail *Papilio polyxenes* feeds almost exclusively on furanocoumarin containing plants. Early studies reported the ability of this species to detoxify xanthotoxin by a P450 monooxygenase-based mechanism (Bull et al., 1986; Cohen et al., 1989). In 1992, one specific P450 was cloned from xanthotoxin-treated larvae, the CYP6B1 (Cohen et al., 1992), and has been shown to be induced by and to metabolize linear but not angular furanocoumarins (Hung et al., 1995b; Ma et al., 1994; Prapaipong et al., 1994). However more recent studies using improved heterologous expression systems containing insect reductases reported that CYP6B1 was also able to metabolize angular furanocoumarins (angelicin) but at a lower level (Li et al., 2003; Wen et al., 2003).

A second CYP6B, CYP6B3, was isolated from *Papilio polyxenes* in 1995 and was shown to be inducible by linear as well as angular furanocoumarins (Hung et al., 1995a). Further CYP6B-like variants were then isolated from related Papilio species: *Papilio brevicauda* and *Papilio glaucus* (Cohen et al., 1992). The latter species is of particular interest as in contrast to *P. polyxenes*, it is a polyphagous species that encounter furanocoumarins only occasionally, but has however the ability to metabolize xanthotoxin through CYP6B metabolism. Moreover, six additional CYP6B have been isolated in *P. canadensis*, another generalist, and have shown to be induced by xanthotoxin (Li et al., 2002). Taken together, these results support the theory of Berenbaum et al. (1996) that high inducible activity toward a specific chemistry can serve as an adaptation of herbivores to a host containing this type of chemistry (Feyereisen, 1999).
3.3.2.3 Insecticide resistance

In addition to plant toxins, insect P450s can also metabolize a large array of insecticides compounds. Back in the early studies, MFO levels were found increased by several insecticides such as carbaryl (Gould and Hodgson, 1980), aldrin, heptachlor or biphenyl (Yu, 1985). P450-based metabolism of such substances has then been reported in some cases of heterologous expression of insect P450s. For example, Sabourault et al. found that over-expressed CYP6A1 from the house fly was able to metabolize diazinon (Sabourault et al., 2001) and CYP6B8 from *Helicoverpa zea* metabolises efficiently three insecticides, cypermethrin, diazinon and aldrin (Li et al., 2004).

This ability of P450 enzymes to metabolize insecticides plays an important role in insecticide resistance as it is now well established. P450 monooxygenase-mediated resistance is mainly due to increased detoxification than can result either from a change in the catalytic activity of the P450 involved and/or a change in the level of expression of the protein (Oppenoorth, 1984). Insecticide resistance associated with the overexpression of specific P450 enzymes have been largely reported in the literature. A good example to illustrate this phenomenon is the expression of Cyp6g1 in the DDT-resistant *Drosophila* strains. In 2001, Daborn and colleagues found that a single gene, Cyp6g1, was found overexpressed in DDT-resistant *Drosophila* strains and conferred cross-resistance to the neonicotinoid insecticide, imidacloprid (Daborn et al., 2001). Thanks to the availability of *Drosophila* complete genome, DNA microarrays were then constructed with all *Drosophila* P450 genes and hydridized with target cDNAs from susceptible and DDT-resistant strains of *Drosophila*. It was shown that Cyp6g1 was the one and only gene showing constitutive overexpression (Daborn et al., 2002). In addition, transgenic flies overexpressing CYP6g1 under the control of tubulin promotor in UAS/GAL4 expression system showed overexpression of only the CYP6g1 and cross-resistance to imidacloprid, acetamiprid and nitenpyram (Le Goff et al., 2003). Similar results were obtained along with 7 other P450s and showed that over-expression of CYP6g1 in transgenic flies increased survival to DDT, nitenpyram and dicyclanil (Daborn et al., 2007).
## 3.4 Regulation of insect P450s

Although less well studied than in vertebrates, insect P450 regulation has been explored in some cases and was found to be undergoing similar regulation mechanisms through transcription factors such as nuclear receptors, which will be discussed in the following chapter, as well as homologs to the AhR receptor as we will now see in the case of CYP6B family.

As it was discussed earlier, CYP6B inducibility by furanocoumarins has been extensively studied in *Papilio* species, in particular in the case of CYP6B1 and CYP6B3 that are inducible by linear (such as xanthotoxin) and angular (such as angelicin) furanocoumarins (see section 3.3.2.2). Analyses of the CYP6B1v3 promotor in transient Sf9 cell expression assay have indicated that xanthotoxin-inducible transcription of this gene requires the presence of a Xenobiotic Response Element to xanthotoxin (XRE-Xan) (Petersen et al., 2003) and further sequence searches have also identified a putative xenobiotic response element to the AhR (XRE-AhR) (Denison et al., 1988). With the cloning of additional CYP6B genes, a number of additional regulatory sequences have been recorded, which are conserved among the different CYP6B members and have similarities to known regulatory sequences (Li et al., 2002; Petersen et al., 2001). For example, in the promotor region of CYP6B1 (Li et al., 2002) and CYP6B4 (McDonnell et al., 2004), an overlapping region of 3 different response elements was identified: a response element to the Ecdysone Receptor (EcRE), the receptor of the major moulting hormone in insects (see figure 12), a XRE-Xan as well as an Antioxidant Response Element (ARE), that has been identified as important for antioxidant-inducible expression of mammalian phase II detoxification genes. In addition to these elements, both of the CYP6B1 and CYP6B4 promoters contain XRE-AhR elements (xenobiotic response element to the aryl hydrocarbon receptor) similar to those found in mammalian P450 promoters that are activated by binding to the activated AhR-ARNT complexes. The fact that CYP6B genes contain response elements similar to those in mammalian genes that are the target of the AhR regulatory cascade suggest that this mechanisms might be conserved in insects.

The bHLH family of proteins has 56 members in Drosophila, and 13 are bHLH-PAS proteins related to AhR and ARNT (Hahn, 2002). The protein most closely related to mammalian AhR
is spineless-aristapedia (Ss), which has been characterised as a transcription factor present in
the imaginal discs of developing Drosophila melanogaster larvae (Duncan et al., 1998). Ss
interacts with different bHLH-PAS proteins (such as sima, sim, trh, dys and cyc) but most
importantly to the ARNT ortholog called Tango (Tgo) that has been first identified as
regulating antennal and tarsal development but shows also xenosensing functions by
activating heterologous XRE-AhR elements (Emmons et al., 1999). In 2005, Brown and
colleagues expressed Ss with and without Tgo proteins in Sf9 cells with wild-type and mutant
CYP6B1:CAT constructs. They showed that Ss and Tgo increase basal expression of the wild-
type CY6B1 promotor in an additive manner, suggesting that AhR/ARNT regulatory
mechanisms of CYP1A1 in vertebrates might be conserved in insects (Brown et al., 2005).

bHLH-PAS proteins represent therefore one of the possible links between ingestion of the
toxin and induction of detoxification genes. However, as signaling mechanisms of the
detoxification machinery have been well studied in vertebrates, it was reported that other
types of transcription factors were also involved in the xenobiotic transduction signal. These
factors are nuclear hormone receptors (NRs) and will be discussed in the following chapter.
# 4. Nuclear Hormone Receptors

## 4.1 General properties

Nuclear hormone receptors (NRs) are one of the most abundant classes of transcriptional regulators in metazoans, in which they regulate functions as diverse as reproduction, differentiation, metabolism, metamorphosis and homeostasis (Escriva et al., 2000). This superfamily is present in all metazoans, and only in metazoans – no nuclear receptors have been found in the complete genome sequences currently available for plants, fungi, or unicellular eukaryotes (Laudet and Bonneton, 2005).

On the historical point of view, before the genes encoding nuclear receptors were cloned, the first NR was identified biochemically in the 1960s. Indeed, Elwood Jensen and his collaborators showed that estradiol was specifically retained in target cells of this hormone, leading to the discovery that its cellular activity is mediated by a specific high-affinity receptor. Subsequently, and only 20 years ago, the human glucocorticoid receptor (GR, NR3C1) was one of the first NRs to be cloned by Ron Evans and his colleagues together with the estrogen receptor (ER) cloned by the Pierre Chambon and Geoffrey Greene laboratories (Germain et al., 2006). Since then, NRs have become recognised as a superfamily of ligand-activated transcription factors, thus providing a direct link between signaling molecules that control these processes and transcriptional responses.

The NR research field has undergone very rapid development and covers areas ranging from structural and functional analyses to the molecular mechanisms of transcription regulation.

## 4.2 Structure

All NR proteins exhibit a characteristic modular structure that consists of five to six domains of homology (designated A to F, from the N-terminal to the C-terminal end) on the basis of regions of conserved sequence and function (Germain et al., 2006; Krust et al., 1986) (Figure 14). This common structural organization contains: a variable N-terminal region (A/B domain) containing the AF-1 transcription activator, a central well-conserved DNA binding domain (DBD, also termed C-domain), a non conserved hinge (D domain), a long and
moderately conserved ligand-binding domain (LBD, E domain) and eventually a C-terminal F domain (Escriva et al., 2000; Robinson-Rechavi and Laudet, 2003).

<figure><img src="image_19.png" /><figcaption>**Figure 14:** graphical representation of the nuclear receptor structure.</figcaption></figure>

### 4.2.1 The A/B domain

The poorly defined N-terminal A/B region contains a transcriptional activation function, referred to as activation function 1 (AF-1), that can operate autonomously (Germain et al., 2006). Deletion of the LBD of steroid hormone receptors results indeed in a protein that is constitutively active in reporter gene assays (Lavery and McEwan, 2005). This led to the identification of the hormone-independent AF1 in the A/B domain. In contrast to the other activation function (AF-2) located in the LBD of liganded NRs, AF-1 can act in a ligand-independent manner when placed outside of the receptor.
The actual three-dimensional structure of the functionally folded AF-1 domain, ideally in the
full-length receptor remains to be determined (Kumar and Litwack, 2009). The A/B domain
and its AF-1 factor are variable in terms of amino acid sequence and length, showing little, if
any, amino acid sequence homology, but has been shown to be important for steroid
hormone receptor-dependent gene regulation, and is a major site for post-translational
modifications such as phosphorylation and sumoylations (Lavery and McEwan, 2005;
McEwan, 2004).

4.2.2 The DBD domain

The central C region of the NRs is the DBD that is the most conserved amino acid
sequence among the members of the nuclear receptor superfamily. The DBD consists of two
zinc-finger motifs, each containing four highly conserved cysteine molecules coordinating
binding of a zinc atom. This results in the formation of a tertiary structure containing helices
that interact specifically with DNA sequences that are organized appropriately in what are
called response elements (RE) (Kumar and Thompson, 1999).

Residues in the first zinc-finger motif determine the specificity of the DNA recognition. It
contains the P-box, which is the highly conserved part in the first zinc finger between the last
two cysteines and determines the sequence specificity of the receptor-DNA binding RE
(Germain et al., 2006).

The second zinc-finger motif is involved in dimerisation. Indeed, nuclear receptors bind to
their REs as either homodimers or heterodimers.

Depending on the type of receptor, the C-terminal extension plays a role in sequence
recognition, dimerisation, or both. The DBD is also the target of post-translational
modifications and is involved furthermore in nuclear localisation and functions in
interactions with transcription factors and coactivators (Claessens and Gewirth, 2004).
4.2.3 The D region

The D region is highly variable in both length and primary among nuclear receptors.
As its name indicates, its main function is to serve a hinge between the DBD and the LBD. It
allows the DBD and LBD to rotate and adopt different types of conformation without
creating any steric hindrance troubles (Giguere, 1999). It also may contain some nuclear
localisation signals (NLS) and contain residues whose mutation abolishes interaction with
nuclear receptor corepressors (Aranda and Pascual, 2001).

4.2.4 The LBD domain

The LBD is a multifunctional domain that, in addition to the binding of ligand, mediates homo- and heterodimerisation, interaction with heat-shock proteins, ligand-dependent transcriptional activity, and in some cases, hormone reversible transcriptional repression (Aranda and Pascual, 2001). The C-terminal domain contains the ligand binding domain (LBD), whose overall architecture is well conserved between the various family members but which is nonetheless different enough to permit the high degree of ligand specificity characteristic of each member of the family. This domain harbours a ligand dependent activation function, called AF-2 that is crucially involved in transcriptional coregulator interaction (Laudet and Bonneton, 2005). The three-dimensional structure of the LBD has been determined for few unliganded (apo) and many liganded (holo) nuclear receptors, which has provided a good understanding of the mechanisms involved in ligand binding and transactivation (Gronemeyer et al., 2004).

Determination of the 3D structure of NRs shown that the LDBs of all nuclear receptors have
a common overall three-dimensional structure that is structured as a three-layered α-helical
antiparallel sandwich of 12 helices forming a hydrophobic pocket (see figure 15). This pocket
is guarded by a twelfth helix (H12), which forms a movable lid over the pocket and contains
residues that are crucial for the function of AF2. Many nuclear receptors are transcriptional
silencers in the absence of ligand (apo-receptor) as a result of interaction with intermediary
factors (i.e. corepressors). Upon ligand binding (holo-receptor), the ligand makes different
contacts with the amino acids residues, promoting a conformational change that closes the
H12 "lid" on the pocket and the corepressor complex dissociates. Thus the activation domain within the H12 (AF2) is able to interact with co-activators and promotes transcription of target genes (Bourguet et al., 2000; Escriva et al., 2000; Laudet and Bonneton, 2005).

<figure><img src="image_20.png" /></figure>

**Figure 15:** comparison of the crystal structures of the apo-retinoid X receptor-α (RXRα) ligand-binding domain (LBD) with the holo-RXRα LBD complexed with 9-cis retinoic acid. The coloured helices (purple in the apo-form; red in the holo form) are relocalised during the conformational change with the H12 helix that comes to close the « lid » of the ligand binding pocket. (from Gronemeyer et al., 2004)

4.2.5 The F region

Some of the nuclear receptors may have a C-terminal region called the F region, which is not conserved and is variable in length. For example the Retinoic Acid Receptor (RAR) and Hepatocyte Nuclear Factor 4 (HNF4) receptors have a region of respectively 36 and 102 aminoacids, when the Peroxysome Proliferator Activated Receptor (PPAR) or the Vitamin D Receptor (VDR) have no F region (Robinson-Rechavi and Laudet, 2003). To date, no exact function has been identified for this region.
4.3 Nomenclature

Given the plethora of names available for the same sequence, it has proved useful to
construct a nomenclature based on the nuclear receptor sequences using molecular
phylogeny.

The Nuclear Receptor Nomenclature Committee has elaborated a system to name NRs
based on the nomenclature system that was developed for cytochrome P450 by Nebert et
al. (1987). This system has proven to be convenient and flexible, allowing for the inclusion of
an ever-increasing number of nuclear receptor genes (Committee, 1999).

Each receptor is described by the letters NR (for “nuclear receptor”) and a three digit nomenclature: the subfamily to which a given receptor is indicated by Arabic numbers, groups by capital letters, and individual genes by Arabic numbers.

This system has proven to be flexible enough to integrate nuclear receptors from
invertebrates as well as sequences generated from genome projects for which no
pharmacological data are yet available.

This nomenclature allows the classification of nuclear receptors based on the sequence
identity of the DBD and the LBD with other members of the same family. In mammals they
are divided into 6 different sub-families (see table 3).

The first (Group I) family includes receptors of thyroid hormones (Thyroid Receptor Tr), RAR, VDR and PPAR as well as the pregnane X receptor (PXR) and the constitutive androstane receptor (CAR).

The second sub-family (Group II) includes retinoid X receptor (RXR), HNF4, testis receptors (TR2) and receptors involved in eye development (TLX and PNR). It is important to point out the particularly important role of the RXR receptor which heterodimerize with some of the group I receptors.

The third sub-family (Group III) contains steroid receptors such as the androgen receptor (AR), the glucocorticoid receptor (GR) and the estrogen receptor (ER).
<table><caption>**Table 3:** different sub-families of human nuclear receptors (adapted from Germain et al., 2006).</caption><thead><tr><th></th><th>Names</th><th>Nomenclature</th><th>Ligand</th></tr></thead><tbody><tr><td rowspan="15">Group I</td><td>TRα</td><td>NR1A1</td><td>Thyroid hormones</td></tr><tr><td>TRβ</td><td>NR1A2</td><td>Thyroid hormones</td></tr><tr><td>RARα</td><td>NR1B1</td><td>Retinoic acid</td></tr><tr><td>RARβ</td><td>NR1B2</td><td>Retinoic acid</td></tr><tr><td>RARγ</td><td>NR1B3</td><td>Retinoic acid</td></tr><tr><td>PPARα</td><td>NR1C1</td><td>Fatty acids, leukotriene E<sub>4</sub>, fibrates</td></tr><tr><td>PPARβ</td><td>NR1C2</td><td>Fatty acids</td></tr><tr><td>PPARγ</td><td>NR1C3</td><td>Fatty acids, prostaglandin J<sub>2</sub>, thiazolidinediones</td></tr><tr><td>Rev-erbα</td><td>NR1D1</td><td>Orphan</td></tr><tr><td>Rev-erbβ</td><td>NR1D2</td><td>Orphan</td></tr><tr><td>RORα</td><td>NR1F1</td><td>Cholesterol, cholesteryl sulfate</td></tr><tr><td>RORβ</td><td>NR1F2</td><td>Retinoic acid</td></tr><tr><td>RORγ</td><td>NR1F3</td><td>Orphan</td></tr><tr><td>LXRα</td><td>NR1H3</td><td>Oxysterols, T0901317, GW3965</td></tr><tr><td>LXRβ</td><td>NR1H2</td><td>Oxysterols, T0901317, GW3965</td></tr><tr><td>LXRγ</td><td>NR1H4</td><td>Bile acids, fexaramine</td></tr><tr><td>FXRα</td><td>NR1H5</td><td>Lanosterol</td></tr><tr><td>FXRβ</td><td>NR1H6</td><td>Vitamin D, 1,25-dihydroxyvitamin D<sub>3</sub></td></tr><tr><td>VDR</td><td>NR1I1</td><td>Xenobiotics, 16α-cyanopregnenolone</td></tr><tr><td>PXR</td><td>NR1I2</td><td>Xenobiotics, phenobarbital</td></tr><tr><td>CAR</td><td>NR1I3</td><td>Orphan</td></tr><tr><td>HNF4α</td><td>NR2A1</td><td>Orphan</td></tr><tr><td>HNF4γ</td><td>NR2A2</td><td>Orphan</td></tr><tr><td>RXRα</td><td>NR2B1</td><td>Retinoic acid</td></tr><tr><td>RXRβ</td><td>NR2B2</td><td>Retinoic acid</td></tr><tr><td>RXRγ</td><td>NR2B3</td><td>Retinoic acid</td></tr><tr><td rowspan="10">Group II</td><td>TR2</td><td>NR2C1</td><td>Orphan</td></tr><tr><td>TR4</td><td>NR2C2</td><td>Orphan</td></tr><tr><td>TLL</td><td>NR3E2</td><td>Orphan</td></tr><tr><td>PNR</td><td>NR2E3</td><td>Orphan</td></tr><tr><td>COUP-TF1</td><td>NR2F1</td><td>Orphan</td></tr><tr><td>COUP-TFII</td><td>NR2F2</td><td>Orphan</td></tr><tr><td>EAR2</td><td>NR2F6</td><td>Orphan</td></tr><tr><td>ERα</td><td>NR3A1</td><td>Estradiol-17β, tamoxifen, roloxifene</td></tr><tr><td>ERβ</td><td>NR3A2</td><td>Estradiol-17β, various synthetic compounds</td></tr><tr><td>ERR α</td><td>NR3B1</td><td>Orphan</td></tr><tr><td>ERR β</td><td>NR3B2</td><td>DES, 4-OH tamoxifen</td></tr><tr><td>ERRγ</td><td>NR3B3</td><td>DES, 4-OH tamoxifen</td></tr><tr><td rowspan="7">Group III</td><td>GR</td><td>NR3C1</td><td>Cortisol, dexamethasone, RU486</td></tr><tr><td>MR</td><td>NR3C2</td><td>Aldosterone, spiro lactone</td></tr><tr><td>PR</td><td>NR3C3</td><td>Progesterone, medroxyprogesterone acetate, RU486</td></tr><tr><td>AR</td><td>NR3C4</td><td>Testosterone, flutamide</td></tr><tr><td>NGFI-B</td><td>NR4A1</td><td>Orphan</td></tr><tr><td>NURR1</td><td>NR4A2</td><td>Orphan</td></tr><tr><td>NOR1</td><td>NR4A3</td><td>Orphan</td></tr><tr><td rowspan="2">Group V</td><td>SF1</td><td>NR5A1</td><td>Orphan</td></tr><tr><td>LRH-1</td><td>NR5A2</td><td>Orphan</td></tr><tr><td rowspan="2">Group VI</td><td>GCNF</td><td>NR6A1</td><td>Orphan</td></tr><tr><td>DAX-1</td><td>NR0B1</td><td>Orphan</td></tr><tr><td rowspan="2">Group 0</td><td>SHP</td><td>NR0B2</td><td>Orphan</td></tr><tr><td></td><td></td><td></td></tr></tbody></table>

The receptors that contain only one of the two functional domains (such as DAX/SHP which lack LBD domain) are artificially clustered in subfamily 0.

The number of NRs identified in sequenced genomes varies considerably. To date 48 members have been identified in humans (Robinson-Rechavi et al., 2001), 21 in the genome of the fruit fly *Drosophila melanogaster* (Adams et al., 2000), over 270 are found in *Caenorhabditis elegans* (Sluder and Maina, 2001), 49 in the mouse and 47 in the rat genome (Zhang et al., 2004).
4.4 Evolution

The structural diversity of the ligands contrasts with the conservation and mode of
action of their receptors. This and the large number of orphan receptors for which no ligand
have been identified, have prompted much speculation on the origins of the signaling
pathway.

Two main theories have been proposed: one says that orphan receptors have evolved as liganded molecules, which through gene duplication reached the present diversity. According to this model, orphan receptors would have lost their ability to bind ligands recently in evolution (Escriva et al., 2000).

The other theory supported by Escriva and colleagues suggest an alternative hypothesis in which ligand binding was acquired during NRs evolution (Escriva et al., 1997). As the presence of NRs for structurally different ligands within the same sub-family, together with the widespread distribution of orphan receptors in the phylogenetic tree, show no correlation between the evolutionary relationships of NRs and the nature of their ligands, this situation suggests independent gain of ligand binding capacity during nuclear receptor evolution (Laudet and Bonneton, 2005).

In their review of ligand binding evolution, Escriva and colleagues suggest that the first NR was an orphan receptor, whose descendants have secondarily, and several times, independently gained the ability to bind a ligand. Based on this theory and considering the known ligands and their respective affinities for their different NRs, they differentiate two types of ligand-binding acquisition: on the one hand, an ancestral orphan receptor recognizes a specific ligand with high affinity and keep that ligand specificity during evolution and on the other hand, an ancestral receptor that recognizes different types of ligands with low affinity and then either specialize for one ligand specificity or keep the ancestral status of low affinity for different related ligands (Escriva et al., 2000).

Taken together, their observations propose that nuclear receptors gained ligand-binding
capacity during evolution from an original orphan receptor.
4.5 NR binding and response elements

Nuclear receptors regulate transcription by binding to specific DNA sequences in
target genes known as hormone response elements (or HREs). These elements are located in
regulatory sequences normally present in the 5-flanking region of the target gene. Although
often the HREs are found relatively close to the core promoter, in some cases they are
present in enhancer regions several kilobases upstream of the transcriptional initiation site
(Aranda and Pascual, 2001). The analysis of a large number of HRE has revealed that 6bp
sequence constitutes the core motif that is recognised by nuclear receptors: 6'Pu-GGTCA
(Escriva et al., 2000). However there is a broad variety of different RE that have been
generated throughout evolution and that are selective for a given class of receptors (Laudet
and Bonneton, 2005).

Two consensus motifs have been identified: steroid hormone receptors typically bind to palindromes of the AGAACA sequence separated by three nucleotides, while the estrogen receptors recognize the consensus AGGTCA motif with the same configuration (Giguere, 1999).

These HREs directly reflect the mode of receptor binding, which can be as heterodimers,
homodimers, or monomers (Mangelsdorf and Evans, 1995). Most receptors bind as homo-
or heterodimers to HREs composed typically of two core hexameric motifs. For dimeric
HREs, the half-sites can be configured as palindromes (Pal), inverted palindromes (IPs),
everted or direct repeats (Ers or DRs). In contrast to steroid receptors that almost exclusively
recognize palindromic elements, nonsteroidal receptors can bind to HREs with different
configurations (Aranda and Pascual, 2001). For monomeric HREs, a single half-site is
preceded by a 5-flanking A/T-rich sequence (Giguere, 1999). At present, only orphan
receptors are known to bind DNA with high affinity as monomers (Escriva et al., 2000) (figure
16).

In the case of heterodimers, the RXR is the promiscuous partner for different receptors. A number of receptors including retinoic acid receptors (RARs), thyroid hormone receptors (TRs) and some orphan receptors heterodimerize with RXR and bind to direct repeats (DR) of the core motif (Aranda and Pascual, 2001). Heterodimerisation among nuclear receptor
superfamily members allows for the fine-tuning of nuclear receptor action by using combinatorial sets of ligands. In this way, RXR ligands can be used to enhance various signaling pathways as demonstrated by the ability of RXR agonists to synergize with RAR ligands, including some RAR antagonists (Bourguet et al., 2000).

Regarding the phylogenetic relationships between NRs, there is a good correlation of the nuclear receptors-HRE binding characteristics with the evolutionary history. For example, all members of the subfamily II are able to form homodimers on direct repeat sequences, whereas all members of subfamily III can form homodimers on palindromic sequences (Laudet and Bonneton, 2005).

<figure><img src="image_21.png" /><figcaption>**Figure 16:** The different types of receptor binding to their response elements. Nuclear receptors can be grouped into 4 classes according to their ligand binding, DNA binding and dimerisation properties. Receptors can bind DNA as homodimers, such as steroid receptors, heterodimers with RXR partner, or monodimers. Homodimers recognize palindromic sequences or direct repeat sequences in the case of orphan homodimers. Heterodimers recognize different HRE arranged as direct, everted or inverted repeats. Monomers recognize their half-site sequence when it is preceded by a 5'-A/T rich sequence.</figcaption></figure>
4.6 Nuclear receptors and P450s regulation: example of CAR and PXR

It is now well established that the regulation of P450s is mainly transcriptional and
involves the activation of nuclear receptors. Indeed, several orphan receptors respond to
xenobiotics in the environment that includes foreign chemicals such as environmental
pollutants and prescription drugs. In response to xenobiotic compounds, these receptors
mediate transcription of a variety of detoxifying enzymes that are members of the
supergene family of cytochrome P450 molecules (Olefsky, 2001).

Progress in understanding the molecular mechanism of induction of P450 enzymes was
made recently when the important roles of the pregnane X receptor (PXR) (Kliewer et al.,
1998) and the constitutive androstane receptor (CAR) (Honkakoski et al., 1998) were
discovered. Both of these receptors belong to the nuclear receptor superfamily and have
been extensively studied for the last decade. They are known to bind an extremely diverse
array of molecules (figure 17) including endogenous compounds such as steroids and bile
acids. However, these two related receptors are most commonly considered to respond to a
wide range of potentially toxic foreign compounds, or xenobiotics, and act as sensors for
these lipophilic xenobiotics, hence their name as xenosensors (Moore et al., 2006).

PXR was first identified by the screening of an EST database derived from a liver cDNA library by Kliewer and his colleagues in 1998. Using a Gal4-PXR chimeric protein to perform an initial search for PXR activators, they found that synthetic pregnanes (C21 steroids) and both glucocorticoid agonists and antagonists were potent inducers of PXR activity. Based on the finding that PXR is best activated by pregnenolone and its derivatives, they proposed that the natural ligand for PXR is likely to be a pregnane; hence, the name pregnane X receptor (PXR). PXR was originally shown to regulate the expression of CYP3A isozymes by binding as a heterodimer with the 9-cis retinoic acid receptor RXR (NR2B1) to xenobiotic response elements located in the regulatory regions of these genes (Kliewer et al., 1998; Lehmann et al., 1998; Maglich et al., 2002).

CAR was originally identified through screening of a cDNA library with a degenerate
oligonucleotide based on a conserved region of the nuclear receptor DBDs. The search for a
ligand revealed that CAR is, in fact, a steroid receptor for androstenol and androstanol and
was therefore named Constitutive Androstane Receptor (CAR) (Giguere, 1999). Like PXR,
CAR regulates the expression of target genes by binding to xenobiotic response elements as
a heterodimer with 9-cis retinoic acid receptor and was originally demonstrated to regulate
CYP2B gene expression (Honkakoski et al., 1998). The CYP2Bs are of special interest with
regard to CAR due to the identification of phernobarbital-responsive elements (PBREM) in
the 5' regions of CYP2B genes followed by the elucidation of CAR as the receptor that is
activated following exposure to phenobarbital (Sueyoshi et al., 1999).

<figure><img src="image_22.png" /><figcaption>**Figure 17 :** PXR and CAR ligands. Examples of PXR ligands (dark and pale orange) and CAR ligands (pale orange only). Many CAR ligands are also PXR ligands (from Willson and Kliewer, 2002).</figcaption></figure>
Since then, both receptors have been extensively studied and are shown to regulate different P450s mainly from the CYP3A and CYP2B families as well as other family members such as CYP1A1 and CYP1A2 for PXR (Maglich et al., 2002) and CYP24 for CAR (Moreau et al., 2007). In addition, they have been shown to regulate the expression of other genes involved in the detoxification machinery, such as GSTs or UDP-glucuronyltransferases (UGTs) (Maglich et al., 2002). A good review of the different genes regulated by CAR and PXR can be found in Honkakoski and Negishi (2000), Handschin and Meyer (2003) and Willson and Kliewer (2002).

PXR and CAR show overlapping but distinct control of the phase II and phase III detoxification pathways. Overall, since CAR and PXR are activated by some of the same ligands and induce specific but overlapping sets of genes, they provide the cell with two overlapping and semi- redundant mechanisms for recognizing and eliminating toxicants (Kretschmer and Baldwin, 2005).

CAR and PXR differ in their cellular localisation and transactivation mechanisms as PXR is exclusively nuclear, when CAR is found in the cytosol in its inactive state and one crucial step in its activation is its translocalisation into the nucleus upon the effect of activators such as phenobarbital (Kawamoto et al., 1999). There is however recent reports suggesting a cytosolic localisation of PXR as well, which then undergoes the same nuclear translocalisation as CAR when activated (Timsit and Negishi, 2007).

CAR and PXR bind as heterodimeric complexes with the RXR receptor to response elements in the regulatory regions of the induced genes (see figure 18 for an overview of CAR and PXR activation). These response elements have been identified as core motifs named Phenobarbital Response Element Modules (PBREM) for CAR and Xenobiotic Response Element Modules (XREM) for PXR (Timsit and Negishi, 2007), which contain direct or everted repeats of hexanucleotides. A review of these different response element motifs can be found in Sueyoshi and Negishi (2001).

Although CAR and PXR response elements or DNA motifs in the regulatory regions of these genes are distinct—it has now been established that CAR can activate CYP3A genes via PXR response elements and that PXR can regulate CYP2B genes via the CAR or phenobarbital response element (Nebert and Russell, 2002).
This crosstalk between CAR and PXR involving overlapping set of induced genes as well as reciprocal recognition of the same response elements has resulted from an adaptive advantage for living organisms to increase their ability to detect and respond to a wide variety of xenobiotics (Pascussi et al., 2008).

<figure><img src="image_23.png" /><figcaption>**Figure 18** : Activation of the mammalian xenosensors PXR and CAR. After entering the cell, xenobiotics trigger cytoplasmic-nuclear translocation of CAR by and directly activate PXR in the nucleus. Subsequently, both PXR and CAR heterodimerize with RXR, bind to their respective response elements, and increase transcription of target genes. (from Handschin and Meyer, 2003)</figcaption></figure>

The regulation of detoxification genes such as P450s in vertebrates is thus well established,
involving a central role of CAR and PXR nuclear receptors. In insects however, the
mechanisms that link the ingestion of the toxin and the expression of detoxification enzymes
has received much less attention (Gatehouse, 2002). AhR-like mechanisms have been
reported in some species as it was discussed in section 3.4, however other nuclear receptor-
based mechanisms might be involved as it was recently suggested and will be discussed in
the following chapter.
5. Nuclear receptors in insects

As it was discussed earlier, nuclear receptors have been identified in all metazoans,
and regulate very diverse and essential functions such as reproduction, development and
metabolism (Escriva et al., 2000). In insects, nuclear receptors are involved in similar
important mechanisms as it will be discussed here, with an emphasis on metamorphosis
regulated by 20-hydroxyecdysone (20E) and Juvenile Hormone (JH) and the detoxification of
xenobiotics.

To date, 21 NRs have been identified in the fruit fly *Drosophila melanogaster* (Adams et al., 2000), 20 in the malaria mosquito *Anopheles gambiae* (Holt et al., 2002), 22 in the honey bee *Apis mellifera* (Velarde et al., 2006), 21 in the silkworm *Bombyx mori* genome (Cheng et al., 2008) 20 in the yellow fever mosquito *Aedes aegypti* (Cruz et al., 2009) and 21 in the red flour beetle *Tribolium castaneum* (Bonneton et al., 2008; Tan and Palli, 2008). Insect NRs have representatives in all 6 sub-families of NRs identified in vertebrates (see table 3) and are represented in figure 19.

The first sub-family of insect NRs (NR1) includes five receptors that are all directly involved
into the ecdysteroid pathway of *Drosophila melanogaster*, such as EcR, which will be
discussed later in this chapter. It also includes the only ortholog of mammalian CAR and PXR,
DHR96. This receptor has been shown to be induced by 20E and is expressed during the
onset of metamorphosis (Fisk and Thummel, 1995) and as its orthology with vertebrate
xenosensors suggests, has also been shown to respond to xenobiotics (King-Jones et al.,
2006). DHR96 functions will be reviewed later in this chapter.

The second family (NR2) contains the receptor Ultraspiracle (USP), which is the ortholog of vertebrate RXR and heterodimerize with EcR to form a fully functional ecdysone (20E) receptor (Yao et al., 1993; Yao et al., 1992). The requirement of heterodimerisation of EcR and USP has been found in all the species studied and it seems therefore that it forms the functional ecdysone receptor for all arthropods (Laudet and Bonneton, 2005).
<figure><img src="image_24.png" /><figcaption>**Figure 19:** classification of identified insect nuclear receptors (from Bonneton et al., 2008; Nakagawa and Henrich, 2009)</figcaption></figure>

The four remaining sub-families of insect NRs contain several nuclear receptors such as steroid receptors (e.g., ERR), an alternative binding partner for USP, DHR38, which plays a role in the ecdysone response (Sutherland et al., 1995), the best known and conserved FTZ-F1 receptor involved in embryogenesis and metamorphosis and atypical receptors from the NRO sub-family, which lacks one of the two DBD or LBD.
5.1. The Ecdysone (20E) receptor: EcR/USP

Holometabolous insects have a typical development cycle that goes through distinct
morphological changes from successive larval instars moults to pupation and
metamorphosis into the adult form. The growth and development from one stage to
another is regulated by two main hormones (see figure 12), the steroidal insect moulting
hormone, 20-hydroxyecdysone (figure 20), and the sesquiterpenoid Juvenile Hormone (JH),
of which there are at least six types (figure 21).

<figure><img src="image_25.png" /><figcaption>**Figure 20:** Chemical structure of the 20-hydroxyecdysone (from Dhadialla et al., 2005)</figcaption></figure>

<figure><img src="image_26.png" /><figcaption>**Figure 21:** Chemical structures of naturally occurring JHs. JHIII is the major JH in most insects when JHI and JHII are the principal ones in Lepidoptera. (from Dhadialla et al., 2005)</figcaption></figure>
Although the mode of action of JH is still not fully understood, 20E signal transduction is well
understood (Riddiford et al., 2001; Thummel, 2002). Steroid hormones exert their effects on
target tissues by activating their respective receptors, which are members of the nuclear
receptor superfamily (Mangelsdorf et al., 1995). The Drosophila ecdysteroid receptor is a
heterodimer of two such proteins: EcR (Ecdysone receptor, NR1H1) and the fly retinoid X
receptor (RXR) homolog, USP (Ultraspiracle, NR2B4). EcR was first identified in Drosophila
nearly 25 years after the structural identification of molting hormones (Koelle et al., 1991),
along with USP (Oro et al., 1990). In *Drosophila melanogaster*, the EcR gene encodes three
different isoforms, EcRA, EcRB1 and EcRB2 that contain unique amino termini but a common
carboxy-terminal region including the characteristic DBD and LBD and the expression of
these isoforms correlates with distinct cell- and tissue-specific responses to ecdysone during
metamorphosis (Talbot et al., 1993). Although Drosophila has only one USP isoform, two
different isoforms have been identified in several insect species that also show different
patterns of induction related to the developmental stage. For example, *Manduca sexta* USP-
1 is expressed during the intermoults periods whereas USP-2 is up-regulated at times of high
ecdysteroid titer during the larval moult, when USP-1 dissapears (Jindra et al., 1997). In
*Bombyx mori*, USP2 appeared also to be coordinate with the pulse of ecdysone during
metamorphosis, and it was suggested that this isoform of USP might be the actual
component of the EcR/USP complex (Cheng et al., 2008).

EcR/USP bind to various ecdysone response elements (EcREs) as a heterodimer to
transactivate several target genes (Yao et al., 1993). The hormone-receptor complex directly
induces a set of « early genes » including the Broad complex (BR-C) and E74 as well as early
puff genes (King-Jones and Thummel, 2005; Sullivan and Thummel, 2003; Thummel, 2002).
These genes encode transcription factors that transduce and amplify the ecdysone signal,
regulating the expression of large batteries of downstream « late genes » leading to the
appropriate stage- and tissue-specific biological responses such as tissue differentiation and
cell death (Thummel, 2002).

The mode of action of 20E through the activation of EcR/USP complex has thus been well
established. In addition, it has been found that USP could also heterodimerize with another
nuclear receptor, DHR38, and interfering with the EcR pathway, suggesting that other
receptor complex might be involved in the ecdysone response as well as the possibility of
USP to bind with alternative receptors (Sutherland et al., 1995). USP has also been suggested as the JH receptor for which the mode of action still remains to be elucidated.

The idea that USP could be the receptor of juvenile hormone (JH) or any of its derivatives is attractive because JH might directly modulate the activity of the EcR / USP complex (Nakagawa and Henrich, 2009). The docking model of JH to USP-LBD, proposed by Sasorith *et al.*, suggested the plausibility of JH binding within the LBD of USP in *Heliothis virescens* (Sasorith et al., 2002). However, the percentage of occupancy of the LBP by these ligands was shown to lie in the bottom range of values for classical nuclear receptors. Even though JH can bind to the USP and stimulate oligomerization of the USP *in vitro* (Jones and Sharp, 1997), further experimentation will be required to establish the function of JH as a ligand for USP *in vivo*. In particular, the Kd for the binding of JH to the Drosophila USP is rather high (approximately 500 nm) (Jones and Sharp, 1997), whereas the typical Kd values for the binding between hormones and NRs are often very low (in the nanomolar range). Further studies conformed the micromolar range of affinity constant of JHIII with USP (Jones et al., 2006) and questions are still raised about the identification of JH receptor.

Recent work has suggested the role of Met (Methoprene tolerant) protein as a potential JH receptor. This protein was cloned and found to be an ARNT-like member of the bHLH-PAS protein family involved in the AhR pathway (see section 3.4) (Ashok et al., 1998) and was found to bind JH with high affinity (Miura et al., 2005). However, the major contention to the hypothesis that Met could be the JH receptor in that the Met mutants are viable in *Drosophila melanogaster* and recent reporter assay using two hybrid system failed in showing an increase of reporter gene activity when JH analog methoprene was added as a ligand for Met homodimers and Met:EcR and Met:USP heterodimers (Bitra and Palli, 2009). The identification of Met as the JH receptor needs therefore further research.

Insects are therefore able to achieve a fine-tuning of their development cycle through high specific signal transduction involving nuclear receptors. As it was discussed earlier, insects are also able to detoxify xenobiotic compounds that they may encounter in their environment thanks to the expression of detoxification enzymes such as P450s. However, little is known about the mechanisms linking the ingestion of the toxin and the expression of the detoxification enzymes. Recent work has nonetheless suggested similar nuclear receptor
pathways than those for vertebrate CAR and PXR with the identification of the Drosophila
receptor, DHR96 as it will be discussed in the next chapter.

5.2 Insect nuclear receptor and xenobiotic metabolism: DHR96

In 1995, Fisk and Thummel have cloned and identify new members of the nuclear hormone receptors that are expressed during the onset of metamorphosis in *Drosophila melanogaster*. They isolated three new receptor genes, including one called DHR96. Sequence analysis showed that the DBD of DHR96 was 64% identical to the human VDR DBD and 52% to EcR (Fisk and Thummel, 1995).

They found that DHR96 encodes two transcripts of 2.8 and 0.6kb that are both expressed
throughout third larval instars and prepupal development. As a subset of 20E regulated
genes are coordinately induced at this time, they also checked for DHR96 induction by 20E
and found that expression of DHR96 was indeed induced by the presence of 20E.

Finally, they found that DHR96 was able to recognize a unique response element, Hsp27-
EcRE, which is also recognised by the EcR/USP receptor. Therefore they suggested that
DHR96 might be involved in 20E response and compete for the binding site of ecdysone
receptor but still no ligand was identified for this orphan receptor.

In 2000, Baker and colleagues studied the transactivation of EcR and the three previously
identified receptors by 37 different insect and plant ecdysteroids and juvenoids. They used
Gal4 chimeras in which the Gal4 DBD is fused to the LBD of the studied receptor, including
DHR96. They found that the measured level of transcript did not exceed those that were
treated with vehicle alone, failing to identify a potential ecdysteroid-derived ligand for
DHR96 (Baker et al., 2000).

No mutations have been yet reported in DHR96. However, its close phylogenetic relationship
to other ligand-regulated members of the sub-family 1, such as EcR, VDR, CAR and PXR (see
figure 22) indicates that this receptor might be regulated by a ligand (Escriva et al., 2000;
Laudet and Bonneton, 2005).
<figure><img src="image_27.png" /><figcaption>**Figure 22:** Phylogenetic tree representing human (blue) and drosophila (purple) nuclear receptor, with the particular cluster of genes coding for DHR96 and its closest human orthologs: CAR, PXR and VDR (from Enmark and Gustafsson, 2001)</figcaption></figure>
DHR96 is represented by an orthologous group of 3 genes in *C. elegans*: nrh-8, nhr-48 and daf-12. Nhr-8 mutant are sensitive to chloroquine indicating a role in xenobiotic pathway (Lindblom et al., 2001), whereas daf-12 controls entry into diapause by monitoring dietary cholesterol levels (Antebi et al., 2000). The function of DHR96 therefore indicates two non-exclusive models: as an insect xenobiotic sensor that control detoxification pathways or as a cholesterol sensor (King-Jones and Thummel, 2005).

As it is shown in figure 23, DHR96 is found clustered with both human CAR and PXR, for which xenosensing functions have been well established (see section 4.6). Such a function can therefore be assumed for DHR96. Palanker et al. have tested this hypothesis in drosophila embryos expressing GAL4-DHR96 treated with the PXR-selective agonist PCN and the CAR- selective agonists TCPOBOP and CITCO. They found that only CITCO gave reproducible, strong activation of the DHR96 ligand sensor, indicating that the activation status of the DHR96 LBD can be regulated by xenobiotic compounds in a manner similar to that of its vertebrate orthologs CAR (Palanker et al., 2006).

The same year, King-Jones and colleagues confirmed the xenosensing function of DHR96 and found that this receptor was involved in phenobarbital (PB)-regulated genes expression. They used DHR96 null mutant flies and found that although DHR96 mutant are viable and fertile, they show nonetheless an increased sensitivity to the sedative effect of PB and lower survival to chronic exposure to high doses of DDT. By using microarray analysis, they compared the pattern of gene expression in PB-treated wild type and DHR96 mutant flies and found that a large portion of genes were affected. However, some PB-responsive genes were superinduced in DHR96 mutant, including many detoxification genes such as P450s and the majority of PB-responsive genes were unaffected by the mutation. Overall, their study showed that DHR96 was involved into xenobiotic metabolism but also suggest that given the massive PB response in Drosophila, it is likely that additional nuclear receptors might feed into this pathway, such as bHLH-PAS proteins involved in the AhR pathway (King-Jones et al., 2006).

HR96 orthologs have been further identified in a small number of other insect species such as *Tribolium castaneum*, where RNA interference studies showed that injection of dsRNA targeted against TcHR96 in larvae had no effects on adult formation and female injected
with the same dsRNA produced viable offspring, suggesting that HR96 is not critical for
molting and metamorphosis (Tan and Palli, 2008). Cheng et al. have also cloned a HR96
ortholog in *Bombyx mori* and found that it was co-ordinately expressed with USP-2 in this
insect (Cheng et al., 2008). DHR96 was also found to be expressed through different
developmental stages along with USP in Drosophila (Sullivan and Thummel, 2003), which in
addition to the confirmation of Fisk's results showing the expression of DHR96 during the
onset of metamorphosis, also points out an interesting feature of DHR96 and USP. Indeed,
DHR96 belongs to the NR1 sub-family, which contains the ecdysone receptor EcR. This
receptor is known to heterodimerize with USP, the insect ortholog of RXR (see section 5.1).
In human, similar binding occurs between CAR and PXR and RXR (see section 4.6). We can
therefore suggest that USP might also be a heterodimer partner for HR96. However, such a
hypothesis has not yet been tested and molecular action of HR96 still remains to be
elucidated.

The most recent work on DHR96 has revealed that this receptor could act as a sterol sensor as its ortholog LXR and as it was suggested in *C. elegans*. Horner *et al*. found that DHR96 is able to bind cholesterol and is required for the coordinate transcriptional response of genes that are regulated by cholesterol and involved in cholesterol up- take, trafficking, and Storage. However, they did not fully identify cholesterol as a ligand for DHR96 as they were unable to detect changes in DHR96 activity in response to exogenous cholesterol and dietary factors (Horner *et al.*, 2009). In another study, they found that DHR96 is indeed indispensable for mediating the transcriptional response to dietary cholesterol and that it acts as a key regulator of genes involved in cholesterol metabolism (Bujold *et al.*).

HR96 seems therefore to exhibit two non-exclusive functions as a xenobiotic sensor and in
the cholesterol metabolism. However, ligands for this receptor are still to be identified and
potential heterodimer partners to be characterised. Moreover, the role of HR96 in the
induction of detoxification genes such as P450s, as it has been demonstrated for CAR and
PXR in mammals, has not yet been reported in insects. It would therefore be particularly
interesting to characterise the role of HR96 as an insect xenosensors, which could bind plant
allelochemicals and xenobiotic compounds and subsequently induce the expression of
P450s. This is one of the objectives of this thesis, as it will be detailed below.
**OBJECTIVES**
The overall objective of this work is to study xenobiotic detoxification mechanisms in the polyphagous lepidopteran pest, *Spodoptera frugiperda* through the identification of the signaling mechanisms that link the ingestion of the toxin to the expression of detoxification enzymes as depicted in figure 23.

<figure><img src="image_28.png" /></figure>

**Figure 23:** the combinatorial problem in insect detoxification mechanism. The chemical space represents a virtually infinite numbers of molecules, from which a small fraction such as insecticides and plant allelochemicals can bind to n nuclear receptors. The number of receptors is limited by the number of known receptors for fully sequenced and annotated insect genomes. These receptors after binding of the xenobiotic will induce typical effectors genes such as P450s for which the number is also limited by the number of their encoded genes present in the insect genome.

This will be done by 1) looking at the regulation of P450s expression in the model insect, *Drosophila melanogaster*, 2) extend this study to the patterns of gene expression and P450s in particular in *S. frugiperda* in response to xenobiotics, 3) studying one of the well characterised receptor-based signaling mechanisms involving EcR and USP in response to hormone agonists and 4) identify potential xenosensors in insects that are able to induce detoxification genes in response to a subset of inducing compounds.
1. In the first chapter, we will present our work on the model species *Drosophila melanogaster* where we have looked at the expression of the specific CYP6A2 cytochrome in response to various natural and synthetic compounds. We will compare this response to the CYP2 family of vertebrate and to the "CYPome" expression data that are available in the literature.

2. In the second chapter, we will present our results on the identification of patterns of gene induction in *S. frugiperda* midgut and in Sf9 cells exposed to different class of chemicals by a genome-wide analysis using a 10k genes microarray and quantitative analysis of P450 gene expression. By choosing a generalist herbivore as in insect model, we maximise our chance to cover many potential detoxification mechanisms. Indeed, in contrast to specialists, polyphagous, generalist herbivores face a more diverse array of chemistries that is directly proportional to the number of different host plants they feed on. Hence these insects have more complex P450-based detoxification mechanisms (Gatehouse, 2002). *S. frugiperda* is a major pest in agriculture in America and has been reported to feed on hosts from >25 plant families, making an excellent model organism to study the molecular mechanisms involved in the plant-insect interactions. Moreover, Sf9 cells derived from ovarian tissues of *S. frugiperda* represent a well-established cellular model that is used widely in thousands of studies, mainly as a heterologous expression system. Our approach will therefore allow a better understanding of *in vivo* mechanisms involved in insect response as well as the study of molecular mechanisms at a finer scale in an *in vitro* environment where we can control many factors. In order to cover as many potential detoxification mechanisms as possible, we will test the response of *S. frugiperda* to different classes of xenobiotic compounds that are presented in figure 24.

Methoprene and Methoxyfenozide are respectively a juvenile hormone analog and an ecdysone agonist. Deltamethrin is a commonly used pyrethrinoid and fipronil is an insecticide that targets GABA receptors. Quercetin is a flavonoid that is encountered in many Lepidoptera host plant species. Indole is a maize defensive compound and indole 3-carbinol a glucobracissin derivative. Xanthotoxin (or 8-methoxypsoralen) is a coumarin derivative that is a potent inducer in many insect species and 2-tridecanone is a secondary compound found in the trichome of wild tomatoes. Finally, the herbicide clofibrate and the barbiturate phenobarbital will be used as model inducers of P450s.
<table><caption>**Figure 24:** chemical structures of the 11 xenobiotic compounds used in *S. frugiperda* induction study.</caption><thead><tr><th>insecticides</th><th>allelochemicals</th><th>model inducer</th></tr></thead><tbody><tr><td><img src="methoprene_structure.png" alt="Chemical structure of methoprene"></td><td><img src="quercetin_structure.png" alt="Chemical structure of quercetin"></td><td><img src="phenobarbital_structure.png" alt="Chemical structure of phenobarbital"></td></tr><tr><td><img src="deltamethrin_structure.png" alt="Chemical structure of deltamethrin"></td><td><img src="indole_structure.png" alt="Chemical structure of indole"></td><td><img src="indole-3-carbinol_structure.png" alt="Chemical structure of indole-3-carbinol"></td></tr><tr><td><img src="methoxyfenozide_structure.png" alt="Chemical structure of methoxyfenozide"></td><td><img src="Indole-3-carbinol_structure.png" alt="Chemical structure of Indole-3-carbinol"></td><td><img src="clofibrate_structure.png" alt="Chemical structure of clofibrate"></td></tr><tr><td><img src="fipronil_structure.png" alt="Chemical structure of fipronil"></td><td><img src="xanthotoxin_structure.png" alt="Chemical structure of xanthotoxin"></td><td><img src="xanthotoxin_structure.png" alt="Chemical structure of xanthotoxin"></td></tr><tr><td></td><td><img src="2-tridecanone_structure.png" alt="Chemical structure of 2-tridecanone"></td><td><img src="2-tridecanone_structure.png" alt="Chemical structure of 2-tridecanone"></td></tr></tbody></table>

3. In the third chapter, we will present our work on the two most characterised nuclear hormone receptors that are essential for insect development and growth, EcR and USP. We will discuss our results on cellular and molecular effects of two hormone agonists, methoprene and methoxyfenozide, in *S. frugiperda* cells.

4. In the last chapter, we will present our results on the characterization of one specific nuclear receptor as a potential xenosensors in *S. frugiperda*, SfHR96 the ortholog of DHR96 in Drosophila.
CHAPTER 1

Article: Regulation of cytochrome P450 in Drosophila : genomic insights.

In press in *Pesticide Biochemistry and Physiology*
<figure><img src="image_29.png" /></figure>

As a first step in the study of detoxification mechanisms in insects, we have chosen to identify gene expression patterns of detoxification enzymes in the model organism, *Drosophila melanogaster*.

Because of its historical importance, large research community, and powerful research tools, as well as its modest genome size (~180 Mb), *Drosophila* was chosen as a model organism in 1990 under the auspices of the Human Genome Project (Adams et al., 2000). The genomic tools available in *Drosophila* now allow both the detailed study of single genes and global approaches on the whole family of the 90 identified P450s in this insect (Tijet et al., 2001). The function of some of these P450s in xenobiotic response of insects has been identified. CYP6A2 for example was one of the first to be characterised as inducible by phenobarbital, over-expressed in resistant strains of *Drosophila* (Brun et al., 1996) and able to metabolize insecticide compounds such as aldrin, heptachlor and diazinon (Dunkov et al., 1997).

Here we focus on one effector (target) gene, *Drosophila* CYP6A2, and we analyze the literature for inducers of this gene, and show that its pattern of induction is similar to that of mammalian CYP2 genes that are induced by PXR/CAR nuclear receptors. Our results are presented below.
<figure><img src="image_30.png" /></figure>

# Regulation of cytochrome P450 expression in *Drosophila*: Genomic insights

Maeva Giraudoª, G. Chandran Unnithanᵇ, Gaëlle Le Goffª, René Feyereisenª,⁺

## ARTICLE INFO

**Article history:**

Received 14 October 2008

Accepted 24 June 2009

Available online xxxx

**Keywords:**

P450 induction

Phenobarbital

DNA microarrays

Cyp6a2

Resistance

## ABSTRACT

Genomic tools such as the availability of the *Drosophila* genome sequence, the relative ease of stable transformation, and DNA microarrays have made the fruit fly a powerful model in insecticide toxicology research. We have used transgenic promoter-GFP constructs to document the detailed pattern of induced *Cyp6a2* gene expression in larval and adult *Drosophila* tissues. We also compared various insecticides and xenobiotics for their ability to induce this cytochrome P450 gene, and show that the pattern of *Cyp6a2* inducibility is comparable to that of vertebrate *CYP2B* genes, and different from that of vertebrate *CYP1A* genes, suggesting a degree of evolutionary conservation for the “phenobarbital-type” induction mechanism. Our results are compared to the increasingly diverse reports on P450 induction that can be gleaned from whole genome or from “detox” microarray experiments in *Drosophila*. These suggest that only a third of the genomic repertoire of CYP genes is inducible by xenobiotics, and that there are distinct subsets of inducers/induced genes, suggesting multiple xenobiotic transduction mechanisms. A relationship between induction and resistance is not supported by expression data from the literature. The relative abundance of expression data now available is in contrast to the paucity of studies on functional expression of P450 enzymes, and this remains a challenge for our understanding of the toxicokinetic aspects of insecticide action.

© 2009 Elsevier Inc. All rights reserved.

## 1. Introduction

Insecticidal action requires the presence of an active form of the insecticide at the target site at an effective concentration and for a sufficient time. The determinants of this bioavailability, transport, metabolism and sequestration, are therefore the toxicokinetic parameters [1] in insecticidal action. They have an equal importance to the molecular details of toxicodynamic, or mode of action, parameters. Efficacy, selective toxicity, and resistance can all be determined by either toxicokinetic or toxicodynamic differences between a sensitive species and a less sensitive species, a target or a non-target organism, and a susceptible or a resistant strain.

The increased use of *Drosophila* as a model insect in toxicology studies over the last decade is a logical consequence of the advantages it offers [2,3]. In the specific area of insecticide metabolism and resistance, the *Drosophila* genome sequence provided a first complete picture of cytochrome P450 diversity and abundance in insects with about 85 active CYP genes [4] while other insect genomes harbor more or fewer CYP genes [5]. The genomic tools available in *Drosophila* now allow both the detailed study of single genes and global approaches on the whole family of P450s in an insect.

Here, we provide examples of both approaches. First we focus on the *Cyp6a2* gene, a gene abundantly expressed in insecticide-

resistant strains [6–9]. The *CYP6A2* enzyme metabolizes organochlorine and organophosphorus insecticides [10] as well as dimethylbenzanthracene and aflatoxin B1 [11]. A mutant form of *CYP6A2* has been reported to metabolize DDT as well [12]. The detoxification function of *CYP6A2* is thus well established. *Cyp6a2* is also known to be inducible by barbiturates [7,9,10,13]. We report here the fine-scale mapping of the tissues in which the *Cyp6a2* gene is induced, and we report the pattern of *Cyp6a2* induction by various classes of chemicals using a screening approach with a transgenic GFP marker.

Secondly, we analyze the literature for induction of other CYP genes in *Drosophila* in order to place *Cyp6a2* in the context of the whole CYP family. The use of DNA microarrays allowed a genome-wide or “CYPome”-wide assessment of transcript abundance, and many important studies have been published since this field was reviewed [14]. Our analysis shows that only a third of the genomic repertoire of CYP genes is inducible by xenobiotics, and that there are distinct subsets of inducers/induced genes, suggesting multiple xenobiotic transduction mechanisms.

## 2. Materials and methods

### 2.1. *Transgenic Drosophila*

The 1.3 kb promoter fragment of the *Cyp6a2* gene [10] was cloned upstream of the Green Fluorescent Protein coding sequence in the

<footnote>ªUMR INRA - CNRS - Université de Nice Sophia Antipolis, 06903 Sophia Antipolis, France</footnote>

<footnote>ᵇDepartment of Entomology, University of Arizona, Tucson, AZ 85721, USA</footnote>

<footnote>ª Corresponding author.
E-mail address: rfeyer@sophia.inra.fr (R. Feyereisen).</footnote>
pCasper P-element vector and this was used to transform the w¹¹¹⁹ line of *Drosophila melanogaster*. The P[w+–1.3-*Cyp6a2*-GFP]-transformed lines were screened for high inducibility of GFP fluorescence following phenobarbital treatment. A double X-chromosome transformant line 1E; 11B; Sco/SM6b was used for further study.

## 2.2. *CYP6A2*-GFP induction

Induction of *Cyp6a2*-GFP by phenobarbital and by the polychlorobiphenyl mixture Aroclor 1254 (PCB) in different tissues/organs in larva, pupa and adult of the transgenic stock 1E; 11B; Sco/SM6b was determined using confocal microscopy. Late first instar or early second instar larvae, and 1-day-old adults were fed for 3–5 days on Ward's *Drosophila* instant diet mixed with phenobarbital (1 g dry instant diet + 2.5 ml 0.4% phenobarbital sodium in distilled water). One-day-old flies were also treated with PCB, by contact for 4–5 days; for this, 15–20 flies were kept in 20 ml glass scintillation vials coated with 0.1 mg (3 µg/cm²) PCB dissolved in acetone. PCB-treated flies were also fed wet instant diet placed at the bottom of the vial. For controls, larvae adults were treated similarly but without phenobarbital sodium or PCB. Whole larvae after molting into second or third instar, adults, and freshly dissected organs from larvae and adults were mounted on microscope slide in 80% glycerin in PBS and were examined for GFP fluorescence using a laser scanning confocal microscope (BioRad 1024 confocal scanning head attached to a Nikon Optiphot 2 microscope with PlanApo objectives). Confocal images were processed, using Lasersharp software (BioRad, Hercules, CA) and Adobe Photoshop. Treatment of flies with other chemicals was either by contact (DDD: 0.1 mg/vial; DDE: 0.01 mg/vial; dicofol: 0.1 mg/vial; aldrin: 1 µg/vial; clofibrate: 10 µg/vial) or by incorporation in the media, i.e. ingestion (*trans*-stilbene oxide: 0.3%; limonene: 0.1%).

# 3. Results

## 3.1. *Tissue specificity of Cyp6a2-GFP induction*

*Cyp6a2*-GFP fluorescence was induced in many but not all the tissues/organs in treated larvae and adults (Table 1). In addition to this non-ubiquitous expression, there were also apparent quantitative differences among different tissues/organs. Yellow/orange auto-fluorescence of chitinous cuticle and cuticular lining of internal structures (trachea, crop, foregut, proventriculus/cardia, hindgut, rectum and salivary gland duct), food content (digestive system), or other inclusions (distal part of the Malpighian tubules, nephrocytes and pericardial cells) sometimes masked or modified the green fluorescence (GF). All muscles/muscle fibers in both larvae and adults showed GF, but in the flight muscles GF was less intense than in other muscular components. Heart also showed only weak GF. In adults, regions of the body covered with only the membranous cuticle showed uniform green fluorescence because of the GFP in hemolymph (hemocytes) and epidermis. The fat body and the entire intestinal wall were intensely green fluorescent, but not the gastric caecum. GFP appeared to be induced throughout the central and peripheral nervous system, and sensory cells and their axons; but GF was weak in the larval brain and ganglion. In adult brain and ganglion, GF was more intense in the neuropile and lamina of the optic lobes than in the neurons. The ring gland and ovary did not show detectable GF, however, accessory glands showed faint GF. In the newly formed pupa, produced from phenobarbital-fed larva, fat body and digestive system were green (Fig. 1L); other internal structures were not distinguishable in whole-mounted pupae examined. None of the tissues/organs in control larvae, pupae and adult flies showed green fluorescence (Fig. 1J, K and M, Fig. 2L–N).

<table><caption>**Table 1**
PCB/phenobarbital-induced GFP in various tissues/organs in larvae and adults.</caption><thead><tr><th>Organs/tissues</th><th>Larva<sup>a</sup></th><th>Adult</th></tr></thead><tbody><tr><td>Cuticle</td><td>–</td><td>–</td></tr><tr><td>Tormogen cells of bristles</td><td>–</td><td>+</td></tr><tr><td>Epidermis</td><td>+</td><td>+</td></tr><tr><td>Imaginal disk</td><td>–</td><td>na</td></tr><tr><td>Skeletal muscles</td><td>+</td><td>+</td></tr><tr><td>Flight muscles</td><td>na</td><td>+</td></tr><tr><td>Fat body</td><td>+</td><td>+</td></tr><tr><td>Heart</td><td>–<sup>b</sup></td><td>–<sup>b</sup></td></tr><tr><td>Pericardial cells and nephrocytes</td><td>–</td><td>+<sup>c</sup></td></tr><tr><td>Hemolymph (hemocytes)</td><td>–</td><td>+</td></tr><tr><td>Pharynx, esophagus, proventriculus/cardia</td><td>+</td><td>+</td></tr><tr><td>Midgut, hindgut, rectum</td><td>+</td><td>+</td></tr><tr><td>Caecum</td><td>–</td><td>–</td></tr><tr><td>Crop muscles</td><td>na</td><td>+</td></tr><tr><td>Salivary gland</td><td>+<sup>d</sup></td><td>+</td></tr><tr><td>Malpighian tubules</td><td>+<sup>e</sup></td><td>+<sup>e</sup></td></tr><tr><td>Tracheal cells/lining</td><td>+</td><td>–</td></tr><tr><td>Spiracular glands</td><td>+</td><td>na</td></tr><tr><td>Brain, thoracic ganglion, peripheral nerves</td><td>+</td><td>+</td></tr><tr><td>Ring gland</td><td>–</td><td>–</td></tr><tr><td>Sense cells/organs</td><td>+</td><td>+</td></tr><tr><td>Compound eye and ocellus</td><td>na</td><td>+</td></tr><tr><td>Ovary/eggs</td><td>na</td><td>–</td></tr><tr><td>Uterus</td><td>na</td><td>+</td></tr><tr><td>Accessory gland</td><td>na</td><td>–<sup>b</sup></td></tr><tr><td>Testis, vas deferens, ejaculatory duct, sperm pump</td><td>na</td><td>+</td></tr></tbody></table>

The results are expressed as presence or absence of green fluorescence and are not meant to be interpreted quantitatively, na, not applicable.

<sup>a</sup> Third instar larva.

<sup>b</sup> Only very weak green fluorescence (GF).

<sup>c</sup> Strong yellow/orange auto-fluorescence masked GF.

<sup>d</sup> GF only in the anterior region of the salivary gland in the larva.

<sup>e</sup> Weak GF in the distal region.

## 3.2. *Cyp6a2* induction by xenobiotics

The clear difference in green fluorescence between induced and non-induced insects allowed us to screen for the ability of various chemicals to induce *Cyp6a2*. Table 2 shows the response of the *Cyp6a2*-GFP transgenic line to treatment by various chemicals, compared to the induction pattern of vertebrate *CYP1A* and *CYP2B* genes [15–17]. Phenobarbital, pentobarbital, organochlorines (DDT and aldrin), *trans*-stilbene oxide and limonene were all inducers of *Cyp6a2* and these are known inducers of the *CYP2B* genes, but not of the *CYP1A* genes. In contrast, *Cyp6a2* was not inducible by β-naphthoflavone or 3-methylcholanthrene, known inducers of the *CYP1A* genes. The case of polychlorobiphenyls is particularly interesting, as a mixture of coplanar and non-coplanar PCBs (Aroclor 1254) induces all three types of genes, but pure isomers show a distinguishable pattern: 2,4,5,2',4',5'-HCB (non-coplanar) induces *Cyp6a2* and the *CYP2B* genes, but 3,4,5,3',4',5'-HCB (coplanar) only induces the *CYP1A* genes [18,19]. Ethanol and clofibrate, typical inducers of the vertebrate *CYP2E* and *CYP4A* genes, respectively, did not induce *Cyp6a2*.

## 3.3. From *Cyp6a2* induction to induction of the CYPome

*Cyp6a2* was initially reported to be phenobarbital (or barbital) inducible, and it is also regulated by ecdysone [20]. It has recently been shown to be also induced by caffeine [21]. Moreover, a number of DNA microarray studies have now identified *Cyp6a2* as being responsive to chemical treatments. We have therefore compared and evaluated the results of ten published studies from 17 datasets on xenobiotic treatment of *D. melanogaster* that used either custom DNA arrays ("detox chips" [22,23]) or whole [24–28] (and partial [29–31]) genome arrays (Table 3). These studies
<figure><img src="image_31.png" /><figcaption>Fig. 1. Phenobarbital-induced GFP in third instar larva and pupa. (A) Early third instar larva. (B) Brain-ganglion complex and anterior part of the digestive system. (C) Abdominal region showing body wall and internal organs. (D and E) anterior and posterior ends of the larva. (F) Anterior part of the salivary gland. (G) Heart and pericardial cells. (H) Anterior spiracle and spiracular glands. (I) Body wall showing muscles. (J and K) Anterior and posterior ends of control larva (without Phenobarbital). (L) Newly formed pupa, treated with Phenobarbital at larval stage. (M) Control pupa. Br, brain; Ed, epidermis; Es, esophagus; Fb, fat body; Gn, ganglion; He, heart; Hg, hindgut; Mg, midgut; Ms, muscles; Pv, proventriculus; Sbr, spiracular branches; So, sense organ; Sp, spiracle; Spg, spiracular glands; Tr, trachea.</figcaption></figure>

covered ten chemicals, with different treatment regimes (concentrations and exposure times), as well as different developmental stages (third instar larvae and adults). Despite the differences in techniques and statistical treatments, we feel that the consensus outcome of this comparison provides an interesting first glimpse of the patterns of xenobiotic response of the fruit fly. Negative results, i.e. chemical treatments that did not induce P450 genes were not considered, although these are also informative [22]. We included the studies on exposure to *Piper nigrum* extracts [30] because the main component piperamides are well known. We did not include the study on dietary shift in larvae (standard cornmeal to bananas) [32] that showed a slight induction of *Cyp9b2* and a moderate decrease in *Cyp6g1* and *Cyp4d2* expression. It is known that P450 genes are involved in the adaptation to diet in other *Drosophila* species, for instance *Cyp28a1* and *Cyp4d10* in *Drosophila mettleri* [33], and that the transcriptome is affected by starvation. Every diet should be considered as a collection of

inducers and inhibitors of various potencies and concentrations, but here we focus on identified xenobiotics.

The results of this survey show that between one and twelve CYP genes are affected by each chemical treatment, and that no more than a third (27 P450s) of the CYPome is responsive to xenobiotics. Different classes of compounds induce different CYP genes, and there is considerable overlap between some chemicals while others stand out (Fig. 3). For instance, paraquat (11 genes) and tunicamycin (7 genes) seem to induce a set of CYP genes with little overlap with the set induced by the other chemicals. Tunicamycin causes a stress at the level of the endoplasmic reticulum and paraquat causes stress through the generation of reactive oxygen species [26]. The CYP genes seen induced in those treatments are perhaps best described as stress-responsive. *Cyp6d4*, *Cyp28a5* and *Cyp4p1* are the common genes in that category. A broader overlap is seen in the response to the other chemicals, with 11 genes retained in the consensus for phenobarbital induction (from six
<figure><img src="image_32.png" /><figcaption>**Fig. 2.** Phenobarbital (Pheno)- or PCB-treated adults. (A) Female fly (Pheno). (B and C) Apex of proboscis (frontal view), surface and interior (PCB). (D) Brain, thoracic ganglion and anterior region of the digestive system (Pheno). (E) Part of midgut and hindgut and proximal region of Malpighian tubules (PCB). (F) Wing (PCB). (G) Abdominal wall showing tergesternal muscles and epidermis (PCB). (H) Hindgut and rectum (Pheno). (I) Abdominal fat body, heart and pericardial cells (Pheno). (J) Part of the salivary gland (Pheno). (K) Part of male reproductive system (PCB). (L-N) Control female, and parts of the digestive system. AcGI, Accessory gland; Br, brain; Cr, crop; Ejd, ejaculatory duct; Fb, fatbody; He, heart; Hg, hindgut; Mg, midgut; Mt, malpighian tubule; Mu, muscles; Pc, pericardial cells; Pr, proventriculus; Pt, pseudotrachea; Re, rectum; Rv, rectal valve; SeCe, sensory cells; SeNe, sensory nerve; Sg, salivary gland Te, testis; Vde, vas deferens. Scale bar:D, 40 µm; E,F, H, K, M and N, 100 µm; B, C, G and I, 50 µm; J, 10 µm.</figcaption></figure>

experimental conditions), 12 genes for piperonyl butoxide and 11 genes for caffeine. *P. nigrum* extracts, atrazine and pyrethrum induced fewer genes, whereas DDT and ethanol had only one major target in the *CYPome* (*Cyp12d1* and *Cyp6a8*, respectively). Those common genes responding to a chemical challenge are *Cyp6a2*, *Cyp12d1*, *Cyp6a8*, *Cyp6d5* and *Cyp6w1*. The wide chemical responsiveness of *Cyp6a2* that we observed using the transgenic fly model is thus confirmed by the various microarray studies.

### 3.4. Induction and insecticide resistance

Several reports in the literature have noted that CYP genes that are constitutively overexpressed in insecticide-resistant strains are also inducible by xenobiotics in susceptible strains, in *Drosophila* and in other insect species. We thus compared the data on CYP gene induction by xenobiotics that were collected in DNA micro-

array experiments with the list of genes that are currently known to be constitutively overexpressed in resistant strains or otherwise related to resistance (Fig. 4). Of the 27 genes known to be inducible by one or the other xenobiotics, and of the 12 genes known to be associated with resistance, eight are overlapping. Of these eight only three (*Cyp6a2*, *Cyp6g1* and *Cyp12d1*) have been shown to be causally related to resistance. Two (*Cyp12a4* and *Cyp6g2*) have been experimentally linked to resistance but have not been shown to be inducible by any of the nine compounds tested to date.

### 4. Discussion and conclusions

#### 4.1. *Tissue-specific expression*

The midgut, Malpighian tubules and fat body are the tissues recognized as the major sites of P450-mediated detoxification in
<table><caption>**Table 2**
Inducers of mammalian CYP1A and CYP2B genes compared with inducers of fruit fly Cyp6a2.</caption><thead><tr><th rowspan="2">Inducer</th><th colspan="2">Mammalian P450</th><th>Drosophila</th></tr><tr><th>CYP1A</th><th>CYP2B</th><th>Cyp6a2</th></tr></thead><tbody><tr><td>Phenobarbital</td><td>—</td><td>+</td><td>+</td></tr><tr><td>Pentobarbital</td><td>—</td><td>+</td><td>+</td></tr><tr><td>OCs</td><td></td><td></td><td></td></tr><tr><td>DDT/DDD/DDE/dicofol</td><td>—</td><td>+</td><td>+</td></tr><tr><td>Aldrin</td><td>—</td><td>+</td><td>+</td></tr><tr><td>trans-Stilbene oxide</td><td>—</td><td>+</td><td>+</td></tr><tr><td>PCBs</td><td></td><td></td><td></td></tr><tr><td>Aroclor 1254</td><td>+</td><td>+</td><td>+</td></tr><tr><td>2,4,5,2',4',5'-HCB</td><td>—</td><td>+</td><td>+</td></tr><tr><td>3,4,5,3',4',5'-HCB</td><td>+</td><td>—</td><td>—</td></tr><tr><td>Limonene</td><td>—</td><td>+</td><td>+</td></tr><tr><td>β-Naphthoflavone</td><td>+</td><td>—</td><td>—</td></tr><tr><td>3-Methylcholanthrene</td><td>+</td><td>—</td><td>—</td></tr><tr><td>Ethanol</td><td>—</td><td>—</td><td>—</td></tr><tr><td>Clofibrate</td><td>—</td><td>—</td><td>—</td></tr></tbody></table>

Induction of Cyp6a2 was estimated *in vivo* by the green fluorescence of treated transgenic 6a2-GFP flies, see Section 2 for details. Induction patterns of CYP1A and CYP2B genes were compiled from the literature.

<figure><img src="image_33.png" /></figure>

Fig. 4. Venn diagram showing the overlap between CYP genes inducible by chemicals and CYP genes associated to resistance in field or laboratory studies. The overlapping genes are identified, and of those, the three genes that have been causally linked to resistance in field-derived strains or experimental studies are in bold.

<table><caption>**Table 3**
Induction of the CYPome.</caption><thead><tr><th>Phenobarbital</th><th>Atrazine</th><th>Caffeine</th><th>Piper nigrum extract</th><th>Piperonyl butoxide</th><th>Pyrethrum</th><th>Ethanol or DDT</th><th>Paraquat</th><th>Tunicamycin</th></tr></thead><tbody><tr><td>6 datasets<br>[22-24,27]</td><td>2 datasets<br>[23]</td><td>1 dataset<br>[22]</td><td>1 dataset<br>[30]</td><td>2 datasets<br>[25]</td><td>1 dataset<br>[31]</td><td>1 dataset<br>[28]</td><td>2 datasets<br>[26,29]</td><td>1 dataset<br>[26]</td></tr><tr><td>Ad-, L-<br>F-, M-</td><td>Ad-<br>F-, M-</td><td>Ad-<br>M-</td><td>Ad-<br>F-</td><td>Ad-<br>M-</td><td>Ad-<br>F-</td><td>Ad-<br>M-</td><td>Ad-<br>M-</td><td>Ad-<br>M-</td></tr><tr><td>Cyp4d4</td><td>Cyp6a2</td><td>Cyp4ae1</td><td>Cyp6a8</td><td>Cyp4ae1</td><td>Cyp9f2</td><td>Cyp6a8 (ethanol)</td><td>Cyp4ac3</td><td>Cyp4e1</td></tr><tr><td>Cyp4e2</td><td>Cyp6d5</td><td>Cyp4d14</td><td>Cyp6d4</td><td>Cyp4e2</td><td>Cyp12d1</td><td>Cyp12d1 (DDT)</td><td>Cyp4p1</td><td>Cyp4p1</td></tr><tr><td>Cyp6a2</td><td>Cyp6g1</td><td>Cyp4e2</td><td>Cyp6d5</td><td>Cyp4p1</td><td></td><td></td><td>Cyp6a13</td><td>Cyp6a23</td></tr><tr><td>Cyp6a8</td><td>Cyp6w1</td><td>Cyp6a2</td><td>Cyp6w1</td><td>Cyp6a2</td><td></td><td></td><td>Cyp6a22</td><td>Cyp6d4</td></tr><tr><td>Cyp6a17</td><td>Cyp12d1</td><td>Cyp6u8</td><td>Cyp9b2</td><td>Cyp6u8</td><td></td><td></td><td>Cyp6a23</td><td>Cyp9b2</td></tr><tr><td>Cyp6a23</td><td>Cyp304a1</td><td>Cyp6a21</td><td>Cyp12d1</td><td>Cyp6a21</td><td></td><td></td><td>Cyp6d4</td><td>Cyp28a5</td></tr><tr><td>Cyp6d5</td><td></td><td>Cyp6d4</td><td></td><td>Cyp6a23</td><td></td><td></td><td>Cyp6d5</td><td>Cyp313a1</td></tr><tr><td>Cyp6g1</td><td></td><td>Cyp6d5</td><td></td><td>Cyp6d5</td><td></td><td></td><td>Cyp28a5</td><td></td></tr><tr><td>Cyp6w1</td><td></td><td>Cyp6g1</td><td></td><td>Cyp6w1</td><td></td><td></td><td>Cyp309a1</td><td></td></tr><tr><td>Cyp9b2</td><td></td><td>Cyp6w1</td><td></td><td>Cyp12b2</td><td></td><td></td><td></td><td></td></tr><tr><td>Cyp12d1</td><td></td><td>Cyp12d1</td><td></td><td>Cyp12c1</td><td></td><td></td><td></td><td></td></tr><tr><td></td><td></td><td></td><td></td><td>Cyp12d1</td><td></td><td></td><td></td><td></td></tr></tbody></table>

Genes listed in bold are the genes most representative of inducibility following chemical challenge or stress response (see Fig. 3). L-, Ad-: third instar larvae or adults. M-, F-: male or female.

insects [34-36]. DNA microarray analyses in *Drosophila* have confirmed that P450 transcripts are enriched in these tissues. In Malpighian tubules of adult flies, the Cyp6a18 transcript was enriched

>25-fold and Cyp6a2 enriched 8-fold [37]. The data of that study reveal that while 1,457 genes (10% of all genes) are expressed more than 2-fold in the Malpighian tubules, 29 P450 genes (or 28% of all P450 genes) are expressed over 2.8-fold. Similarly, 36 P450 genes (40%), including Cyp6a2, are represented in the larval midgut transcriptome [38]. A distillation of FlyAtlas data on tissue expression of P450 genes [39] shows that Cyp6a2 is not expressed at high levels in normal flies, with Malpighian tubules, hindgut and head being the major sites of expression. Our results also show that Cyp6a2 basal expression is very low, although our technique does not measure the Cyp6a2 transcript levels but the accumulation of GFP protein driven by the Cyp6a2 promoter, and is therefore qualitatively different and perhaps less sensitive. Following induction, however, the gene is highly expressed both in larvae and in adults, in the midgut, Malpighian tubules and fat body. These results confirm and expand in detail the early results obtained by in situ hybridization in adult flies [7]. They also clearly show that Cyp6a2, after induction, is expressed in many other tissues as well as in the recognized sites of detoxification. The expression in the nervous system, in both larvae and adults is of significance, as the majority of insecticides have nervous system targets, such as the

<figure><img src="image_34.png" /></figure>

Fig. 3. Venn diagram summarizing the inducibility of *Drosophila* CYP genes by various chemicals. 1: ethanol; 2: pyrethrum; 3: phenobarbital; 4: caffeine; 5: piperonyl butoxide; 6: Piper nigrum extract; 7: atrazine; 8: tunicamycin; 9: paraquat. The limited overlap between the CYP genes induced by paraquat and tunicamycin and those induced by the other chemicals led us to draw an arbitrary line to separate the genes that we designate as representing the response to stress from those representing the response to chemical challenge.

Please cite this article in press as: M. Giraudo et al., Regulation of cytochrome P450 expression in *Drosophila*: Genomic insights, Pestic. Biochem. Physiol. (2009), doi:10.1016/j.pestbp.2009.06.009
organophosphorus insecticides that are metabolized by CYP6A2 [10]. Expression of *CYP6D1* in the thoracic ganglia of adult house flies has also been reported to contribute to cypermethrin resistance in the Lpr strain [40]. The metabolism of xenobiotics by P450 enzymes is highly "uncoupled", costing much NADPH and generating considerable amounts of reactive oxygen species, as shown recently in a detailed study of house fly CYP6A1 [53]. The uninduced, low basal production of P450 enzymes is therefore probably an optimal state in the absence of chemical challenge. The observation that induced *Cyp6a2* should be found in a wide variety of tissues, beyond those in the front line of defense against xenobiotics such as the midgut and Malpighian tubules, may be an evolved response of insects who lack a closed circulatory system and therefore have all their tissues exposed to xenobiotics, once these have crossed the midgut or integument barriers. *Cyp6a2* has a pattern of expression that is similar to that of *Cyp6g1*. Chung et al. [50] have used in situ hybridization as well as GFP reporter constructs to study the spatial expression of the *Cyp6g1* gene. They showed that the wild-type *Cyp6g1* is expressed in parts of the midgut and in the Malpighian tubules. In wandering larvae, expression is also detected in the fat body. In resistant strains carrying a fragment of the Accord transposable element, the expression is higher in those tissues, the whole midgut, and also includes the gastric caeca. Manipulation (RNAi or overexpression) of *Cyp6g1* transcript levels in transgenic flies under the control of a Malpighian tubule promoter is sufficient to affect DDT resistance, whereas RNAi suppression of *Cyp6g1* expression in fat body or brain is without effect [39]. Thus, the tissue expression pattern is critical in determining the toxicodynamics of insecticides (or more generally xenobiotics) that are metabolized by P450 enzymes.

## 4.2. Specificity of induction

The *Cyp6a2* gene appears to be responding to multiple chemical challenges, but not to all chemical treatments. Other genes in the CYPome also appear to have a specific range of response. It is obvious that the detailed study of one gene or the cursory analysis of the whole CYPome as seen through various microarray experiments does not allow definitive conclusions to be drawn, and that more detailed analyses should focus on the time course of induction and its dose response (see [22] for time courses). Nonetheless, it seems that about a third of the CYPome is inducible, while the majority of P450 genes is not regulated by xenobiotic exposure. The non-inducible genes probably include the Halloween genes encoding enzymes of ecdysteroid biosynthesis [41] but also include many genes of unknown function, that are expressed at low levels in the whole insect and may in fact have a very restricted physiological function and pattern of expression. Among the inducible genes, there are obvious subsets, evidenced in our survey as stress-responsive genes and genes responding to chemical challenge. This suggests that the chemical diversity of xenobiotics is perceived by different, probably overlapping transduction mechanisms, and this leads to an adapted response. In vertebrates, this is achieved by two types of receptors, the members of the nuclear receptor family including the xenosensors PXR and CAR, and the bHLH-PAS proteins such as the Ah receptor. Cross-talk between these receptors and with receptors for endogenous hormones probably allows a fine-tuning of the response [42].

The inducibility of certain P450 genes in bacteria, plants, and insects by phenobarbital remains somewhat mysterious. However, the remarkable parallel between *Cyp6a2* induction and the known induction pattern of mammalian *CYP2B* genes strongly suggests that the "phenobarbital-type" induction in insects and mammals may share some characteristic that has been conserved during evolution. The nuclear receptor CAR is an essential component of the mouse response to the drug [43,44]. Analysis of the evolutionary

relationships between *Drosophila*, *Caenorhabditis elegans* and vertebrate nuclear receptors revealed that the orphan receptor DHR96 was most closely related to the nematode NHR-8 and DAF-12 receptors and to vertebrate CAR, vitamin D receptor (VDR) and Pregnane X receptor (PXR) [45,46]. The nematode NHR-8 receptor is essential for wild-type resistance to colchicine and chloroquine [45]. Recently, King-Jones et al. [24] presented evidence for the role of DHR96 in the response of *Drosophila* to phenobarbital. They obtained DHR96 null mutants and showed that the expression of some but not all phenobarbital-inducible genes was affected in the mutants.

We attempted to silence the DHR96 gene by RNAi, using the UAS-GAL4 system wherein a transgenic line with a promoter-driven GAL4 expression was crossed to a transgenic line with UAS-driven expression of DHR96 dsRNA (line 11783R-2 kindly provided by Dr. Ueno, NIG-Fly, Japan). The heat shock promoter (hsp70-GAL4) or the promoter with moderate expression from the *Cyp6g1* gene (*Cyp6g1*–GAL4; line kindly provided by Dr. Ffrench-Constant, Univ. Exeter) were insufficient as drivers to obtain significant downregulation of DHR96, whereas a stronger, ubiquitous promoter (tubulin-GAL4) led to high mortality in both male and female crosses (results not shown). These negative results nonetheless indicate a complex role for DHR96 that may be involved in essential physiological or developmental process in addition to its reported implication in the response to xenobiotics.

## 4.3. Resistance and induction

*Cyp6a2* and *Cyp12d1* are both inducible by at least four compounds and their constitutive overexpression has been causally linked to resistance. *CYP6A2* metabolizes organophosphorus insecticides [10] and *CYP12D1*, when overexpressed in transgenic flies, confers resistance to dicyclanil and to DDT [47]. We note here that *Cyp12d1* is a recently duplicated gene (i.e. *Cyp12d1* and *Cyp12d2* that differ by only three non-neutral nucleotide changes (http://p450.sophia.inra.fr), *Cyp12d2* having been masked by a gap in the initial release of the *Drosophila* genome sequence). A third gene, *Cyp6g1*, is inducible by phenobarbital and by caffeine, and its overexpression in natural populations of *Drosophila* [48,49] as well as in transgenic flies [48,50,47,39] causes resistance to DDT and to neonicotinoids. Five other genes, *Cyp4e2*, *Cyp4p1*, *Cyp6a17*, *Cyp6a8* and *Cyp6w1* are both inducible and overexpressed in some resistant strains, but there is no evidence for their causal relationship to resistance. The case of *Cyp6a8* is noteworthy, as the recombinant enzyme metabolizes aldrin to a modest degree but not heptachlor or DDT [51], and its overexpression in transgenic flies does not confer resistance to DDT, nitenpyram, dicyclanil or diazinon [47]. Two genes are known to be linked to resistance but have not been shown to be inducible in any of the studies that we analyzed. *Cyp12a4* confers a selective resistance to lufenuron in field strains and in laboratory, transgenic strains [52]. *Cyp6g2* is a gene expressed at very low levels in the head [39] but its transgenic overexpression in gut and Malpighian tubules confers diazinon and dicyclanil resistance [47]. These discrepancies highlight the danger of associating resistance with constitutive overexpression, and resistance with induction. We have argued [23] that inducibility of a P450 gene implies the presence of cis-regulatory sequences in addition to the sequences controlling the correct tissue and developmental expression. The broader target (cis-sequences of the induced gene and genes for the trans-factors that are necessary for its induction) for mutational events is therefore a risk factor for resistance when the gene encodes a xenobiotic-metabolizing enzyme. It is well known that inducibility is under genetic control with different insect strains showing different induction patterns (see e.g. [54–56]). Hällström [57] has compared a resistant strain in which the resistance mutation has caused increased constitutive
expression and has abolished further inducibility to a "constitutive mutant". Any other link between induction and resistance is probably coincidental rather than causal, because the time needed for induction and its dose-response would not sufficiently improve the toxicokinetics of most fast-acting neurotoxins used as insecticides to explain resistance.

While this paper was under review, Chung et al. [58] reported on the constitutive expression of CYP genes in *Drosophila* larvae. They obtained expression patterns for about two thirds of the CYPome, of which half were expressed in the fat body, midgut and Malpighian tubules, including all the genes that we highlighted in Fig. 3. Many other genes had a more discrete expression pattern, sometimes specific to small organs or cell types. Their results and our survey lead to a similar conclusion, that only a restricted subset of P450 enzymes are likely to be involved in detoxification. Many endogenous functions remain to be discovered for the other P450 enzymes.

Acknowledgments

Work on *Cyp6a2*-GFP was supported by National Institutes of Health Grants GM39014 and ES06694. We thank Dr. Ueno and Dr. Ffrench-Constant for the UAS or GAL4 transgenic lines. We thank B. Dunkov for help with the GFP transgenic lines and the Synthetic Core Facility, Southwest Environmental Health Sciences Center, University of Arizona, for the hexachlorobiphenyls.

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CHAPTER 2

Manuscript: Xenobiotic gene induction response in *Spodoptera frugiperda* midgut and Sf9 cells.
<figure><img src="image_35.png" /></figure>

Now that we have identified patterns of gene induction in the model species *Drosophila melanogaster*, we would like in this second chapter to extend our study to a polyphagous lepidopteran pest, *Spodoptera frugiperda*.

As it was discussed earlier (see section 3.3.2.2 of the introduction), because the majority of phytophagous insects have a restricted host range (three or fewer families of plants; (Dicke, 2000)), they encounter large amounts of predictable chemistries, and have characteristically high P450-based metabolism towards such chemistries. This is clearly the case for CYP6B1, a P450 from the black swallowtail, *Papilio polyxenes*, that is induced by and metabolizes toxic host plant furanocoumarins (review in Berenbaum, 2002). Polyphagous, generalist herbivores on the other hand face a more diverse array of chemistries. It is well known that plant chemicals, pesticides and other xenobiotics can induce the activity of insect P450 enzymes and the expression of some specific genes (review in Feyereisen, 2005). It is not yet known which types of chemicals induce which type of P450 genes. It is not known how relevant concentrations of plant-derived inducers affect the transcriptome in the cells exposed to, and responding to the inducer - principally the midgut in the case of Lepidoptera.

Thanks to the Génoscope BAC sequencing project "comparative genomics of Lepidoptera" and the access to ESTs from 8 different *S. frugiperda* libraries built from different tissues and development stages as well as from treated individuals, we were able to print long oligonucleotide microarrays containing 9773 60-mers oligonucleotides from *S. frugiperda*. In
addition, we were able to identify 33 sequences of P450 genes in *S. frugiperda*, out of which 9 could be efficiently tested in qRT-PCR reactions in Sf9 cells and 18 in *S. frugiperda* midgut. The remaining P450s were either not detectable in the midgut or in Sf9 cells or did not show acceptable efficiency in qRT-PCR and were thus not tested.

We have used both approaches, microarray and qRT-PCR on specific P450 genes to study the xenobiotic response in the polyphagous pest *S. frugiperda* both *in vivo* in the larvae midgut and *in vitro* in Sf9 cells exposed to a battery of xenobiotic compounds containing plant allelochemicals, insecticides and potent inducers, as described earlier in the objectives. By studying the effect of different types of xenobiotics, we wish to span different paths of the chemical space that insects have to navigate through. As we said in the first chapter of this manuscript, chemical space encompasses every possible existing molecules, with only a small part being biologically relevant, meaning synthesised and/or used by living organisms. These molecules can be characterised by different descriptors from their molecular weight to the partition coefficient in octanol relative to water. We have summarised in table 4 the main descriptors of the xenobiotic compounds that we have used in this study. They range from 117 to 505 in MW and from somewhat polar to liphophilic.

We will now present our results where we have identified patterns of gene induction in response to xenobiotics by microarray analysis and specific induction of P450 genes by qRT-PCR.

<table><caption>**Table 4**: chemical properties of the xenobiotic compounds used as inducers in S. frugiperda larvae and Sf9 cells</caption><thead><tr><th>compound</th><th>logP<br>octanol/<br>water</th><th>MW</th><th>source</th><th>Molar<br>volume<br>(cm³)</th><th>H bond<br>donors</th><th>H bond<br>acceptors</th><th>Molar<br>refractivity<br>(cm³)</th><th>Polar surface<br>area<br>(angström)</th></tr></thead><tbody><tr><td>Deltamethrin</td><td>6.20</td><td>505.2</td><td>Synthetic</td><td>316.7</td><td>0</td><td>4</td><td>116.0</td><td>59.32</td></tr><tr><td>Quercetin</td><td>1.82</td><td>304.2</td><td>Many</td><td>178.6</td><td>5</td><td>7</td><td>73.7</td><td>72.45</td></tr><tr><td>Methoxyfenozide</td><td>3.72</td><td>368.5</td><td>Synthetic</td><td>335.3</td><td>1</td><td>5</td><td>108.2</td><td>49.85</td></tr><tr><td>Xanthotoxin</td><td>1.93</td><td>216.2</td><td>Apiaceae</td><td>158.0</td><td>0</td><td>4</td><td>56.6</td><td>48.67</td></tr><tr><td>Indole</td><td>2.14</td><td>117.1</td><td>Many</td><td>101.8</td><td>1</td><td>1</td><td>38.5</td><td>4.93</td></tr><tr><td>Indole 3-carbinol</td><td>0.96</td><td>147.2</td><td>Brassicaceae</td><td>115.3</td><td>2</td><td>3</td><td>45.0</td><td>14.16</td></tr><tr><td>Clofibrate</td><td>3.32</td><td>242.7</td><td>Synthetic</td><td>211.4</td><td>0</td><td>3</td><td>62.7</td><td>35.53</td></tr><tr><td>2-tridecanone</td><td>5.16</td><td>198.3</td><td>Solanaceae</td><td>240.2</td><td>0</td><td>1</td><td>62.3</td><td>17.07</td></tr><tr><td>Fipronil</td><td>3.21</td><td>437.1</td><td>Synthetic</td><td>233.6</td><td>2</td><td>5</td><td>81.8</td><td>81.13</td></tr><tr><td>Methoprene</td><td>5.63</td><td>310.5</td><td>synthetic</td><td>339.8</td><td>0</td><td>3</td><td>93.4</td><td>35.53</td></tr></tbody></table>
Xenobiotic gene induction response in *Spodoptera frugiperda* midgut and Sf9 cells.

Maeva Giraudoª, René Feyereisenª*, Frédérique Hilliouª, Trang Tranᵇ and Gaëlle Le Goffª.

*Corresponding author: René Feyereisen, UMR 1301 IBSV, INRA-CNRS-Université de Nice
Sophia-Antipolis, 400 Route des Chappes, 06903 Sophia-Antipolis, FRANCE; phone : 33
492386450, fax : 33 492386401, e-mail : rfeyer@sophia.inra.fr

short title: *Spodoptera frugiperda* xenobiotic response

<footnote>ª UMR 1301 IBSV, INRA-CNRS-Université de Nice Sophia Antipolis, 400 Route des Chappes, 06903 Sophia-Antipolis, France</footnote>

<footnote>ᵇ Lanaud Gestion-Pôle de Lanaud, 87220 Boisseuil, France</footnote>
**Abstract**

Phytophagous insects have developed sophisticated mechanisms to metabolically inactivate the potentially toxic plant allelochemicals and insecticides that they ingest. These mechanisms are mainly based on the induction of detoxification genes such as cytochrome P450s. We have studied the xenobiotic response in the polyphagous lepidopteran pest, *Spodoptera frugiperda* and in the derived Sf9 cells in response to different chemical compounds. Microarray analysis showed that the xenobiotic response was different *in vitro* and *in vivo*, with a common pattern involving the regulation of detoxification genes. qRT-PCR analysis showed that the plant allelochemicals indole and xanthotoxin were the strongest inducers of a small subset of P450s in the midgut (CYP321A7, CYP321A9, CYP9A32) whereas insecticides induced P450 expression mostly *in vitro* (CYP9A30, CYP9A31). Our results suggest that the xenobiotic response in a polyphagous pest rely on detoxification gene regulation involving the induction of different subset of P450s.

**Keywords:** P450, Spodoptera frugiperda, microarray, xenobiotics.
# 1. Introduction

The chemical space, which encompasses all possible existing small molecules (>10⁶⁰), comprises only a small fraction of biological relevant molecules that are synthesized and/or used by living organisms (Dobson, 2004). Plant metabolome represents a large part of that space as plants synthesize a broad range of secondary metabolites with some 200 000 compounds well known to pharmacologists (Hartmann, 2004). These plant allelochemicals are essential for their growth and development as well as defensive compounds against their natural enemies, such as herbivorous insects. Indeed, plants and insects have co-evolved for more than 350 millions years, entering an evolutionary arms-race in which plants evolved allelochemicals-based mechanisms to fend off their natural enemies and herbivorous insects in turn have developed sophisticated mechanisms to counteract these plant defences (Gatehouse, 2002; Rausher, 2001).

Some of the most important mechanisms that have allowed herbivorous insects to persist in a modern agriculture are their ability to metabolize and detoxify xenobiotics that they encounter as constituents in their host plants (Ferry et al., 2004; Li et al., 2007). This ability has been shown to grant phytophagous insects with the capacity of developing cross-resistance to insecticide compounds used in pest management. This is the case for example in the Lepidoptera *Helicoverpa zea*, which shows increased tolerance to the insecticide carbaryl after being exposed to 2-tridecanone, a toxic allelochemicals from trichomes of wild tomatoes (Riskallah et al., 1986). Lepidoptera represent a diverse and important group of agricultural insect pests that cause widespread economic damage on food and fiber crop plants, fruit trees, forests, and stored grains. Understanding detoxification machinery in these species has therefore become critically important in order to develop efficient pest control strategies.
This detoxification machinery includes different classes of enzymes, the most abundant and the best known being the cytochrome P450 enzymes. Insects have about 100 P450 genes (Ranson et al., 2002; Tijet et al., 2001) that play numerous functional roles, including growth, development, feeding, resistance to pesticide and tolerance to plant toxins (Feyereisen, 1999). Given the importance of insect P450 in metabolism, this versatile metabolic system has been a main topic in insect research for many decades. It is now well known that plant chemicals, pesticides and other xenobiotics can induce the activity of insect P450 enzymes and the expression of some specific genes as it has been reviewed in (Feyereisen, 2005; Yu, 1986). For example, members of the CYP4 family in the tobacco hornworm, *Manduca sexta* (Snyder et al., 1995) of the CYP9 family in *Helicoverpa armigera* (Zhou et al., 2009) and the CYP6 family in *Helicoverpa zea* (Li et al., 2002) have all been reported to be inducible by host plant chemicals and xenobiotics. However, the pattern of response within the whole multigene families or the selectivity of the inducer has yet to be documented.

With the recent advent of complete, annotated genomes of different insect species such as Drosophila (Adams et al., 2000) and Tribolium (Richards et al., 2008), the study of xenobiotic response in insects has been taken to the whole transcriptome level thanks to the development of post-genomic tools such as DNA microarrays and more recently to the pyrosequencing. DNA microarray hybridization has rapidly become a method of choice to investigate genome wide response to xenobiotics. For example, this technique has been used to study the molecular response of *Drosophila* to chemical stressors such as H₂O₂, paraquat and tunicamycin (Girardot et al., 2004). Moreover, the design of specific “detox” chips that represent all members of the detoxification gene families have provided an excellent tool to identify inducible genes. For example, detox chips have been used in *Drosophila* to study P450 gene induction in response to phenobarbital and atrazine (Le Goff
et al., 2006) or to measure the induction of detoxification gene families such as P450s, GST sans esterases in response to insecticides, caffeine and phenobarbital (Willoughby et al., 2006). Similar approaches have also been widely used in mosquitoes such as *Anopheles gambiae* (Vontas et al., 2005) and *Aedes aegypti* (Poupardin et al., 2008) to identify detoxification response to insecticide exposure.

In Lepidoptera, a relatively lower number of studies have used post-genomic tools to investigate patterns of gene expression in these insects. For example, cDNA libraries were constructed from pheromone glands of *Heliothis virescens* and compared to other available moths cDNA databases to identify genes involved in the biosynthesis of pheromones (Vogel et al., 2010) and the recently developed pyrosequencing technique was applied to *Manduca sexta* to define larval midgut transcriptome and revealed the identification of new genes, among which 36 new P450s (Pauchet et al., 2010). With the sequence of *Helicoverpa armigera* on its way, there is no doubt that lepidopteran transcriptomic studies will grow, using recent advances in high-throughput genomic analysis such as pyrosequencing.

Among Lepidoptera, the fall armyworm *Spodoptera frugiperda* (JE Smith) is a serious polyphagous pest present mostly in the American continent that causes severe damages on several economically important crops such as corn, soybeans, peanuts, sorghum, Bermuda grass and cowpeas. This pest has developed resistance to major classes of insecticides in many areas as a result of chemical control (Negre et al., 2006; Yu and McCord, 2007). In addition to being important agricultural pests these noctuids are biological models studied for several purposes. For example, *S. frugiperda* is well known through its famous Sf21 cell line and its Sf9 subclone (Vaughn et al., 1977) which is used for numerous heterologous protein productions (Negre et al., 2006).
We have constructed a *Spodoptera frugiperda* microarray consisting of 9773 60-mers
oligonucleotides designed from ESTs databases obtained from eight different tissues,
developmental stages and treated insects. Here we report the use of this microarray to
study the detoxification response in this polyphagous lepidopteran pest exposed to different
plant allelochemicals and insecticides. By choosing a generalist species, we have increased
our chance to cover many different detoxification pathways as in contrast to specialist
insects, they encounter large amounts of predictable chemistries, and have therefore
characteristically high P450-based metabolism towards such chemistries (Stevens et al.,
2000). We have therefore also monitored P450 gene induction in response to the different
compounds tested to identify their role in the xenobiotic response of *S. frugiperda*.

**2. Material and Methods**

2.1 Reagents

The analytical grade plant allelochemicals xanthotoxin, quercetin, 2-tridecanone, indole, indole 3-carbinol and the herbicide clofibrate used in this study, as well as the drug phenobarbital were purchased from Sigma-Aldrich (St Quentin Fallavier, France). The insecticides deltamethrin, methoxyfenozide and fipronil were purchased from Cluzeau (CIL, France). Methoprene was a gift from the former Zoecon Corp., Palo Alto, California.

2.2 Insect rearing and treatments

Larvae of *Spodoptera frugiperda* were fed *ad libitum* on artificial medium adapted
from Poitout and Bues (Poitout and Bues, 1974) and were reared at 25,5°C ±1°C, and 70%
relative humidity under a L 14: D 10 photoperiod. Under these conditions, the 5th (penultimate) and 6th (ultimate) larval stage last about 2 to 3 days each, with additional two pre-pupal days characterized by the arrest of wandering and beginning of weight lost towards pupal formation.

For xenobiotic induction of gene expression, newly molted 6th instar larvae were fed for 24h on artificial medium containing doses of different xenobiotic compounds chosen according to the literature for plant allelochemicals and/or to toxicity testing for insecticide compounds (table 2). Briefly, newly moulted 6th instar larvae were fed for 24h with increasing concentrations of each of the xenobiotic compound and mortality was recorded for 10 larvae for each of the concentrations. Induction doses were chosen the highest dose that causes no mortality in 10 larvae.

Newly molted 6th instar larvae were then fed either 0.5% phenobarbital, 0.05% xanthotoxin, 0.25% 2-tridecanone, indole 3-carbinol, indole, clofibrate or quercetin, 20nM methoxyfenozide, 2µM methoprene, 10nM deltamethrin, 2µM fipronil or the equal amount of DMSO for control experiments.

## 2.3 *cell culture and treatments*

Sf9 cell line derived from the pupal ovarian tissue of *Spodoptera frugiperda* was purchased from Invitrogen. The cells were cultured and maintained in insect-Xpress serum free medium (Lonza) at 27°C in suspension spinner flasks with an agitation rate of 100 rpm and passaged routinely every third day. Cell density was determined by Malassez hemocytometer counts and cell viability was evaluated by methylene blue (1mg/mL, v/v)
staining. Prior to experiments, cells were sowed in 6 well plates (TPP) at 5.10⁵ cells/ml and left at 27°C for adhesion. Attached cells were treated for 24h with doses of different xenobiotics chosen after 24h MTT cytotoxicity testing. Briefly, Sf9 cells were seeded in 96-well culture plates and treated in triplicates for 24 hours with increased concentrations of each of the different compounds. Cells in culture were then loaded with MTT (5mg/ml) and incubated at 27°C for 2 hours. Cell homogenates were used to measure absorbance at 570nm using a microplate reader (SpectraMax, Molecular Devices). Induction doses were chosen at the DL10, which represents the dose lethal to 10% of the cells (table 3). Sf9 cells were treated for 24h with 500µM 2-tridecanone, 250µM phenobarbital, indole and indole 3-carbinol, 100µM xanthotoxin, clofibrate, and deltamethrin, 50µM quercetin, 25µM methoprene, 1µM fipronil, 0.1µM methoxyfenozide or the same volume of DMSO.

## 2.4 RNA extraction

For xenobiotic induction of gene expression studies in *S. frugiperda* larvae, midguts were dissected from larvae fed for 24h on artificial medium containing each of the tested compounds at the concentration stated above.

Total RNA was extracted from the midgut using TRIzol Reagent (Invitrogen Life Technologies) according to the manufacturer's protocol. After quantification of RNA concentration at 260nm and verification of RNA quality on 2% agarose gels, RNAs were stored at -80°C until use. RNA extractions were performed on 3 independent biological replicates of 5-pooled midguts.

Total RNA was extracted from cells of a well of the 6 wells plate also using Trizol Reagent and RNA were analysed and stored the same way than for tissue samples. Extractions were performed on three independent biological replicates.
## 2.6 Quantitative real-time PCR

Total RNA (1µg) was reverse transcribed using the iScript cDNA Synthesis kit (Biorad). qRT-PCR reactions were carried out on an Opticon monitor 2 (Biorad) using the qPCR Mastermix plus for SYBR Green I no ROX (Eurogentec). PCR primer pairs for *S. frugiperda* P450s and the three control genes (G6PD, L18 and Rpl4) are described in table 1. Final concentration of primers was 20nM. Efficiency of each primer pair was determined by absolute standard curves for the different gene transcripts and their controls by serial (5x) dilutions of 6th instar midgut and Sf9 cells cDNA. The PCR conditions were as follows: 50°C for 2 min, 95°C for 10min, followed by 40 cycles of 95°C for 30 sec, 60°C for 30 sec and 72°C for 30 sec. Each reaction was performed in duplicate and the mean of the three independent biological replicates was calculated for midgut and Sf9 cells. All results were normalized using mRNA level of three control genes (RpL4, L18 and G6PD) and relative expression values were calculated in R using the RqPCRBase package developed in our laboratory (Hilliou and Tran, manuscript in preparation).

## 2.6. Microarray experimental design

Our oligonucleotides were designed from 79148 ESTs sequences of eight different cDNA libraries of *Spodoptera frugiperda* representing many tissues including midgut and Sf9 cells (see http://bioweb.ensam.inra.fr/spodobase/). Using the assembly analysis (programme CAP3), we obtained 10092 contigs and singletons from these ESTs. Our *Spodoptera frugiperda* microarray consist of 9773 60-mers oligonucleotides synthesised by Sigma-Aldrich that were designed to match unique contigs or singletons and to suit our hybridization conditions (GC content average 46% and average Tm of 46°C). Each
comparison consisted in six microarrays, three biological replicates hybridized with dye swap (fully balance dye swap design) and duplicate spots. cDNA were synthesized from 7µg of total RNA and labelled with the dyes Cy3-dCTP and Cy5-dCTP (Amersham) using the ChipShot direct labelling system Promega according to the manufacturer's instructions. The microarrays were hybridized with cDNA prepared as described in Le Goff *et al* (2006) and scanned using GenePixPro scanner (Axon, version 3.01). Experimental data and associated microarray designs have been deposited in the NBCI Gene Expression Omnibus (GEO) (http://www.ncbi.nlm.nih.gov/geo/) under serie X and platform record X using Mediante database for data transfer (Le Brigand and Barbry, 2007).

## 2.7. Microarray data analysis

We used the Bioconductor suite of statistical packages (Gentleman et al., 2004): *limma* (Wettenhall and Smyth, 2004) for our data analysis. The expression intensities were obtained by subtracting the background intensity from the foreground intensity for each non-flag spot (all flagged spots were eliminated). The expression data were normalised by the use of the within-array normalization with the “loess method” and the between-array normalization using the “quantile method” (Yang and Thorne, 2003). The linear model for series of arrays and empirical Bayes method were then applied for assessing differential expression (Smyth, 2004). The false discovery rate of the p-value for multiple tests was controlled by using Benjamini-Hochberg method. Differentially expressed genes were selected if the absolute value of log2-fold-change greater than 1 and adjusted p-value below 0.01 and if the average intensity is greater than two time of average background. In order to provide an overall measure of evidence of differential expression, we used the Fisher's
method for combining adjusted p-values from independent tests of significance of duplicate
spots (Hess and Iyer, 2007).

**3. Results**

3.1 Choice of compounds and doses used

In order to study the xenobiotic response in *S. frugiperda*, last instar larvae and Sf9 cells were exposed for 24h to different plant allelochemicals: indole-3-carbinol, a glucobracissin derivative that is a known inducer in *S. frugiperda* and an Ah receptor activator in vertebrates; xanthotoxin (8-methoxypsoralen), a coumarin derivative that is a potent inducer in many insect species; 2-tridecanone, a secondary compound found in the trichomes of the wild tomato; indole as maize defensive compound and quercetin, a flavonoid that is encountered in many Lepidoptera host plant species. Some commonly used insecticides have also been tested: methoprene and methoxyfenozide, respectively juvenile hormone analog and ecdysone agonist; deltamethrin, a commonly used pyrethrinoid and fipronil, an insecticide that targets GABA receptors. In addition, we used phenobarbital as archetypal inducer and the PPAR-type inducer clofibrate.

The different xenobiotics were used at doses similar to the ones reported in the literature
and known to induce detoxification genes when it was possible (table 2), as this was the case
for plant allelochemicals in *in vivo* experiments. However, induction doses for insecticides in
*S. frugiperda* larvae and for all xenobiotic tested in Sf9 cells (table 3) were chosen from
mortality monitoring assay and cytotoxicity testing respectively. That way we could chose
doses that would be high enough to potentially induce molecular xenobiotic response in
both systems without causing toxic-related effects that could interfere with the
identification of detoxification machinery response. Our results show that insecticides had
quite different toxic effects in *S. frugiperda* larvae as some compounds such as fipronil and
methoprene that start to cause mortality at doses within the micromolar range, when
methoxyfenozide and deltamethrin seem to exert toxic effects at a much lower nanomolar
range (table 2).

In Sf9 cells, some compounds showed no cytotoxicity at the highest doses tested such as deltamethrin and 2-tridecanone, when compounds such as methoxyfenozide were toxic for the cells at concentrations as low as 0.1 µM (table 3). Interestingly, methoprene was the only compound that showed a more important toxic effect in larvae than in the cells.

3.2 Microarray results

These doses were then used in 24h exposure of *S. frugiperda* larvae and Sf9 cells to monitor
gene expression patterns in both systems by microarray analysis. The number of genes for
which levels of expression were significantly affected by each of the tested compounds in
both systems is depicted in figure 1. Our results indicate that *S. frugiperda* larvae and Sf9
cells show a striking different gene expression response to xenobiotics. Plant allelochemicals
have the most important effect in vivo, such as indole, quercetin and xanthotoxin that
regulate the expression of 227, 277 and 117 genes respectively, but have a moderate effect
in vitro compared to clofibrate, which affected 173 genes in Sf9 cells. In addition, the
number of commonly affected genes in both systems is extremely low, not exceeding 6
common genes in response to indole 3-carbinol and none in response to deltamethrin. Table
4 summarizes the genes that are affected both in larvae and cells. Expression values are
represented as the log of the fold change for each of the genes and its associated *p* value.
Sequences of each gene were submitted to blastx to retrieve gene functions. The sequences for which no homology was found by blastx were submitted to a blastn in Butterflybase and classified in "hypothetical protein" category if homology was found with the sequence of another lepidopteran transcript, while the sequences restricted to *Spodoptera frugiperda* were put in a "hypothetical transcripts" category. As we can see, some genes are found up-regulated in both cells and larvae for most of the compounds such as carboxyl/choline esterase, which is known to be involved in detoxification metabolism of xenobiotic in insects and insecticide resistance (Yu et al., 2009). However, some genes show opposite regulation patterns in larvae and cells as it is the case in response to clofibrate and methoprene, where all commonly affected genes are found up-regulated in Sf9 cells and down-regulated in larvae. *In vivo* and *in vitro* systems show therefore a different response to xenobiotics, which do not seem to be correlated as shown in figure 2. The number of gene affected by each of the treatments shows indeed no correlation between the larvae and the cell line.

When looking at the number of genes found up- and down-regulated in *S. frugiperda* larvae and Sf9 cells in response to xenobiotics, our results show once again that both systems display different regulation patterns of gene expression as it can be seen in figure 3. We found that the plant allelochemicals xanthotoxin, quercetin and indole were the stronger inducers of gene expression in *S. frugiperda* larvae with 46, 100 and 88 genes up-regulated by these compounds respectively. In Sf9 cells, it seems that the herbicide clofibrate was the stronger inducer with 84 genes up-regulated by this xenobiotic when only 35 were induced in the larvae.

Among the genes that respond significantly to the tested xenobiotic, we have found that a number of them were involved in the detoxification response as it is shown in table 5 for *in*
vivo experiments and in table 6 for Sf9 cells. Genes such as carboxylesterases and glutathione S-transferases (GSTs) were up-regulated in response to indole, indole 3-carbinol, methoxyfenozide, phenobarbital and xanthotoxin in larvae, in addition we also found a glucosyltransferase induced by fipronil and phenobarbital and two P450s, CYP6B40 and CYP4M18 that were induced by methoxyfenozide and quercetin respectively. Genes involved in oxidative stress response were also found up-regulated in larvae such as a peroxiredoxin in response to clofibrate, a catalase and a peroxidase were up-regulated by indole and methoxyfenozide induced the expression of a superoxide dismutase. Interestingly, a number of the detoxification genes were also down-regulated in response to the different xenobiotics. This is the case for several GSTs in response to indole, indole 3-carbinol, quercetin and xanthotoxin and for two P450s: CYP6B40 in significantly down-regulated by indole and CYP9A25 by xanthotoxin. In addition, one CYP9A9 related gene and CYP6B6 was down-regulated by quercetin and a P450 similar to CYP321A1 from *Helicoverpa armigera* also showed a reduced expression in response to xanthotoxin. We found similar detoxification gene response in Sf9 cells with the induction of carboxylesterases in several treatments, including indole, indole 3-carbinol, methoprene, phenobarbital and xanthotoxin as it was found in larvae. Oxidative stress related genes such as peroxiredoxin and peroxidase were also found up-reglated in response to clofibrate and indole respectively, in concordance to the larvae response. We also found that CYP4M18 was induced by deltamethrin, as well as the *Helicoverpa* related CYP321A1 and CYP4M15 showed an increased expression in response to 2-tridecanone.

3.3 P450 expression monitored by qRT-PCR
We have further explored specific P450 gene expression in *S. frugiperda* larvae and Sf9 cells by quantitative-real time PCR analysis and results are presented in table 7 and table 8. We found 33 P450 genes in *S. frugiperda* databases built from different tissues, developmental stages and treated individuals. Out of these 33 genes, we were able to measure with acceptable qRT-PCR efficiency 19 of them in *S. frugiperda* midgut and 9 in Sf9 cells. The remaining P450s were either not detectable in one or the other system or did not show qRT-PCR efficiency in the 90-110% range. Except for CYP321A7 and CYP333B2, which are both induced in larvae and cells in response to indole 3-carbinol and fipronil respectively, none of the compounds showed the same patterns of gene induction in both systems. Plant allelochemicals were the stronger P450 inducers in larvae, with CYP321A7 being induced by 162 fold in larvae treated with xanthotoxin and by 135 fold in response to indole. Insecticides such as methoprene, deltamethrin and the herbicide clofibrate did not induce significantly P450 gene expression. Different results were observed in Sf9 cells, with these latter 3 compounds showed an increase expression of several P450s and xanthotoxin being unable to induce any of them. We were not able to validate microarray results that showed specific induction or repression of P450 genes as for example, CYP6B40 that was found down-regulated by indole in microarray analysis but up-regulated in qRT-PCR experiments.

Taken together, our results show that Sf9 cell response to xenobiotics does not match *in vivo* response and that both system show different patterns of gene induction. However different these patterns are, they still involve detoxification gene expression and P450s in particular.
Discussion

We have studied gene expression patterns in the midgut of *S. frugiperda* larvae exposed to various plant allelochemicals and insecticides and used the same battery of chemicals on the Sf9 cell line derived from this species. Insect cell culture have been used in numerous studies as research tools in virology, in studies of signaling mechanisms to study insect immunity, hemocyte migration, and to test hypotheses about gene expression, and in screening programs designed to discover new insecticide chemistries (for a review see Smagghe et al., 2009). Sf9 cells have however mainly been used as a heterologous expression system through baculovirus mediated expression. We report here the first evidence of the use of Sf9 cells as a model system to study xenobiotic response of insects. Our results show that *S. frugiperda* larvae and Sf9 cells display gene expression response to xenobiotic exposure involving several detoxification genes. We found that plant allelochemicals such as xanthotoxin, indole and indole 3-carbinol were able to induce carboxylesterases in both systems, an enzyme class that has been shown to be involved in the detoxification of xenobiotics and insecticide resistance such as carbamate and pyrethroids (Yu et al., 2009).

We also found that GSTs were found up-regulated in response to many different xenobiotics, which is consistent with the literature, for example GSTs induction was reported in Drosophila in response to atrazine and phenobarbital (Le Goff et al., 2006; Willoughby et al., 2006) and to indole and xanthotoxin (Li et al., 2007). We have found one GST induced in the midgut exposed to phenobarbital and to xanthotoxin and two GSTs over-expressed in response to indole. We also showed that some GSTs were down-regulated by some compounds such as the plant allelochemicals quercetin. This is not surprising as quercetin has been shown to inhibit the expression of some GSTs in *Trichoplusia ni* (Ahmad and Pardini, 1990). Glucosyltransferases (UGTs) were found mostly down-regulated in the
midgut by compounds such as clofibrate, fipronil and quercetin and similarly in response to clofibrate, indole 3-carbinol and 2-tridecanone in Sf9 cells. UGTs have been identified in many insect species and although their function is not yet clearly established, that have been involved in detoxification of xenobiotic and insecticide resistance (Luque et al., 2002). The fact that these genes were found mostly down-regulated in both systems could account for the toxic effect of the corresponding compounds. In addition, we found a number of antioxidant enzymes coding genes that were affected by the different xenobiotic treatments in both larvae and cells. Oxidative stress is a major deleterious mechanism that occurs in every aerobic living organisms by the generation of reactive oxygen species (ROS), and occurs in insects through respiration and the ingestion of ROS generating compounds such as phenols (Barbehenn, 2002). We found that clofibrate was able to induce one peroxiredoxin *in vivo* and in Sf9 cells. This enzyme was also found to respond to H₂O₂ and temperature stress in *Bombyx mori* (Wang et al., 2008) and the bumble bee (Hu et al., 2010).

We also found that indole and methoxyfenozide were inducers of a catalase and a superoxide dismutase respectively in the midgut, both being active antioxidant enzymes found in many insect species, such as in the grasshopper in response to tanic acid diets (Barbehenn, 2002). Our results show therefore a common response in *S. frugiperda* larvae and Sf9 cells involving detoxification genes. However, the regulation of these genes showed different patterns in both systems, suggesting different sensitivity of midgut and cells to xenobiotics. This is confirmed by the number of genes found up- and down-regulated that differs remarkably in both systems in response to the same xenobiotic compounds. Cell lines have always been a model system in toxicological studies in vertebrate and have been successfully used in insect to understand endogenous regulatory pathways involved in ecdysteroid signalling mechanisms and the effects of hormone agonists such as methoprene
and methoxyfenozide (Dhadialla et al., 1998). Our results indicate that although this system
might be useful to study detoxification response in insects, it does not represent what is
happening in insect tissues such as insect midguts. We have indeed found that P450 gene
induction was quite different in larvae and in Sf9 cells, with compounds such as xanthotoxin
and indole being strong P450 inducers in midguts and inversely having no or moderately
effects in the cells. Similarly, insecticides such as methoprene and deltamethrin were only
able to induce P450s in the cells. Sf9 cells have been shown to be a reliable system to study
the induction of an heterologous CYP6B1 by xanthotoxin (Brown et al., 2005). However, we
were unable to show a significant induction of any of the 9 P450s tested in Sf9 cells by
xanthotoxin, thus indicating that additional P450 not yet identified in the cells might be
involved. P450 gene induction was also found different in microarray analysis and qRT-PCR
studies, with CYP6B40 being induced by methoxyfenozide in midgut by specific qRT-PCT
measurement but the corresponding gene in the microarray analysis was found down-
regulated. Discrepancies between microarray and real-time quantitative PCR data can result
from biological variation or technical issues during microarray statistical analysis, which
might have eliminated genes that did not reach normalization and statistical reproducibility
requirements, as it was also the case in P450 gene expression studies in *Aedes aegypti*
(Poupardin et al., 2008). Our results confirm that microarray experiments require cross-
validation with other gene-expression profiling techniques. We have therefore explored the
available catalogue of P450 gene expression in both system in response to plant
allelochemicals and xenobiotics. P450s are the best known enzymes involved in the
detoxification machinery of xenobiotics in insects (Feyereisen, 2005). We found that the
CYP9A and CYP321A families were the most induced by the plant toxins xanthotoxin and
indole. Our results confirm the inducibility of these P450 that is found in the literature.
Indeed, CYP321A1 from *Helicoverpa zea* is induced by indole and xanthotoxin (Zeng et al., 2007), and CYP9A members are actively induced in *Manduca sexta* by xanthotoxin, phenobarbital, indole 3-carbinol, 2-tridecanone and clofibrate (Stevens et al., 2000). CYP9A30, 31 and 32 were inducible in Sf9 cells by deltamethrin as it was found for CYP9A12 and CYP9A17 in *Helicoverpa armigera* (Zhou et al., 2009), however we could not detect any induction of these genes in the midgut nor in response to phenobarbital as it is the case for *H. armigera*. In addition, we found that CYP4M14 was induced by 2-tridecanone and methoprene in Sf9 cells and CYP4M members have already been shown to respond to 2-tridecanone, clofibrate and phenobarbital in *Manduca sexta* (Snyder et al., 1995).

Our results according to the literature indicate that detoxification mechanisms in insects are complicated, hence in a polyphagous species such as *S. frugiperda*, which encounters a large array of xenobiotic compounds, and is mainly based on P450 gene induction. We have identified numerous P450s induced by the different xenobiotics, with genes from the CYP9A and CYP321A families that respond particularly to plant allelochemicals.