Ecdysone, Structures and Functions [1 ed.] 1402091117, 9781402091117

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Ecdysone: Structures and Functions

Guy Smagghe Editor

Ecdysone: Structures and Functions

Editor Guy Smagghe Laboratory of Agrozoology Faculty of Bioscience Engineering Ghent University Belgium

ISBN 978-1-4020-9111-7

e-ISBN 978-1-4020-9112-4

Library of Congress Control Number: 2008938015 © 2009 Springer Science + Business Media B.V. No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Printed on acid-free paper springer.com

Preface

The 16th International Ecdysone Workshop took place at Ghent University in Belgium, July 10–14, 2006 and drew some 150 attendees, many of these young students and postdoctoral associates. These young scientists had the opportunity to discuss their work with many senior scientists at meals, breaks and during the several social events, and were encouraged to do so. This book resulting from the meeting is more up-to-date than might be expected since manuscripts were not delivered to the editor until 2007. The workshop itself had 54 oral presentations as well as many posters. This book, and the meeting itself, is comprised of 23 contributed chapters falling into five general categories: Fundamental Aspects of Ecdysteroid Research: The Distribution and Diversity of Ecdysteroids in Animals and Plants; Ecdysteroid Genetic Hierarchies in Insect Growth and Reproduction; Role of Cross Talk and Growth Factors in Ecdysteroid Titers and Signaling; Ecdysteroid Function Through Nuclear and Membrane Receptors; Ecdysteroids in Modern Agriculture, Medicine, Doping and Ecotoxicology. Clearly, all that was presented at the meeting and in this volume cannot be summarized here in a single page, but the reader is cordially invited to explore this relatively large volume that attempts to synthesize the most current knowledge base for this important category of steroid hormones. I use the plural because the nomenclature has indeed undergone a metamorphosis akin to what our experimental animals undergo. In the 1950s when the Butenandt and Karlson laboratory first crystallized and characterized ecdysone, it was thought to be the insect molting hormone since when injected into experimental insects the result was molting. Later, it was found that ecdysone was converted to 20-hydroxyecdysone in tissues peripheral to the prothoracic glands through the mediation of an ecdysone monooxygenase and we then believed that ecdysone was only the precursor for 20-hydroxyecdysone. However, a reading of the older literature as well as new work has demonstrated convincingly that ecdysone does have regulatory roles of its own, and perhaps joins with 20-hydroxyecdysone to initiate the molting process. Further, it must be remembered that the research of Ulrich Clever and Peter Karlson on Chironomus was the very first to demonstrate that a steroid hormone acts at the nuclear level and v

vi

Preface

indeed Karlson wrote several theoretical papers subsequently urging endocrinologists working on mammals to accept that view. In time they did! Despite the molecular focus of this meeting there remains much research to do at the physiological and biochemical levels, let alone in the areas of chemistry and crystallography, to further define the roles of these hormones that serve more organisms on this planet than any other steroid hormone. For at least current usage, I suggest that we use the term principal molting hormone for 20-hydroxyecdysone and the terms substrate for the molting hormone and/or hormone for ecdysone (although many Drosophila geneticists still use the term ecdysone for 20-hydroxecdysone; but that is another problem). The Ghent meeting was beautifully organized by Professor Smagghe who seemed to be around 24 h a day insuring that everything proceeded correctly and on schedule. He deserves our thanks for that, and also for this volume that he labored over for many months. I personally believe that this was among the very best of the many ecdysone workshops I have attended and the book is certainly among the most comprehensive outcomes of any of the workshops. I was most honored to have been chosen to give the Karlson lecture on our work on the Halloween genes. I realize that this is not the usual preface in that I haven’t discussed any of the contributions, but they are so good that I would do them a disservice by summarizing each in a sentence – they deserve sustained reading and I believe that they will be cited for many years to come. As I approach my 80th birthday after more than half century in this field, I am very pleased to have my new work published here, that I had the chance to attend this very successful meeting in such a very beautiful city, and the opportunity to enjoy the amazing varieties of such fine beers offered to the ‘ecdysonists’ in Ghent. Chapel Hill, North Carolina, USA September 2008

Lawrence I. Gilbert

Contents

Preface .............................................................................................................

v

Color Plates ....................................................................................................

xi

Part I

1

Fundamental Aspects of Ecdysteroid Research: The Distribution and Diversity of Ecdysteroids in Animals and Plants

Phytoecdysteroids: Diversity, Biosynthesis and Distribution ....................................................................................... Laurence Dinan, Juraj Harmatha, Vladimir Volodin, and René Lafont

3

2

Diversity of Ecdysteroids in Animal Species ......................................... René Lafont and Jan Koolman

47

3

Crustacean Ecdysteroids and Their Receptors..................................... Penny M. Hopkins

73

4

Synthetic Ecdysteroidal Compounds ..................................................... Josep Coll Toledano

99

Part II

5

6

In the Post-Genomic Era, Ecdysteroid Genetic Hierarchies in Insect Growth and Reproduction

Ecdysteroids and Ecdysteroid Signaling Pathways During Insect Oogenesis ........................................................ Luc Swevers and Kostas Iatrou Regulation of Sciarid DNA Puffs by Ecdysone: Mechanisms and Perspectives................................................................. Nadia Monesi, Juliana Aparecida Candido-Silva, Maria Luísa Paçó-Larson, and Jorge Cury de Almeida

127

165

vii

viii

7

8

Contents

The Ecdysteroids’ Effects in the Control of Cell Proliferation and Differentiation ............................................. David Siaussat, Patrick Porcheron, and Stépahne Debernard Applications of RNA Interference in Ecdysone Research ................. Garry N. Hannan, Ronald J. Hill, Skarlatos G. Dedos, Luc Swevers, Kostas Iatrou, Anjiang Tan, R. Parthasarathy, Hua Bai, Zhaolin Zhang, and Subba R. Palli

185 205

Part III Role of Cross Talk of Genes and Growth Factors in Ecdysteroid Titers and Signalling 9

10

The Function and Evolution of the Halloween Genes: The Pathway to the Arthropod Molting Hormone............................. Lawrence I. Gilbert and Kim F. Rewitz

231

Recent Studies on Prothoracic Gland Cell Growth and Ecdysteroidogenesis in the Silkworm, Bombyx mori .................. Shi-Hong Gu and Ju-Ling Lin

271

11

Diversity in Factors Regulating Ecdysteroidogenesis in Insects ....... Sandrien Van de Velde, Liesbeth Badisco, Elisabeth Marchal, Jozef Vanden Broeck, and Guy Smagghe

12

20-Hydroxyecdysone, Juvenile Hormone and Biogenic Amines: Mechanisms of Interaction in Control of Drosophila Reproduction Under Normal and Stressful Conditions..................... Nataly Gruntenko and Inga Rauschenbach

Part IV

283

317

Ecdysteroids Function Through Nuclear and Membrane Receptors

13

The Structure and Function of Ecdysone Receptors ......................... Isabelle M.L. Billas, Christopher Browning, Michael C. Lawrence, Lloyd D. Graham, Dino Moras, and Ronald J. Hill

335

14

The Multidimensional Partnership of EcR and USP ......................... Vincent C. Henrich, Josh Beatty, Heike Ruff, Jenna Callender, Marco Grebe, and Margarethe Spindler-Barth

361

15

Functional Analysis of Ecdysteroid Receptor from Drosophila melanogaster “In Vitro” ............................................ Anca Azoitei, Heike Ruff, Christian Tremmel, Sabine Braun, and Margarethe Spindler-Barth

377

Contents

16

Intracellular Localization of the Ecdysteroid Receptor..................... Klaus-Dieter Spindler, Katarzyna Betan´ska, Claudia Nieva, Tomasz Gwóz´dz´ , Joanna Dutko-Gwóz´dz´ , Andrzej Oz˙ yhar, and Margarethe Spindler-Barth

17

Genomic and Nongenomic Actions of 20-Hydroxyecdysone in Programmed Cell Death ................................................................... Masatoshi Iga and Sho Sakurai

18

Rapid, Non-Genomic Responses to Ecdysteroids and Catecholamines Mediated by a Novel Drosophila G-Protein-Coupled Receptor............................................. Peter D. Evans, D.P. Srivastava, and V. Reale

ix

389

411

425

Part V Ecdysteroids in Modern Agriculture, Medicine, Doping and Ecotoxicology 19

Ecdysone Receptors of Pest Insects – Molecular Cloning, Characterisation, and a Ligand Binding Domain-Based Fluorescence Polarization Screen ......................................................... 447 Lloyd D. Graham, Wynona M. Johnson, Donya Tohidi-Esfahani, Anna Pawlak-Skrzecz, Marianne Bliese, George O. Lovrecz, Louis Lu, Linda Howell, Garry N. Hannan, and Ronald J. Hill

20

SAR and QSAR Studies for In Vivo and In Vitro Activities of Ecdysone Agonists ............................................................ Yoshiaki Nakagawa, Robert E. Hormann, and Guy Smagghe

21

22

Ecdysone Receptor-Based Gene Switches for Applications in Plants ...................................................................... Venkata S. Tavva, Randy D. Dinkins, Glenn B. Collins, and Subba R. Palli Ecdysteroids and Their Importance in Endocrine Disruption Research .............................................................................. Thomas Soin, Tim Verslycke, Colin Janssen, and Guy Smagghe

475

511

539

Innovative and Future Applications for Ecdysteroids ....................... René Lafont and Laurence Dinan

551

Index ................................................................................................................

579

23

Color Plates

Fig. 3.3 Comparison of the growth of a regenerating limb bud from Uca pugilator to amounts of protein in the limb bud and total circulating ecdysteroids, as well as predominant circulating ecdysteroid. At top are molt cycle stages (see legend for Fig. 3.1). Solid line represent growth curve of limb buds as R-values (see legend for Fig. 3.1 for description). PA = ponasterone A and 20E = 20-hydroxyecdysone (Redrawn from Hopkins, 1993) (see also page 82)

xi

Fig. 3.4 Comparative binding affinities of Uca pugilator ecdysteroid receptor (UpEcR) to two isoforms of the retinoid-X-receptor (UpRXR). Surface Plasmon binding assay used. Sf9 expressed UpEcR (100nM) interacted with verious concentrations of UpRXR isoforms – larger and smaller. Response is relative amount of binding compared to control flow cells (see also page 86)

xii Color Plates

Color Plates

xiii

20E

(A) (1-2 h) (1-

EcR/USP

FTZ-F1 -F1

Early

(6 h)

HR3B HR3C

(24 h) (48-72 h)

Late

VMP30 ((vitellin membrane)

HR3 E75A

FTZ-F1

EcR (B1) E75A E75C Early-Late

(B)

HR3 EcR,USP E75C GATAβ SH3 ESP

(6 h) (12 h) (18 h)

HNF-4a

Chorion genes (24 h)

HR3A ESP

Choriogenesis

Vitellogenesis

Fig. 5.1 Regulation of silkmoth (Bombyx mori) oogenesis during pharate adult and adult development by 20E produced by the prothoracic glands. Panel a: Induction of vitellogenesis by rising titers of 20E in the hemolymph. Indicated are different phases in the hormone response: early (repression of FTZ-F1, transient induction of B1-EcR and E75C, permanent induction of E75A), early-late (induction of HR3B and HR3C) and late (induction of HNF-4A, HR3A and ESP). Expression of ESP (egg-specific protein, a yolk protein precursor produced by the follicular epithelium; Sato and Yamashita, 1991) marks the initiation of vitellogenesis. Panel b: Induction of choriogenesis by declining titers of 20E in the hemolymph. At the top of the cascade is shown the nuclear receptor FTZ-F1, which plays a pivotal role in the regulation of developmental events during low titers of ecdysone (Broadus et al., 1999). As in Drosophila, induction of FTZ-F1 may be triggered by changes in the relative levels of the HR3 and E75 receptors (Swevers et al., 2002). Expression of FTZ-F1 is followed by the repression of HR3 and EcR/USP (at 6–12 h), induction of E75C, GATAβ and SH3 (at 12 h), repression of ESP (at 12–18 h) and induction of chorion gene expression (at 18–24 h). Because the vitellin membrane protein VMP30 is co-expressed with FTZF1, it was hypothesized that FTZ-F1 is a positive regulator of the expression of VMP30 (Kendirgi et al., 2002). Note that the deduction of the regulatory cascades that trigger vitellogenesis and choriogenesis in the silkmoth is based on expression patterns of mRNAs and remains to be investigated by functional analysis (see also page 130)

xiv

Color Plates

Fig. 5.2 Model to explain the hierarchy of ecdysone response genes regulating apoptosis of stage 8 and 9 follicles in Drosophila melanogaster. Upper Panel: Complete nutrition induces normal development of follicles during oogenesis. In this case, just the Z1 isoform of BR-C is expressed in the follicle cells at stage 8. E75B suppresses E75A expression to prevent apoptosis. Middle Panel: Injection of 20E induces apoptosis in stage 8 and 9 follicles. 20E injection results first in induction of the Z2 and Z3 isoforms of BR-C which in turn decrease E75B and increase E75A expression. While E75B is an apoptosis inhibitor, E75A is an apoptosis inducer. Lower Panel: Starvation induces apoptosis in stage 8 and 9 follicles. During starvation, ecdysone concentrations increase and the Z2 and Z3 isoforms of BR-C become expressed in the follicle cells to suppress E75B and activate E75A expression. The increase in E75A results in induction of apoptosis. JH can counteract the effects of starvation by interference with the increase in ecdysone concentration and by stimulation of the expression of E75B (Reprinted from Terashima and Bownes, (2006). E75A and E75B have opposite effects on the apoptosis/development choice of the Drosophila egg chamber. Cell Death Differ. 13, 454–464. With permission from Macmillan.) (see also page 140)

Fig. 6.4 The BhEcR colocalizes with RNA polymerase II at DNA puff forming sites. Chromosome C from Bradysia hygida larvae at age E7 labeled with antibodies anti-BhEcR (red) and anti-RNA Pol II (green). Yellow signals indicate colocalization of both antibodies. From left to right, images of the same field were captured by confocal microscopy at increasing magnifications. The bars at the bottom left of each picture correspond, from left to right, to 10 µm, 5 µm and 1 µm, respectively (see also page 175)

Color Plates

xv

Fig. 7.2 Protocol of synchronization by hydroxyurea in the IAL-PID2 cell line. A high degree of synchrony was reached when cells were exposed to two consecutive hydroxyurea treatments at 1 mM for 36 h spaced 16 h apart leading to a massive arrest of cells at the transition G1/S (90%). (a) Only 2 h after the removal of the drug, 70% of cells were recovered in S. This cohort of cells began to enter into G2 at 10 h then kept on progressing through the cell cycle to re-enter progressively into G1 after 18 h. (b) Under these conditions, 20E at physiological concentration induced an inhibition of cell growth by an arrest of 90% of the cells in G2/M phase. In the bubble of each figures, corresponding cytometry profiles are presented (see also page 190)

xvi Fig. 7.10 Schematic representation of control of cell proliferation and differentiation by ecdysteroïds. At the transition S/G2, the 20E binds to EcR/USP heterodimeric complex and induces a maximum induction of these two partners then HR3. This 20E signaling pathway is responsible for G2/M arrest of cells by inhibiting the expression of B cyclin and is also involved long term in the morphological differentiation of cells through an increase in the synthesis of β tubulin (see also page 198)

Color Plates

S

20E EcR USP

HR3

Period of sensitivity to 20E

G2/M

B Cyclin β Tubulin Arrest in G2/M Differentiation

Fig. 8.3 Dose-dependent decrease in expression of BmEcR at increasing concentrations of BmEcR dsRNA. Panel A: CAT reporter assay. CAT activities are compared with that observed for the reporter construct in the absence of cotransfected dsRNA (100%). Panel B: western blot assay. The membrane was probed for the presence of BmEcR (upper) and BmUSP (lower) proteins. MW markers are indicated at the right (see also page 215)

Color Plates

xvii

Fig. 8.4 Dose-dependent decrease in expression of BmUSP at increasing concentrations of BmUSP dsRNA. Panel A: CAT reporter assay. CAT activities are compared with that observed for the reporter construct in the absence of cotransfected dsRNA (100%). Panel B: western blot assay. The membrane was probed for the presence of BmEcR (upper) and BmUSP (lower) proteins. MW markers are indicated at the right (see also page 215)

xviii

Color Plates

Fig. 8.5 Phenotypes observed after knock-down of EcR, TcE75, TcHR4, TcHr39 or Tcbr. dsRNA was injected at 24 h after ecdysis into the final instar larval stage and the phenotypes observed are shown in comparison with control. The larvae injected with TcEcR (b), TcE75 (b) or TcHR4 (c) dsRNA died during quiescent stage. In general, the dsRNA injected larvae were smaller and darker than the control larvae injected with malE dsRNA. About 50% of the larvae injected with TcHR39 successfully pupated but had problems with wing development (d). The larvae injected with Tcbr had problems undergoing larval-pupal ecdysis, the exuvium remained attached to the body (e) and these insects showed antennae, wings, legs and compound eyes similar to those found in adults (f). Scale bar: 1 mm (see also page 220)

Color Plates

xix

Fig. 9.1 Scheme of 20-hydroxyecdysone (20E) biosynthesis in Drosophila. Multiple arrows indicate an uncharacterized pathway, the Black Box. Yellow shade delineates the areas of the sterol that are involved in the transformation to the next compound (see also page 236)

xx

Color Plates

Fig. 9.2 In situ expression of the Halloween genes disembodied, shadow and phantom during late embryonic and larval stages. Shown are stage 17 embryos (a, e and i), late second instar (b, f and j), and both early (c, g and k) and late (d, h and l) third instar brain-ring gland complexes. Note the down regulation of the expression of all three genes between the late second and early third instars and their subsequent up regulation between the early and late third instars. RG, ring gland; Br, brain; VG, ventral ganglion (Data on embryonic dib expression from Chavez et al., 2000. Data on larval dib and sad expression from Warren et al., 2002. Data on phm expression from Warren et al., 2004) (see also page 239)

Color Plates

xxi

Fig. 9.3 Scheme of 20-hydroxyecdysone (20E) biosynthesis in Drosophila with gene names, gene products (enzymes) and CYP designations (see also page 241)

xxii

Color Plates

E

W

Fat Body

4

25 2

2 Commitment Peak

0

0 1

2

3

4

5

6

7

8

9

0

Day of fifth larval instar

E

W

100

0

1 2 3 4 5 6 Day of pupal-adult development

Midgut 600

400 50 Commitment Peak

25

200

4

2

E quivalents (mg/ml hemolymph)

6 75 E20MO activity

shade expression (arbitray units)

4

E quivalents (mg/ml hemolymph)

6

50 E20MO activity

shade expression (arbitray units)

6

0

0 1

2 3 4 5 6 7 8 Day of fifth larval instar

9

0

1 2 3 4 5 6 Day of pupal-adult development

Fig. 9.4 Developmental changes in Manduca shade expression in the fat body and midgut during the fifth instar and through day 6 of pupal-adult development. Expression was analyzed by qPCR and values are means +/−S.E.M. (Data from Rewitz et al. 2006b). Ecdysteroid titer data and enzyme activity are from the Gilbert and Smith laboratories with references given in the above cited publication. W, wandering; E, ecdysis (see also page 244)

Color Plates

xxiii

Fig. 9.6 Conservation of microsynteny for phm in Drosophila melanogaster (Diptera), Apis mellifera (Hymenoptera) and Daphnia pulex (Crustacea). In these species, CYP18A1 and phm are paralogs closely linked on the same chromosome (see also Claudianos et al., 2006). Arrows indicate transcriptional orientation of genes on the genome DNA sequence (horizontal lines). The size of the chromosomal regions is shown in parenthesis but distances are not to scale (see also page 255)

Fig. 9.8 Conservation of microsynteny for spo-like (CYP307A) genes in Drosophila species of the Sophophoran subgenus and Drosophila subgenus (see Fig. 9.7), Anopheles gambiae and Daphnia pulex. The intron-containing gene (spo in Daphnia and Anopheles and spok in Drosophilidae) is adjacent to neverland (nvd), a gene believed to be involved in the conversion of cholesterol to 7-dehydrocholesterol (7dC) which is the first step in the formation of 20E (Yoshiyama et al., 2006). Symbols otherwise as in Fig. 9.6 (Data on Drosophilids modified with permissionfrom Sztal et al. 2007) (see also page 258)

xxiv Color Plates

Color Plates

Fig. 9.11 Conservation profiles of ecdysteroidogenic CYP products of the Halloween genes and CYP6 proteins with the predicted substrate recognition sites (SRSs) 1–6. Note that the area encompassing SRS2 and SRS3 is shown as 2/3. n = number of aligned sequences. — indicates the position of the conserved heme-binding domain (Reproduced from Rewitz et al. 2007) (see also page 264)

xxv

xxvi

Color Plates

Fig. 10.5 Immunoperoxidase-stained whole mounts of silkworm salivary glands (a, b), and corpora allata (c, d) showing the incorporation of BrdU. (a) Salivary gland from day 0 last instar larvae incubated in 50 µl Grace’s medium containing BrdU. (b) Salivary gland from day 0 last instar larvae incubated in 50 µl Grace’s medium containing BrdU and 10% hemolymph. (c) Corpus allatum from day 0 last instar larvae incubated in 50 µl of control medium containing BrdU. (d) Corpus allatum from day 0 last instar larvae incubated in 50 µl prothoracic gland-preconditioned medium. Each incubation was maintained for 2 days, and the number of BrdU-labeled cells was counted after a 2-day incubation. BrdU-labeled cells are identified by the intense black precipitation of diaminobenzidine. Scale bar, 75 µm (Adapted from Gu, 2006) (see also page 279)

Fig. 13.2 Two types of USP LBD structures. Depicted are the LBD structures of HvUSP (in blue) and TcUSP (in brown). (a) The activation helices H12 (in red and orange for HvUSP and TcUSP, respectively) adopt an antagonist conformation. The global fold of the two types of USP structures is conserved, but significant structural differences occur, consistent with the ligand-binding status of the domain (phospholipid-bound HvUSP and ligand-free TcUSP). The phospholipid bound to HvUSP is shown in a ball representation with atoms coloured as follows: carbon atoms magenta, oxygen atoms red and phosphorus atom orange. The ligand-binding pocket is shown as a light blue semi-transparent surface. Helices H6 and H11 of HvUSP are replaced by loops in TcUSP. An important shift in the positioning of helix H3, and to a lesser degree of helix H1, can be observed. (b) Enlarged view of the region corresponding to helices H6 and H11 of HvUSP that in the case of TcUSP are reorganized into two loops (L6 and L11, respectively) that are folded onto the surface the LBD, stabilizing the apo conformation. The colour scheme of the activation helices and the bound ligand atoms follows that of Panel (a) (see also page 341)

xxviii

Color Plates

Fig. 13.3 The ligand-binding domains of EcR/USP. (a) The overall structures of the EcR/USP LBD heterodimer in complex with an ecdysteroid are depicted in two orientations for Heliothis virescens (Hv) and Tribolium castaneum (Tc), insect species belonging to the orders Lepidoptera and Coleoptera, respectively. The EcR/USP heterodimerization interface is conserved between the different species. However, differences are seen between that of the lepidopteran/dipteran EcR/ USP and those of other insects. Globally, the EcR/USP LBD heterodimer is more compact and more symmetric in lepidopteran and dipteran species (exemplified here by H. virescens) as compared to that of other insects (exemplified here by T. castaneum) as explained in the text. HvEcR and HvUSP are shown as dark green and blue ribbons respectively, and TcEcR and TcUSP as light green and brown ribbons respectively). The activation helices H12 of HvEcR and TcEcR are depicted as a red and orange helical ribbon, respectively. (b) Enlarged view of the L9-10 loop region of HvEcR and TcEcR LBDs in the context of their heterodimer with the corresponding USP LBD partner. The differences both in sequence (a two amino-acid insertion) and in conformation are indicated for the loop L9-10 of lepidopteran/dipteran EcRs (exemplified here by HvEcR) as compared to that of EcRs of other insects (exemplified here by TcEcR). Residues of loop L9-10 and a few residues of loop L7-8 that interact with this element are shown in stick representation, with carbon atoms in dark green and light green for HvEcR and TcEcR respectively, oxygen atoms in red and nitrogen atoms in blue. Yellow dotted lines indicate hydrogen bonds between residues of the protein and the colour scheme of the ribbons follows that of Panel (a) (see also page 343)

Color Plates

xxix

Fig. 13.4 Opened-out view of the major and ancillary pockets of BtEcR LBD. In these Panels the pocket surface (grey) has been open out into two “shells” by slicing in a plane parallel to that of the page. Residues lying beyond the plane of slicing are shown in each Panel, with carbon atoms in orange if they highly conserved across all insect species, otherwise with carbon atoms in green. PonA, bound in the major pocket of the LBD, is shown in stick representation with carbon atoms shown in black. Oxygen atoms in both PonA and BtEcR are shown in red and nitrogen atoms in blue. Red dotted lines indicate hydrogen bonds between the protein and the ligand. For clarity, side chain or main chain groups of individual residues are omitted from the Figure where these are not involved in the formation of the pocket surface (see also page 346)

Fig. 13.5 Residues in the region of the ancillary pocket of the EcR LBD. (a) The existence of an ancillary pocket, as observed in the BtEcR LBD, is intimately linked to the conformation of the side chains of residues (in particular HvEcR-E309/BtEcR-E199 and HvEcR-Y437/BtEcR-Y325) that fill or do not fill the volume between helices H1 and H8. HvEcR and BtEcR LBDs are depicted as dark green and yellow ribbons, respectively. Residues are shown in a stick representation, with carbon atoms in grey and yellow for HvEcR and BtEcR, respectively, oxygen atoms in red and nitrogen atoms in blue. The ligandbinding pocket of HvEcR and TcEcR are represented by a blue and yellow translucent molecular surface, respectively. In HvEcR the entrance to the cavity corresponding to the ancillary pocket of BtEcR is occluded by the side chains of HvEcR-E309 and HvEcR-Y437. (b) In the TcEcR LBD, the volume corresponding to the ancillary pocket of BtEcR is occupied by water molecules located between helices H1 and H8 and is not occluded by side chain atoms. The ligand-binding pocket was calculated without the three water molecules and is represented by a light green translucent molecular surface. The TcEcR LBD is depicted as a light green ribbon. The residues E330 and Y456 and the bound ponA ligand are shown in a stick representation with carbon atoms in green, oxygen atoms in red and nitrogen atoms in blue. The three water molecules are represented by red balls (see also page 347)

Fig. 13.6 Stereoviews of water-mediated interaction network between the ecdysteroid and EcR residues. (a) In the case of 20E bound to the HvEcR LBD, the hydrogen-bond interaction network involves the 20-, 22-, and 25-hydroxyl groups of the 20E alkyl side-chain and three structural water molecules. (b) In the case of ponA bound to the TcEcR LBD, the interaction network involves the 20- and 22-hydroxyl groups of ponA alkyl side chain and up to five structural water molecules. The HvEcR and TcEcR LBDs are depicted as dark green and light green ribbon, respectively, with individual atoms within the LBD and ligand shown in stick representation and with carbon atoms coloured in light blue and green, respectively, oxygen atoms in red, nitrogen atoms in blue and sulphur atoms in yellow. The water molecules are represented by red balls. Hydrogen bonds are indicated by yellow and pink dotted lines in (a) and (b) respectively (see also page 349)

xxxii

Color Plates

Fig. 13.7 Inter-species comparison of side-chain packing in walls of the EcR ligand-binding pocket. (a) B. tabaci EcR with ponA bound, (b) H. virescens EcR with ponA bound, (c) H. virescens EcR with BYI06830 bound and (d) T. castaneum EcR with ponA bound. The orientations of the EcR-LBDs in Panels (a–d) are identical, with the dashed line indicating a common line of in-plane register. Panel (a) shows (in rod representation) the side chains of B. tabaci residues I230, M272, T304, L308, M389, T393 and V404, whilst Panels (b) and (c) show their counterparts of these residues in H. virescens EcR (namely M342, V384, V416, L420, Q503, M507 and L518, respectively), and Panel (d) their counterparts in T. castaneum (namely I361, M403, T435, L439, Q520, M524 and L535, respectively). Atoms in these residues are coloured as follows: carbon atoms green, oxygen atoms red, nitrogen atoms blue and sulphur atoms yellow, with the position of the Cα atom of each residue being highlighted by a sphere for clarity. Also shown in each panel is the ligand-binding pocket (coloured transparent gold and calculated with a 1.2 Å probe radius), encapsulating the respective bound ligand. Ligand atoms are coloured according to the same scheme as that of the EcR side chains, except that carbon atoms are shown in black. Panels (e), (f), (g) and (h) show the atomic packing of the ligand-binding cavity wall in the vicinity of the residues highlighted to the left of the ligand in Panels (a–d) respectively, but viewed from the interior of the binding pocket and looking in the direction of the arrows. Atoms are shown in CPK representations and include both main and side chain atoms from all residues contributing to the immediate wall of the binding pocket in the direction of the view. The site of the protrusion into the protein atomic volume of the ligand-binding pocket walls of H. virescens EcR and T. castaneum EcR in the vicinity of the alkyl tail of bound ponA (see text) is indicated by an asterisk in Panels (b), (d), (f) and (h). The site of deeper protrusion into the protein atomic volume of ponA-bound HvEcR LBD in the vicinity of the C4 atom of the ligand is indicated by a solid star in Panel (b); this deeper protrusion with respect to the pocket walls of ponA-bound BtEcR LBD and TcEcR LBD is a consequence of the substitution of BtEcR I230 and TcEcR I361 by methionine in HvEcR (M342, see text) (see also page 350)

Color Plates

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Fig. 13.8 Stereoviews of the differential conservation of the pocket residues HvEcR-V384 and HvEcR-V416 in Lepidoptera. (a) HvEcR-V384, which is replaced by methionine in non-lepidopteran EcRs, allows the specific side chain conformation of HvEcR-L420 and a closer van der Waals’ interaction with the ecdysteroid 22-hydroxyl group. (b) HvEcR-V416, which is replaced by threonine in TcEcR and BtEcR, is also closer to the ecdysteroid 22-hydroxyl group due to the concomitant shift of helix H7 towards the interior of the receptor pocket. The stereoviews depict HvEcR in blue, TcEcR in green and BtEcR in yellow. Amino acid residues of HvEcR, TcEcR and BtEcR are shown in stick representation colour with carbon atoms in light blue, green and yellow for the three species respectively and oxygen atoms in red. The ligands 20E (for HvEcR) and ponA (for TcEcR and BtEcR) are shown in a stick representation with carbon atoms in blue (20E), green (ponA in TcEcR) or wheat (ponA in BtEcR) and oxygen atoms in red. The activation helix H12 is also shown as a red ribbon (see also page 351)

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Color Plates

Fig. 13.9 Flexible region in the EcR LBD exploited by DBH compounds. (a) Ribbon diagram showing the superimposition of the structures of HvEcR-LBD in complex with 20E (dark green ribbons) and with BYI06830 (orange ribbons). The view is restricted to the region differing the most between the two EcR-LBDs that includes helices H2, H6, H7 and the β-sheet. 20E and BYI06830 are shown in stick representation with carbon atoms in cyan and light grey, respectively, oxygen atoms in red and nitrogen atoms in blue. (b) Superimposition of 20E and BYI06830 bound to HvEcR. Oxygen atoms are shown in red, nitrogen atoms in blue, 20E carbon atoms in cyan and BYI06830 carbon atoms in olive. The three structural water molecules observed in the structure of 20E-bound HvEcR, shown by red balls, superimpose well with the region of the B-ring of the DBH compound (see also page 353)

Color Plates

xxxv

Fig. 13.10 The steric relationship between the alkyl side chain of ponA in the binding pocket of the B. tabaci ecdysone receptor LBD and the side chain sulphydryl group of BtEcR C394. The distance between the van der Waals’ surfaces of the terminal side chain carbon atom of the steroid and the sulphur atom of the cysteine residue C394 in the pocket wall is 0.5 Å. This cysteine residue is strictly conserved among all known EcR sequences (see also page 356)

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Color Plates

Steroid c

a b

Nuclear receptor at plasma membrane Nuclear receptor

GPCR

cyclic AMP Calcium MAPK

5 4 TMTM 3 TM TM 6 TMTM 1 2 TM 7

α GDP

γ β

mRNA

Protein Synthesis Fig. 18.1 Steroid hormone signaling. (a) Conventionally steroid hormones are thought to be lipophylic hormones and to be able to enter the cell easily. They then bind with a range of ‘nuclear’ receptors which transport them into the nucleus where the complex binds to the DNA and initiates changes in gene expression. Non-genomic actions of steroids are very rapid and mediate changes in second messenger levels without any changes in gene expression or protein synthesis. (b) It has been postulated that these rapid non-genomic effects could be mediated via steroid interactions with the nuclear binding proteins that migrate to become closely associated with the plasma membrane. Once activated the complex is then proposed to bring about rapid changes in second messenger levels. (c) It has also been proposed that steroids may interact with specific seven transmembrane-spanning GPCRs to activate second messenger pathways via the activated G-proteins (see also page 426)

Color Plates

xxxvii Rapid Non-Genomic Effects

E

DA CG18314

G1 G2

Cyclic AMP AKT (PI3Kinase )

MAPKinase

E

Early Genomic Effects Development

ER

USP

Late

Fig. 18.3 The orphan Drosophila GPCR (CG18314) responds to both ecdysteroids and to the catecholamine dopamine and has been renamed DmDopEcR (Srivastava et al., 2005). DmDopEcR shows ‘agonist-specific coupling’ whereby the catecholamine dopamine can couple the receptor to an increase in cyclic AMP levels together with an activation of the PI3Kinase pathway, as judged by the increased phosphorylation of Akt. Ecdysteroids can produce rapid non-genomic effects through this receptor by coupling it to the activation of the MAPKinase pathway and by inhibiting the actions of dopamine. The receptor has a much higher affinity for ecdysteroids compared to dopamine (see also page 435)

Fig. 20.4 CoMFA electrostatic and steric field contour plots for DAH toxicity toward C. suppressalis, S. exigua, and L. decemlineata. The diacylhydrazine with X = 2-Cl and Y = 2,3-di-Cl is depicted. Green/yellow polyhedra represent sterically-favored/disfavored regions; blue/red polyhedra represent regions where positive charge is favored/disfavored (Reproduced with the permission of Wiley-VCH Verlag GmbH & Co. KGaA) (see also page 489)

Fig. 20.5 CoMFA hydrogen-bond field contour plots for toxicity toward C. suppressalis, S. exigua, and L. decemlineata. The diacylhydrazine with X = 2-Cl and Y = 2,3-di-Cl (Fig. 20.3) is depicted. Green/yellow polyhedra represent hydrogen-bond acceptor-favored/disfavored regions; blue/red polyhedra represent hydrogen-bond donor-favored/disfavored regions (Reproduced with the permission of Wiley-VCH Verlag GmbH & Co. KgaA) (see also page 490)

Fig. 20.7 Stereoviews of CoMFA fields with tebufenozide. (a) The contours are shown to surround regions where a positive (blue) and a negative (red) electrostatic potential increases the activity. (b) The contours are shown to surround regions where a higher steric bulk increases (green) or decreases (yellow) the activity (Reproduced with the permission of Elsevier) (see also page 499)

Color Plates

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Fig. 20.8 The docking of tebufenozide to the cavity of BmEcR -LBD. Hydrophobicity gradient is shown from blue (lowest/–0.1) to brown (highest/0.1). Brown indicates receptor surface that is most hydrophobic, and blue indicates the surface area that is least hydrophobic. Hydrophobic amino acid residues (M409, M503, L507, L515) are surrounding the t-butyl group (Reproduced with the permission of Elsevier) (see also page 501)

Fig. 20.9 The docking of tebufenozide to the LBD of BmEcR with CoMFA steric fields (Reproduced with the permission of Elsevier) (see also page 501)

xl

Color Plates

Fig. 20.10 Stereoview of the tebufenozide in the binding cavity of modeled BmEcR with the neighboring hydrophilic amino acids (Reproduced with the permission of Elsevier) (see also page 502)

Color Plates

xli

Fig. 21.3 Methoxyfenozide-inducible AtZFP11 phenotype in Arabidopsis and tobacco. Seed collected from the transgenic Arabidopsis and tobacco plants expressing AtZFP11 gene under the control of CfEcR + LmRXR two-hybrid gene switch were plated on agar media supplemented with different concentrations of methoxyfenozide. (a) Wild-type Arabidopsis seedlings growing on agar media. (b) T3 transgenic Arabidopsis seed germinated on agar media containing DMSO and 50 mg/l kanamycin. (c) T3 transgenic Arabidopsis seed germinated on agar media containing 400 nM methoxyfenozide and 50 mg/l kanamycin. (d–g) Microscopic pictures of the T3 transgenic Arabidopsis seedlings collected from different methoxyfenozide treatments. (d) 0 (DMSO), (e) 16, (f) 80 and (g) 400 nM. (h–m) Photographs of the T2 transgenic tobacco seedlings and plants treated with different concentrations of methoxyfenozide. (h) Wild-type tobacco seedlings growing on agar media. (i) T2 tobacco seed germinated on agar media containing DMSO and 300 mg/l kanamycin. (j) T2 tobacco seed germinated on agar media containing 400 nM methoxyfenozide and 300 mg/l kanamycin. (k–m) Transgenic tobacco plants growing in the greenhouse. (k) DMSO, (l) and (m) 400 nM methoxyfenozide (see also page 533)

xlii

Color Plates

crude plant extracts (elixirs or powders)

purified ecdysteroids (or ecdysteroid mixtures)

ecdysteroids + other substances (e.g. isoflavones) and/or proteins

transdermal applicator

and cosmetic uses

Fig. 23.2 More or less pure phytoecdysteroids as dietary supplements for farm and domesticated animals and for humans as food supplements for healthy or sick people for use as additives in cosmetics (see also page 571)

Chapter 1

Phytoecdysteroids: Diversity, Biosynthesis and Distribution Laurence Dinan, Juraj Harmatha, Vladimir Volodin, and René Lafont

Abstract We review the current status of knowledge on phytoecdysteroids, with particular emphasis on their occurrence, diversity, functions, biosynthesis and biotechnological production. We also consider evolutionary aspects with regard to their probable role in the deterrence of phytophagous invertebrates and in respect to their relationship to other classes of phytosteroids. Keywords Allelochemical • biosynthesis • cell culture • deterrence • distribution • diversity • ecdysteroid • insect-plant relationships • phytosteroid Abbreviations 2dE: 2-deoxyecdysone; 22dE: 22-deoxyecdysone; 2,22dE: 2,22-dideoxyecdysone; 2,22,25dE: 2,22,25-trideoxyecdysone; 20E: 20-hydroxyecdysone; BR: Brassinosteroid; DAH: Diacylhydrazine; E: Ecdysone; LBD: Ligand-binding domain; makA: Makisterone A; NMR: Nuclear magnetic resonance; poA: Ponasterone A; polB: Polypodine B.

1.1

Introduction

The discovery of ecdysteroids in plants (phytoecdysteroids) followed soon after the elucidation by X-ray analysis of the structure of ecdysone (Huber and Hoppe, 1965). Almost simultaneously, four independent laboratories reported L. Dinan and R. Lafont () Laboratoire Protéines: Biochimie Structurale et Fonctionnelle, Université Pierre et Marie Curie, CNRS FRE 2852, F-75252 Paris cedex 05, France e-mail: [email protected] J. Harmatha Academy of Sciences of the Czech Republic, Institute of Organic Chemistry and Biochemistry, CZ-16610 Prague, Czech Republic V. Volodin Biochemistry and Biotechnology Laboratory, Biology Institute, Komi Science Centre, Ural Department of the Academy of Science of the Russian Federation, Syktyvkar, Russia G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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the isolation of closely related molecules from two gymnosperms (Galbraith and Horn, 1966; Nakanishi et al., 1966), one fern (Jizba et al., 1967) and one angiosperm (Takemoto et al., 1967a). Such an unexpected finding was the impetus for extensive investigations of many plant species to examine the distribution of these molecules in the plant kingdom (see Ecdybase [Lafont et al., 2002] for an extensive bibliographic survey of >2,350 species; Dinan and Lafont, 2007). At that time, there was great hope that ecdysteroids would form the basis for a new class of safe and specific endocrine disruptors for controlling insect pests. Indeed, these molecules are able to interfere with insect development/reproduction (see Chapter 6), but for various reasons (the complexity of the molecules, the difficulty of their chemical synthesis, environmental and metabolic lability, penetration problems) this has not led to any practical use of ecdysteroids per se in the area of pest control (which is not the case for synthetic ecdysteroid agonists [e.g. diacylhydrazines], which have much simpler, and unrelated, structures). These extensive investigations allowed the isolation of a wide array of molecules (see Ecdybase), i.e. more than 300 different phytoecdysteroids, and showed the surprisingly wide distribution of these molecules in the plant kingdom (Dinan, 2001). The presence of ecdysteroids is not restricted to terrestrial species, and several aquatic plants also contain ecdysteroids (Chadin et al., 2003), which may be ecologically relevant (see Chapter 23 of this book). Ecdysteroids are also present in fungi (reviewed in Kovganko, 1999), and closely related molecules (pinnasterols) have been isolated from a red alga (Fukuzawa et al., 1981, 1986). Ecdysteroids are somewhat related to brassinosteroids (BRs), i.e. plant growth steroid hormones, in their chemical structures, although it is apparent that this similarity is not so strong when one considers 3D-representations of the molecules. While BRs may act as weak ecdysteroid antagonists in insects (Dinan and Hormann, 2005), the reverse is not true, since ecdysteroids do not appear to be active on BR receptors. This makes biological sense when considering (i) the uneven distribution of phytoecdysteroids in the plant world and (ii) the huge concentrations of ecdysteroids reached in certain plant species, when compared with the minute amounts of BRs (Adam and Marquardt, 1986). This does not however preclude other biological activities of ecdysteroids in the plant itself (Lehmann et al., 1988), although convincing experimental evidence for this is lacking (Dinan, 2001). In this chapter, we shall first consider the diversity and the distribution of ecdysteroids. We will then discuss in detail the biosynthesis of these molecules and identify the remaining open questions. We will then consider the in vitro systems developed for the production of ecdysteroids. The putative allelochemical functions of phytoecdysteroids will then be examined. Finally, we shall consider phytoecdysteroids in relation to the diversity of non-ecdysteroidal plant sterol derivatives.

1 Phytoecdysteroids: Diversity, Biosynthesis and Distribution

1.2 1.2.1

5

Distribution and Diversity of Phytoecdysteroids General

The literature concerning this aspect has been comprehensively reviewed recently (Lafont, 1997; Dinan, 2001). Thus, the main features of the distribution of phytoecdysteroids within the world flora and their distribution within plants will be just briefly summarised, with emphasis on publications appearing since 2001, and attention will be drawn to the larger surveys of plant species to aid those wishing to ascertain if particular species have already been investigated for the presence of ecdysteroids. A good resource in this respect is Ecdybase (http://www.ecdybase.org), which provides chemical and biological data for ecdysteroid analogues and listings of the literature concerning the occurrence of ecdysteroid in plants, their effects on various organisms and their applications. Ecdysteroids or ecdysteroid-like compounds have been found in gymnosperms, angiosperms, fungi, algae and certain marine organisms, in addition to arthropods and other invertebrate groups (see Chapter 2). The most extensive studies of ecdysteroid distribution within these categories of organisms have focussed on terrestrial plants, but even here only a very small percentage of the world’s 250,000 species have been analysed, and, of course, the published literature tends to report only the ecdysteroid-positive species, rather than those not found to contain them. Additionally, one needs to take the method and protocol used to detect ecdysteroids into account, since they can possess very different thresholds for detection, as plants can contain vastly different levels of ecdysteroids, and the method may detect only particular types of ecdysteroids. Thus, a new class of ecdysteroid conjugate (glucosyl-ferulate) has recently been detected in the fern Microsorum membranifolium (Ho et al., 2008), which could easily have been missed in earlier studies on other species. Also, one has to ask if the ecdysteroids are naturally produced by the organism. For example, do fungi produce mycoecdysteroids de novo, or do they represent modifications of phytoecdysteroids taken up by the fungus? Only in a few terrestrial plants has the biosynthetic capacity properly been demonstrated (see later).

1.2.2

Phytoecdysteroid Distribution Within the Natural World

Within the higher plants, ecdysteroids are found in gymnosperms and angiosperms (mono- and di-cotyledonous plants). It appears that 5–6% of terrestrial plant species contain significant levels of ecdysteroids (Imai et al., 1969; Dinan, 1995). However, use of sensitive immunoassays has suggested their presence in the leaves of 40% of randomly selected species

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(Dinan et al., 2001a). Additionally, individual ecdysteroid-positive plants in species generally regarded as ecdysteroid-negative (e.g. Arabidopsis thalliana; Dinan et al., 2001a) can be found. Put together, this suggests that most, if not all, plants retain the genetic capacity to produce ecdysteroids, but that the accumulation is suppressed in most species. It was originally thought that there was a preponderance of ecdysteroid-containing species amongst the ferns, but it has come to be realised that this just reflected the large number of ferns in early surveys (Hikino et al., 1973; Yen et al., 1974). Few plant families do not contain at least some ecdysteroid-positive species; where none has been found, it is probably because the family is very small, or that too few species have been examined. Levels of phytoecdysteroids in ecdysteroid-positive plant species vary enormously, from the barely detectable to the staggeringly high, where ecdysteroids can make up 2–3% of the dry weight (e.g. seeds of Rhaponticum [Leuzea] carthamoides [Koudela et al., 1995] or stems of Diploclisia glaucescens [Bandara et al., 1989]). Even low concentrations may be ecologically significant in the deterrence of invertebrate predators, since they may act in synergy with other classes of secondary compounds to bring about effective deterrence. Equally, there is not a simple relationship between the frequency of occurrence of ecdysteroid-positive species in a genus/family and the levels found in individual species. Thus, there are a few individual species in the Compositae (Leuzea carthamoides, Serratula coronata), which contain very high levels of ecdysteroids, but in general the vast majority of composites are ecdysteroid-negative (Dinan, unpublished). On the other hand, in the genus Silene, where there is a high preponderance of ecdysteroid-positive species, there are several ecdysteroid-rich species (Zibareva et al., 2003). As more species are examined within certain large families/genera, patterns are beginning to be discerned between their presence/absence and the taxonomic position of the species e.g. in the Chenopodiaceae (Dinan et al., 1998) or the genus Silene (Zibareva et al., 2003).

1.2.3

Phytoecdysteroid Distribution Within Plants

Phytoecdysteroid levels are not uniform in ecdysteroid-positive species. Rather, they vary from organ to organ, and can undergo changes according to season or geographical location. Further, the ecdysteroid profiles of different organs may vary. It is not known where in the plant cell or tissue that ecdysteroids accumulate, although it is often speculated that it is in the vacuole. Data on the relative biosynthetic capacity of different plant organs are also scarce, as is information about the kinetics and mechanisms of synthesis and distribution around the plant. It has been suggested that the qualitative and quantitative aspects of ecdysteroid presence or profiles could be used for chemotaxonomic purposes (Zibareva, 2000). However, the fluctuations associated with organ type, season and geographical

1 Phytoecdysteroids: Diversity, Biosynthesis and Distribution

7

location may confound this. In spite of this, levels and profiles in seeds may truly have some potential in this regard (Dinan et al., 1998; Zibareva, 2000). The taxonomic value of ecdysteroids in the chemosystematics of mushrooms, particularly in genera Paxillus and Tapinella, has been discussed and used as an additional chemosystematic character (Vokácˇ et al., 1998a, b). Evidence is gradually accumulating that ecdysteroid concentrations are highest in tissues which are most important for the survival of the plant, or, in the case of annuals, of the species into the next generation. Thus, in the windpollinated annual Chenopodium album, high levels of ecdysteroids are found in the anthers protecting the developing pollen, in the seeds and in young leaves (Dinan, 1992).

1.2.4

Large-Scale Surveys of Phytoecdysteroid Distribution

It is estimated that there are more than 250,000 species of terrestrial plants, but only a small percentage of these has been surveyed for the presence of phytoecdysteroids. Further, since the possibility for varietal, developmental, ecological or geographical variation of ecdysteroid content exists within a species, very few species indeed have been examined under different circumstances, and certainly not enough to be able to state categorically that a particular species does not accumulate ecdysteroids under any circumstances. The first large-scale survey was conducted by Imai et al. (1969) who assessed methanolic extracts of 1,056 taxonomically diverse species and 351 crude drug preparations by means of the lepidopteran Chilo suppressalis ‘dipping’ test. They found that 24 (from 47) pteridophytes, 7 (from 42) gymnosperms and 30 (from 967) angiosperms were positive. The publication names and presents data only for the positive species. Hikino et al. (1973) examined methanolic extracts of 283 species (871 specimens) of Japanese ferns by means of the dipteran Sarcophaga peregrina test, whereby the plant extract is injected into the abdomen of ligated last instar larvae to see if pupariation is induced (positive response). 170 species proved positive, of which 51 gave high activity. The paper reports the data for all positive and negative species. The Sarcophaga test was also employed by Yen et al. (1974), who assessed methanolic extracts of 115 species (164 specimens) of Taiwanese ferns. Sixty-four species were positive, of which 18 were highly so. Again, all the tested species are named. During the period 1995–2002, the Insect Biochemistry Group at Exeter University conducted a survey for the presence of ecdysteroid agonist and antagonist activities in a large number of plant extracts. The plant material (25–30 mg dw) was extracted with methanol, partially purified by addition of water (30%) and partition of the aq. methanol phase against hexane. The aq. methanol phase was then assessed by means of the dipteran Drosophila melanogaster BII cell-based microplate assay for ecdysteroid agonist and antagonist activities (Clément et al., 1993) and in two or three ecdysteroid-specific immunoassays (Dinan, 1995).

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Phytoecdysteroid-containing extracts were positive in the agonist bioassay and in the immunoassays. The plant material surveyed consisted of: • • • • • • • •

2,454 randomly selected seed samples 2,111 targeted seed samples Parts of plants grown from 180 randomly selected species (Dinan et al., 2001a) Seeds of 290 species assessed for the presence of hydrolysable ecdysteroid conjugates (Dinan et al., 2001a) Circa 200 species of the Chenopodiaceae (Dinan et al., 1998) Seeds and plant parts of 128 species of the Solanaceae (Savchenko et al., 2000) Circa 110 species of the Caryophyllaceae (Zibareva et al., 2003) 470 species of the Compositae

Some of this data has been published over the years in the scientific literature (e.g. Savchenko et al., 1997, 1998, 2001; Whiting et al., 1998; Dinan et al., 2001b, c, 2002), and detailed summary tables of the findings have now being incorporated into the Ecdybase website (http://www.ecdybase.org), which permits one, with significant certainty, to identify the species which contain (i) phytoecdysteroids (agonist bioassay positive, RIA-positive; 5–6% of species based on seed extracts), (ii) ecdysteroid antagonists (antagonist bioassay positive, RIA-negative; 2% of species), or (iii) non-steroidal agonists (agonist bioassay positive, RIA-negative; very rare), and to use the data to consider the taxonomic significance of the distribution of ecdysteroid agonists/antagonists. The approach of the Exeter Survey has been extended by the Biochemistry and Biotechnology Laboratory of the Komi Research Centre to examine the ecological and geographical distribution of ecdysteroid-containing plant species in a specific geographical region (European North-east Russia). The findings of this study have been published (Volodin et al., 2002; Volodin, 2003). Seven hundred samples collected from eight geographical locations and representing 411 species were investigated. Four percent of the species tested positive for phytoecdysteroids. All the data are presented and analysed for the impact of ecological/geographical factors on phytoecdysteroid distribution.

1.2.5

Diversity of Phytoecdysteroids

To date, somewhere in the region of 300 different phytoecdysteroids have been identified (Lafont et al., 2002). Ecdybase provides an up-to-date listing of all their structures, together with spectral and biological data, where available. The diversity of ecdysteroid analogues has also been reviewed previously (Lafont and Horn, 1989; Lafont, 1997, 1998; Dinan, 2001). They differ in the number of C-atoms present (24C–29C), the number, position and location of hydroxyl and keto groups on the steroid skeleton, whether the ecdysteroid is free or conjugated and, if conjugated, whether the conjugating moiety is polar or non-polar. Multiple

1 Phytoecdysteroids: Diversity, Biosynthesis and Distribution

9

conjugates also exist. This diversity is summarised in Fig. 1.1. By far the most commonly occurring phytoecdysteroid is 20-hydroxyecdysone, followed by polypodine B. Typically, a phytoecdysteroid-containing species will contain 1–3 major ecdysteroids, which together make up ca. 95% of the total ecdysteroid, together with a plethora of minor ecdysteroids, forming a diverse ‘cocktail’ of ecdysteroid structural analogues. The number of minor ecdysteroids may be very large, since in few cases have only the most major of them been isolated and identified. In Leuzea carthamoides, which is rich in ecdysteroids, has been extensively studied and where very large amounts of plant material have been extracted to provide a commercial source of the major ecdysteroids present, the total number of ecdysteroid analogues so far identified is >20 (Budeˇšinský et al., 2008). Figure 1.2 demonstrates the ecdysteroid profile present in fronds of the fern Microsorum membranifolium (Ho et al., 2008), including the presence of novel ecdysteroid glucosylferulate conjugates. This species is characterised by high ecdysteroid concentrations (0.65% of the dry weight of the fronds) and, unusually, by significant proportions of a number of ecdysteroid analogues, where 20E is not the major component. Although far fewer ecdysteroids have been isolated from fungi (mycoecdysteroids) than from plants, it would appear that, although there is some overlap in the structural analogues found (20E, ajugasterone C etc.), some structural variations (e.g. side-chain epoxide groups) appear to be unique to fungi. It is currently not clear if fungi possess the ability to generate ecdysteroids from sterols, or whether they take in phytoecdysteroids from their substratum and modify them. OH

a

OH

OH OH

OH

OH

29C

28C

27C

HO

HO

HO OH

OH HO

OH

OH

OH

OH HO

HO H O

H O

Makisterone C

Makisterone A

H

O

20-Hydroxyecdysone

O

24C

O

HO

O

21C

HO OH HO H O

Sidisterone

O

HO

OH HO

19C

OH HO

H O

Poststerone

H O

Rubrosterone

Fig. 1.1 Summary of the diversity of phytoecdysteroid structures, show examples of variety in steroids (panel a)

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Fig. 1.1 (continued) and conjugating moieties (panel b).The numbers in brackets in panel B refer to the C-atoms of the steroid which can be modified

1 Phytoecdysteroids: Diversity, Biosynthesis and Distribution

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50000

UV Absorbance

2d20E

2dE

20E

2

E

3

1

0

10

20

30 time (min) 40

Fig. 1.2 Ecdysteroid profile in fronds of Microsorum membranifolium as determined by C18 RP-HPLC (15 cm × 4.6 mm; 5 µm particle size), eluted at 1 ml/min with a gradient from 15–30% acetonitrile in water over 40 min; peak 1 = 2dE 25-rhamnoside, peak 2 = 2d20E 3-glucosylferulate and peak 3 = 2dE 3-glucosylferulate

1.3

1.3.1

Pathways and Regulation of Phytoecdysteroid Biosynthesis Introduction

Studies on phytoecdysteroid biosynthesis started ca. 30 years ago, i.e. soon after their discovery, and, in fact, at about the same time as in insects. Given the large amounts of edysteroids found in some plant species, it could be thought that elucidation of the pathway could be easily addressed. In reality, the high amounts are the result of ecdysteroid accumulation over a long period, which does not require such high rates of synthesis when compared to that in insect moulting glands; moreover, the sites of ecdysteroid production have not been defined, and we do not know whether biosynthesis takes place in all, or only is some, specialized cells (yet to be identified). It will become possible to address such a question only with a molecular biological approach, when genes encoding some of the biosynthetic enzymes have been identified, which is not yet the case for the following reasons: (1) none of the plant species for which genomic sequence information is available accumulate ecdysteroids and (2) unlike brassinosteroids, we cannot expect that mutants will

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display a specific phenotype. Thus, very few attempts have been made so far to purify biosynthetic enzymes (e.g. ecdysone 20-monooxygenase) using classical biochemical approaches (Grebenok et al., 1996; Canals et al., 2005).

1.3.2

Methods

Ecdysteroids are derived from sterols, and different strategies can be used to elucidate their biosynthetic pathway, which are aimed at providing different types of information: 1. Use of radioactive (3H or 14C) molecules and testing whether they are converted into ecdysteroids (mainly 20-hydroxyecdysone); this can be done with very early sterol precursors (acetate, mevalonate), sterols (cholesterol) or any available putative intermediate; radioactive intermediates/substrates can also be used to characterize enzymes (organ distribution, subcellular localization, cofactors etc.) involved in some biosynthetic steps (e.g. hydroxylases). 2. Use of doubly-labelled (3H + 14C) molecules, where tritium is introduced at specific positions of the A/B-rings with the aim of understanding the mechanism of the formation of the (5β−H)14αOH-7-en-6-one structure, or the stereochemistry of a given reaction (e.g. 7-dehydrogenation). 3. Use of molecules labelled with stable isotopes (2H, 13C) followed by NMR analysis of 20-hydroxyecdysone; this gives information about effective incorporation and, in the case of 2H, it allows the establishment of a possible migration of deuterium atoms to another position.

1.3.3

Biological Systems

In addition, labelling experiments are faced with the problem of compound delivery to/uptake by the plant. Topical applications have a limited efficiency, owing to the low permeability of plant cuticle. Whole plant experiments can allow uptake by the roots. Excised leaves can be efficiently loaded through their petiole thanks to evaporation. Hairy roots (from Serratula tinctoria [Delbecque et al., 1995; Corio-Costet et al., 1996] or Ajuga nipponensis [Fujimoto et al., 2000]) and ecdysteroid-producing cell cultures from Ajuga reptans and Serratula coronata (Filippova et al., 2003) should provide more efficient biological systems. Polypodium vulgare prothalli also represent a good model for metabolic studies (Reixach et al., 1996, 1997, 1999).

1.3.4

Biosynthetic Pathway(s)

Early on, it was shown that cholesterol was a precursor of C27 ecdysteroids, but this may not be true for all species; for instance, Spinacia oleracea contains mainly Δ7-sterols and lathosterol (a Δ7-sterol) is a more likely ecdysteroid precursor (Grebenok

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13

and Adler, 1993). Acetate and mevalonate are also converted into ecdysteroids, and acetate may give C27-, C28- and C29-ecdysteroids (Tomás et al., 1993) (Table 1.1). It is conceivable that C28- or C29-sterols are the precursors for the corresponding C28and C29- ecdysteroids (e.g. clerosterol for cyasterone; Okuzumi et al., 2003). The use of hydroxylated cholesterol derivatives (25-hydroxycholesterol, 22R-hydroxycholesterol) has been essentially restricted to Polypodium vulgare prothalli and these molecules were very efficiently converted into ecdysone and 20-hydroxyecdysone in this fern (Reixach et al., 1999), whereas 25-hydroxycholesterol labelling experiments with Podocarpus elata (Joly et al., 1969), Serratula tinctoria hairy roots (J.-P. Delbecque, personal communication), or Silene otites seedlings (R. Lafont, unpublished data) were negative. In P. vulgare, this means that early hydroxylation at the 22R- or 25-positions does not prevent further reactions taking place. The terminal steps have been investigated by using available tritium-labelled intermediates previously used with insects and crustaceans (5β-diketol, 5β-ketodiol, 2,22dE, 2dE, E etc.), but the limited number of species investigated does not allow general conclusions to be drawn. In P. vulgare, 5β-ketodiol and 2,22dE were converted only up to 22dE, but no further, which means that 22-hydroxylation must take place at an early stage (in this species at least, but not in Achyranthes fauriei, Ajuga reptans or Spinacia oleracea). On the other hand, 25-hydroxylation may take place at any time in P. vulgare (where both 25-hydroxycholesterol and ponasterone A [25-deoxy-20-hydroxyecdysone] can be converted to 20E), whereas 25-hydroxycholesterol is not converted in other species. These two examples illustrate possible differences in the biosynthetic pathway among plant species (pteridophytes vs. spermatophytes), possibly connected with the narrow substrate-specificity of some hydroxylases. In fact, similar differences have also been recorded in the case of arthropods. 20-Hydroxylation may also be a point of divergence between plants and animals; this step is the last one in insect larvae, because it takes place outside the moulting glands (e.g. in the fat-body), but there is no reason to think that the whole plant pathway to 20E should not take place in the same cell, thus allowing 20-hydroxylation to take place at an earlier stage (hence the presence of 22-deoxy20-hydroxyecdysone [taxisterone] in several plant species. However, it is possible that in some animal systems the same situation exists, e.g. in the case of Bombyx ovaries where, for instance, 2,22d20E has been isolated (Ikekawa et al., 1980), or the presence of taxisterone in a marine arachnid, Pycnogonum litorale (Bückmann et al., 1986). The few enzymatic studies performed with subcellular fractions (mitochondria, microsomes, cytosol) have shown differences with insects concerning the localization of some hydroxylases or oxidoreductases (Reixach et al., 1999; Bakrim, 2007). During metabolic studies, special attention was given to the, as yet unsolved (but in insects too), early-step problem, i.e. the formation of the (5β−H)14αOH7-en-6-one chromophore characteristic of ecdysteroids. This question was initially addressed in parallel experiments performed on plants and insects, at first by using putative intermediates (cholest-4-en-3-one, cholesterol 5α,6α-epoxide, cholesterol 5β,6β-epoxide), then doubly-labelled cholesterols: [4-14C,3α-3H] to test whether a 3-oxo-Δ4 intermediate was formed (Lloyd-Jones et al., 1973), [4-14C,7α-3H], [4-14C,7β-3H] to understand the stereochemistry of Δ7-bond formation

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Table 1.1 Biosynthetic studies performed with radioactive or non-radioactive labelled precursors LABELLED ECDYSTEROIDS PRECURSOR PLANT SPECIES PRODUCED REFERENCE 20E + Cyasterone + Tomás et al., 1993 Acetate Ajuga reptans 29-Norcyasterone Sipahimalani et al., 1972 Sesuvium E + 20E portulacastrum Grebenok and Adler, 1993 20E + Polypodine B Spinacia oleracea Mevalonate

only sterols, no Cyasterone E + 20E + Polypodine B 20E 20E E + 20E

Boid et al., 1975

20E + Polypodine B, E + E conjugate 20E + Ponasterone A

Grebenok et al., 1994

Taxus baccata Ajuga reptans

20E

Nagakari et al., 1994a, b; Yagi et al., 1996; Fujimoto et al., 1997; Nakagawa et al., 1997; Nomura and Fujimoto, 2000

Podocarpus elata

20E

Sauer et al., 1968; Joly et al., 1969

Podocarpus macrophyllus

20E + Ponasterone A

Hikino et al., 1970

Polypodium vulgare

20E

Cook et al., 1973; Lockley et al., 1975; Davies et al., 1980; Reixach et al., 1996

Serratula tinctoria

E + 20E

Delbecque et al., 1995

Sesuvium portulacastrum

E + 20E

Sipahimalani et al., 1972

Spinacia oleracea

20E + Polypodine B

Grebenok et al., 1994

Taxus baccata

20E

Lloyd-Jones et al., 1973; Cook et al., 1973

Zea mays

20E conjugate

Devarenne et al., 1995

Podocarpus elatus

no conversion

Sauer et al., 1968

Sesuvium portulacastrum

no conversion

Sipahimalani et al., 1972

7-Dehydrocholesterol

Ajuga reptans

20E (low)

Ohyama et al., 1999

25-OH-Cholesterol

Podocarpus elata Polypodium vulgare

None E + 20E

Joly et al., 1969 Reixach et al., 1999

Cyathula capitata Polypodium vulgare Serratula tinctoria Sesuvium portulacastrum Spinacia oleracea

Cholesterol

Cholest-4-en-3-one

De Souza et al., 1970 Reixach et al., 1996 Delbecque et al., 1995 Sipahimalani et al., 1972

De Souza et al., 1969

(continued)

1 Phytoecdysteroids: Diversity, Biosynthesis and Distribution Table 1.1 (continued) LABELLED PRECURSOR

PLANT SPECIES

ECDYSTEROIDS PRODUCED

15

REFERENCE

22R-OH-Cholesterol

Polypodium vulgare

E + 20E

Reixach et al., 1999

22S-OH-Cholesterol

Polypodium vulgare

none

Reixach et al., 1999

Clerosterol

Ajuga reptans

Cyasterone + Okuzumi et al., 2003 Isocyasterone + 29-Norcyasterone

Lathosterol

Ajuga reptans Spinacia oleracea

20E 20E

Ohyama et al., 1999 Grebenok and Adler, 1993

3β-HydroxyAjuga reptans 5β-cholestan-6-one

20E

Hyodo and Fujimoto, 2000

2β,3β-Dihydroxy-5βcholestan-6-one

Ajuga reptans

20E

Hyodo and Fujimoto, 2000

3β-Hydroxy-5βAjuga reptans cholest-7-en-6-one

20E

Hyodo and Fujimoto, 2000

3β,14α-Dihydroxy-5β- Ajuga reptans cholest-7-en-6-one

no 20E

Hyodo and Fujimoto, 2000

20E

Fujimoto et al., 2000

5β-Cholest-7-ene-3,6- Polypodium vulgare dione Spinacia oleracea

22,25dE + 22dE 20E

Reixach et al., 1999 Bakrim, 2007

22,25Dideoxyecdysone

Achyranthes fauriei

20E + Inokosterone

Tomita and Sakurai, 1974

2,22-Dideoxyedysone

Polypodium vulgare

22dE

Reixach et al., 1999

2-Deoxyecdysone

Polypodium vulgare Spinacia oleracea

E + 20E 2d20E + E + 20E

Reixach et al., 1999 Bakrim et al., 2008

Ponasterone A

Polypodium vulgare

20E

Reixach et al., 1999

Ecdysone

Polypodium vulgare

20E (+ polypodine B Reixach et al., 1996 + abutasterone)

Sesuvium portulacastrum

20E

Spinacia oleracea

20E + polypodine B Grebenok and Adler, 1993; + E conjugate Grebenok et al., 1994

Sipahimalani et al., 1972

(Cook et al., 1973), then [4-14C,3α-3H] or [4-14C, 4α-3H] or [4-14C,4β-3H] for understanding the formation of the chromophore (Polypodium vulgare; Davies et al., 1980) (Fig. 1.3). More recently, molecules labelled with stable isotopes have allowed a more convenient and exact approach with the A. reptans hairy root system. After labelling with [3α-2H]-, [4α-2H]- or [4β-2H]cholesterol, it was shown that, in

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Fig. 1.3 Proposed early steps of ecdysteroid biosynthesis in Polypodium vulgare (Davies et al., 1980)

these three cases, deuterium atoms remained in the same positions, suggesting a direct mechanism involving carbons 5 and 6, and no oxidation at C-3 (Nagakari et al., 1994a). Further labelling experiments with [6-2H]- or [3α,6-2H2]cholesterol showed that the 6-H proton of cholesterol was found in position-5β in 20E (Fujimoto et al., 1997). These data were consistent with the intermediate formation of a 5α,6α–epoxide. However, the mechanism is probably not so simple, because only 70% of the deuterium is conserved, and this non-stoichiometric behaviour has received no explanation. Labelling experiments with a Δ7-sterol, [3α,6α-2H2]- or [3α,6β-2H2]lathosterol, have shown that the 6β-proton migrates to the 5β–position, whereas the 6α-H is eliminated (Ohyama et al., 1999) (Fig. 1.4). These data, taken together with the above, suggest that cholesterol and lathosterol are converted into 20E through the formation of 7-dehydrocholesterol. Here again, 30% of the deuterium is lost during the migration of the 6β-2H to C-5. However, this means that direct oxidation of C-6 of lathosterol is not involved. According to this scheme, 14α-hydroxylation would represent an independent step, which is confirmed by the conversion of 3β-hydroxy5β-cholest-7-en-6-one (5β-ketol) into 20-hydroxyecdysone (Hyodo and Fujimoto, 2000). In this respect, we are reminded that 14β-hydroxylation is a separate step in cardenolide biosynthesis in plants (Kreis et al., 1998). In the same way, the origin

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17

Fig. 1.4 Proposed early steps of ecdysteroid biosynthesis in Ajuga reptans (Fujimoto et al., 1997; Ohyama et al., 1999)

of polypodine B, and more generally of 5βOH-ecdysteroids, might result from a late 5β-hydroxylase, as evidenced by the (low) conversion of labelled ecdysone into polypodine B in Spinacia oleracea (Grebenok and Adler, 1993) and Polypodium vulgare (Reixach et al., 1996). Finally, and most unexpectedly, [3α-2H]3β-hydroxy-5β-cholestan-6-one and [5β,7α,7β-2H]2β,3β-dihydroxy-5β-cholestan-6-one are efficiently incorporated into 20E, which means that the introduction of the Δ7-bond may take place at a late stage in the biosynthetic sequence (Hyodo and Fujimoto, 2000). These data permit the conclusion that formation of the (5β−H)14α−OH-7-en6-one chromophore may proceed by at least two different routes in plants (both differing from that in animals?). Whether this means that the capacity to synthesize

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ecdysteroids has appeared several times independently remains to be established. To address this question, it will be absolutely necessary to develop molecular tools. Further steps (mainly hydroxylations) may also occur in different sequences; so, like the biosynthesis of brassinosteroids (Bishop, 2007), we should perhaps consider the biosynthetic pathway for phytoecdysteroids as a grid rather than a linear sequence, with possibly privileged ways in any given species. Finally, we wish to underline the absolute need for molecular biological approaches in this area in order to address the following basic questions: (1) In which plant cells (all or specialized ones?) does ecdysteroid biosynthesis take place? (2) Does the whole pathway take place in the same cells or does it involve some ‘cooperation’, as is often the case for animal steroids? (3) How conserved are the concerned genes in the plant kingdom and in fungi? (4) Are these genes present in all plants species, but silent (or very poorly expressed) in most of them?

1.3.5

Biogenetic Diversity of Phytoecdysteroids

The current biosynthetic studies have so far not explained the formation of the main feature of ecdysteroid structural specificity, i.e. the 14α-hydroxy-7-en-6-one moiety, but have provided some information on the sequence of hydroxylations. Some evidence exists for the occurrence of multiple or branching pathways (see Figs. 1.3 and 1.4). In some studies the main effort has concentrated on the elucidation of the formation of the 5β−H configuration, which is characteristic for ecdysteroids, or the 5β-OH substituent present in certain analogues. However, the structural variability of ecdysteroids, even if maintaining the above indicated characteristic features, is much higher. Some variability is associated with the basic ecdysteroid carbocyclic part of the skeleton, e.g. the presence of hydroxyls at positions C-1, C-11, and less frequently also at C-9 or C-15. Further variability is elicited by 3-epi-OH, or a 2,3-diepi-OH configuration, C-2 deoxy or C-2 dehydro formation, occasionally also in combination with a C-9(11) double bond. However, the 9(11)-double bond may be an artefact arising from the ready dehydration of 11α-hydroxyecdysteroids (Szendrei et al., 1988). Much more variability, however, is found in the side-chain. The C27/C28/C29 homology, which has already been mentioned, has been studied at various levels and types of sterols and steroids, including ecdysteroids, as reviewed by Goad and Akihisa (1997) and Brown (1998). This homology together with various hydroxy, oxo, oxy, isopropyloxy, acyloyl, aryloyl, glycosyl and other substituents, significantly enhances the structural variability. Moreover, the oxygenated side-chain derivatives are further appropriate for generating five- or six- membered cyclic ethers or lactones, depending on the side-chain homology (C28 or C29) and on the oxygenation level and positions of participating substituents. Different configurations of the participating alkyloxy- and oxygen-containing groups induce additional increase

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19

of variability. Formation of such a structural variability has so far only been very marginally studied by biosynthetic methods. Often, the biogenetic relations of similar structural types can only be inferred from tentative chemical relationships between the compounds. The assumed structural relationship of selected minor ecdysteroids from Leuzea carthamoides (Vokácˇ et al., 2002; Budeˇšínský et al., 2008) can serve as an illustration. This tentative biosynthetic scheme (Fig. 1.5) is based on the structural relations of the constituents so far identified from L. carthamoides, and on feasible biochemical reaction sequences. Such a sequence of reactions could be proposed only because an unusually large amount of plant material was processed (Vokácˇ et al., 1999), enabling identification of a large number and variety of minor constituents, which would be unattainable from amounts customarily used for routine laboratory extractions. This is why it has not been possible to derive such relationships for other rich ecdysteroid-containing genera, e.g. Ajuga, Silene or Serratula (see Ecdybase; Lafont et al., 2002). On the other hand, the biosynthesis of some side-chain lactones, e.g. cyasterones in the genus Ajuga, has already been experimentally explored (Okuzumi et al., 2003). Even when a large range of constituents has been identified from certain species, a tentative biogenetic relationship for the so far identified ecdysteroid analogues could not be proposed, since some of the identified ecdysteroids might only be artefacts obtained by simple oxidation, dehydration or other reactions performed during plant processing or compound isolation; especially the cyclic ethers might be formed in such a way. Indirect evidence is provided by the formation of shidasterone and congeners by catalytic thermal reactions (Harmatha et al., 2002a,b), or by specific anhydride-induced dehydration (Odinokov et al., 2002). Many more biosynthetic studies are required to explain the large, and still growing, structural diversity of natural ecdysteroids, especially those of plant origin, which are collated and thoroughly characterised in Ecdybase (Lafont et al., 2002).

Fig. 1.5 Assumed biogenetic relations of Leuzea carthamoides ecdysteroids 1. leuzeasterone, 2. carthamosterone, 3. 24-hydroxy-dihydroleuzeasterone, 4. 24-hydroxy-dihydrocarthamosterone, 5. hypothetical precursor, 6. makisterone C, 7. 24(28)-dehydro- 29-hydroxymakisterone C

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The biological activities of natural ecdysteroids, or those artificially formed from the main natural metabolites, or those structurally modified by chemical transformations, are all interesting and important for structure-activity relationship studies. Almost all types have been tested for their activity at the ecdysteroid receptor (Harmatha and Dinan, 1997; Dinan et al., 1999) and the summed data evaluated by appropriate computational methods (Ravi et al., 2001), which permits mapping of the ecdysteroid receptor ligand-binding domain. Both natural ecdysteroid metabolites and the artefactual analogues derived from them serve as source compounds or leads for the preparation of new structural types by targeted chemical transformations (Bourne et al., 2002) or by unspecific, and rather unconfined, phototransformations (Harmatha et al. 2002a,b, 2006). In this way, new and unusual structural classes have been obtained, e.g. epimeric 14-epi analogues or dimeric derivatives (Harmatha et al., 2002a,b). The synthetic or transformed derivatives of native ecdysteroids with unusual configurations, conformations or constitutions (not currently included in Ecdybase) significantly increase the structural variability of ecdysteroids, as well as their bioactivity value and potential. They represent new possibilities and unconventional outlooks for ecdysteroid research and for practical utilization.

1.3.6

Regulation of Biosynthesis

Any biosynthetic pathway requires regulatory mechanisms in order to control the accumulation of terminal metabolites. In the present case, we know that ecdysteroids are stable molecules which undergo, at best, a very slow turnover (Schmelz et al., 2000). As a consequence, ecdysteroids accumulate during plant life, but, owing to their possible migration within the plant, their distribution may vary greatly during ontogeny; thus, in developing spinach, ecdysteroids are produced in older leaves, but transported and accumulated in the young apical leaves (which, on the other hand, are unable to synthesize them) (Grebenok et al., 1991; Grebenok and Adler, 1993). Removal of the apical leaves (= the sink) will result in a cessation of ecdysteroid production in the older leaves (Bakrim et al., 2008). Further, loading excised leaves with unlabelled ecdysteroids will also induce a cessation of ecdysteroid production (Grebenok et al., 1991; Grebenok and Adler, 1993; Adler and Grebenok, 1995), which might result from the formation/accumulation of ecdysteroid (poly)phosphates. Similar evidence for feed-back mechanisms which do not involve phosphate conjugates has been obtained with Polypodium vulgare prothalli (Reixach et al., 1997, 1999). The prothalli actively synthesize ecdysteroids (mainly 20-hydroxyecdysone) up to a certain concentration. By immersing the prothalli in hot water (45 °C for 60 min; i.e. heat shock), it is possible to induce an almost complete release of ecdysteroids, which will result in the stimulation of de novo ecdysteroid production up to their initial level (Reixach et al., 1997). This treatment was used

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to enhance the conversion of 25-hydroxycholesterol into ecdysteroids (Reixach et al., 1999) (Fig. 1.6). It is possible to observe both quantitative and qualitative changes during development/ageing, as described for shoots of Taxus cuspidata (Ripa et al., 1990) (Fig. 1.7). Phytoecdysteroids are believed to represent a chemical protection of plants against phytophagous insects (and soil nematodes). It is therefore logical to expect that their production will be enhanced by mechanical wounding or insect attack. Indeed, this was demonstrated in Spinacia oleracea, but, unexpectedly, only in roots (Schmelz et al., 1998, 1999). Similarly, it was shown that jasmonate, a signalling lipid produced by plants in response to insect attack, was able to increase ecdysteroid production (Schmelz et al., 1998).

Fig. 1.6 Comparative ability of control (panel a) and heat-shocked (panel b) prothalli of Polypodium vulgare to convert 25-hydroxycholesterol (25C) into ecdysteroids (E and 20E). Separation occurred on a C18 RP-HPLC column (25 cm × 4.6 mm; 5 µm particle size) eluted at 1 ml/min with a gradient of acetonitrile/isopropanol (5:2 v/v; Solvent B) in 0.1% TFA in water (Solvent A), pre-equilbrated at 15% B and eluted as follows from the time of injection: linear gradient from 15–25% B over 2 min, isocratic at 25% B for 6 min, linear gradient from 25–75% B over 20 min, isocratic at 75% B for 22 min, linear gradient from 75–90% B over 5 min, linear gradient from 90–100% B over 5 min and isocratic at 100% B for 15 min (Reixach et al., 1999)

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Fig. 1.7 Variation in ecdysteroid content with ageing in Taxus cuspidata shoots (Redrawn from Ripa et al., 1990)

Could feed-back mechanisms be responsible for the inhibition of ecdysteroid production in non-accumulating plant species? This question was raised by the data obtained by Devarenne et al. (1995) working with maize plants (Zea mays). This plant does not accumulate detectable amounts of ecdysteroids; however, after long-term labelling with [14C]mevalonate, the authors isolated labelled ecdysone and 20-hydroxyecdysone conjugates which released the free ecdysteroids upon glycosidase treatment. If confirmed, these data would mean that possibly every plant species is able to produce ecdysteroids and explain the surprising (uneven?) distribution of ecdysteroid-accumulating species in the plant kingdom. Another possible inhibition may be explained by assumedly preferential biosynthesis of steroid saponins, which can bind simple sterols into water insoluble complexes, and thus eliminate them from the possibility of being ecdysteroid precursors. Such a mechanism was already suggested at a plant-insect interaction level (Harmatha, 2000) for Allium porrum (Arnault et al., 1986), belonging to Liliaceae family, known as a generally ecdysteroid-negative plant group.

1.4

1.4.1

Plant Cell Cultures as Model Systems for Ecdysteroid Production Introduction

The transition from intact plant to cultivated plant cells in vitro, a process involving repression of cell differentiation as well as specialization, is accompanied, as a rule, by a sharp decline in the biosynthesis of specialized secondary metabolites

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23

(Zaprometov, 1981). This is accounted for by de-differentiated plant cells partly losing the ability to express genetic information related to secondary metabolism. The loss is not irreversible, but certain conditions are required for its continued expression. Understanding these conditions is an important challenge. Ecdysteroid production by plant cell cultures is of both scientific and practical interest. On the one hand, plant cell cultures are an amenable experimental model for the elucidation of ecdysteroid biosynthetic pathways and their regulation mechanisms; this is also important for understanding ecdysteroid function in plants. On the other hand, cell cultures could be used for the production of specific ecdysteroids by bioengineering techniques. Moreover, the possibility of obtaining significant amounts of 20-hydroxyecdysone (20E), which has potential in human medicine (Lafont and Dinan, 2003), as well as other rarer ecdysteroid analogues, which may also possess interesting biological properties (Báthori and Pongrácz, 2005), is promising.

1.4.2

Callus and Suspension Cultures

Ecdysteroid formation in vitro was originally demonstrated in callus cultures of Achyranthes fauriei (Amaranthaceae; Hikino et al., 1971) and Trianthema portulacastrum (Aizoaceae; Ravishankar and Mehta, 1979), as well as in gametophytes of the fern Pteridium aquilinum cultivated in liquid medium (McMorris and Voeller, 1971). More intensive research into secondary metabolism in tissue and cell cultures of ecdysteroid-containing plants started in 1990s. Species with high phytoecdysteroid content were introduced into culture in vitro. Prothalli of the fern Polypodium vulgare cultivated under aseptic conditions produced the phytoecdysteroids characteristic of intact plant roots (20E, polypodine B [polB] and ecdysone [E]); the 20E content in prothalli (as high as 0.8% of the dry mass) was twice that found in the roots and the 20E:polB ratio varied from 7:1 to 10:1 (in wild plants, the ratio is between 1:1 and 2:1) (Camps et al., 1990). Five further phytoecdysteroids (inokosterone, pterosterone, abutasterone, 24-hydroxyecdysone and 5β-hydroxyabutasterone) previously unknown in the fern P. vulgare were released from prothalli in culture (Coll et al., 1994). The high biosynthetic quotient of prothallus cultures permitted the development of a method for obtaining 20E (J.-J. Bonet, IRTA Institute, Barcelona, Spain, personal communication). P. vulgare prothallus and callus cultures are good experimental systems for ecdysteroid biosynthetic studies. Calluses, however low their ecdysteroid content may be, efficiently transformed E added to the medium into 20E (Irrure-Santilari et al., 1996a). Tracer precursors (mevalonate, cholesterol and E) added to the calluses were incorporated into all ecdysteroids synthesized by this culture (IrrureSantilari et al., 1996b). Compounds less polar than E were also investigated as possible precursors; poA and 2dE were transformed in both prothallus and callus cultures, whereas 2,22dE and 2,22,25tE were transformed into ecdysteroids atypical for P. vulgare (Reixach et al., 1999).

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Callus and suspension cultures were obtained from Pteridium aquilinum fern spores (Vanek et al., 1990; Macˇek and Vanek, 1994). The concentration of ecdysteroids in suspension culture exceeded that in the intact plant by 20-fold in a medium optimized for nutrient components and phytohormones. The synthetic ability of callus cultures was much lower. PoA, 20E, polB, E and five unidentified compounds were found in the cell cultures. Allowing for the ecdysteroid accumulation rate by suspension culture during the cultivation cycle, a procedure was developed which increased the yield of the target products by elimination of the lag-phase from the cultivation cycle and by maintaining the cells at the optimal growth rate (Svatos and Macˇek, 1994); the cells accumulated up to 0.89 mg ecdysteroid per gram dry weight, poA being the major ecdysteroid. Cell cultures of Serratula coronata (Karnachuk et al., 1991) and S. tinctoria (Corio-Costet et al., 1996) are of great interest as a source of ecdysteroids. Wild plants of these species have high contents of various bioactive ecdysteroids. In S. coronata suspension culture, the total cell ecdysteroid content amounted to 0.074% of the dry matter, while the ecdysteroid concentration in the callus tissue was an order of magnitude lower (Saad et al., 1992a, b). The highest concentration of ecdysteroids was present during the exponential phase of cell growth. A comparative study of the composition and content of ecdysteroids and their precursors (i.e. sterols) in intact S. tinctoria plants and callus and suspension cultures showed that plant roots contain 20E, 20E 3-acetate and polB at a total concentration of 1.2–1.5% of the dry weight, whereas cell cultures contained only 20E, the content being 0.003–0.01% (Corio-Costet et al., 1993a). Based on the data obtained, a biosynthetic scheme from expected ecdysteroid precursors could be proposed. A steroidogenetic study (including the detection of sterols which are most likely to be ecdysteroid precursors) was carried out on a suspension culture of Chenopodium album (Corio-Costet et al., 1993b, 1998). The cells produced 20E in much lower quantities than the intact plants. However, the possibility of increasing the level of ecdysteroid biosynthesis by using some cultivation procedures was noted. Cell cultures of different species of Ajuga have been obtained. Synthesis of 20E and turkesterone was established in both callus and suspension cultures of A. turkestanica, the 20E concentration in the cells being several times higher than that in the roots and leaves of the plants, whereas the turkesterone concentration was somewhat lower than in the intact plant (Lev et al., 1990). A number of studies has been carried out on sterile culture and A. reptans shoots and roots cultivated separately (Tomás et al., 1992, 1993; Camps and Coll, 1993). The plants cloned in vitro retained the ecdysteroid composition inherent to both intact wild plants and glasshouse plants. Ajugalactone and cyasterone (C29-ecdysteroids), 29-norsengosterone and 29-norcyasterone (C28-ecdysteroids) and 20E (C27-ecdysteroid) were found. PolB was not found in plants in vitro, but was detected in intact plants. The ecdysteroid content in the roots of the plants in vitro amounted to 0.4% of the dry weight and was 1.5–2.5 times higher than that in intact plant roots. The ratio of C28/C29-phytoecdysteroids in intact and in in vitro cloned plants was substantially different. A. reptans roots were established to be the site of ecdysteroid biosynthesis. Based on the overall data obtained both under in vivo and in vitro conditions, the existence of two major metabolic pathways in A. reptans was pro-

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posed (Camps and Coll, 1993), involving side-chain dealkylation and differing in hydroxylation at C-5, and resulting in two groups of biogenetically related C29/C28/ C27 compounds (cyasterone/29-norcyasterone/20E and sengosterone/29-norsengosterone/polB), which differ in the absence or presence of an OH-group at C-5. Dealkylation occurs mainly in the roots of the plant. The presence of ecdysteroids was not detected in A. reptans callus cultures by Tomás et al. (1992). However, trace amounts of 20E were detected in A. reptans callus and suspension cultures by Mboma et al. (1986). Recently, cell suspension cultures of Vitex glabrata have been used to obtain evidence that 7-dehydrocholesterol and ergosterol are ecdysteroid precursors since supplementation with these two sterols enhanced 20-hydroxecdysone production by the cells, whereas addition of cholesterol did not (Sinlaparaya et al., 2007).

1.4.3

‘Hairy-Root’ Cultures

Hairy root culture models, i.e. a genetically modified “bearded” root culture resulting from the treatment of plants by Agrobacterium rhizogenes, have several advantages over normal isolated root cultures (Kuzovkina, 1992). A hairy-root culture with stable and sufficiently high ecdysteroid synthesis level (0.1–0.2% of dry weight) was obtained from a sterile sprout of S. tinctoria by means of modification with A. rhizogenes. The roots of intact plants, as well as the hairy roots, produced 20E and its 3-acetate. In both cases, a concentration gradient increasing toward meristematic zones was detected. After inclusion of labelled precursors, such as cholesterol and mevalonate, into S. tinctoria hairy roots, they synthesized labelled ecdysteroids, making this system an appropriate model for research into ecdysteroid metabolism (Delbecque et al., 1995; Corio-Costet et al., 1996). Hairy root cultures have also been obtained for a number of other ecdysteroidproducing plant species: Ajuga reptans var. artropurpurea, Achyranthes fauriei, Pfaffia iresinoides and Vitex strickeri (Matsumoto and Tanaka, 1991). All of the A. reptans hairy root clones were shown to synthesize 20E, norcyasterone, cyasterone and isocyasterone, which are characteristic ecdysteroids of intact plant roots. The component ratio was also similar to that in intact plant roots, with 20E prevailing. There was a positive correlation between the ecdysteroid content and the clone growth-rate. Selection resulted in the isolation of the rapidly growing Ar-4 clone, where the 20E concentration amounted to 0.14% of the dry weight. When growing the Ar-4 clone in an Airlift-type fermenter for 45 days, the weight of the culture increased 230-fold, and the 20E content was as much as 0.12% of dry weight. Regenerated plants cultivated from high-producing hairy root clones of A. reptans var. atropurpurea were shown to have higher ecdysteroid accumulation than untransformed regenerants (Tanaka and Matsumoto, 1993). The transformed plants had root/total plant weight ratios of 68–75%, compared to 50% for normal regenerant plants, the ecdysteroid content being as high as in the parent clones. Thus, the possibility of obtaining microclonal reproduction of modified plants with

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an increased ability to produce ecdysteroids was demonstrated, with the associated possibility of creating phytophagous insect-resistant plants. Genetically modified rhizogenic culture of A. reptans var. atropurpurea also proved to be a boon for ecdysteroid biosynthesis research (see Section 1.3.4). Transformed Rhaponticum carthamoides (Willd.) Iljin root cultures were obtained by modifying sterile shoots with A. rhizogenes, the cultures differing qualitatively and quantitatively in ecdysteroid content from that in the intact plant roots (Orlova et al., 1998). These authors also developed an effective system for regeneration from leaf explants and carried out genetic modification of this plant species with a A. rhizogenes GV 3101 recombinant strain containing rolC-gene plasmid under a 35S CaMV promoter and with the inherent dwarf phenotype (Orlova et al., 2000). In the authors’ opinion, the known multiple effects of rolCgene, e.g. changing the hormonal status of modified plants (Estruch et al., 1991), are the probable cause of the changes in the qualitative and quantitative content of secondary metabolites, i.e. ecdysteroids.

1.4.4

General Conclusions and Prospects for Future Research

The potential and convenience for ecdysteroid biosynthesis research has been demonstrated with various experimental systems in vitro, including nondifferentiated cells, morphogenic structures, isolated organs and sterile plant culture. The data obtained show the possibility of regulating phytoecdysteroid synthesis both at cell and organ levels. In vitro cultures with high biosynthetic activity are of interest from the point of view of developing bioengineering techniques for obtaining valuable bioactive ecdysteroids, few of which are commercially available. A comparative study of ecdysteroid formation in cell cultures of three species of plants has been conducted with Rh. carthamoides, S. coronata (both belonging to the Asteraceae) and Ajuga reptans (Lamiaceae). They differ in both the composition and levels of ecdysteroids in the intact plants (Volodin, 2003). Rh. carthamoides callus cultures obtained from different organs of the plant were found to be incapable of ecdysteroid synthesis or to synthesize them only in trace quantities. The level of ecdysteroid synthesis in Rh. carthamoides cell cultures increased relative to that of the intact plant only by utilizing cell cultivation in suspension culture. Of major importance was a correlation between the highest ecdysteroid synthesis level and the expotential growth phase of suspension cultures. In contrast to Rh. carthamoides, young S. coronata callus cultures retain the ability to synthesize ecdysteroids. However, there is a significant decrease in their concentration in the callus in comparison with the intact plant (five-fold, on average). A. reptans plants are characterized by a low ecdysteroid content. However, in contrast to Rh. carthamoides callus cultures which are incapable of synthesizing significant amounts of ecdysteroids, strains of A. reptans cultures were obtained which surpassed both plants cultivated in vitro and wild plants in their ecdysteroid content. The content of 20E was found to increase significantly during long-term

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(for 9 years) cultivation of S. coronata and A. reptans callus cultures. This is likely to be a result of cell differentiation over time. Thus, ecdysteroid-producing cell cultures were shown to retain their biosynthetic abilities to different extents. In young callus cultures with high or moderate ecdysteroid content (S. coronata and Rh. carthamoides), a decrease in ecdysteroid synthesis occurs. After being introduced into culture, species with low ecdysteroid content, such as A. reptans match, or even surpass, intact plants as far as ecdysteroid accumulation is concerned. Moreover, ecdysteroid accumulation dynamics in cell cultures of phylogenetically distant species is similar in spite of the difference in morphological features. The data obtained are of considerable importance for a better understanding of ecdysteroid functions in plants. If ecdysteroids act as toxins/deterrents towards phytophagous insects in plants with high or moderate content of the former, the repression of ecdysteroid synthesis occurring in de-differentiated-cell cultures could be accounted for by the fact that, in the absence of organismal control, there is no longer a need for these compounds as eco-regulators (Volodin, 2003). In future studies, attention should be given to the regulation of ecdysteroid synthesis in plant cell cultures. An opportunity was opened up by the isolation of ecdysone-20-monooxygenase from the leaves of spinach, Spinacia oleracea L., and demonstration of the induction of ecdysteroid accumulation in plants attacked by insects. The effect demonstrated in spinach subjected to attack by the fungus gnat Bradysia impatiens (Johannsen) (Schmelz et al., 2002) is triggered by methyl jasmonate, which is a plant pathogen signal transducer. Encouraging results have been achieved from the use of cytochrome-P450 inducers (manganese salts, exogenic sterols, phytohormones, methyl jasmonate) for promoting ecdysteroid synthesis in cell cultures of ecdysteroid-containing plants (Alexeeva, 2004, 2005, 2006).

1.5

Do Phytoecdysteroids Have a Physiological Role in Plants?

Based on the hormonal role of ecdysteroids in arthropods, it was suggested that phytoecdysteroids might regulate physiological processes in plants. The early evidence has been summarised previously (Lafont, 1998; Dinan, 2001) and it is not convincing for such a role. Basically, ecdysteroids have been employed in a series of standard plant bioassays used for determination of the activities of phytohormones (auxins, brassinosteroids, cytokinins, ethylene formation, giberellins: Hendrix and Jones, 1972; Dreier and Towers, 1988; Machácˇková et al., 1995; Golovatskaya, 2004). Activities were either absent or slight. A priori, it seems unlikely that ecdysteroids could possess a hormonal role in plants, since their occurrence is not universal and yet, when they do occur, the levels can be very high, far surpassing those expected of hormonal molecules. The varying distributions within ecdysteroid-containing plants and seasonal variations also accord better with an allelochemical function (see below), and it is, of course, now known that the hormonal steroids in plants are the brassinosteroids. However, ecdysteroids do, by an

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as yet unexplained mechanism, appear to significantly affect growth, cell size and biochemical properties both in the cyanobacterium Nostoc 6720 (Maršálek et al., 1992) and in Chlorella vulgaris (Bajguz and Dinan, 2004). Also, it is possible that phytoecdysteroids could be released by plants to have allelochemical effects on other plants or microbes in their environment, since 20E has been shown to affect seed germination and seedling growth (DellaGreca et al., 2005; Bakrim et al., 2007), and ecdysteroids have been shown to possess antimicrobial activity (Ahmad et al., 1996).

1.6

Allelochemical Functions of Phytoecdysteroids

1.6.1

Probable Function of Phytoecdysteroids

The most generally accepted hypothesis for the function of phytoecdysteroids in plants is that they act, either alone or in conjunction with other classes of secondary metabolites or even together with physical defence mechanisms, to protect the plant against non-adapted invertebrate predators, bringing about reduced consumption of the plant or even endocrine disruption and death of the phytophagous invertebrate. In general, it has been the effects on phytophagous insects which have been considered, but plant nematodes and even crustaceans have also been examined to a lesser extent.

1.6.2

Ecdysteroid Effects on Insects

1.6.2.1

Effects of 20-Hydroxyecdysone on Insects

Ingested ecdysteroids (predominantly 20E) have been shown to have a range of detrimental effects on the development and survival of a number of insect species (Bombyx mori [Kubo et al., 1983], Pectinophora gossypiella [Kubo et al., 1981, 1983], Spodoptera frugiperda [Kubo et al., 1981], Acrolepiopsis assectella [Arnault and Sláma, 1986; Harmatha, 2000], Agrius convolvulus [Tanaka and Naya, 1995]), including inhibition of growth, supernumerary larval instars, death without moulting, death associated with promoted moulting and prothetely. However, certain insect species (e.g. Heliothis virescens, H. armigera, Locusta migratoria, Manduca sexta, Spodoptera littoralis, Lacanobia olereaceae, Acherontia atropos; summarised in Dinan, 1998) are remarkably tolerant to ecdysteroids in their diet, showing no apparent ill-effects when fed 400 ppm or more 20E in their diet (although S. littoralis larvae are affected by 1,000 ppm 20E; Ufimtsev et al., 2006a). However, several of these species (H. virescens, H. armigera, S. littoralis, L. oleracea) and Ostrinia nubilalis (Rharrabe et al., 2007) detoxify the ecdysteroid intake by conjugating them to fatty acids, thereby blocking the C-22 hydroxyl group, which

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is important for the biological activity of ecdysteroids (Dinan and Hormann, 2005). Such a detoxification mechanism has a considerable energy cost, such that although the insect may develop normally when food it plentiful, the energy demand will be detrimental to insect development when food is more scarce. In addition to conjugation with fatty acids, insects use several other detoxification mechanisms for ingested ecdysteroids (22-glucosides, 2/22-phosphates, 3-acetates, 3-oxo/3-epi derivatives and side-chain cleavage; Rharrabe et al., 2007; see Fig. 1.8), or even excreting ingested 20E unmetabolised (e.g. Acherontia atropos [Blackford and Dinan, 1997c]). Other insect species are partially tolerant to ecdysteroids in their diet, being able to cope with low levels (e.g. Cynthia cardui, Tyria jacobaeae; Blackford and Dinan, 1997b), while higher levels are toxic. Yet other species are extremely sensitive to ecdysteroids in the diet, such that they eat and succumb at very low concentrations (e.g. Aglais urticae), or are so deterred from eating an ecdysteroid-containing diet that they die of hunger rather than consume ecdysteroid-containing food (e.g. Inachis io; Blackford and Dinan, 1997b). There seems, as might be expected, to be a relationship between the tolerance of the phytophagous insect species and the probability of it encountering ecdysteroids in its diet, such that highly polyphagous species are ecdysteroid-tolerant, while oligophagous and polyphagous species are semi-tolerant and monophagous species are ecdysteroid-sensitive. Recently, Malausa et al. (2006) have examined the genetic variability of the response of six clones of the peach-potato aphid (Myzus persicae) to moderate (10−6M) and high (10−3M) concentrations of 20E by measuring fecundity (number of offspring produced) and found that they differ very considerably. The fecundity was decreased, unaffected or increased, depending on clone, at the moderate 20E concentration relative to controls (no 20E), whereas it was the same or reduced at the high 20E concentration.

phosphorylation acylation glycosylation OH OH 22 20

25 OH glycosylation

phosphorylation HO acetylation oxidation HO epimerization phosphorylation

14

2 3

side-chain cleavage

OH dehydroxylation 6 H

O

Fig. 1.8 Summary of the detoxification pathways for ingested phytoecdysteroids in insects

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Effects of Other Purified Ecdysteroids on Insects

Most studies have used ecdysteroid preparations where 20E is the sole or major component. 20E is without doubt the most commonly occurring phytoecdysteroid, but an important question concerns the raison d’être of the many other phytoecdysteroid analogues found in plants; Do they possess differential activities? Are they active against different insect species? Are they detected differentially? Are they subject to different metabolic rates or fates? Are the minor components there to provide evolutionary flexibility for when potential predators become tolerant of the major ecdysteroids? There are few studies which compare the activities of a significant number of ecdysteroid analogues, and where such studies have been performed in vivo (Sláma et al., 1993), the ecdysteroids have been injected into the test species, which does not mimic the natural route of exposure to phytoecdysteroids.

1.6.2.3

Effects of Serratula coronata Extracts and Its Ecdysteroids on Insects

Long-term investigations on the impact of Serratula coronata L. (Asteraceae) leaves, extracts and total ecdysteroids, or of isolated major and minor ecdysteroids of the plant, incorporated into artificial nutrient media on the viability and behaviour of different age caterpillars of three species of polyphagous insects, namely Ostrinia nubilalis Hb., Mamestra brassicae L. and Spodoptera littoralis Boisd., have been performed. Significant antifeedant effects of ecdysteroids in different artificial nutrient media were observed for first instar larvae of O. nubilalis and M. brassicae. This was associated with mass migration of the caterpillars from the feedstuff and their death. A difference was found in the effect of ecdysteroid-containing nutrient media effect on third and fourth instar larvae of M. brassicae, when compared to that on the first instar larvae. Diets of different ecdysteroid contents first stimulated nutrition of the older larvae, and then rejection of the feedstuff occurred, resulting in an outburst of cannibalism and mass caterpillar lethality. Caterpillars kept on the ecdysteroidcontaining diet revealed developmental defects, significantly impaired pupation and formation of nonviable pupae with various abnormalities. With O. nubilalis caterpillars of all age groups, the ecdysteroid-containing diet was more toxic than for M. brassicae, causing total lethality (Volodin, 2003; Ufimtsev et al., 2001). A detrimental effect (reduced caterpillar weight, delayed pupation and reduced adult fecundity) of a diet containing 1,000 ppm 20E on sixth instar larvae of S. littoralis, previously considered to be resistant to high 20E concentrations (400 ppm; Blackford et al., 1996), was observed. Ecdysone at 1,000 ppm had no effect. When fourth instar larvae were fed on nutrient media containing 10% powdered parts of S. coronata (containing 100–2,200 ppm ecdysteroids), the detrimental effects were shown to occur in the absence of a pronounced antifeedant effect. However, distinct correlations between the expressivity of the effects and the ecdysteroid content in different parts of the plants (roots, stems, leaves and buds) were not found

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(Ufimtsev et al., 2003, 2006a, b), which imply that other components of the plant are responsible for, or contribute to, the effects. In comparative experiments on wild-type and the mutant line 147 Drosophila virilis (with enhanced levels of E and 20E and diminished JH levels) an effect of exogenous ecdysteroids on the fecundity and a delay in egg-laying by wild-type imagos was shown to occur. Female flies with artificially enhanced 20E level were found to undergo a decrease in JH degradation (and consequently, an increase in its titre), mediated through the dopamine metabolism system (Rauschenbach et al., 2005). The development of last instar larvae of Ephestia kühniella, after having been submerged in ecdysteroid-containing solutions, revealed diverse effects, depending both on the structure and the concentration of ecdysteroids, as well as on the nature of the solvent and the duration of submersion. Methanolic 20E solutions were less toxic than pure methanol (which caused total caterpillar lethality) or aqueous 20E solutions. This was attributed to the adaptogenic properties of 20E when in the presence of the damaging agent (methanol in this case). 20E 22-acetate and 25S-inokosterone do not possess this property. After submersion of caterpillars in methanolic 20E 22-acetate or methanolic or aqueous 25S-inokosterone solutions, pupation occurred much more rapidly than after submersion into 20E solutions. However, the number of abnormal pupae was far higher (Volodin, 2003; Ufimtsev et al., 2002a, b).

1.6.2.4

Ecdysteroid Taste Receptors in Insects

Where phytoecdysteroids act as deterrents, the insect must possess taste receptors to be able to recognise the presence of these compounds in the food. The location and properties of such taste receptors are now starting to be investigated (Ma, 1969; Tanaka et al., 1994; Descoins and Marion-Poll, 1999; Marion-Poll and Descoins, 2002). Initially, it was shown for Pieris brassicae (Ma, 1969) and Bombyx mori (Tanaka et al., 1994) that a reduction in food intake is mediated by specialised sensory perception of ecdysteroids. With the earlier studies on these two species with specialised diets (containing no or low levels of ecdysteroids), Descoins and Marion-Poll examined taste detection of three ecdysteroids (E, 20E and poA) in three polyphagous (Mamestra brassicae, Spodoptera littoralis and Ostrinia nubilalis) and one monophagous (B. mori) species (Descoins and Marion-Poll, 1999; Marion-Poll and Descoins, 2002). Electrophysiological studies demonstrated that all four species possess contact chemoreceptor cells on the maxilla, which respond to ecdysteroids (but not necessarily to all three), indicating that perception of ecdysteroids by phytophagous lepidopterans is a common feature, even if the toxicity or antifeedant activity of phytoecdysteroids differs between the species, such that, when given a choice, larvae of these species will avoid ecdysteroid-containing food and show preference for an ecdysteroid-free diet. Since the survival of first instar lepidopteran larvae is highly dependent on the quality of food available to them on hatching, the ability of the adult female to detect good host plants for oviposition is crucial. For both Lobesia botrana (Calas

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et al., 2006) and Ostrinia nubilalis (Calas et al., 2007), it has been shown that 20E deters oviposition by adult females and that this is mediated by tarsal sensilla, which demonstrate a sensitivity similar to that of the first instar larvae to the same compound. In the European Grapevine moth (Lobesia botrana), 20E deters not only larval feeding, but also oviposition by adult females (Calas et al., 2006), the taste sensilla being located on the last tarsus of the prothoracic leg of the adult female. This is clearly an important strategy since neonate larvae of this species have limited dispersal capacities, such that the eggs should be deposited by the adult female on to an adequate food source for larval development.

1.6.2.5

Do High Levels of Phytoecdysteroids Deter Phytophagous Insects?

Although all parts of L. carthamoides contain very high levels of ecdysteroids (300–1,000 ppm 20E equivalents in the leaves), there is an extensive diversity of insect species which associate with this plant. Zeleny et al. (1997) found 126 species of arthropods on plants of this species in the Czech Republic (to where it had been introduced from Asia) over 2 years, of which 74 fed on the leaves and 34 could complete their development on the plant without any apparent problems. The most abundant arthropods on the plant were all oligo- or phytophagous and belonged to groups of arthropods (collembolans, psocopterans etc.) which have so far been poorly studied with regard to their response to phytoecdysteroids; extensively studied groups which are known to be, at least partially, susceptible to ecdysteroids e.g. lepidopterans were essentially absent from the plants. This study underlines how inadequate our current knowledge is in terms of plants being protected by even exceedingly high levels of phytoecdysteroids and our ability to extrapolate knowledge concerning the susceptibility of certain groups of holometabolous insects to arthropods in general.

1.6.2.6

A Commercial Application of Phytoecdysteroids

In the specific case of silkworms, particularly Bombyx mori, the response of last instar larvae to exogenous ecdysteroids has been exploited to hasten maturation and synchronise cocoon spinning. It is important to stress that the doses used to bring about the beneficial effects on silkworms are low relative to those having detrimental effects on insects (see Section 1.6.2.1). The ecdysteroids are extracted from ecdysteroid-containing plants (see Chandrakala et al., 1998) and sprayed on to mulberry leaves, which are then given to the larvae as the first ones in the batch are about to spin their cocoons (Ninagi and Maruyama, 1996; Maribashetty et al., 1997, 2002; Trivedy et al., 2003a, b, c). It is not certain whether mulberry (Morus nigra or M. alba) contains endogenous ecdysteroids (Takemoto et al., 1967b; Blackford and Dinan, 1997c), but if it does, the levels are low. Recently, the consequences of controlled ecdysteroid application (2 µg/larva) at other times during the fifth

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instar have been determined (Nair et al., 2005), showing that the quantity of silk can be significantly increased (by about 10%) by treating the insects 48 h into the fifth instar, while the time taken for cocoon formation in the batch can be reduced from 60 to 36 h by treating when the first insects in the batch begin to spin. Even more recently (Trivedy et al., 2006), an extract of ecdysteroid containing Silene gallica has been used for the induction of spinning and uniform maturation in B. mori. Spraying of last instar larvae with ecdysteroid to induce uniform maturation does not affect silk yield or quality, but spraying twice during the instar to control both spinning and maturation does reduce cocoon traits, although it does shorten development time, thereby saving on mulberry leaves and reducing the possibility of crop loss owing to disease in the last phase of rearing. Ecdysteroids have also been proposed as a treatment for the enhancement of fecundity of honey bees in apiculture (cited in Kholodova, 2001).

1.6.3

Ecdysteroid Effects on Crustaceans

As with insects, most studies of the effects of exogenous ecdysteroids on crustaceans have been performed by injection of the compounds. Where the animals have been externally exposed to the compounds, it has generally been with smaller crustaceans, such as shrimps, where injection would be more difficult. With the snapping shrimp Alpheus heterochelis, for example, exposure to 20E at 5 µg/ml in the seawater was able to reduce the length of the moult cycle by up to 65% (Mellon and Greer, 1987). The possibility of improving prawn/shrimp aquaculture by exposure of the animals to ecdysteroids has been examined. In an early study (Kanazawa et al., 1972), Penaeus japonicus was exposed to the phytoecdysteroids 20E (0.5–2.5 mg%), inokosterone (0.1–12.5 mg%) or cyasterone (0.5–2.5 mg%), incorporated into an artificial diet. The ecdysteroids were found to enhance moulting rate, but were also found to reduce survival and growth rates in a concentration-dependent manner. In a more recent study (Cho and Itami, 2004), an ecdysteroid-containing extract of Achyranthes spp. (composition and ecdysteroid content not revealed) was incorporated into diet and fed to Marsupenaeus japonicus. The treated shrimps showed improved weight gain (29%) over the control animals. The effects of exposure of crustaceans to ecdysteroid agonists (steroidal or non-steroidal) and antagonists is of increasing interest and concern because of the potential of such compounds as endocrine disruptors in the environment and the suitability of various crustacean species as signal species to monitor the quality of the environment (Hutchinson, 2002). Certain decapod crustaceans (e.g. Carcinus maenas, Cancer pagurus, Homarus americanus, Astacus astacus) have been shown to reject ecdysteroid-containing food, indicating the presence of taste receptors for ecdysteroids associated with the mouthparts (Tomaschko, 1995). The origin of this discovery goes back to

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the finding that the pantopod Pycnogonum litorale, in addition to using 20E to regulate its moulting, produces and accumulates in the gut and cuticle very large amounts of seven other ecdysteroids, which are released to the exterior when the animal is attacked (Bückmann et al., 1986; Bückmann and Tomaschko, 1992; Tomaschko and Bückmann, 1993; Tomaschko, 1994, 1995). This raised the question as to whether these ecdysteroids were acting as defensive chemicals and the demonstration that the crab Carcinus maenas was indeed deterred from feeding on the pantopod by the ecdysteroids excreted from the pantopod. Further studies developed a bioassay based on the crab’s rejection mechanism (Tomaschko et al., 1995) and this was used to examine the ecdysteroid specificity of the taste receptors (Tomaschko, 1995). The available evidence indicates that the ligand specificity is very different to that of nuclear ecdysteroid receptors (Dinan and Hormann, 2005), both in terms of structural specificity for ecdysteroids and recognition of diacylhydrazines. Since certain aquatic plants contain significant amounts of phytoecdysteroids (e.g. Potamogeton spp.; Chadin et al., 2003), these may also be protected against crustacean predators.

1.6.4

Ecdysteroid Effects on Plant Nematodes

Whether ecdysteroids possess a hormonal role in nematodes is uncertain (Chitwood, 1999), but studies on several species of nematodes have shown effects of exogenous ecdysteroids upon them, including plant nematodes which could be exposed to phytoecdysteroids. Soriano et al. (2004) demonstrated that cereal cyst nematodes (Heterodera avenae) exposed to exogenous 20E (>ca. 10−6M) were far less able to invade roots of Triticum aestivum, and that exposure of H. avenae, H. schachtii (sugarbeet cyst nematode), Meloidogyne javanica (root-knot nematode) and Pratylenchus neglectus (root lesion nematode) to 20E at 5.2 × 10−5M brought about abnormal moulting and/or mortalilty. The authors also demonstrated that Spinacia oleracea (spinach) plants in which ecdysteroid levels had been enhanced (Schmelz et al., 1999) by treatment with methyl jasmonate suffered reduced damage after inoculation with H. schachtii, M. javanica and P. neglectus. The spraying of tomato plants with a solution of ecdysone reduced infestation by the root-knot nematode Meloidogyne incognita (Udalova et al., 2004).

1.7

Possible Relationships Between Ecdysteroids with Other Phytosteroids

The available evidence, although far from complete or conclusive, indicates that ecdysteroids have arisen independently in arthropods and plants, and possibly several times in plants and fungi. The generally accepted explanation for this is the need for sessile plants to protect themselves against predation after the evolution of

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the insects during the Devonian Period and the subsequent ensuing chemical race between plants and arthropods. This can, on the one hand, account for the large amounts of ecdysteroids found in certain plant species, the wide occurrence of phytoecdysteroids in the plant world and the large diversity of analogues present, and, on the other hand, for the diversity of biochemical and behavioural strategies found in insects to avoid or overcome phytoecdysteroids when present in their host plants (see Fig. 1.8 for a summary of the metabolic strategies). Neither side in the battle will be fully successful, since this would be too costly either in limiting necessary interactions or in energetic terms. Plants’ interactions with insects are ambivalent, since many are dependent on insects for pollination. It suffices for plants to reduce predation and phytophagous insects to consume enough plant material to a level where the species are able to maintain themselves from generation to generation. The complex cocktail of phytoecdysteroids typically found in ecdysteroid-containing species can be viewed as a resource for the generation of new potent analogues to be selected for when, and if, the current major ecdysteroids become ineffective, owing to a change of predator or its ability to overcome the major ecdysteroids. Further, ecdysteroids may not act alone in the defence of the plant, but in conjunction with other classes of secondary compound or even physical defence mechanisms to give synergistic interactions, such that the level of phytoecdysteroids required to provide effective protection may be low. This appears to be the case for Kochia scoparia (burning bush; Dinan, 1994) and Pteridium aquilinum (bracken; Jones and Firn, 1978), which contain only low levels of ecdysteroids together with many other defensive components to provide good general resistance to insect attack. In fact, it appears that plants can use phytoecdysteroids in the full spectrum of defence possiblities ranging from full emphasis on one class of defensive chemicals (‘all eggs in one basket’) to a complex mixture of chemically highly diverse components (‘hedging one’s bets’), depending on species and presumably a consequence of the range, aggressivity and susceptibilities of predators they have been/are exposed to. There are chemical and structural similarities between ecdysteroids and other classes of phytosteroids (brassinosteroids, withanolides, etc.) beyond the fact that they are all steroidal. These similarities include the types and locations of functional groups. The most marked similarities are between the ecdysteroids and the brassinosteroids, where (i) a full sterol side-chain is retained, (ii) diols are found on the A-ring and the side-chain and (iii) an oxygen-containing functional group is generally associated with ring-B. However, these similarities are more superficial than real, deriving from the 2D-representations of the molecules, rather than through thorough consideration of their 3D-structures. It is the 3D-structure which reveals particular and individual biochemical features specific to the steroid class, and these then impart specific biological properties. Thus, the stereospecific orientation of the hydroxyl groups is different, as is the nature of the A/B-ring junction (cis in ecdysteroids and trans in brassinosteroids). Admittedly, a combination of 2α,3α-diol and trans-ring junction in brassinosteroids or a 2β,3β-diol and cis-ring junction in ecdysteroids put O-2 and O-3 in similar (but not identical) spatial locations if the C- and D-rings are superimposed, but given that even small changes can significantly alter

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chemical reactivity and biological potency, even within a class of steroids (e.g. just changing the stereochemical orientation of the 3β-hydroxyl group of 20E to give 3-epi-20-hydroxyecdysone is associated with a 20-fold reduction in biological activity; Dinan, 2003), the summed structural differences between classes of phytosteroid would be expected to reduce activity in heterologous assays to low or non-existent levels. Thus, ecdysteroids do not appear to show activity in brassinosteroid bioassays and, where activity (agonist or antagonist) of brassinosteroids in ecdysteroid assays has been described, it only occurs at very high concentrations (>10−5M) i.e. it is not truly specific and could even be a consequence of impurity of the test compound or metabolism in the assay system. Also, in this specific case, any activity has no biological relevance, since the levels of brassinosteroids found in plants are so low that no phytophagous arthropods would be naturally exposed to adequate levels to bring about any possible effects. Several groups have synthesised steroids which are chemically hybrid between ecdysteroids and brassinosteroids and determined their activities in ecdysteroid- and brassinosteroid-specific bioassays to determine which features are responsible for specific activities and to determine which are essential to obtain a cross-over in activity (Voigt et al., 2001; Watanabe et al., 2004). In contrast to the brassinosteroids, most classes of phytosteroid do not possess phytohormonal roles and, although not ubiquitous, where they do occur, they occur at much higher concentrations than the brassinosteroids. Predominantly defensive roles have been ascribed to these other classes and their associated biological activities are manifold, acting against a wide range of organisms and at a wide range of biochemical sites. It is therefore perhaps not surprising then that some of these can interfere with the hormonal actions of ecdysteroids, even if they do not generally mimic the ecdysteroids as agonists (Dinan et al., 2001a). The ecdysteroid receptor antagonist activities of cucurbitacins (triterpenoids, but not strictly steroids) and withanolides are cases in point (Dinan et al., 1996, 1997), where the features of size and general shape and polarity with certain chemical similarities are adequate to permit interaction with the LBD to prevent ecdysteroid binding, but not to bring about the conformational changes associated with agonism i.e. these compounds act as weak antagonists. It should be borne in mind that agonism is not compulsorily associated with an ecdysteroidal structure, since DAHs and other classes of non-steroidal agonists and antagonists exist, which bear no obvious chemical or biochemical similarity to ecdysteroids (Dinan and Hormann, 2005). One is struck by the wide array of natural analogues in each phytosteroid class and how the profile can vary between plant species. The continuing ability to find new analogues suggests that many analogues still remain to be discovered. For example, over 300 phytoecdysteroid analogues are currently known and 10–20 new ones are described each year in the literature. Given this vast array of analogues in each phytosteroid class, it is perhaps not surprising that the same chemical functional groups occur across the classes of phytosteroids in at least some of the analogues. One is left wondering if this represents a biosynthetic relatedness between the different classes of phytosteroids, such that at least some of the biosynthetic enzymes might

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be common to the pathways for the different classes. Circumstantial evidence exists that most, if not all, plant species have the genetic capacity to produce phytoecdysteroids (see above). Perhaps this is also the case for the other classes of defensive steroids/triterpenoids and certainly seems highly probable for the phytohormonal brassinosteroids. Whether a particular plant species produces a particular class of defensive steroid would then depend on whether the particular pathway is activated or repressed. It is conceivable that these pathways are not mutually exclusive, since some may use enzymes with lesser specificity to modify intermediates in similar locations. In the extreme case, one could envisage that each class might have a distinct key early intermediate, which is acted on by a number of common enzymes (hydroxylases etc.) to generate the cocktail of molecules of that class. We currently know very little about the genetic organisation of the defensive steroid pathways in plants, so it is not possible to answer this question at the moment. However, progress in the elucidation of the brassinosteroid pathway(s) (Bishop, 2007) is providing some of the biological tools to begin to investigate possible commonalities in the pathways.

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Chapter 2

Diversity of Ecdysteroids in Animal Species René Lafont and Jan Koolman

Abstract Arthropods contain a significant diversity of ecdysteroids which are used to control their development and reproduction. This diversity is for one part connected with the diversity of the sterol precursors used. Outside Arthropods, ecdysteroids or/and closely related molecules have been found in most other Invertebrates and also in Tunicates. Definite proof for their endogenous origin is however still lacking, and their physiological functions (if any) remain to be established. It is hoped that thanks to the availability of whole genome sequencing of an increasing species number, tools will become available for answering these questions. Keywords Ecdysone • 20-hydroxyecdysone • makisterone • insect • arthropod • crustacean • arachnid • mollusc • echinoderm • tunicate • annelid • nematode • nemertine • platyhelminth • cnidarian • porifera Abbreviations BIO: Bioassay; GC: Gas chromatography; HPLC: High-performance liquid chromatography; ISOL: Isolation and complete chemical identification; MS: Mass spectrometry; RIA: Radioimmunoassay; Ecdysteroids AjC: Ajugasterone C; C: Cholesterol; 3DE: 3-dehydroecdysone; 2dE: 2-deoxyecdysone; 2d20E: 2-deoxy20-hydroxyecdysone; 22d20E: 22-deoxy-20-hydroxyecdysone (taxisterone); 25dE: 25-deoxyecdysone; 20dMakA: 20-deoxy-makisterone A; 2,22dE: 2,22dideoxyecdysone; 20,23E: 20,23-dihydroxyecdysone (gerardiasterone); 20,26E: 20,26-dihydroxyecdysone; E: Ecdysone; 20E: 20-hydroxyecdysone; K: Ketodiol (2,22,25-trideoxyecdysone); MakA: Makisterone A; PonA: Ponasterone A.

R. Lafont () Université Pierre et Marie Curie, Laboratoire de Biochimie Structurale et Fonctionnelle des Protéines, CNRS FRE 2852, F-75252 Paris cedex 05, France e-mail: [email protected] J. Koolman Physiologisch-Chemisches Institut, University of Marburg, D-35033 Marburg, Germany

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Introduction

The first ecdysteroid, ecdysone (termed for some time α-ecdysone), was isolated in 1954 by Butenandt and Karlson (500 kg silkworm pupae yielded 25 mg ecdysone) using a bioassay-directed purification based on the induction of pupation in ligated Calliphora erythrocephala (C. vicina) larvae. Previous experiments on this blowfly had demonstrated that pupation was induced by a blood-borne factor, i.e. a hormone (Fraenkel, 1935). A second minor hormone, β-ecdysone (now termed 20-hydroxyecdysone), was also isolated, which later proved to be the major moulting hormone (ecdysone accumulation is a peculiarity of young lepidopteran pupae). At that time, structural elucidation of biomolecules was a difficult task and required large amounts of pure molecules. For this purpose, Karlson et al. (1963) isolated about 250 mg ecdysone, the structure of which was finally elucidated by crystallographic analysis (Huber and Hoppe, 1965); for more historical details, see Horn (1989). This was the starting point of extensive studies which rapidly showed the general occurrence of ecdysteroids as moulting hormones in all arthropods (Hampshire and Horn, 1966; Horn et al., 1966; Galbraith et al., 1967; Krishnakumaran and Schneiderman, 1968; Spindler, 1989; Jegla, 1990), and to the isolation of a significant number of “zooecdysteroids” (Rees, 1989; Koolman, 1990). The development of more convenient analytical procedures, i.e. gas chromatography and radioimmunoassays, and the improvement of mass and nuclear magnetic resonance spectrometers then allowed many laboratories to search for the presence of ecdysteroids in various invertebrate groups, in particular those which undergo periodical moulting (nematodes, nemertines, leeches), and ultimately almost all investigated groups have been shown to contain ecdysteroids by more or less sophisticated methods (Table 2.1; see Rees and Mendis, 1984; Franke and Käuser, 1989; Barker et al., 1990; Lafont, 1991; Lafont et al., 1995 for a more extensive survey of available literature). As will be discussed later, an endogenous origin of ecdysteroids in animals has not been established outside arthropods, which raises questions about the physiological relevance of the presence of these molecules in non-arthropod invertebrates. Indeed, owing to the presence of ecdysteroids in many plant species (see Chapter 1; e.g. Lafont, 1997; Dinan, 2001), a possible dietary origin of animal ecdysteroids must always be borne in mind.

2.2

Diversity of Arthropod Ecdysteroids: Carbon Number

Ecdysone is synthesized from cholesterol, and more generally ecdysteroids are synthesized from the sterols that arthropods, which are auxotrophs for these molecules (unless they host symbionts), take up from their food (Clark and Bloch, 1959). The major sterols differ between animals, plants and fungi (Nes and McKean, 1977). Therefore, depending on whether arthropods feed on plants or on other animals, they will ingest different types of sterols, i.e. mainly cholesterol, a C27 sterol

Table 2.1 Presence of ecdysteroids in non-arthropod invertebrates (Updated from Lafont, 1991) Phylum/class Occurrencea Reference Protozoa Porifera

— ISOL

Cnidarians Hydrozoa Anthozoa

— ISOL

Scyphozoa Ctenophora Platyhelminths Turbellaria Trematodes Cestodes Nemertines Nematodes

Annelids Polychaetes Oligochaetes Achaetes

Molluscs Gastropods

— 20E, AjC, 20E Diop et al., 1996; Cafieri et al., 1998; 22Ac, PoA, … Costantino et al., 2000

2d20E, 20E, AjC, 20,23E, …

— — — HPLC-RIA GC-MS GC-MS

E, 20E E, 20E E, 20E, 20,26E

HPLC-RIA GC-MS GC-MS

E, 20E 20E E, 20E, 20,26E

ISOL RIA HPLC-MS

20E none detected MakA, 20,26E

HPLC-RIA HPLC-RIA GC-MS HPLC-RIA ISOL

E, 20E, 2dE-like E, 20E, 2dE-like 20E, 2dE-like E Oxysterols

Bivalves

GC-MS HPLC-RIA ISOL BIO

Cephalopods Echinoderms

— RIA

Tunicates

ISOL

E, 20E 2dE-like Diaulusterolsb no structural identification no structural identification Diaulusterol Bb Hyousteronesb

— Sturaro et al., 1982; Guerriero and Pietra, 1985; Guerriero et al., 1986; Searle and Molinski, 1995; Shigemori et al., 1999; Parameswaran et al., 2001; Suksamrarn et al., 2002 — — — — Nirdé et al., 1983 Foster et al., 1992 Mendis et al., 1984; Mercer et al., 1987a Okazaki et al., 1988 Okazaki et al., 1998 Hitcho and Thorson, 1971a,b; Dennis, 1977a,b; Mendis et al., 1983 Horn et al., 1974 Chitwood et al., 1987 Soriano et al., personal communication (2007 Porchet et al., 1984; Gaillet, 1985; Reuland et al., 1987 Sauber et al., 1983; Reuland et al., 1987 Sauber et al., 1983; Reuland et al., 1987 Kalarani et al., 1995 Zipser et al., 1998 Romer, 1979; Whithehead and Sellheyer, 1982 Gomot, 1984 Williams et al., 1986 Takemoto et al., 1967 — Franke and Käuser, 1989 Miyata et al., 2007

BIO: bioassay; GC: gas chromatography; HPLC: high-performance liquid chromatography; ISOL: isolation and complete chemical identification; MS: mass spectrometry; RIA: radioimmunoassay. Compounds: AjC: ajugasterone C; E: ecdysone; 2dE: 2-deoxyecdysone; 2d20E: 2-deoxy20-hydroxyecdysone; 20E: 20-hydroxyecdysone; 20,23E: gerardiasterone; 20,26E: 20,26-dihydroxyecdysone; 22d20E: taxisterone; MakA: makisterone A; PonA: ponasterone A. a This column refers to the most conclusive methodology used, either by the first authors or in subsequent reports by other authors working on the same group. b Diaulusterols and hyousterones are not « standard » ecdysteroids (see Fig. 2.4).

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(in animals), various C28 and C29 phytosterols (in plants), or specific C28 sterols (in yeast and fungi). The majority of phytophagous species can convert C28 and C29 sterols to cholesterol, a process termed dealkylation, thereby producing mainly ecdysone and 20-hydroxyecdysone, whereas those unable to dealkylate phytosterols will produce C28 and C29 analogues (Fig. 2.1). In insects, for instance, some Hemiptera, Hymenoptera and Diptera belong to the second category (Feldlaufer, 1989), and they produce either makisterone A (Kaplanis et al., 1975; Redfern, 1984; Feldlaufer and Svoboda, 1986), makisterone C (Feldlaufer et al., 1991; Mauchamp et al., 1993) or 24-epi-makisterone A (Feldlaufer et al., 1993; Maurer et al., 1993). Some species unable to dealkylate produce exclusively C28 ecdysteroids, and their receptors have evolved a high affinity for makisterone A (Aldrich et al., 1981), whereas others appear more flexible and are able to switch from C27 to C28 ecdysteroids according to their sterol diet; this is the case for e.g. Drosophila melanogaster (Redfern, 1986) and Bombus terrestris (Regali, 1996). As expected, the affinities of the Drosophila ecdysteroid receptor for 20-hydroxyecdysone and makisterone A are in the same range (Dinan and Hormann, 2005).

2.3

Diversity of Arthropod Ecdysteroids: Position and Number of Substituents

The synthesis of moulting hormones is a two-step process; classically, moulting glands (or more generally steroidogenic organs/cells) secrete an inactive « prohormone » which is then converted by peripheral tissues to the ‘active hormone’. If we consider the whole arthropod phylum, diversity concerns both the steroidogenic organs (moulting glands, epidermis and possibly oenocytes; see Redfern, 1989) and their secretory products (Lafont, 1997). Moulting glands (i.e. epidermal endocrine glands) have not been identified in all arthropods, thus the general epidermis (and/or associated oenocytes) has often been proposed as an/ the ecdysteroid source in more ‘primitive’ arthropod species (Romer and Gnatzy, 1981; Zhu et al., 1991; Lachaise et al., 1993), or even in some insect species/ stages when moulting glands are not functional (or have degenerated) (Delbecque et al., 1990; Jenkins et al., 1992). There is also some diversity of the secretory products of the steroidogenic tissues (Table 2.2), but after peripheral metabolism the major circulating ecdysteroid is 20-hydroxyecdysone (or a C28/C29 homologue), or ponasterone A in a few species (Apterygota and some Crustacea; Lachaise, 1989, 1990). Some species may not follow this rule, hence the presence, for instance, of high levels of circulating 2-deoxyecdysone in the coleopteran Zophobas atratus (Aribi et al., 1997). At a later step, the active moulting hormones undergo degradation through several pathways (Fig. 2.3) which result in a complex array of biologically inactive molecules: the metabolic pathways include 26-oxidation to ecdysonoic acids of general

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Diversity of Ecdysteroids in Animal Species

51

Compound

R1

R2

R3

R4

R5

R6

R7

R8

Ecdysone

OH

OH

H

H

H

H

OH

H

20-Hydroxyecdysone

OH

OH

H

OH

H

H

OH

H

3-Dehydroecdysone

OH

H

H

H

OH

H

=O

2-Deoxyecdysone

H

OH

H

H

H

H

OH

H

25-Deoxyecdysone

OH

OH

H

H

H

H

H

H

Ponasterone A

OH

OH

H

OH

H

H

H

H

Inokosterone

OH

OH

H

H

H

H

H

OH

20-Deoxymakisterone A

OH

OH

H

H

CH3

H

OH

H

Makisterone A

OH

OH

H

OH

CH3

H

OH

H

24-Epi-makisterone A

OH

OH

H

OH

H

CH3

OH

H

Makisterone C

OH

OH

H

OH

C2H5

H

OH

H

Fig. 2.1 Structures of the major circulating ecdysteroids in arthropods

occurrence, epimerization of the 3-OH (in lepidopteran larvae) and the formation of various kinds of polar and/or apolar conjugates (Lafont et al., 2005). All these classes of metabolites can be unambiguously characterized by their chromatographic behaviour and susceptibility to hydrolysis (Table 2.3). Some of these reactions (formation of 3-epimers or of 26-oic derivatives) constitute an irreversible inactivation process. On the other hand, conjugation is a potentially reversible process as conjugates can be converted back to active hormones: this is the case for maternal conjugates, which are deposited in the eggs and later used (at least in some species) during early embryonic development, until moulting glands become functional (Isaac and Slinger, 1989; Sonobe and Yamada, 2004; but see also Connat et al., 1988).

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Table 2.2 The major secretory products of arthropod moulting glands (Modified after Lafont, 1997) Type of ecdysteroid(s) Examples References E E + 2dE 3DE

Kiriishi et al., 1990 Locusta migratoria Zophobas atratus Aribi et al., 1997 Manduca sexta Warren et al., 1988 Orconectes limosus Böcking et al., 1993 Cancer antennarius Spaziani et al., 1989 E + 25dE Carcinus maenas Lachaise, 1989 Uca pugilator Hopkins, 1992 E + 3DE Pieris brassicae Blais and Lafont, 1991 25dE + 3DE Menippe mercenaria Rudolph and Spaziani, 1992 E + 25dE + 3DE Carcinus maenas Dauphin-Villemant et al., 1994 E + 20dMakA Drosophila melanogaster Redfern, 1984 E: ecdysone; 2dE: 2-deoxyecdysone; 3DE: 3-dehydroecdysone; 25dE: 25-deoxyecdysone; 20dMakA: 20-deoxy-makisterone A.

Fig. 2.2 Metabolic modifications of ecdysone or its higher homologues. (Modified after Lafont et al., 2005.) R = H, Me, or Et

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Fig. 2.3 Some unusual ecdysteroids. (a) Zoanthusterone from the zoanthid Zoanthus sp. (Suksamrarn et al., 2002); (b) palythoalone B from the zoanthid Palythoa australiae (Shigemori et al., 1999); (c) ajugasterone C from the Zoanthid Gerardia savaglia (Guerriero and Pietra, 1985) and from the sponge Agelas dispar (Cafieri et al., 1998); (d) 5α-Hydroxy-cholest-7-en-6-one from the sponge Oscarella lobularis (Aiello et al., 1991); (e) gymnasterone B, isolated from a fungal strain (Gymnascella dankaliensis) from the sponge Halichondria japonica (Amagata et al., 1998); (f) gymnasterol from the same fungus Gymnascella dankaliensis (Hayakawa et al., 2003)

2.4

Diversity of Ecdysteroid Functions in Arthropods

The first established function of ecdysteroids was their role in the control of moulting processes, which is logical, as they were isolated through a pupation bioassay-directed procedure (Butenandt and Karlson, 1954), a role which then

54 Table 2.3 Characterization of the different classes of zooecdysteroids Susceptibility Electrically to hydrolysis charged pKa (when applicable) (Yes/no) Polarity (Yes/no)

R. Lafont and J. Koolman

Conclusion

High

Y 6.5–7.0 Y Phosphate esters Y 4.5 N 26-Oic acids Y 2.0 Y Sulfate estersa N Y Glucosides Medium N N 26-OH derivatives N N E, 20E and related Low N N (Deoxy) precursors N N 14-Deoxy derivatives N Y Acetyl/acyl esters a Synthetic conjugates, only found in insects during in vitro tissue incubations (Shampengtong and Wong, 1989; Matsumoto et al., 2003).

proved general to all arthropods. Although there was early evidence for the presence of ecdysteroids in adult insects (Karlson and Stamm-Menéndez, 1956), their significance was not considered until Hagedorn et al. (1975) demonstrated that mosquito ovaries are a source of ecdysone. The role of ecdysteroids in female Aedes aegypti – i.e. the stimulation of vitellogenin synthesis by the fat body – is rather similar to that of ovarian steroids in oviparous vertebrates, but this scheme does not apply to all insects (Hagedorn, 1989; Lafont, 1991; Bellés, 1998). Ovarian ecdysteroids may indeed fulfil different (one or several) functions among the following: maternal supply of hormone to the eggs (general, i.e. the primitive function?), triggering of vitellogenesis (mosquitoes + other diptera), negative feed-back control of gonadotropins (for long-lived insect species which undergo several reproductive cycles, e.g. cockroaches), triggering of pheromone biosynthesis. This diversity connected with that of insect reproductive strategies has been considered as a good example of “hormone capture”, i.e. the acquisition by various tissues of a competence (receptors) to respond to the production/secretion of ovarian ecdysteroids, which would provide a selective advantage in a given situation (Hagedorn, 1989). As concerns males, the situation is not fully clear. The testes of adult blowflies contain significant amounts of ecdysteroids (Koolman et al., 1979). Source of these ecdysteroids in some species of Lepidoptera are the interstitial cells of larval testes (Loeb et al., 1982; Jarvis et al., 1994), but any generalization would be premature, since, by contrast, in Aedes aegypti male adults no ecdysteroid biosynthesis was detected (Sieglaff et al., 2005), whereas in Anopheles gambiae it was observed to take place in male accessory glands, and not in testes (Pondeville et al., 2007). The role of these ecdysteroids (e.g. in the control of germ cell proliferation, spermatogenesis, accessory glands differentiation) needs more detailed investigation. So what were the primitive functions of ecdysteroids? An attractive hypothesis was proposed by Hagedorn (1989). He suggested that the original source of ecdysteroids were the ovarian follicle cells, involved in the large maternal supply of ecdysteroids for supporting both embryonic and post-embryonic development. We

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can propose that this role was later restricted to embryonic development, together with the development of the steroidogenic capacity of (larval) epidermis, from which moulting glands progressively differentiated (Bückmann, 1984; Lachaise et al., 1993). Finally, an allelochemical function has also to be considered. Pycnogonum litorale (a pantopod) accumulates huge amounts of ecdysteroids in its integument (Bückmann et al., 1986) which are released into the surrounding seawater upon stress and act as a feeding deterrent against crabs (Tomaschko, 1994, 1995, 1999). Among insects, the chrysomelid Chrysolina carnifex possesses exocrine glands which secrete a highly concentrated solution of 20-hydroxyecdysone 22-acetate, able to deter predatory spiders (Laurent et al., 2003). These examples are reminiscent of plant ecdysteroids (phytoecdysteroids) acting as feeding deterrent against phytophagous insects (Dinan, 2001).

2.5

Ecdysteroids in Non-arthropod Invertebrate Species: Nature and Origin

The presence and putative functions of ecdysteroids in non-arthropod invertebrates has been reviewed several times (see e.g. Hoffmann and Charlet, 1985; Spindler, 1988; Walgraeve and Verhaert, 1988; Käuser, 1989; Koolman, 1990; Lafont, 1991; Lafont and Mathieu, 2007). These molecules were found thanks to analytical tools and not from homologous bioassay-directed isolation. Thus, we are faced with a typical situation of “reverse endocrinology” (Lafont, 1991, 2000). Different situations are possible: 1. Either ecdysteroids (or ecdysteroid-related molecules) have been unambiguously identified (i.e. isolated and fully characterized by MS and NMR), or their presence relies on strong (e.g. GC-MS or HPLC-MS) or weak (e.g. RIA alone) experimental evidence (see Table 2.1). 2. They are present either in large (e.g. in zoanthids) or minute amounts. 3. Their distribution shows (or does not show) significant tissue differences. 4. Their concentrations show (or do not show) fluctuations correlated with developmental (e.g. moulting in nematodes or leeches) and/or, more often, reproductive (e.g. vitellogenesis) events. 5. Exogenously applied ecdysteroids (or agonists, or anti-ecdysone antibodies) disturb these processes (but is this physiological or pharmacological?). 6. There is evidence for (or against) an endogenous production of these molecules by the whole animal or by individual organs e.g. gonads or endocrine glands (e.g. dorsal bodies of snails). This last point has been addressed by many authors and investigated in several ways: 1. In vitro analysis of the release/production of immunoreactive material can be investigated with individual organs (gonads), but this cannot provide a proof of de novo synthesis.

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2. In vivo/in vitro labelling experiments can be performed with either cholesterol or the biosynthetic intermediates already used with insects (Table 2.4), but only the use of cholesterol can demonstrate the presence of a full biosynthetic pathway. 3. In vitro enzymatic studies can provide evidence that a given reaction may (or may not) take place (Table 2.5: this supposes that the biosynthetic pathway, if present, is the same as that in insects), but as such they do not demonstrate that the observed reactions are specific (or involve enzymes which use totally different endogenous substrates). With the above criteria, there is presently no definitive evidence for the endogenous synthesis of ecdysteroids outside arthropods: in particular, all investigated species lack the 2- and 22-hydroxylases, i.e. the two mitochondrial cytochrome P450 enzymes crucial for ecdysone biosynthesis. It must be noted that this approach concluded that there is an absence of 2-hydroxylase in Lithobius forficatus, a myriapod (Descamps and Lafont, 1993), so we cannot exclude that these negative data result from the use of inadequate substrates: at the moment, it is premature to state that there is a single biosynthetic pathway for animal ecdysteroids. To exclude a direct/indirect dietary origin is not so easy: many animals feed on plants, 5% of which contain significant amounts of phytoecdysteroids (Dinan, 2001; see also Ecdybase – Lafont et al., 2002); other are predators (e.g. zoanthids, nudibranchs) or filter feeders (e.g. tunicates) and may feed on arthropods: moreover, all aquatic animals (especially filter feeders) might accumulate ecdysteroids Table 2.4 Search for an endogenous origin of ecdysteroids in non-arthropod invertebrates and in myriapods (After Lafont et al., 1995) Species Precursor Products Reference Nematodes: Caenorhabditis elegans Nematodes: Litomosoides carinii Nematodes: Parascaris equorum Nematodes: D. immitis, B. pahangi Annelids: Perinereis cultrifera Annelids, Gastropods

C, 22,25dE

No E or 20E

C

No E or 20E

Chitwood and Feldlaufer, 1990 Koolman et al., 1984

C, K, 2dE

No E or 20E

O’Hanlon et al., 1987

C, K, 2dE E K

No E or 20E No 20E 2,22dE, (2dE, E, 20E ??) No E or 20E No E or 20E

Mercer et al., 1989

2dE C K

Myriapods: Lithobius forficatus

C K

2,22dE, 2,22d20E, no E or 20E No E or 20E

Gaillet, 1985

Garcia et al., 1986, 1989, 1995

Descamps and Lafont, 1993

2,22dE, 2dE, no E or 20E E 20E C: cholesterol; K: ketodiol (2,22,25-trideoxyecdysone); 2,22dE: 2,22-dideoxyecdysone; 2dE: 2-deoxyecdysone; E: ecdysone; 20E: 20-hydroxyecdysone; ?? means tentative identification

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Table 2.5 Presence/absence of some key hydroxylases involved in ecdysteroid biosynthesis (or metabolism) in Invertebrates (Modified after Lafont et al., 1995, 2005) Hydroxylations at positions C2 C20 C22 C25 C26 Other Animals Arthropods Insects + + + + + 16β, 23 Crustaceans + + + + + Ticks + + + + + Myriapods − + + + ? Non-arthropods Molluscs − + − + ? 16β (gastropods) Annelids Achaetes − + − + ? 16β Polychaetes − + − + + Oligochaetes − − − + ? 18 Nematodes − − ? ? + Nemertines ? − ? ? ? ?: Does not mean that the reaction is absent, but only that there is at the moment no evidence for it (with the substrates tested).

even present in minute amounts in their environment. The endogenous origin may become even more difficult to define/assess when animal (e.g. sponges, zoanthids, nudibranchs) hosts possess algal or fungal symbionts, some of which may produce ecdysteroids or ecdysteroid-related molecules (Amagata et al., 1998; Hayakawa et al., 2003). Of course, an exogenous origin does not exclude a vitamin-like biological activity, and indeed there are many observations that exogenously applied ecdysteroids act like endocrine disrupters, thus supporting the idea that ecdysteroids (or at least closely related molecules) may fulfil a hormonal function in the concerned animals. This last idea is however questionable, if we consider that plenty of plant molecules structurally unrelated to ecdysteroids may act as weak agonists or antagonists in the Drosophila BII cell bioassay (Dinan et al., 2001; Dinan and Hormann, 2005).

2.6

Ecdysteroids in Non-arthropod Invertebrate Species: Do They Have a Biological Significance?

Porifera contain many ecdysteroids or ecdysteroid-related molecules. They are remarkable for the great diversity of steroid ring oxidation patterns which gives rise to hundreds of different oxysterols, many of which are cytotoxic (Sarma et al., 2005). The contribution of symbionts to this diversity is an open question (e.g. Amagata et al., 1998; Hayakawa et al., 2003), and it is generally considered that they serve as a chemical defence. Anthozoa also may contain very large amounts of ecdysteroids, the dietary (or symbiotic) origin of which has not been

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investigated, and a defensive role has also been proposed (e.g. Guerriero and Pietra, 1985; Suksamrarn et al., 2002). A similar role was proposed for the diaulusterols A and B of the nudibranch Diaulula sandiegensis (which feeds on sponges) (Williams et al., 1986) and for the hyousterones of the tunicate Synoicum adareanum (Miyata et al., 2007), which contains also diaulusterol B (Fig. 2.4). The latter case is of particular interest, because it represents the first evidence for the presence of ecdysteroids in a Deuterostome species. Among molluscs, only snails were investigated in detail (Romer, 1979; Whitehead and Sellheyer, 1982; Gomot, 1984). It was shown that dorsal bodies (endocrine ganglia close to the brain which are known from extirpation experiments to control growth and reproduction of snails) secrete immunoreactive ecdysteroids (Nolte et al., 1986; Mukai et al., 2001). Exogenous ecdysteroids are metabolised by original pathways (Garcia et al., 1986), they slightly stimulate growth (Garcia et al., 1995), and they promote regeneration (Whitehead, 1977; Whitehead and Saleuddin, 1978) and reproduction (egg maturation, polysaccharide synthesis by the albumen gland: Mukai et al., 2001). These data, although encouraging, do not suffice to prove an endocrine function of ecdysteroids in gastropods. The case of annelids is rather similar (Porchet et al., 1984). Early studies were performed with leeches, because these animals undergo regular moulting, and a correlation was established between moults and the fluctuation of ecdysteroid titres (Sauber et al., 1983). Evidence was obtained for the presence of 2-deoxyecdysonelike molecules (not isolated) in all classes of annelids (Gaillet, 1985; Reuland et al., 1987), and on the effect of ecdysteroids on gametogenesis (Kalarani et al., 1995), but these experiments were not further developed. More recently, several oxysterols were isolated and fully characterized from the leech Hirudo medicinalis (Zipser et al., 1998); they show a limited relationship to ecdysteroids (see Fig. 2.4), and their structure is in agreement with the reported absence of 2- and 22-hydroxylases in this species (Garcia et al., 1989). Helminths: are ecdysteroids of any diagnostic use? The observation of ecdysteroids in a few nematode and platyhelminth species (Table 2.1) raised the question whether helminths (parasitic worms belonging to the taxa of nematodes, trematodes and cestodes) could be detected in their hosts by the analysis of ecdysteroids secreted by these parasites. This would enable the physician to diagnose a parasitosis (i.e. an infection by helminths) on the basis of an analysis of a blood or urine sample taken from a patient (Nirdé et al., 1984a, b; Koolman et al., 1984; Koolman and Moeller, 1986). For detection of these ecdysteroids sensitive radioimmunoassays were employed. Unfortunately a low background of immunoreactive ecdysteroids was detected in the serum of healthy mammals as well (Simon and Koolman, 1989). This background of immunoreactive ecdysteroids precluded the detection of parasites infecting a human host because of their low ecdysteroid content (and, possibly, secretion). More specific analytical tools with a similar or even better sensitivity should allow resumption of the approach. The method will work only if the background ecdysteroids in humans, probably a result of ecdysteroids absorbed from the food, differ chemically from the ecdysteroids secreted by the helminths.

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Fig. 2.4 Examples of the many Δ4-steroids present in animals, plants or fungi. (a) 4-Dehydroecdysterone from the zoanthid Parazoanthus sp. (Searle and Molinski, 1995); (b) 14-Hydroxyergosta-4,7,22-triene-3,6-dione from the fungus Phellinus igniarius (Honda et al., 1996); (c) hyousterones A (R = 14α-OH) and B (R = 14β-OH) from the tunicate Sinoycum adareanum (Miyata et al., 2007); (d) Δ4-Diketol, a putative ecdysone biosynthesis intermediate (Blais et al., 1996); (e) diaulusterol B from the nudibranch Diaulula sandiegensis (Williams et al., 1986); (f) pinnasterol from the red alga Laurencia pinnata (Fukuzawa et al., 1986); (g) oxysterol from the leech Hirudo medicinalis (Zipser et al., 1998); (h) ergosta-4,7,22-triene-3,6-dione from the marine sponge Raphidostila incisa (Malorni et al., 1978)

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Nematodes have been extensively studied, in particular because many species are harmful parasites, but also because they undergo moulting and, with arthropods, phylogenetically belong to the Ecdysozoa (reviewed in: Barker and Rees, 1990; Chitwood, 1999). Early on, 20-hydroxyecdysone was isolated from Ascaris lumbricoides (Horn et al., 1974), but in minute amounts only (0.3 µg/kg). Although the presence of ecdysteroids was confirmed by several other studies, it was not possible to demonstrate that nematodes are able to de novo synthesize ecdysteroids from cholesterol or even from later precursors (Table 2.4). On the other hand, many effects of exogenous ecdysteroids have been described (Dennis, 1977b; Barker et al., 1991; Goudey-Perrière et al., 1992; Warbrick et al., 1993). A totally different approach to this problem was developed using Cænorhabditis elegans, taking profit of the availability of its entire genome sequence and of the possibility to study mutants where some genes were inactivated. This work rapidly demonstrated that a steroid (the ligand of DAF-12, a nuclear receptor) was involved in the control of reproduction, development and longevity (see Beckstead and Thummel, 2006), but the endogenous ligands of this receptor were identified as novel steroids, dafachronic acids (Fig. 2.5), i.e. neither ecdysteroids nor vertebrate-type steroids (Motola et al., 2006; Rottiers et al., 2006). However, this story is perhaps not finished, as other authors showed that vertebrate-type steroids, including pregnenolone, may also act through the same receptor DAF-12 (Broué et al., 2007). Nemertines contain ecdysteroids, and the presence of 20-hydroxyecdysone has been conclusively demonstrated (Okazaki et al., 1988, 1998). They also belong to the Ecdysozoa, and they are predatory animals feeding on polychaetes or crustaceans, which are expected to ingest ecdysteroids with their food, especially in the latter case. Metabolic studies with [3H]ecdysone failed to demonstrate a 20-hydroxylation (Snyder et al., 1992), thus there is presently no evidence for their ability to endogenously produce ecdysteroids. Finally, it may be worth signalling that some protozoa (symbionts or parasites of insects) undergo sexual reproduction (Cleveland and Burke, 1960; Cleveland et al., 1960) or sporulation (Lord and Hall, 1983) as a response to their host’s ecdysteroids. Similar examples of the use of host hormones have also been described for vertebrate parasites.

2.7

Ecdysteroid-Related Molecules and “Protoecdysteroids” vs. “Vertebrate-Type” Steroids?

Ecdysteroids bear several specific structural features: they usually keep the whole carbon skeleton of the sterol from which they are formed, they bear a 7-en-6-one chromophore, a cis-A/B-ring junction (5β-H), and several hydroxyl groups (2, 3, 14, 20, 22, 25). The formation of the chromophore is not yet fully understood, but we know that it starts from a Δ5,7-sterol (7-dehydrocholesterol, which occupies a key position) and probably proceeds through the formation of a 4,7-diene-3,6-dione;

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Diversity of Ecdysteroids in Animal Species

A

61

B

OH

OH

O

OH

O

HO OH

D

C

OH

OH

HO

O OH OH

Fig. 2.5 Some very unusual invertebrate steroids. (a) Δ4-Dafachronic acid from the nematode C. elegans (Motola et al., 2006); (b) geodisterol, an oxysterol from the marine sponge Geodia sp. (Wang and Crews, 1996); (c) bombycosterol from Bombyx mori ovaries (Fujimoto et al., 1985); (d) cybisterone, a defensive steroid from Cybister lateralimarginalis (Schildknecht et al., 1967)

the 5β-H would then be formed by a 5β-reductase (Blais et al., 1996), as is the case for bile acids. Slight modifications of this process (Fig. 2.6) may lead to molecules still bearing a Δ4-double bond, and/or a hydroxyl instead of an oxo-function at C-6. A good picture of the possible diversity is given by sponges (Sarma et al., 2005), as this group may even include hybrid molecules like geodisterol (Fig. 2.5) a C29-molecule which bears an aromatic A-ring (Wang and Crews, 1996). Concerning the different hydroxylations (in addition to the 3β-OH already present in sterols), it is probable that many combinations have been “tried” (consider again the diversity of sponge oxysterols; Aiello et al., 1999), and that like in vertebrates, the total number of possible hydroxylases has progressively increased by gene duplication and diversification. From the available metabolic studies (Tables 2.4 and 2.5), we assume that the two mitochondrial enzymes (2- and 22-hydroxylases) were the last to appear, whereas the 26-hydroxylase (which is involved in hormone inactivation in insects) appeared early (it is already present in C. elegans [DAF-9] and is involved in dafachronic acid biosynthesis), and is widespread among animals and plants (Meaney, 2005). In the same way, 20-hydroxylase (i.e. the last biosynthetic step of insects) has appeared earlier than the 2- and 22-hydroxylases. All this may help us to imagine which (2,22-dideoxy-)steroid hormones could be found in different groups of invertebrates. The insect steroid story is however not so simple. More than 20 years ago, a strange compound, bombycosterol (Fig. 2.5) was isolated from Bombyx ovaries

Fig. 2.6 Suggested scenario for the evolution of A/B-ring functionalization starting from 7-dehydrocholesterol. This scenario would possibly make 14α-hydroxylation an individual reaction

62 R. Lafont and J. Koolman

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together with various ecdysteroids (Fujimoto et al., 1985). It has not yet been found in any other insect species, and its biological relevance is unknown. Whether this 2,22-dideoxy compound is biogenetically related to ecdysteroids is not clear, but this question would merit further investigation. Another question is: do insects/arthropods produce other types of steroids beyond ecdysteroids (e.g. “vertebrate-type” steroids)? Three facts will be mentioned here, in order to show that this might well be the case: 1. During ecdysteroid metabolic studies, side-chain cleavage between carbons 20 and 22 has been observed in Calliphora (Galbraith et al., 1969) and in Bombyx (Hikino et al., 1975) larvae, which means that a 20,22-desmolase is present in these species (or in associated microorganisms?). 2. During in vivo cholesterol labelling experiments with Manduca sexta, a labelled pregnenolone glucoside was isolated unexpectedly (Thompson et al., 1985). 3. Several species of aquatic Coleoptera have steroid-synthesizing exocrine glands which produce large amounts of vertebrate-type steroids (cybisterone [Fig. 2.5] Schildknecht et al., 1967; see also Chadha et al., 1970; Chapman et al., 1977).

2.8

Conclusion: Future Approaches Should/Will Take Profit of Our Knowledge of Animal Genomes

Molecular biology presently offers new tools for reconsidering the evolution of steroid hormones in animals. Biosynthetic enzymes belong essentially to two large families, i.e. hydroxysteroid dehydrogenases (HSDs, which include also several reductases, e.g. the 5α/5β-reductases), and cytochrome P450 monooxygenases (CYPs). Every species contains many genes encoding such enzymes, which are also involved in plenty of other metabolic processes, e.g. xenobiotic detoxification. With the increasing number of genome sequences available, it is possible to search in a given species for orthologue genes to those identified in another species, and this applies well to insects as concerns ecdysteroid biosynthetic enzymes (Rewitz et al., 2007). Owing to the rather rapid evolution of such genes, it is advisable to use a protein expression system for assessing the catalytic activities of the cloned enzymes. Indeed, a single amino acid change suffices to change the substrate specificity for a CYP enzyme (Ramaro and Kemper, 1995). Testing the enzyme activities requires the availability of adequate substrates, only a few of which are commercially available, thus it is crucial that organic chemists are interested in such projects. The rapid evolution of enzymes prevents the identification of homologous genes directly from insects to lower invertebrates, but it is hoped that moving from insects to crustacea and then to arachnida will prove feasible. Thereafter, it will perhaps be possible to move to lower invertebrates. Of interest is to note that molecular probes will unambiguously allow the determination of the tissue expression of biosynthetic

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genes (which is of fundamental importance for arthropod species where endocrine glands have not been identified), and to ascertain which cells are steroidogenic within a given organ. Such a strategy can be illustrated by “vertebrate-type” steroid hormones: tunicates seem to be unable to produce such steroids, as they lack several key genes (Campbell et al., 2004). On the other hand, they contain (produce?) the ecdysteroid-related hyousterones (Miyata et al., 2007). By contrast, Cephalochordates (Amphioxus) contain the whole set of genes required for the production of vertebrate-type steroids and are therefore closer to vertebrates than urochordates (Mizuta and Kubokawa, 2007). The same strategy applies also to steroid hormone receptors, which belong to the family of nuclear receptors. Ecdysone receptors (EcRs) have been cloned from insects and crustaceans, and phylogenetic reconstructions show that EcRs are close to vertebrate farnesoid X receptor (FXR) (Enmark and Gustafsson, 2001; Laudet and Bonneton, 2005). Again, such studies will allow the definition of which species possess receptors close to those of insects, and also which tissues are targets of steroid hormones (De Mendonça et al., 2000). That this task is not so easy is illustrated by the observation that the genome of C. elegans contains >280 nuclear receptor genes (Gissendanner et al., 2004). That this strategy may have its limits can be drawn from the fact that the “estrogen receptor” cloned in Aplysia californica has lost the ability to bind estrogens (Thornton et al., 2003). Clearly molecular approaches open new ways to answer old questions, e.g. do some animals outside arthropods produce ecdysteroids and use them as hormones? Which tissues (outside moulting glands and female gonads) are able to produce ecdysteroids? Do Insects produce/use other types of steroids? No doubt an exciting new era is opening up now.

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Porchet M, Gaillet N, Sauber F, Charlet M, Hoffmann JA (1984) Ecdysteroids in Annelids. In: Hoffmann J, Porchet M (eds), Biosynthesis, Metabolism and Mode of Action of Invertebrate Hormones. Springer, Berlin, pp 346–348. Ramaro M, Kemper B (1995) Substitution at residue 473 confers progesterone 21-hydroxylase activity to cytochrome P450 2C2. Mol Pharmacol 48:417–424. Redfern CPF (1984) Evidence for the presence of makisterone A in Drosophila larvae and the secretion of 20-deoxymakisterone A by the ring gland. Proc Natl Acad Sci USA 81:5643–5647. Redfern CPF (1986) Changes in patterns of ecdysteroid secretion by the ring gland of Drosophila in relation to the sterol composition of the diet. Experientia 42:307–309. Redfern CPF (1989) Ecdysiosynthetic tissues. In: Koolman J (ed), Ecdysone, from Chemistry to Mode of Action. Georg Thieme Verlag, Stuttgart, pp 182–187. Rees HH (1989) Zooecdysteroids: structures and occurrence. In: Koolman J (ed), Ecdysone, from Chemistry to Mode of Action. Georg Thieme Verlag, Stuttgart, pp 28–38. Rees HH, Mendis AHW (1984) The occurrence and possible physiological significance of ecdysteroids during nematode and cestode development. In: Hoffmann JA, Porchet M (eds) Biosynthesis, Metabolism and Mode of Action of Invertebrate Hormones. Springer, Berlin, pp 338–345. Regali A (1996) Contribution à l ‘étude des besoins alimentaires en stéroïdes de Bombus terrestris (L.), Thesis, University of Mons-Hainaut, Belgium, 143 pp. Reuland M, Sauber F, Boilly Y, Charlet M (1987) Further investigations on ecdysteroids in Annelids: predominance of 2-deoxyecdysone-like ecdysteroids in the three classes of Annelids, Polychaeta, Oligochaeta and Hirudinea. Communication presented at the VIII Ecdysone Workshop, Marburg, Germany. Rewitz KF, O’Connor MB, Gilbert LI (2007) Molecular evolution of the insect Halloween family of cytochrome P450s: phylogeny, gene organization and functional conservation. Insect Biochem Mol Biol 37:741–753. Romer F (1979) Ecdysteroids in snails. Naturwissenschaften 66:471. Romer F, Gnatzy W (1981) Arachnid oenocytes: ecdysone synthesis in the legs of harvestmen (Opilionidae). Cell Tissue Res 216:449–453. Rottiers V, Motola DL, gerisch B, Cummins CL, Nishiwaki K, Mangelsdorf DJ, Antebi A (2006) Hormonal control of C. elegans dauer formation and life span by a Rieske-like oxygenase. Dev Cell 10:473–482. Rudolph PH, Spaziani E (1992) Formation of ecdysteroids by Y organs of the crab, Menippe mercenaria. II. Incorporation of cholesterol into 7-dehydrocholesterol and secretion products in vitro. Gen Comp Endocrinol 88:235–242. Sarma NS, Sri Rama Krishna M, Ramakrishna Rao S (2005) Sterol ring system oxidation pattern in marine sponges. Mar Drugs 3:84–111. Sauber F, Reuland M, Berchtold JP, Hétru C, Tsoupras G, Luu B, Moritz ME, Hoffmann, JA (1983) Cycle de mue et ecdystéroïdes chez une Sangsue, Hirudo medicinalis. C R Acad Sci Sér III 296:413–418. Schildknecht H, Siewerdt R, Maschwitz U (1967) Über Arthropodenabwehrstoffe, XXIII. Cybisteron, ein neues Arthropoden-Steroid. Liebigs Ann Chem 703:182–189. Searle PA, Molinski TF (1995) 4-Dehydroecdysterone, a new ecdysteroid from the zoanthid Parazoanthus sp. J Nat Prod 58:264–268. Shampengtong L, Wong KP (1989) An in vitro assay of 20-hydroxyecdysone sulfotransferase in the mosquito, Aedes togoi. Insect Biochem 19:191–196. Shigemori H, Sato Y, Kagata T, Kobayashi J (1999) Palythoalones A and B, new ecdysteroids from the marine zoanthid Palythoa australiae. J Nat Prod 62:372–374. Sieglaff DH, Adams Duncan K, Brown MR (2005) Expression of genes encoding proteins involved in ecdysteroidogenesis in the female mosquito, Aedes aegypti. Insect Biochem Mol Biol 35:471–490. Simon P, Koolman J (1989) Ecdysteroids in Vertebrates : pharmacological apsects. In: Koolman J (ed), Ecdysone, from chemistry to mode of action, Georg Thieme Verlag, Stuttgart. pp 254–259.

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Snyder MJ, Okazaki RK, Chang ES (1992) Nemertean ecdysteroids: relationship to reproduction. Invertebr Reprod Dev 21:7–13. Sonobe H, Yamada R (2004) Ecdysteroids during early embryonic development in silkworm Bombyx mori: metabolism and functions. Zool Sci 21:503–516 Spaziani E, Rees HH, Wang WL, Watson RD (1989) Evidence that Y-organs of the crab Cancer antennarius secrete 3-dehydroecdysone. Mol Cell Endocrinol 66:17–25. Spindler KD (1988) Parasites and hormones. In: Mehlhorn H (ed), Parasites. Springer, New York, pp 465–476. Spindler KD (1989) Hormonal role of ecdysteroids in Crustacea, Chelicerata and other Arthropods. In: Koolman J (ed), Ecdysone, from Chemistry to Mode of Action. Georg Thieme Verlag, Stuttgart, pp 290–295. Sturaro A, Guerriero A, Declauser R, Pietra F (1982) A new, unexpected marine source of a moulting hormone: isolation of ecdysterone in large amounts from the zoanthid Gerardia savaglia. Experientia 38:1184–1185. Suksamrarn A, Jankam A, Tarnchompoo B, Putchakarn S (2002) Ecdysteroids from a Zoanthus sp. J Nat Prod 65:1194–1197. Takemoto T, Ogawa S, Nishimoto N, Hoffmeister H (1967) Steroide mit Häutungshormon4ktivität aus Tieren und Pflanzen. Z Naturforsch B 22: 681–682. Thompson MJ, Svoboda JA, Lusby WR, Rees HH, Oliver JE, Weirich GF, Wilzer KR (1985) Biosynthesis of a C21 steroid conjugate in an insect: the conversion of [14C]cholesterol to 5-[14C]pregnen-3α,20α-diol glucoside in the tobacco hornworm, Manduca sexta. J Biol Chem 260:15410–15412. Thornton JW, Need E, Crews D (2003) Resurrecting the ancestral receptor: ancient origin of estrogen signalling. Science 301:1714–1717. Tomaschko K-H (1994) Defensive secretion of ecdysteroids in Pycnogonum litorale (Arthropoda, Pantopoda). Z Naturforsch 49c:367–371. Tomaschko K-H (1995) Autoradiographic and morphological investigations of the defensive ecdysteroid glands in adult Pycnogonum litorale (Arthropoda: Pantopoda). Eur J Entomol 92:105–112. Tomaschko K-H (1999) Nongenomic effect of ecdysteroids. Arch Insect Biochem Physiol 41:89–98. Walgraeve HRMA, Verhaert PDEM (1988) Presence and function of ecdysteroids in Invertebrates. ISI Atl Sci 1:164–172. Wang YS, Crews P (1996) Geodisterol, a novel polyoxygenated sterol with an aromatic A ring from the tropical marine sponge Geodia sp. Tetrahedron Lett 37:8145–8146. Warbrick EV, Barker GC, Rees HH, Howells RE (1993) The effect of invertebrate hormones and potential hormone inhibitors on the third larval moult of the filarial nematode, Dirofilaria immitis, in vitro. Parasitology 107:459–463. Warren JT, Sakurai S, Rountree DB, Gilbert LI (1988) Synthesis and secretion of ecdysteroids by the prothoracic glands of Manduca sexta. J Insect Physiol 34:571–576. Whitehead DL (1977) Steroids enhance shell regeneration in an aquatic gastropod (Biomphalaria glabrata). Comp Biochem Physiol 58C:137–141. Whitehead DL, Saleuddin ASM (1978) Steroids promote shell regeneration in Helix aspersa (Gastropoda: Pulmonata). Comp Biochem Physiol 59C:5–10. Whitehead DL, Sellheyer K (1982) The identification of ecdysterone (20-hydroxyecdysone) in 3 species of molluscs (Gastropoda: Pulmonata). Experientia 38:1249–1251. Williams DE, Ayer SW, Andersen J (1986) Diaulusterols A and B from the skin extracts of the dorid nudibranch Diaulula sandiegensis. Can J Chem 64:1527–1529. Zhu XX, Oliver JH Jr, Dotson EM (1991) Epidermis as the source of ecdysone in an argasid tick. Proc Natl Acad Sci USA 88:3744–3647. Zipser B, Bradford JJ, Hollingsworth RI (1998) Cholesterol and its derivatives, are the principal steroids isolated from the leech species Hirudo medicinalis. Comp Biochem Physiol 120C:269–282.

Chapter 3

Crustacean Ecdysteroids and Their Receptors Penny M. Hopkins

Abstract Ecdysteroids in crustaceans differ substantially from those of their fellow arthropods, the insects. Crustacean ecdysteroids and ecdysteroid nuclear receptors are similar to those of insects, but differ in the number of hormones and in the number and structure of the receptor isoforms. Moreover, the control(s) of ecdysteroid synthesis by crustacean Y-organs is primarily inhibitory - through molt-inhibiting hormone (MIH) - whereas in insects ecdysteroid synthesis is positively stimulated by a very different neurosecretory hormone. The in vivo effects of ecdysteroids are less understood in crustaceans than in insects but appear to have some concordance. Ecdysteroid-responsive genes in crustaceans are just beginning to be uncovered and may have some identities to insect genes. The differences in ecdysteroid control between insects and crustaceans are thought to have evolved to accommodate the differences in life-histories seen in these diverse arthropod groups. Keywords Crustaceans • Y-organs • ecdysteroids • invertebrate nuclear receptors

3.1

Introduction

During the course of evolution the utilization of ecdysteroids by arthropods has undergone much divergence. Crustaceans and insects both draw on ecdysteroids to control a variety of important physiological events especially growth and differentiation. It is becoming increasingly obvious, however, that the overall control by ecdysteroids in crustaceans is very different from that of insects. Unlike most insects, crustaceans have a rather complex adulthood, where control of growth and reproduction must be alternated. The predominant hormones coordinating insect larval molts and metamorphosis differ from the number and variety in crustaceans. Differences are also seen in the number and structure of functional crustacean ecdysteroid receptor isoforms when compared to insect receptors. P.M. Hopkins () Department of Zoology, University of Oklahoma, Norman, 73019, USA e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Ecdysteroids in Crustaceans

Levels of circulating crustacean ecdysteroids differ from animal to animal and from researcher to researcher because of natural variations but also because of variations in methods used. RIA remains the most sensitive method to measure total circulating ecdysteroids and RIA coupled with HPLC and/or LC-MS is used to identify individual circulating ecdysteroids. Results, however, disagree due to differences in the sensitivity and specificity of the antibodies. Levels of total ecdysteroid in the hemolymph of crustaceans remain very low during the intermolt portion of the molt cycle – a time when animals are feeding and mating. Lowest reported levels of circulating ecdysteroids during intermolt range from undetectable levels in many crustaceans to 3.3 ng/ml in the crab, Callinectes sapidus (Lee et al., 1998) and 16 ng/ml in the shrimp, Panaeus vannamei (Chan et al., 1988). Ecdysteroid levels rise rapidly as crustaceans approach molt (or ecdysis -E) and then abruptly fall just prior to E itself (Fig. 3.1a; Hopkins, 1992; Nakatsuji et al., 2000; Martin-Creuzburg et al., 2007). Highest reported levels of circulating ecdysteroids occur just prior to ecdysis and range from 600 ng/ml in the lobster, Homarus americanus (Chang, 1985) to 120 ng/ ml in the crayfish, Procambarus clarkii (Nakatsuji et al., 2000). Ecdysteroids in malacostracans are produced in hypertrophied strips of hypodermis called Y-organs. Y-organs are found in varying locations in the anterior body

Fig. 3.1 Schematic representations of circulating levels of ecdysteroids in the hemolymph of the crab, Uca pugilator during a molt cycle. (a) Total RIA active ecdysteroids in the hemolymph (pg/ul) from an eyestalk-intact, non-regenerating crab. (b) Total RIA active ecdysteroids in hemolymph from a regenerating crab. Dotted line represents the R-value (= length of regenerating limb bud divided by the width of carapace times 100). C4 is intermolt stage of molt cycle, D0 is early proecdysis stage, D1–4 is late proecdysis stage, E is ecdysis (Redrawn from Chung et al.,1998a)

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cavity of these crustaceans and Y-organ anatomy is extremely variable (see Lachaise et al., 1993). Y-organs are currently considered the major source of ecdysteroids but within the microrcrustaceans (copepoda, phyllopoda, and cladocera) Y-organs probably do not exist – the hypodermis may be the major source of ecdysteroids in these smallest crustaceans. Most arthropods depend on dietary sterols and cholesterol for ecdysteroid synthesis. A major intermediate in ecdysteroid biosynthesis is 7-dehydrocholesterol which is converted to 2,22,25-trideoxyecdysone-ketodiol (Rees, 1985). In the insect prothoracic gland, there are sequential hydroxylations at the 2, 22 and 25 positions. The primary product in many insects is ecdysone (Meister et al., 1987; Smith and Sedlmeier, 1990) which is subsequently converted to 20-hydroxyecdysone in peripheral tissues (Gilbert et al., 2002). The Halloween enzymes that are responsible for the multiple hydroxylations of ketodiol in insects are discussed elsewhere in this volume. Steroid metabolism in decapod Y-organs is different from that of insect prothoracic glands (Fig. 3.2): The 25-deoxy form of ecdysone (25dE), as well as ecdysone (E), are produced and released by Y-organs of the crab, Carcinus maenas maintained in vitro (Lachaise et al., 1986, 1989). A third compound (3-dehydroecdysone) has been shown to be produced by in vitro Y-organs of the crab Cancer antennarius

Fig. 3.2 The major ecdysteroid products of crustacean Y-organs. On the left are the structures of the three major secretion products from crustacean Y-organs. On the right is a selection of references reporting the secretion products either in vivo or in vitro

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the freshwater prawn, Macrobrackium rosenbergii (Spaziani et al., 1989; Okumura et al., 1989) and the crayfish, Procambarus clarkii and Orconectes limosus (Sonobe et al., 1991; Okumura et al., 2003). 25dE and E released by the crab Y-organ are further hydroxylated at the 20-position by tissues other than the Y-organ. 25dE and its 20-hydroxylated metabolite ponasterone A (PA), as well as E and its metabolite 20 hydroxyecdysone (20E), have been found in the blood of three species of crabs (McCarthy, 1979; Lachaise and Lafont, 1984; Hopkins, 1992). Hydroxylation at the 25 position of the side chain must be the first step in formation of ecdysone in Drosophila because the 2,22-dihydroxy form of the ecdysteroid cannot serve as a substrate for 25 hydroxylase enzyme (Warren et al., 2004). This P450 hydroxylase may be a control point for the selective production of ecdysteroids in the crustacean Y-organ. The 25 hydroxylase is not necessary for the production of 25dE but it is necessary for the synthesis of E and 3dE. Due to the lack of an ecdysone oxidase in Y-organs, 3dE cannot be a metabolic product of E and its production and release by some Y-organs argues for a separate pathway (Lachaise et al., 1993). 3dE, E, and 25dE are all found in the hemolymph of crustaceans but in varying ratios and amounts during the molt cycle. Since both E and 25dE are found simultaneously in the hemolymph of the crab Uca pugilator (Hopkins, 1992), the 25 hydroxylase activity must be closely regulated in this crab Y-organ. Indeed, down regulation of the 25 hydroxylase gene (phantom- phm) appears to be a mechanism used to control circulating levels of ecdysteroid in Bombyx. In Bombyx fourth larval instar prothoracic glands the expression of Bmphm decreases 90% from its peak value and does not rise to previous activity until late in the fifth larval instar (Warren et al., 2004). Down regulation of the 25-hydroxylase in U. pugilator during late PE (D1–4) could account for the relatively higher levels of PA seen at that time (Hopkins, 1992). The variety of ecdysteroids found in the hemolymph of some crustaceans is, therefore, more diverse than that found in most insects. This difference may be a function of the differences in insect and crustacean life histories. The insect life strategy is, in effect, compartmentalized into larval and adult stages. Larval insects do not reproduce and most adult insects do not molt. Many crustaceans, on the other hand, continue to molt and reproduce well into adult stages. Because insects partition growth and reproduction into different stages, fewer hormones may be able to serve more functions. Whereas the more complex adult stage of crustaceans may require a greater variety of ecdysteroids.

3.3

Control of Ecdysteroid Synthesis/Release in Crustaceans

The in vivo control of Y-organ ecdysteroid synthesis has long been thought to be via neurosecretory hormones secreted by ganglia located in the stalked eyes of crustaceans (Carlisle, 1954). There are three ganglia located in the eyestalk that are extensions of the brain. These eyestalk ganglia contain a cluster of large neurosecretory cell bodies (the X-organ) that send axons to a single neurohemal site on the surface of the eyestalk ganglia (the sinus gland –Bliss and Welch, 1952). The brain controls and

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integrates environmental and physiological input that regulates the production/release of a variety of neuropeptides from the X-organ/sinus gland complex. At least one of these neuropeptides is thought to control ecdysteroid synthesis (Watson et al., 2001; Lago-Leston et al., 2007). The control of ecdysteroid synthesis in vivo, however, is more complex than a simple on/off control mechanism: During the course of the molt cycle, the blood of U. pugilator shows complicated patterns of various steroids and their metabolites. These patterns appear to be the result of a carefully controlled, complex program that seems to involve more than a single factor (Hopkins, 1983). The in vivo control of ecdysteroid production in crustaceans differs substantially from that in insects. In insects a neurosecretory hormone from the brain (PTTH) stimulates the release of ecdysteroids (Gilbert et al., 2002) while in crustaceans control of ecdysteroid production/release appears in some aspects to be an inhibitory control. It has been known for over a century that eyestalk removal (ESX) leads to molting (Zeleny, 1905). ESX also leads to elevated levels of circulating ecdysteroids in some crustaceans and re-implantation of eyestalk ganglia can reverse these preparations for molt (see Skinner, 1985).

3.3.1

Molt-Inhibiting Hormone (MIH)

Evidence that eyestalk extracts contain a putative molt-inhibiting hormone (MIH) capable of inhibiting the onset of molt was first presented for the crab Ocypode macrocera in 1965 (Rao, 1965). Since then a series of workers have shown, using in vitro techniques, that eyestalk or sinus gland extracts may affect molt by controlling production of steroids by the Y-organ (Soumoff and O’Connor, 1982; Watson and Spaziani, 1985; Webster and Keller, 1986; Naya et al., 1988; Lachaise et al., 1988; Sonobe et al., 1991). ESX crabs that proceed into a successful ecdysis, produce about 1.5X more total RIA-active ecdysteroids than do eyestalk-intact crabs demonstrating the inhibitory effect the eyestalks have on total ecdysteroid production (Hopkins, 1992). Eyestalk extracts also inhibit in vitro conversion of ketodiol to 25dE and E in Y-organs from the crab C. maenas (Lachaise et al., 1988). The pattern of circulating ecdysteroids during stage D is of low levels followed by a large peak four to five days prior to E. This pattern is exactly the same in eyestalk intact and ESX crabs. The large increase in ecdysteroids days after eyestalk removal cannot be due to further reductions in MIH titers because the half life of MIH in the hemolymph is only 10–20 min (Chung and Webster, 2005). It would seem that the patterns of circulating ecdysteroids must be controlled by more than eyestalk-derived inhibiting factors. Moreover, ESX in crustaceans does not always result in molting (Carlisle, 1954; Flint, 1972; Lachaise, 1992). In the crab, U. pugilator, half of the crabs undergoing ESX fail to molt (Hopkins, 1992). The total RIA-active ecdysteroids in the hemolymph of these non-molting crabs fails to reach the very high late PE levels observed in molting crabs (Hopkins, 1989a). Apparently these crabs never prepare

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for E since the Y-organs are not activated, even though the eyestalks are removed. Both the large PE peak of ecdysteroids in ESX crabs that molt and the failure of a large number of other ESX crab to enter PE argues very strongly for extraeyestalk source(s) of Y-organ control. One such extra-eyestalk control could be feedback effects of ecdysteroids on the Y-organ itself. The Y-organs of the shrimp Marsupenaeus japonicus contain the highest levels of ecdysteroid receptors of any tissue examined (Asazuma et al., 2007). A mixture of ecdysteroids extracted from the hemolymph have been shown to have a stimulatory effect on in vitro release of ecdysteroids from Y-organs in U. pugilator (Hopkins, 1986) while in the crayfish, Orconectes limosus, exogenous 20E and RH-5849 had inhibitory effects on both in vivo circulating levels and in vitro Y-organ release of ecdysteroids (Dell et al., 1999). It is hard to determine what holds the Y-organs in abeyance for such a long time after ESX or why Y-organs in some ESX crabs fail to produce ecdysteroids in preparation for molt but it is clearly more complicated than first envisioned. Purified MIHs have been isolated and sequenced from the crabs, Carcinus maenas and Charybdis teriatus, (Webster, 1991; Chan et al, 1998), the shrimps, Penaeus japonicus, Penaeus vannamei, Penaeus monodon and Metapenaeus ensis (Yang et al., 1996; Wang et al., 2000; Krungkasem et al., 2002; Gu et al., 2002), and the lobster, Jasus lalandii (Marco et al., 2000). Endogenous MIH levels in vivo are quite low ranging from 4–6 fmoles/ul of hemolymph (Nakatsuji and Sonobe, 2004; Chung and Webster, 2005). A number of different MIHs have been identified with 74–78 amino acids in length and six highly conserved cysteine residues (Webster, 1991; Yang et al., 1996; Marco et al., 2000). Based on similarities in sequences, MIH is considered to be part of a larger peptide hormone family that includes the crustacean hyperglycemic hormone (CHH), vitellogenesis-inhibiting hormone (VIH), mandibular organ-inhibiting hormone (MOIH), and other peptides from other organisms (Wainwright et al., 1996; Soyez, 1997; Chen et al., 2005). Peptides from the CHH family have overlapping pleiotropic activities. Some of the members of this family are produced in the X-organs of the eyestalk but many of them are produced and released from multiple sites throughout the Cancer nervous system where they may be released into hemolymph in response to a variety of clues and thus may serve as extra-eyestalk sources of Y-organ controlling hormones (Hsu et al., 2006). In the crab, Charybdis feriatus, levels of MIH mRNA in the eyestalk do not change during the molt cycle suggesting that secretion of MIH may be post-transcriptionally regulated Chan et al., 1998. This is supported by the fact that immunopositive MIH in the sinus gland correlated negatively with circulating levels of ecdysteroids (Lee et al., 1998; Nakatsuji et al., 2000). These observations suggest that there is a constant MIH production in and release from the X-organ/sinus gland during intermolt but intermittent release with concomitant storage in the sinus gland during PE. This could account for the constant level of inhibition of the Y-organs during the molt cycle and the episodic release of ecdysteroids during PE. Crustacean MIH cDNAs have been cloned, sequenced and expressed from the shrimps, Panaeus vannamei, Penaeus japonicus, Metapenaeus ensis, Marsupenaeus japonicus (Sun, 1994; Ohira et al., 1997; Gu and Chan, 1998; Okumura et al., 2005),

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the prawn, Macrobrachium rosenbergii (Yang and Rao, 2001), and the crabs, Cancer magister, Callinectes sapidus, Gecarcinus lateralis (Klein et al., 1993; Umphrey et al., 1998; Watson et al., 2001; Chen et al., 2007; Lee et al., 2007a). The presence of MIH in the circulation in vivo and its ability to lower circulating ecdysteroids in vitro have been determined (Lee et al., 1998; Nakatsuji et al., 2000; Chung and Webster, 2005). Circulating levels of MIH in vivo, however, are lowest in the hemolymph during intermolt in the crab, C. maenas (Chung and Webster, 2005) when levels of circulating ecdysteroids are lowest. MIH levels in hemolymph of P. clarkii and C. maenas are highest during late proecdysis, when circulating levels of ecdysteroid are also highest (Nakatsuji and Sonobe, 2004; Chung and Webster, 2005). Levels of MIH in the hemolymph of these crustaceans are, therefore, inconsistent with MIH as the sole inhibitor of Y-organ activity. These data also argue for a more complex control system of the crustacean Y-organ than a simple on/off system. An MIH receptor protein has been isolated from Y-organs in the crab C. sapidus (Lee et al., 1995). This receptor is a membrane-bound receptor that regulates intracellular cGMP (and possibly cAMP) levels in the Y-organ tissue (Nakatsuji et al., 2006). A recombinant MIH was able to maximally suppress ecdysteroid production by only 65% (Nakatsuji et al., 2006). MIH may function to suppress Y-organ activity during specific molt stages and not exert any effect during other stages. There is some evidence that the receptor for MIH may be down-regulated during most of proecdysis in the crayfish, P. clarkii (Nakatsuji and Sonobe, 2004). The exact function of MIH in controlling the production of circulating ecdysteroids awaits further clarification.

3.3.2

Methyl Farnesoate

Methyl farnesoate (MF) is a sesquiterpenoid – an unepoxidated form of insect juvenile hormone (JHIII). It has been implicated in a number of hormonal activities in crustaceans (see Laufer and Biggers, 2001; Borst et al., 2001). MF is secreted by the mandibular organs (MOs) of crustaceans (Borst et al., 1987; Tobe et al., 1989; Nagaraju et al., 2005, 2006). The MOs – which are analogous to the insect copora allata – are located in the anterior body cavity of crustaceans in very close proximity of the mandibles (Byard et al., 1975) and at one time were often confused with the Y-organs (Sochasky et al., 1972). In the crab, Scylla serrata, the MOs secrete farnesoic acid (FA) in much greater amounts than MF (Tobe et al., 1989). Moreover, the enzyme that converts FA to MF (O-methyl transferase) is found in many peripheral crustacean tissues (Kuballa et al., 2007). MF has been implicated in crustacean reproduction control and may enhance ovarian maturation (Laufer et al., 1998; Jo et al., 1999). MF has also been shown to stimulate ecdysteroid synthesis in crustacean Y-organs (Borst et al., 1987; Tamone and Chang, 1993). Thus, MF may serve as another extra-eyestalk source of Y-organ control. There is also some evidence that

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methyl-farnesoate (MF) may have anti-ecdysteroid actions (Yu et al., 2002; Mu and Leblanc, 2004; Tuberty and McKenney, 2005). Exogenous MF administration results in the formation of larval intermediates and retards larval development in freshwater prawns (Abdu et al., 1998). Due to its chemical similarities to molecules known to function as ligands for nuclear receptors, it is possible that MF could interact with a crustacean nuclear receptor in the same way as it does with an insect nuclear receptor (Jones et al., 2006). It has been suggested that the anti-ecdysteroid actions of MF may be effected through its binding to a nuclear receptor and that by doing so in some way inactivates the functional ecdysteroid receptor (Mu and LeBlanc, 2004). MF binds with low affinity to UpRXR, the diner partner of the ecdysteroid receptor in the crab, U. Pugilator, and may modulate the action of the functional ecdysteroid receptor in this crab (Hopkins et al., 2008).

3.4

In Vivo Effects of Ecdysteroids in Crustaceans

Ecdysteroids in crustaceans have been reported to control many physiological processes. Most of the processes under the sway of ecdysteroids are related to the molt cycle. There is some evidence that ecdysteroids may play a role in reproduction (Subramoniam, 2000) but the effects of ecdysteroids on reproduction in male and female crustaceans remains contradictory and poorly documented (Lafont and Mathieu, 2007).

3.4.1

Growth in Crustaceans

To grow, crustaceans must first shed (molt) the old exoskeleton, then synthesize, expand and harden a new one. This is followed by growth in muscle and organ mass (Skinner, 1985). The crustacean molting cycle is divided into five stages (Drach, 1939) on the basis of changes in cuticle morphology. Subsequently other parameters (such as rate of limb regeneration and levels of circulating ecdysteroids) have been added to further refine Drach’s stages (Hopkins, 1989a, 1992). The old exoskeleton is shed at Drach’s stage E, ecdysis (Fig. 3.1a). During stages A and B, metecdysis or postmolt, the new exoskeleton expands and hardens. During stages A/B, levels of total circulating ecdysteroids in the crab U. pugilator are low ( E (1.6 × 10−6M). These data suggest that the binding kinetics of UpEcR to ligand and to DNA differ significantly depending upon the UpRXR isoform with which it is paired. Whereas the insect EcR/USP heterodimer has been shown to bind ecdysteroids (see Riddiford et al., 2003), it is presently unclear whether the insect EcR is a

Fig. 3.4 Comparative binding affinities of Uca pugilator ecdysteroid receptor (UpEcR) to two isoforms of the retinoid-X-receptor (UpRXR). Surface Plasmon binding assay used. Sf9 expressed UpEcR (100nM) interacted with verious concentrations of UpRXR isoforms – larger and smaller. Response is relative amount of binding compared to control flow cells (See Color Plates)

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permissive or non-permissive binding partner for insect USP ligand binding. The LBD of the lepidopteran and dipteran USP has diverged significantly from vertebrate RXRs (which bind 9-cis retinoic acid and fatty acids). This together with the crystal structure studies has led to the speculation that USP has no cognate ligand or a different ligand-binding specificity (Kapitskaya et al., 1996; Guo et al., 1998; Chung et al., 1998b; Hayward et al., 1999). Ligand-binding and transcriptional activation studies have tested a variety of ecdysteroid, retinoid and juvenile hormone (JH) analogues for USP binding activity with negative results (Oro et al., 1990; Yao et al., 1993; Harmon et al., 1995). Low affinity binding of Drosophila USP to JH III, however, and USP-mediated transactivation in response to JHIII, has been reported (Jones and Sharp, 1997; Jones et al., 2001; Xu et al., 2002), as has JHIII mediated repression in transactivation studies (Maki et al., 2004). Methyl farnesoate (a product of the Drosophila ring gland – Richards et al., 1989) may also be a ligand for dmUSP (Jones et al., 2006). There is no question that JH activity is involved in the modulation of the ecdysteroid response (see Dubrovsky, 2005; Riddiford et al., 2003), but the mechanism of how crosstalk is modulated is still unclear. The anti-ecdysteroid effects of juvenoids in Daphnia magna (which has an EcR partner that is more like insect USP than crustacean RXR) were strongly synergistic indicating that they modulate ecdysteroid signaling by a mechanism that may sequester the EcR binding partner (RXR/ USP) into other partnerships that prevents heterodimerization with EcR and thus leads to reduced EcR-mediated activity (Mu and LeBlanc, 2004). One of the best-studied ligands involved in the control of vertebrate limb morphogenesis is retinoic acid (RA; Oro et al., 1992; Means and Gudas, 1995; Brockes, 1997). RA derivatives function in many vertebrate tissues to induce differentiation by controlling the production of morphogenic signals and exogenous RA has profound, disruptive effects on the regeneration of vertebrate limbs (see Brockes, 1997; Maden, 2000 for reviews). All-trans retinoic acid (atRA) disrupts the normal regeneration of limbs in U. pugilator (Hopkins and Durica, 1995). There are two episodes of cuticle secretion observed during early limb regeneration. The first episode is unaffected by the presence of RA but the second episode is disturbed, leading to excessive cell proliferation but no subsequent differentiation (Hopkins and Durica, 1995). The effects of retinoids on limb bud differentiation may be similar to the anti-ecdysteroid effects of juvenoids in Daphnia (Mu and LeBlanc, 2004) in that retinoids may promote UpRXR homodimer (or other heterodimer) formation that reduces or eliminates UpEcR/UpRXR formation and thus normal limb bud differentiation (Hopkins and Durica, 1995). Retinoids may be endogenous components of the regenerating limb blastema. There is evidence of endogenous retinoid production in blastema tissue and that retinoids can synergistically affect PA binding by UpEcR (Hopkins et al., 2008). An aldehyde-hydrogenase enzyme necessary for endogenous retinoid synthesis is present during early blastema development in U. pugilator (Hopkins, 2001) and the U. pugilator EST library indicates the presence of cellular retinoic acid binding protein and retinaldehyde-dehydrogenase – both of which have active roles in retinoid synthesis and signaling.

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Tissue Distribution of Receptors

Probes derived from these clones have identified putative ecdysteroid target tissues throughout the molt cycle in both U. pugilator and M. japonicus (Chung et al., 1998b; Hopkins, 2001; Asazuma et al., 2007). Virtually all tissues examined express EcR and RXR receptors but in variable patterns that sometimes do and sometimes do not correlate with circulating levels of ecdysteroids. Expression of the UpEcR and UpRXR genes monitored with Northern blots, ribonuclease protection assays (RPA), quantitative reverse transcription PCR (Q-RTPCR) and immunocytochemistry show that UpEcR and UpRXR are widely expressed in a large number of non-regenerating crab somatic tissues, in ovarian tissue, and in regenerating limb buds (Durica and Hopkins, 1996; Chung et al., 1998a, b; Durica et al., 2002; Wu et al., 2004). The pattern of expression of UpEcR mRNA is greatest in regenerating limb buds when 20E is present in low titers and is the predominant ecdysteroid in the hemolymph. UpEcR mRNA in the large cheliped is upregulated during late PE when total ecdysteroids in the hemolymph are reaching their maximum peak and PA is the predominant ecdysteroid (Chung et al., 1998b). These differences may, in turn, lead to implementation of different genetic programs in crustacean tissues throughout the molt cycle. UpRXR variants and UpEcR are present throughout the two phases of limb regeneration but clearly each variant is subserving different transactional activities.

3.6

Ecdysteroid Responsive Genes in Crustaceans

Arthropod models were one of the first biological systems used to investigate the molecular mechanisms of steroid hormone action. Although experiments demonstrating ecdysteroid regulation of gene transcription were historically at the frontiers of classical endocrinology, we still know comparatively little about how circulating ecdysteroids are capable of programming differential gene expression and ultimately distinct physiological responses. The function of ecdysteroids in the regulation of gene transcription during insect metamorphosis has been the object of intense study. Variations in 20-hydroxyecdysone (20E) titers have long been associated with molting and metamorphosis in insects (see Karlson, 1996), and a hierarchy of transcription factor gene expression mediated by ecdysteroid exposure has been characterized in Drosophila and Manduca (Talbot et al., 1993 see Riddiford et al., 2003). The Drosophila E75 gene was one of the first loci suspected to be under ecdysteroid control (Segraves and Hogness, 1990). Studies in several insect systems indicate that both EcR and E75 are primary ecdysteroid-responsive genes (Karim and Thummel, 1992; Wang et al., 2002; Riddiford et al., 2003; Siaussat et al., 2004). During early blastema development (8 days after autotomy – A + 8), short intervals of 20E in vitro exposure was ineffective in increasing EcR gene transcript

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levels in limb bud explants (Durica et al., 2006). Exposure of more developed limb buds (A + 12) to 0.5–1 µM 20E for periods greater than 2 h led to decreases in EcR transcript levels. Bud explants taken from animals in late proecdysial stages (when circulating levels of PA are highest) appear refractory to 20E exposure; neither Q-RTPCR nor RPA indicate significant changes in EcR transcript abundance. The limb buds used in these induction studies were removed from animals with very low levels of total circulating ecdysteroids. During this time 20E is present in the circulation at about 20 pM concentration and injection of high levels of 20E in vivo inhibited blastema development (Hopkins et al., 1979). In the crab G. lateralis there are strong correlations between elevated circulating ecdysteroids and GIRXR, E75 and intracellular cGMP expression suggesting that these loci may be ecdysteroid-responsive genes (Kim et al., 2005; Lee et al., 2007b). The appearance of a growth arrest-specific protein (GAS7) from the X-organs/Sinus gland complex of the white shrimp Fenneropenaeus indicus and phosphorylation events for the signaling protein ERK2 shows molting stage specific correlations (Devaraj and Natarajan, 2006) that suggest that these gene/proteins may be controlled by ecdysteroids. Crustacean tissue-specific EST data bases are currently available on line. Data from U. pugilator, C. sapidus, C. maenas, and H americanus can be found in EST libraries that are BLAST and keyword searchable; the data set is also available through GenBank (Durica et al., 2006). The first characterized library from U. pugilator was made from mRNA isolated 4 days post-autotomy (A + 4), when the first sign of morphological differentiation, cuticle secretion, is observed. Analysis of 2,030 individual cDNA clones led to assignment of 473 contigs and 417 singlets, for a total of 890 different transcripts. Of these, approximately 63% showed no BLAST homology on database searching, while approximately 11% could be assigned to a known ortholog in the COG (Clusters of Orthologous Groups) database. The U. pugilator EST database screened against the Gene Ontology (GO) database. tBLASTx searches (Blast2Go, Conesa et al., 2005) produced a significantly greater number of biological functional assignments than for the COG database at the same expect criterion (10−5); approximately 40% of the domains identified among the assembled sequences could be assigned a GO annotation (see website for assignments). Nevertheless, a majority of ESTs from this library are as yet unassignable as to molecular function, cellular process or cellular component. Sequence similarity searches between other crustacean EST databases produced hits between 13–30% of the U. pugilator query sequences (Pymood program; see Durica et al., 2006 for example of program output). Comparisons indicate that approximately 65% of the U. pugilator sequences are not found in any of the other three crustacean databases. There are also clear differences between the libraries in terms of BLAST hits to the U. pugilator database. The H. americanus database, which represents an olfactory-organ-specific library, contains hits to only 14% of U. pugilator ESTs, while the C. sapidus database, derived from hypodermal tissue, contains hits to 30% of U. pugilator ESTs. Although this comparison is influenced

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by the number of ESTs present in the target databases, it also undoubtedly reflects similarities in gene-expression profiles for subsets of genes not related to “housekeeping” functions. The U. pugilator early blastema library represented a tissue that is undergoing both proliferation and remodeling. An EST screening from a library constructed from a later proecdysial stage of limb bud regeneration, where the individual limb segments are already formed, and are undergoing an increase in mass due to muscle tissue synthesis (Hopkins, 1989b). From this library, 1,337 sequence reads resulted in 239 contigs and 399 singlets for a total of 638 different transcripts. Pymood comparisons indicate that the early and late U. pugilator regenerating limb bud databases show a greater degree of similarity (33% BLAST hits) than to the other crustacean databases, although tBlastX analysis indicates that many of the shared sequences from the U. pugilator regenerating limb databases are still undefined as to biological function. The most abundant sequences (i.e. greatest number of times a specific sequence is detected among the ESTs) in the two different U. pugilator regenerating limb bud mRNA populations show profound differences. Constitutive ‘house-keeping’ genes were represented in both populations, the 25 most abundant sequences in the two libraries are distinctly different. In early blastemal library, there is a group of proteins (TNFRSF1a modulator protein; macropain; cyclophorin; TGFβ inducible nuclear protein) that has been linked to proliferative or apoptotic decisions tied to TNF signaling pathways (Gu et al., 1998; Li et al., 2004; Bouwmeester et al., 2004). For the proecdysial library, proteins involved in muscle biogenesis and arthrodial cuticle deposition (i.e. differentiation, not determination) are among the most abundant (muscle LIM protein; muscle actin; myosin three light chain; cuticle protein LCP18; arthrodial cuticle protein AMP9.3). This suggests that, even with limited sample size screening, information on gene networks and genes involved in important biological processes relevant to a particular physiological state can be obtained from these studies.

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Laufer, H., Biggers, W.J., and Ahl, J.S., 1998. Stimulation of ovarian maturation in the crayfish Procambarus clarkii by methyl farnesoate. Gen. Comp. Endocrinol. 111:113–118. Lee, K., Elton, T., Bej, A., Watts, S., and Watson, D. 1995. Molecular cloning of a cDNA encoding putative molt-inhibiting hormone from the blue crab, Callinectes sapidus. Biochem. Biophys. Res. Commun. 209:1126–1131. Lee, K., Watson, D., and Roer, R. 1998. Molt-inhibiting hormone mRNA levels and ecdysteroid titer during a molt cycle of the blue crab, Callinectes sapidus. Biochem. Biophys. Res. Commun. 249:624–627. Lee, K., Kim, H.-W., Gomez, A., Chang, E., Covi, J., and Mykles, D. 2007a. Molt-inhibiting hormone from the tropical land crab, Gecarcinus lateralis: cloning, tissue expression, and expression of biologically active recombinant peptide in yeast. Gen. Comp. Endocrinol. 150:505–513. Lee, S., Bader, B., Chang, E., and Mykles, D. 2007b. Effects of elevated ecdysteroid on tissue expression of three guanylyl cyclases in the tropical land crab Gecarcinus lateralis; possible roles of neuropeptide signaling in the molting gland. J. Exp. Biol. 210:3245–3254. Li, L., Thomas, R.M., Suzuki, H., De Brabander, J.K., Wang, X., and Harran, P.G. 2004. A small molecule Smac mimic potentiates TRAIL- and TNFα-Mediated cell death. Science 305: 1471–1474 Maden, M. 2000. The role of retinoic acid in embryonic and post-embryonic development. Proc. Nutr. Soc. 59:65–73. Maestro, O., Cruz, J., Pascual, N., Martin, D., and Belles, X. 2005. Differential expression of two RXR/ultraspiracle isoforms during the life cycle of the hemimetabolous insect Blattella germanica (Dictyoptera, Blattellidae). Mol. Cell. Endocrinol. 239:27–37. Maki, A., Sawatsubashi, S., Ito, S., Shirode, Y., Suzuki, E., Zhao, Y., Yamagata, K., Kouzmenko, A., Takeyama, K., and Kato, S. 2004. Juvenile hormones antagonize ecdysone actions through co-repressor recruitment to EcR/USP heterodimers. Biochem. Biophys. Res. Commun. 320:262–267. Marco, H., Stoeva, S., Voelter, W., and Gade, G. 2000. Characterization and sequence elucidation of a novel peptide with molt-inhibiting activity from the South African spiny lobster, Jasus lalandii. Peptides 21:1313–1321. Martin-Creuzburg, D., Westerlund, S., and Hoffmann, K. 2007. Ecdysteroid levels in Daphnia magna during a molt cycle: determination by radioimmunassay (RIA) and liquid chromatography-mass spectrometry (LC-MS). Gen. Comp. Endocrinol. 151:66–71. Mattson, M.P. and Spaziani, E. 1986. Regulation of Y-organ ecdysteroidogenesis by molt-inhibiting hormone in crabs: involvement of cyclic AMP-mediated protein synthesis. Gen. Comp. Endocrinol. Sept; 63:414–423. McCarthy, J.F. 1979. Ponasterone A: a new ecdysteroid from the embryos and serum of brachyuran crustaceans. Steroid 34:799–806. McCarthy, J.F. and Skinner, D.M. 1977. Interruption of proecdysis by autotomy of partially regenerated limbs in the land crab, Gecarcinus lateralis. Dev. Biol. 61:299–310. McVean, A. 1984. Autotomy. In D.E. Bliss (ed.), The Biology of Crustacea, pp. 107–132. Academic, New York. Means, A.L. and Gudas, L.J. 1995. The roles of retinoids in vertebrate development. Annu. Rev. Biochem. 64:201–233. Meister, M.F., Brandtner, H.M., Koolman, J., and Hoffmann, J.A. 1987. Conversion of a radiolabelled putative ecdysone precursor, 2,22,25-trideoxyecdysone (5B-ketodiol) in larvae and pupae of Calliphora vicina. Int. J. Invertebr. Reprod. Dev. 12:13–28. Mu, X. and LeBlanc, G. 2004. Cross communication between signaling pathways: juvenoid hormones modulate ecdysteroid activity in a crustacean. J. Exp. Zool. 301A:793–801. Nagaraju, G., Prasad, G., and Reddy, P. 2005. Isolation and characterization of mandibular organ inhibiting hormone from the eyestalks of freshwater crab, Oziotelphusa senex senex. Int. J. Appl. Sci. Eng. 3:61–68. Nagaraju, G., Reddy, P.R, and Reddy, P.S 2006. In vitro methyl farnesoate secretion by mandibular organs isolated from different molt and reproductive stages of the crab Oziotelphusa senex senex. Fish. Sci. 72:410–414.

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Nakatsuji, T. and Sonobe, H. 2004. Regulation of ecdysteroid secretion from the Y-organ by moltinhibiting hormone in the American crayfish, Procambarus clarkii. Gen. Comp. Endocrinol. 135:358–364. Nakatsuji, T., Keino, H., Tamura, K., Yoshimura, S., Kawakami, T., Aimoto, S., and Sonobe, H. 2000. Changes in the amounts of the molt-inhibiting hormone in sinus glands during the molt cycle of the American crayfish, Procambarus clarkii. Zool. Sci. 17:1129–1136. Nakatsuji, T., Han, D.-W., Jablonsky, M., Harville, S., Muccio, D., Watson, D. 2006. Expression of crustacean (Callinectes sapidus) molt-inhibiting hormone in Escherichia coli: characterization of the recombinant peptide and assessment of its effects on cellular signaling pathways in Y-organs. Mol. Cell. Endocrinol. 253:96–104. Naya, Y., Kishida, K., Sugiyama, M., Murata, M., Miki, W., Ohnishi, M., and Nakanishi, K. 1988. Endogenous inhibitor of ecdysone synthesis in crabs. Experientia 44:50–52. Ohira, T., Watanabe, T., Nagasawa, H., and Aida, K. 1997. Molecular cloning of a molt-inhibiting hormone cDNA from the kuruma prawn Penaeus japonicus. Zool. Sci. 14:785–789. Okumura, T., Kamba, M., Sonobe, H., and Aida, K. 2003. In vitro secretion of ecdysteroids by Y-organ during molt cycle and evidence for secretion of 3-dehydroecdysone in the giant freshwater prawn, Macrobrackium rosenbergii (Crustacea: Decapoda: Caridea). Invertebr. Repro. Dev. 44:1–8.Okumura, T., Ohira, T., Katayama, H., and Nagasawa, H. 2005. In vivo effects of a recombinant molt-inhibiting hormone on molt interval and hemolymph ecdysteroid levels in the kuruma prawn, Marsupenaeus japonicus. Zool. Sci. 22:317–320. Oro, A.E., McKeown, M., and Evans, R.M., 1990. Relationship between the product of the Drosophila ultraspiracle locus and the vertebrate retinoid X receptor. Nature 347:298–301. Oro, A.E., Mckeown, M., and Evans, R.M. 1992. The Drosophila retinoid X receptor homolog ultraspiracle functions in both female reproduction and eye morphogenesis. Development 115:449–462. Rao, K.R. 1965. Isolation and partial characterization of the moult-inhibiting hormone of the crustacean eyestalk. Experientia 21:593–596. Rees, H.H. 1985. Biosynthesis of ecdysone. In G.A. Kerkut and L.I. Gilbert (eds.), Comprehensive insect physiology, biochemistry and pharmacology, vol. 7, pp. 249–293. Pergamon Press, Oxford. Renaud, J.P. and Moras, D. 2000. Structural studies on nuclear receptors. Cell. Mol. Life Sci. 57:1748–1769. Riddiford, L.M., Cherbas, P., and Truman, J.W., 2001. Ecdysone receptors and their biological actions. Vitam. Horm. 60:1–73. Richards, D., Applebaum, S., and Gilbert, L. 1989. Developmental regulation of juvenile biosynthesis by the ring gland of Drosophila. J. Comp. Physiol. B 159:383–387. Riddiford, L.M., Hiruma, K., Zhou, X., and Nelson, C.A. 2003. Insights into the molecular basis of the hormonal control of molting and metamorphosis from Manduca sexta and Drosophila melanogaster. Insect Biochem. Mol. Biol. 12:1327–1338. Segraves, W.A. and Hogness, D.S. 1990. The E75 ecdysone 75B early puff in Drosophila codes two new members of the steroid receptor superfamily. Gene Dev. 4:204–219. Siaussat, D., Bozzolan, F., Queguiner, I., Porcheron, P., and Debernard, S. 2004. Effects of juvenile hormone on 20-hydroxyecdysone-inducible EcR, HR3, E75 gene expression in imaginal wing cells of Plodia interpunctella Lepidoptera. Eur. J. Biochem. 14:3017–3027. Skinner, D.M. 1985. Molting and regeneration. In D.E. Bliss (ed.), The Biology of Crustacea, pp. 43–146. Academic, New York. Smith, W.A. and Sedlmeier, D. 1990. Neurohormonal control of ecdysone production: comparison of insects and crustaceans. Invertebr. Reprod. Dev. 18:77–90. Sochasky, J.B., Aiken, D.E., and Watson, N.H.F. 1972. Y-organ, molting gland, and mandibular organ. A problem in decapod crustacean. Can. J. Zool. 50: 993–997. Sonobe, H., Kamba, M., Ohta, K., Ikeda, M., and Naya, Y. 1991. In vitro secretion of ecdysteroids by Y-organs of the crayfish, Procambarus clarkii. Cell. Mol. Life Sci. 47:948–952. Soumoff, C. and O’Connor, J.D. 1982. Repression of Y-organ secretory activity by molt-inhibiting hormone in the crab Pachygrapsus crassipes. Gen. Comp. Endocrinol. 48:432–439.

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Soyez, D. 1997. Occurrence and diversity of neuropeptides from the crustacean hyperglycemic hormone family in arthropods. Ann. N.Y. Acad. Sci. 814:319–323. Spaziani, E., Rees, H.H., Wang, W.L., and Watson, R.D. 1989. Evidence that Y-organs of the crab Cancer antennarius secrete 3-dehydroecdysone. Mol. Cell. Endocrinol. 66:17–25. Subramoniam, T., 2000. Crustacean ecdysteroids in reproduction and embryogenesis. Comp. Biochem. Physiol. 125:135–156. Sun, P.S. 1994. Molecular cloning and sequence analysis of a cDNA encoding a molt-inhibiting hormone-like neuropeptide from the white shrimp Penaeus vannamei. Mol. Mar. Biol. Biotechnol. 3: 1–6. Swevers, L., Cherbas, L., Cherbas, P., and Iatrou, K. 1996. Bombyx EcR (BmEcR) and Bombyx USP (BmCF1) combine to form a functional ecdysone receptor. Insect Biochem. Mol. Biol. 26:217–221. Talbot, W.S., Swyryd, E.A., and Hogness, D.S., 1993. Drosophila tissues with different metamorphic responses to ecdysone express different ecdysone receptor isoforms. Cell 73:1323–1337. Tamone, S.L. and Chang, E.S. 1993. Methyl fanesoate stimulate ecdysteroid secretion from Y-organs in vitro. Gen. Comp. Endocrinol. 89:425–432. Thomas, H.E., Stunnenberg, H.G., and Stewart, A.F. 1993. Heterodimerization of the Drosophila ecdysone receptor with retinoid X receptor and ultraspiracle. Nature 362: 471–475. Tobe, S.S., Young, D.A., Khoo, H.W. 1989. Production of methyl farnesoate by the mandibular organs of the mud crab, Scylla serrata: validation of a radiochemical assay. Gen. Comp. Endocrinol. 73(3):342–353. Tuberty, S.R., and Mckenney, Jr., C.L. 2005. Ecdysteroid responses of estuarine crustaceans exposed through complete larval development to juvenile hormone agonist insecticides. Int. Comp. Biol. 45: 106–117. Umphrey, H., Lee, K., Watson, D., and Spaziani, E. 1998. Molecular cloning of a cDNA encoding molt-inhibiting hormone of the crab, Cancer magister. Mol. Cell. Endocrinol. 136:145–149. Wainwright, G., Webster, S., Wilkinson, M., Chung, J., and Rees, H. 1996. Structure and significance of mandibular organ-inhibiting hormone in the crab, Cancer pagurus. Involvement in a multihormonal regulation of growth and reproduction. J. Biol. Chem. 271:12749–12754. Wang, Y., Hayes, T., Holman, G., Chavez, A. and Keeley, L. 2000. Primary structure of CHH/MIH/ GIH-like peptides in sinus gland extracts from Penaeus vannamei. Peptides 21:477–484. Wang, S-F., Li, C., Sun, G., Zhu, J., and Raikhel, A.S. 2002. Differential expression and regulation by 20-hydroxyecdysone of mosquito ecdysteroid receptor isoforms A and B. Mol. Cell. Endocrinology 196: 29–42. Warren, J.T., Petryk, A., Marques, G., Parvy, J.-P., Shinoda, T., Itoyama, K., Kobayashi, J., Jarcho, M., Li, Y., O’Connor, M., Dauphin-Villemant, C., and Gilbert, L. 2004. Phantom encodes the 25-hydroxylase of Drosophila melaongaster and Bombyx mori: a P450 enzyme critical in ecdysone biosynthesis. Insect Biochem. Mol. Biol. 34:991–1010. Watson, D. and Spaziani, E. 1985. Biosynthesis of ecdysteroids from cholesterol by crab Y-organs and eyestalk suppression of cholesterol uptake and secretory activity in vitro. Gen. Comp. Endocrinol. 59:140–148. Watson, D., Lee, K., Qui, S., Luo, M., Unphrey, H., Roer, R., and Spaziani, E. 2001. Molecular cloning, expression, and tissue distribution of crustacean molt-inhibiting hormone. Am. Zool. 41L:407–417. Webster, S.G. and Keller, R. 1986. Purification, characterization and amino acid composition of the putative moult-inhibiting hormone (MIH) of Carcinus maenas (Crustacea, Decapoda). J. Comp. Physiol. B 156:617–624/Mol. Cell. Endocrinol. 66:17–25. Webster, S. 1991. Amino acid sequence of putative moult-inhibitng hormone from the crab Carinus maenas. Proc. Biol. Sci. 244:247–252. Wu, X., Hopkins, P.M., Palli, S.R., and Durica, D.S. 2004. Crustacean retinoid-X receptor isoforms: distinctive DNA binding and receptor-receptor interaction with a cognate ecdysteroid receptor. Mol. Cell. Endocrinol. 218:21–38.

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Chapter 4

Synthetic Ecdysteroidal Compounds Josep Coll Toledano

Abstract This review compiles ecdysteroids (usually but not always) less readily available and ecdysteroid derivatives synthesized, either for structure-biological activity studies or as part of their structure elucidation processes, as well as synthetic intermediates showing the features of “true” ecdysteroids. Keywords Ecdysteroid • chemical • synthesis Abbreviations Recommended standardized abbreviations for common ecdysteroids [1,2] have been adapted, and one single letter is used for most substituents. Schemes 4.1 and 4.2 show structures of reference compounds and examples of abbreviations use. Reference compounds BNC: Bis-nor-cholane ecdysteroid; C: Cholane ecdysteroid; CCL: Cholane-24, 22-carbolactone ecdysteroid; E: Ecdysone; MaA: Makisterone A (campestane steroid); MaC: Makisterone C (stigmastane steroid); MNC: Mono-nor-cholane ecdysteroid; Pan: Panuosterone, 24-epi-25-deoxy-24-hydroxyMaA (ergostane steroid); PoA: Ponasterone A, 25-deoxy-20-hydroxyE; Pos: Poststerone (pregnane steroid); Rub: Rubrosterone (androstane steroid); Tax: Taxisterone, 22-deoxy-20-hydroxyE. Hydroxyl groups Additional: locant as prefix, using comma if several. Configuration 1β, 9α, 11α, 24α implied (configuration not mentioned). Opposite configuration indicated as epimer: (e.g∴1' = 1α, etc). Lack of: d (deoxy) after locant(s) as suffix, using comma if several. Carbonyl groups D (dehydro) after locant as suffix in parent-structure hydroxyl-bearing positions. Other substituted positions: k (ketone) or al (aldehyde). Carbonyl reduction (locant implied in the name): locant as suffix with H. J. Coll Toledano() Departament de Química Orgànica Biològica, Institut d’Investigacions Químiques i Ambientals de Barcelona “Josep Pascual Vila”, Consejo Superior de Investigaciones Científicas, c. J. Girona 18. 08034-Barcelona (España) e-mail: [email protected]

G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Acetals First locant as suffix (2 for 2,3; 20 for 20,22) with a: acetonide; et: ethylidene; etme: butan-2-ylidene; fu: furfurylidene; f2c: furan2-carboxylate; hbu: 4-hydroxybutylidene; hbz: 4-hydroxybenzylidene; hbz:4hydroxybenzylidene; meop: 4-oxopentan-2-ylidene; PB: phenylboronate Acyl conjugates Locant as suffix with A: acetate; apa: p-azidophenylacetate; B: benzoate; cin: cinnamate; ibt: p-iodobenzoylthioisocyanate; mal: malonate; malOBz: benzyloxymalonate; Ms: mesylate; N: nitrate; P: phosphate; p2c: pyrrole-2-carboxylote; S: sulphate; Tf: trifluororomethanesulphonate, tfa: trifluoroacetate; t2c: thiophene-2-carboxylate, Other acyl esters: (chain length:number of double bonds if any) [examples: (18) = stearyl; (18:1) = oleyl; (18:2) = linoleyl; (18:3) = linolenyl]. Alkyl conjugation/substitution EE:1-ethoxyethyl; G: glucoside; hpo: hydroperoxide; hpp: p-hydroxyphenylpropyl; SEM: (2-trimethylsilyl-ethoxy)methyl. As commonly used: Et, Me, TBDMS, THP, TMS, F, CL, Br, I. Multiple substitution: number*substituent abbreviation (5*TMS = PentaTMS; 4*A = tetraacetate, etc). Oxygen bridges (locant-O-locant). CL: Carbolactone. Double/triple bond Additional: locant(s) as suffix and en/in as required. Double bond reduction: 7H (= 7,8-dihydro). Epimers Primed locant as suffix in parenthesis (trans A/B ring fusion = 5a). Suffix order Substitution follows the parent reference abbreviation as: d,D,acetal,acyl,alkyl,H, (x-O-y),en,in,k,al,(epi). In a table, compounds with only stereochemical changes are listed first.

Introductory Remarks All ecdysonists know well the wealth of information contained in the “Ecdybase” [i], and envisage the effort behind it to provide general data on all natural ecdysteroids. In the forward it is written “as presently available, is still considered by the authors as a developing resource, which can be continuously improved and extended with the help of all other ecdysonists”. In its predecessor, “The Ecdysone Handbook” [ii], with 196 compounds listed, “Chemical synthesis” or “Chemical and enzymatic synthesis” were displayed occasionally as a further information, along with two main entries for (first) isolation source (Animals; Plants), in the OCCURRENCE box.

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Scheme 4.1 Parent structures

“Chemical and enzymatic synthesis” was first to appear (3-dehydroecdysone) but it was not immediate to find out the reference providing such information. The next entry showed again the same words without parenthesis but the title of the reference for “First isolation” (from animals) was “Enzymatic and chemical synthesis of a metabolite of the moulting hormone of insects”. “Chemical synthesis” was found next (22,25-dideoxyecdysone, misplaced alphabetically) and the reference heading was exceptionally “First synthesis” (instead of “First isolation” as in all previous entries). Chemical synthesis appeared again (14-deoxyecdysone) wherein “First isolation” quoted a 1988 paper on ecdysone catabolism, and two papers dealt with synthesis (1966 and 1987 under “General” heading). And so on.

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Scheme 4.2 Examples of abbreviations use

The ecdysteroid family increased to a list of 262 entries in the second edition, “still considered as a draft”, but providing all information for an entry very conveniently “at a glance”. Chemical synthesis moved first to appear with calonysterone (a correction of this entry) in this second (and last printed) edition of “The Ecdysone Handbook”. The Ecdybase new design “lost” a couple of features. It does not display directly a whole list of compounds, but a simple “trick” solves the problem: just entering C at formula searching (and you find out 360 listed compounds). On the other hand, chemical (synthesis) is no longer present after Occur(r)ence in plants and Occur(r)ence in animals. However, a search of the term “chemical” under organism species returns six headings (using “synthesis” you get only five and the sixth with “syntheses”) and 52 listed compounds. Remarkably, two of the entries do not show any matches. The goal of the present review is to complement the literature reports on “true” ecdysteroids from animal or plant origin, in the same mood as stated in the

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Ecdybase as far as future improvement and extension. No matter how simple chemistry is involved, compounds prepared by synthetic procedures displaying a “true” ecdysteroid structure are compiled. Such information is sometimes not apparent in a paper title but (hopefully) may appear in the abstract or remain hidden in the text and/or experimental.

4.1

Introduction

Ecdysteroid is a generic term introduced to apply to all compounds structurally related to 2β,3β,14α,22R,25-pentahydroxy-5β-cholest-7-en-6-one, simply and better known as ecdysone and E its recommended standard abbreviation [1–3]. Such a definition should imply a few features present in the parent molecule steroidal skeleton. However, since the limits for such “structural relationship” were rather arbitrary, a compound was more precisely distinguished as a “true” ecdysteroid when displaying a cis-fused A/B ring junction and the 14α-hydroxy-7-ene-6-one system, whereas other compounds showing just a partial share of those features (lack of the 7-ene, or 14α-OH; trans-fused A/B ring junction,…) were considered as ecdysteroid-related substances [4]. Despite 20-hydroxyecdysone (20E), the second isolated member of this group, was rapidly established as the major moulting hormone of all arthropods, ecdysone became the reference compound and the major parent name in semi-systematic ecdysteroid nomenclature. A few other trivial names have been used occasionally also as parent names and specific three-letter abbreviations were suggested [2]. The terms “zooecdysteroid” and “phytoecdysteroid” were coined to point out the isolation source and since the present review is focused in compounds prepared by synthesis (mainly chemical) displaying the “true” ecdysteroid features, the term “synthecdysteroid” (although cumbersome) seems appropriate. The reactivity of ecdysteroids (dehydration, isomerisation, oxidation, reduction, side-chain cleavage or derivative formation prepared for structural elucidation purposes [such as acetates and/or acetonides], trimethylsilylethers, etc…) as well as detailed descriptions of early synthetic schemes leading to ecdysteroids have been already reviewed comprehensively [5,6]. E and 20E synthesis from common sterols was a major objective in the early period of ecdysteroid research [5]. A number of intermediates prepared while solving the problems of steroidal nucleus construction and control of side chain stereochemistry display “true” ecdysteroid features and therefore should be considered as (synthetic) ecdysteroids. The same is true for derivatives prepared for characterisation purposes. A few have been recognised as such, later on, after isolation as natural product but most of them were not included as synthetic ecdysteroids in [6], a thorough revision for that period being needed. Only “a few chemical reactions that….do not require an experienced chemist” with no details about chemical synthesis procedures were included in a recent coverage of ecdysteroid chemistry and biochemistry [7]. Here again, chemical synthesis procedures will not be treated in detail but presented in abbreviated flow-chart style. Furthermore, inclusion of structurally related compounds will be restricted to those either closely-related biogenetically or requiring very few synthetic steps (exceptionally more than one). Abbreviations have been selected to point out structural correlation.

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Precedings

The sate-of-the-art in the mid eighties [6] is summarised in Tables 4.1 and 4.2. This “must” reference lists 87 naturally occurring ecdysteroids, wherein “true” ecdysteroids are actually 73 and ecdysteroid-related substances 14. Table 4.1 displays compounds isolated from natural sources also prepared by synthetic procedures. It is worth pointing out that in a few instances synthesis preceded the isolation as a natural product or was reported almost simultaneously. Taxisterone (22-deoxy20-hydroxyecdysone) provides a particular example. It was isolated in 1982 [8] just in time to be appropriately mentioned in the addendum of [6]. However, since the compound had been synthesized earlier, it appears listed as a synthetic ecdysteroid (the synthesis was reported in 1968 [9] but this fact was not recorded in the isolation paper). Now it has been included in Table 4.1.

Table 4.1 Synthesized natural ecdysteroids listed in [6] [(S); zoo (Z); phyto (P)] Name (number) in [6] S Z P 1977 Calonysterone [7] Ecdysone [24] 1966 Kaladasterone [42] 1973 Polypodine B [54] 1977 Polypodine B 2-cinnamate [55] 1972 Ponasterone A [58] 1968 Poststerone [63] 1967 Rubrosterone [67] 1968 Stachysterone C [75] 1971 Taxisterone, 20E22d 1968c 3-Dehydroecdysone [15] 1977 3-Dehydro-20-hydroxyecdysone [16] 1978 2-Deoxyecdysone [18] 1975 2-Deoxy-20-hydroxyecdysone [19] 1982 22-Deoxyecdysone 1969 2,25-Dideoxyecdysone 1975 3-Epi-ecdysone [29] 1970 3-Epi-20-hydroxyecdysone [30] 1978 20-Hydroxyecdysone [32] 1967 20-Hydroxyecdysone 2-acetate [35] 1969f 20-Hydroxyecdysone 2-cinnamate [36] 1972 2,14,22,25-Tetradeoxyecdysone [77] 1973 2,22,25-Trideoxyecdysone [78] 1973 a first isolation in crystalline form b structure based on X-ray analysis c depicted as 20S but reported as 20R d referenced only as synthetic ecdysteroid e partial identification and labelled metabolite f not referenced as synthetic ecdysteroid

1954a, 1965b

1979

1986d 1972 1974e 1977 1968 1972d 1978d 1979 1974 1966 1980

1973 1967 1973 1967 1971 1966 1970 1968 1970 1982d

1970 1970

1966 1971

1978 1978

4

Synthetic Ecdysteroidal Compounds Table 4.2 Synthetic ecdysteroids listed in [6]a–c S INPd E(5a) 1966 5E2,14,22,25d(5a) E(22') 1966 5E2,22,25d E2,3,22,25d 1976 5E2,22,25d(5a) E2,3,22,25d(5a) 1976 5E2,22,25d3A(5a) E2,3,22,25d14TMS(5a) 1976 5E14,22,25d(5a) E2,14,22,25d3A 1978 5E22,25d E2,14,22,25d3A(5a) 1978 5E22,25d(5a) E2,22,25d(5a) 1974 E2,22,25d(5a,3’) 1978 20E2d(3') E2,22,25d3A 1981 20E22,25d E2,22,25d3A(5a) 1970 20E14hpo E2,22,25d3A(5a,14’) 1978 E2,22,25d3A(14’) 1981 PoA14hpo E2,22,25d3Et(Me) 1981 PoA26oic E2,22,25d14hpo 1968 E2,25d 1975 5,11PoA14d (muristerone A)14d E14d 1966 1988Z E22d(5a) 1972 Rub2d E22,25d 1966e 1985Z E22,25d(5a) 1970 E22,25d2,3A(5a) 1970 E22,25d2,3N 1976 E22,25d2,3TMS 1970 E22,25d3Me 1970 E22,25d24REt 1970 E22,25d24RMe 1970 E25d 1972 1986Z E2-hemisuccinate 1975f E22-hemisuccinate 1976 a Only earliest reference quoted b Labeled compounds not included c Except compounds included in Table 4.1 d Isolation as natural product: z zooecdysteroids, P phytoecdysteroids e Quoted as first isolation f Actually 20E derivative (but not characterized)

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S

INP

1978 1971 1978 1978 1976 1971 1972 1969 1972 1980

1989Z

1968 1976 1982

1976

Other synthetic ecdysteroids listed in [6] are now compiled in Table 4.2 (excluding those already mentioned in Table 4.1, labelled analogues prepared for biosynthetic studies, and some intermediates). As already mentioned, reference to “(earlier) synthesis” or “new compounds” should be carefully checked for correct crediting since, as already mentioned, the reported lists may be far from comprehensive.

106

4.3

J.C. Toledano

Synthecdysteroids

Synthetic compounds (reported from 1980 on to overlap the last years collected in [6]) are presented in the following tables along with some convenient additions. Derivatives of the parent ecdysteroid E are listed in Table 4.3. Tables 4.4a–d list those 20E-related, Table 4.5 those based on hydroxylated derivatives of 20E, Table 4.6 those related to ponasterone A or taxisterone, Table 4.7 compounds showing shorter (or no) side-chain (rubrosterone, poststerone, etc.) and Table 4.8 compounds based on C-24 branched (C-28 or C-29) ecdysteroids. Acetonides and acetates, particular examples of acetal and acyl derivatives respectively, have been routinely involved as protecting groups or in structural elucidation (sometimes conveniently combined). It is no surprise to find very early references reporting their preparation but with limited structural information owing to the technological development at the time.

4.3.1

Ecdysone Group

A multistep sequence reported a convenient conversion of diosgenin to CCL2, 3A(5a) [10], and this compound used to prepare labeled E. The intermediate was previously obtained from stigmast-22-en-3,6-dione (in turn prepared from stigmasterol) and this lactone used to synthesize E [11,12]. E(22') was prepared from the epimeric CCL2,3A(5a,22'). diosgenin →→ CCL2,3A(5a) → CCL → E

(4.1)

As shown in Scheme 4.3, E2d and E2,22d were also prepared from ergosterol [13–16]. A small amount of E2,22,25d was formed upon hydrogenation. The sequence was useful to prepare labeled substrates for biosynthetic studies. Cholesta-6,8(10)-dien-3-ol was obtained from cholest-7-en-3-ol, and then converted to E2,22,25d(5a) [17]. E2,22,25d3A(5a) and E2,22,25d3Ms(5a) were also prepared. Similar treatment applied to ergosterol afforded the corresponding 24-methyl-22-en analogues. The chemical synthesis of ecdysone 22-(acyl)esters of long-chain fatty acids from E2a, through the corresponding acetonide intermediates, has been reported [18]. From conveniently protected E, monosulfates (E2S, E22S, E2,22,25A3S, E2,3,22A25S) and one disulfate (E2,22S) were prepared and used as references for HPLC analysis of polar conjugates [19]. The following compounds have been synthesized: E3D (from E) [20]; E3D (+ E3,22D from E), and E3D14d (from E14d) [21]; E2,22,25d3D (diketol) from E2,22,25d3D1en [22]. Similarly E25d and E2,25d were prepared from E and E2d, respectively [23] as shown in Eqs. 4.2 and 4.3:

4

Synthetic Ecdysteroidal Compounds

107

Table 4.3 E derived synthecdysteroids and related natural compounds Ref. INP E(22'-O-25) E(22-O-25)

[24] [24]

E2d E2d3D (silenosterone) E2d3A E2d3A25THP23in(5a) (+22') E2d3A23in(5a) E2d3,22A E2d3,22A24 + 25en E2d22A E2d22A25B(5a), (tomentesterone A) E2d22B E2d22P E2d22en E2d23in (+5a) E2d24 + 25en E2d25B(5a), (tomentesterone B)

[13,14]

[14] [14] [23] [23] 1990 1995 1987 1982 [15] [14] [23] 1996

E2,22d

[15,16]

E2,22,25d(5a) E2,22,25d3D E2,22,25d3D1en E2,22,25d3A22,24en (E + Z) E2,22,25d3A(5a) E2,22,25d3Ms(5a) E2,22,25d22en(E + Z) (24-O-25) E2,22,25d22,24en(E + Z)

[17] [22] [22] [16] [17] [17] [16]

E2,25d E14d3D E25d E25d7H, (cheilantone B)

1970 1979 1986

1978

[16] [13,23] [21] [23]

1986a

1970

E3D [20,21] 1972 E3,22D [21] E22D2a [24] a As a labeled intermediate was prepared in 1972

E2a E2a(22') E2a(22'-O-25) E2a22,25A E2a22(12) E2a22S E2a22(14) E2a22(16) E2a22(16:1) E2a22(18) E2a22(18:1) E2a22(18:2) E2a22(18:3) E2a22(20) E2A E2,3,22A E2,3,22A25S E2,3,22A24 + 25en E2,22,25A E2,22,25A3S E2S E2,22S E3A E22A E22,25A E22(2A) E22(12) E22(14) E22(16) E22(16:1) E22(18) E22(18:1) E22(18:2) E22(18:3) E22(20) E22G E22S E7H, cheilantone A E24 + 25en

Ref. [18,19, 24] [24] [24] [19] [18] [19] [18] [18] [18] [18] [18] [18] [18]

INP

1986 1986 1986 1986 1986

[25] [19,23] [19] [23] [19] [19] [19] [19] 1981 2005 [19] 1986 [18] [18] [18] [18] [18] [18] [18] [18] [19]

1986 1986 1986 1986 1986

1991 1984 1970

[23]

108

J.C. Toledano

Scheme 4.3 Synthesis of 2-deoxy- and 2,22-dideoxyecdysone from sterol precursors

E → E2,3,22A → [POCl3] → E2,3,22A24 + 25en → [a: DIBAL-H; b: DDQ] → E25d24 + 25en → [H/Pd-C] → E25d

(4.2)

E2d → E2d3,22A → [POCl3] → E2,25d3,22A24 + 25en → [a: DIBAL-H; b: DDQ] → E2,25d24 + 25en → [H/Pd-C] → E2,25d (4.3)

4

Synthetic Ecdysteroidal Compounds

109

Ecdysone was the starting material to prepare the corresponding side chain 22,25 cyclic oxides [24]: E → E2a → [Tf2 O] → E2a(22'-O-25) → E(22'-O-25) E2a → E22D2a → E2a(22') → [Tf2 O] → E2a(22-O-25) → E(22-O-25)

(4.4) (4.5)

Regioselective 2-acetylation by Candida antarctica lipase B catalysis afforded E2A amongst other substrates [25].

4.3.2

20-Hydroxyecdysone Group

20E derivatives are presented in separated sub-tables according to functional group, owing to the large number of compounds. 20E 2-deoxy, 2-dehydro, 3-deoxy, 3-dehydro and 22-dehydro derivatives are shown in Table 4.4a. 20E acetal derivatives are listed in Table 4.4b. 20E 2,3;20,22-diacetonide (20E2,20a) has been widely used owing to its simple preparation. 20E 20,22acetonide (20E20a) is also easy to prepare by selective 2,3-acetonide hydrolysis, whereas the 20E 2,3-acetonide (20E2a) has been prepared by partial non-selective hydrolysis or through E20PB. E20PB and acetylation were combined to prepare 20E monoglycosides [55]. A variety of acetals was prepared to improve deprotection conditions and recovery yields [46]. 20E acyl and alkyl derivatives are collected in Table 4.4c. Acylation rates of 20E hydroxyl functions were established very early, and 20E2A, 20E22A, 20E2,22A, 20E2,3A, 20E2,3,22A and 20E2,3,22,25A were isolated [52], although none of these derivatives was “claimed” as a synthetic ecdysteroid in [6]. The faster acylation rate of C-2 hydroxyl and specific protection allowed the synthesis of mono acyl derivatives, mainly C-2 (directly or after 20,22-diol protection) [25,26,43] and C-22 (previous 2,3-diol protection) [26,44] but also all mono, di, tri and tetraacetates [43]. A recent claim of first synthesis of 20E22B [45] overlooked the existing previous one [26]. Viticosterone E, 20E25A, was another selectively acetylated derivative of 20E through the diacetonide [43] and improved yield was reported when using other protecting acetals [46]. 20E2mal and 20E2 + 3cin were prepared from 20E2malOBn [25]. Table 4.4d lists miscellaneous and C-14 modified (Δ14 (=14en), 14-deoxy and 14-O-R) 20E derivatives. Ecdysteroids displaying one extra hydroxyl than 20E are listed in Table 4.5.

110

J.C. Toledano Table 4.4a 20E deoxy, dehydro, derivatives (2 or 3-deoxy, 2 or 3 or 22-dehydro) Non dehydro derivates Ref. INP Dehydro derivates Ref. INP 20E2d 20E2d(3') 20E2d(5a) 20E2d20a 20E2d20a(3') 20E2d20a(5a) 20E2d20alen 20E2d20alen(5a) 20E2d20a3A 20E2d20a3A(3') 20E2d20a3A(5a) 20E2d20a3Alen 20E2d20a3Alen(5a) 20E2d3A 20E2d3A(3') 20E2d3A(5a) 20E2d3B 20E2d3G 20E2d3TBDMS(5?) 20E2d3,22A 20E2d3,22A(5a) 20E2d22A 20E2d22B 20E2d25A 20E2dlen 20E2dlen(5a)

[21,26,31] [34] [31,34] [26] [34] [34] [33] [33] [26] [34] [34] [33] [33] [26] [34] [34]

20E3d 20E3d(2',5a) 20E3d20a 20E3d20a(2',5a)

[35] [35] [35] [35]

[31] [31] [31] [21,26] [26]

1968 T2

20E2d3D 20E2d3D(5a) 20E2d3D20a 20E2d3D20a(5a) 20E2d3D22A

[34] [34] [34] [34] [21]

20E3d2D 20E3d2D20a

[35] [35] [30,32] [21] [30] [21] [32]

1988

1985

20E2D(3') 20E2D(5a) 20E2D(3α9α-cyclo) 20E2D3,22A 20E2D20a3A(3')

1997 2000 1991

20E3D 20E3D2,22A 20E3D20a2A

[21] [21] [32]

1977

1997

20E22D 20E22D5*TMS

[27-29] [27-29]

1992

1991

1987 1990 2002

[33] [33]

Table 4.4b 20E acetal derivatives 20E2a 20E2a(2') 20E2a22A 20E2a22B 20E2a22Ms 20E2a(20-O-22) 20E2a(20-O-25) 20E2a,20PB 20E2a,20(RS-OH, CF3) 20E2,20a

Ref. [24,26,36, 43–45,51] [52] [26] [26,45] [24,51] [24,51] [24,51] [24,36,51,55] [46] [38-43,45,4749,53,54]

20E20a 20E20a(2') 20E20a(2',3') 20E20a(2',3',5a) 20E20a2A 20E20a2Ms 20E20a2Ms3A 20E20a2,3A 20E20a2,3Ms 20E20a2,3,25A(2',3') 20E20a2 + 3(1)

Ref. [26,32–35,43– 45,47,48,50] [52] [50] [50] [26,33,30] [33] [43] [50] [50] [35] (continued)

Table 4.4b (continued) Ref. 20E2,20a(2') 20E2,20a14,25TMS 20E2,20a25A 20E2,20a25A14TMS 20E2,20a25tfa 20E2,20(R-et) 20E2,20(R-et)25A 20E2(RS-etme) 20(R-etme) 20E2(RS-etme) 20(R-etme)25A 20E2(RS-fu)20(R-fu) 20E2(RS-meop) 20(R-meop)

[52] [48,54] [37,43,46] [37] [47] [46] [46] [46]

20E2,20a6H(5a,6?) 20E2,20a6H(5a,6a/b) 20E2,20a6H14TMS (5a,6?) 20E2,20a6H14,25TMS(5a,6?) 20E2,20a7H(8a)

[53] [54] [54]

[46] [46] [46]

20E20a3A 20E20a25A 20E20a25tfa 20E20a4*TMS

[43] [46] [47] [48]

20E20(R-hbu) 20E20(R-et) 20E20(R-et)25A 20E20(R-etme) 20E20(R-etme)25A 20E20(R-fu) 20E20(R-meop) 20E20(RS-et)

[46] [46] [46] [46] [46] [46] [46] [46]

20E20PB 20E20PB2,3A 20E20PB2(G4*A) 20E20PB3(G4*A) 20E20PB25(G4*A)

[24,36,51,55] [55] [55] [55] [55]

[54] [53]

Table 4.4c 20E acyl and alkyl derivatives Ref. 20E2A [25,43,56] 20E2apa [57] 20E2mal [25] 20E2malOBz [25] 20E2(12) [25] 20E2(16) [25] 20E2 + 3cin [25] 20E2,3A [43,55,56] 20E2,3,22A [21,23,43,55,56] 20E2,3,22A(2') [52] 20E2,3,22A25(G4*A) [55] 20E2,3,22,25A [43,56] 20E2,3,22,25A(2') [52] 20E2,3,25A [43] 20E2,22A [21,43,56] 20E2,22A(2') [52] 20E2,3A22(G4*A) [55] 20E2Ms22A [45] 20E2Ms22B [45] 20E2,22,25A [43] 20E2,25A [43]

[56] first synthesis 1969

Ref.

20E3A 20E3,22A 20E3,22A(2') 20E3,22,25A 20E3,25

Ref. [43,56] [21,43,56] [52] [43] [43]

20E22A 20E22B 20E22(16) 20E22(A2Cl) 20E22(f2c) 20E22(p2c) 20E22(t2c) 20E22,25A

[43,56] [26,45] [44] [44] [44] [44] [44] [43]

20E25A (viticosterone E)

[43,46]

20E2G 20E3G 20E22G 20E22(OEt) 20E22(OMe) 20E25G 20E6*TMS

[55] [55] [55] [44] [44] [55] [29,48]

112

J.C. Toledano

Table 4.4d Miscellaneous and C-14 modifieda 20E derivatives Ref. INP 20E(2',3') 20E(2',3',5a) 20E(5a) 20E(22-O-25) (shidasterone) 20E(22') 20E2,3,14,20,25TMS 20E7H 20E7,7'-dimer 20E8(14)en 20E9(11)en 20E24en 20E25en 20E26al (=20, 26E26D) a 14 Δ (14en), 14-deoxy, 14-O-R

[50] [50] [24-51] [27–29] [27–29] [58] [60] [61] [58,59] [58] [58] [68]

1971 1968 1998

Ref.

INP

20E14en (stachysterone B) 20E14en2,20a6H(5a,6a) 20E14en2,20a 20E14en2,20a25tfa 20E14en20a 20E14en20a25tfa

[47]

1970

20E14d 20E14d(14,18-cyclo) 20E14hpo 20E(14') calonysterone

[60,61] [60] [60] [60] [62]

1990

Ref.

INP

Table 4.5 Hydroxylated 20E derivatives Ref. INP 1,20E (integristerone A) 1,20E(1',2') 1,20E(1',2',5a) 1,20E3d1,2,22A 1,20E3d1,2,22,25A 1,20E5*A 1,20E25A 5,20E (polypodine B) 5,20E7*TMS 5,20E9(11)en (herkesterone) 9,20E

[33] [33] [33] [67] [67] [69]

[65]

9,20E(9') 9,20E3D20a2A(3-O-9)

[32]

1977

1999 1967 [48] 2004 2003 2004

11,20E (turkesterone) 11,20E(3) 11,20E(4) 11,20E(6) 11,20E(10) 11,20E(12) 11,20E(14) 11,20E(20) 11,20E2A 11,20E2,11A 11,20E11A 11,20E11,22A 20,23E(23S) (gerardiasterone) 20,23E(23R) 20,23E(22S,23R) 20,24E (abutasterone) 20,24E(24') 20,24E2,20a 20,24E2,20a(24') 20,26E 20,26E2,20a 20,26E2,20a26Ms 20,26E26Ms 20,26E(22-O-26S)(25,26 t diol) 20,26E(22-O-26R)(25,26 c diol) 20,26E25a(22-O-26)(25,26 t diol) 20,26E25a(22-O-26)(25,26 c diol)

[54] [47] [47] [47] [47]

1973

1975 [66] [66] [66] [66] [66] [66] [66] [66] [66] [66] [66] [63,64] [64] [63] [41,49] [41,49] [49] [49] [49] [65,49] [65] [65] [68] [68] [68] [68]

1986

1983 1997

4

Synthetic Ecdysteroidal Compounds

4.3.3

Ponasterone A and Taxisterone Derivatives

4.3.3.1

Ponasterone A Group

113

PoA has been synthesized from 20E diacetonide, through dehydration, catalytic reduction and deprotection [38]: 20E → 20E2,20a → [MsCl/Py/DMAP] → PoA2,20a(24 + 25)en (3:2 mixture) → [5% Pd/C H 2 ] → PoA2,20a → [deprotection] → PoA

(4.6)

Double bond reduction of the deprotected 20E24en or 20E25en to PoA or PoA7H, as well as PoA to PoA7H, was selective depending on reaction time [58]. PoA2,20a(24 + 25)en mixture obtained in the dehydration step, was (a) Hydrated to afford pterosterone (24PoA), 24-epi-pterosterone [24PoA(24')] and two 25-epimers of 26PoA2,20a mixture [38,39] (b) Treated with m-CPBA to afford the corresponding epoxide mixture [40], en route to 24-epi-pterosterone [24PoA(24')] or (c) Deprotected to a mixture of PoA24en (stachysterone D) and PoA25en [49], from which each one was obtained pure (a) PoA2,20a(24 + 25)en → [BH 3 -THF;H 2 O2 /NaOH] → 24PoA2,20a +24Poa2,20a(24′ ) + 26PoA2,20a (two 25-epimers)

(4.7)

(b) PoA2,20a(24 + 25)en → PoA2,20a(24-O-25) + PoA2,20a(24'-O-25) + PoA2,20a(25-O-26) (two isomers)

(4.8)

(c) PoA2,20a(24 + 25)en → PoA24en (stachysterone D) + PoA25en (4.9) PoA was biotransformed by Curvularia lunata to PoA2D(3') and PoA2D(3α9αcyclo) [30] whereas mild 14-OH trimethylsilylation of PoA27nor2,20a25k was achieved (→ PoA27nor2,20a25k14tms) (Scheme 4.4) [37].

Scheme 4.4 Biotransformations by Curvularia lunata

114

J.C. Toledano

Halide derivatives PoA26Br(25R), PoA26Br(25S), PoA26Cl(25R), PoA26Cl(25S), PoA26I(25R), and PoA26I(25S) were prepared from each 26PoA2,20a epimer [38]. The synthesis of 26-iodoponasterone A from inokosterone [65], and the preparation of PoA2,20a25F, PoA20a25F and PoA25F from 20E2,20a [42] were reported previously (Scheme 4.5).

Scheme 4.5 Synthesis of 25-fluoroponasterone A

PoA9(11)en (dacryhainansterone) from 11PoA (ajugasterone C), 5PoA9(11)en (kaladasterone) from 5,11PoA (muristerone A) were prepared as potential photoaffinity labels [59], whereas 5,11PoA was regioselectively acetylated by Candida antarctica lipase B catalysis to 5,11PoA2A [25]. Table 4.6 PoA and tax derived synthecdysteroids Ref. INP PoA2d(5a) (cyanosterone A) PoA14d7H(2',3',5a)

[76]

PoA2D(3') PoA2D(3α9α-cyclo) PoA3,22D2A4en

[30] [30] [75]

PoA2,20a6H(5a,6?) PoA2,20a6H24/25en(6?) PoA2,20a7H24/25en(8a) PoA2,20a14TMS24/25en PoA2,20a14TMS6H24/ 25en(5a,6?) PoA2,20a24/25en

[54] [53] [53] [54] [54]

2002 5PoA9(11)en (kaladasterone) 5,11PoA2A 5,26PoA2,20a 5,26PoA2,20a26MTPA 9PoA2A4en(3')

[38–41,49]

24PoA (pterosterone) 24PoA(24') 24PoA2D(3') 24PoA2D(3α9α-cyclo) 24PoA2,20a

Ref.

INP

[59]

1973

[25] [70] [70] [75] [39] [39,40] [30] [30] [38]

(continued)

4

Synthetic Ecdysteroidal Compounds

115

Table 4.6 (continued) Ref. PoA2,20a(24-O-25) PoA2,20a25F PoA2,20a25F14en PoA2,20a26I PoA20a9(11)en =[kaladasterone]5d,20a PoA20a25F

[42]

PoA2A4en(3') PoA2,3,22A4en(3') PoA7H PoA25F

[75] [75] [58] [42]

PoA26Br PoA26Cl PoA26I

[38] [38] [38,65]

PoA9(11)en PoA24en PoA25en

[59] [41,49] [41,49]

4.3.3.2

[40] [42] [42] [65]

INP

Ref. 24PoA2,20a(24') 24PoA2,20a25en(24') 24PoA7H

INP

[38,40] [40] [58]

2001 26PoA-I (inokosterone-I) [39] 26PoA-II (inokosterone-II) [39] 26PoA2,20a [38] 26PoA26apa [57] 26PoA26hpp [36,71] 26PoA26hpp23in [36] PoA27nor2,20a25k [37] PoA27nor2, [37] 20a25k14TMS Tax2a25THP22en 11Tax (scabrasterone) 24Tax (pinnatasterone) 24Tax2,3,22A(24?) 24Tax2,3,22,25A(24?) 26Tax(3')

[63,64] 2002 1993 [73,74] [73,74] 2000

Pterosterone (24PoA) Group

Pterosterone and 24-epi-pterosterone [24Poa(24')] were prepared from the already mentioned mixture of 24PoA2,20a, 24Poa2,20a(24'), and 26PoA2,20a [38,39] (obtained from PoA2,20a24en + PoA2,20a25en), and from the epoxide mixture of PoA2,20a(24-O-25), PoA2,20a(24'-O-25), PoA2,20a(25-O-26) [40]. As above, 24PoA was similarly biotransformed by Curvularia lunata to 24PoA2D(3') and 24PoA2D(3α9α-cyclo) [30] (Scheme 4.4), whereas the 7,8-double bond was also selectively reduced (→ 24PoA7H) [58].

4.3.3.3

Inokosterone (26PoA) Group

The C-25 epimer mixture of 26PoA2,20a [34,35] was best separated prior to deprotection and the absolute configuration of the two epimers of inokosterone (26PoA-I, inokosterone-I; 26PoA-II, inokosterone-II) was determined. PoA26halide derivatives preparation has been already mentioned [38]. 5,26PoA was isolated as palythoalone B, and the chemical shift differences of the (+) and (−)-MTPA esters revealed the 25R configuration [70]. The esters were prepared according to Eq. 4.10: 5,26PoA → 5,26PoA2,20a → 5,26PoA2,20a26MTPA

(4.10)

116

J.C. Toledano

From the protected Pos2a14SEM or Pos2a14EE (preferred) 26-alkoxy derivatives of PoA were also prepared [36,71]: Pos2a14EE → MNC2a14EE22en → BNC2a14EE22al → → 26PoA26hpp23in → 26PoA26hpp

4.3.3.4

(4.11)

Taxisterone Group

As mentioned, a preliminary account reported the synthesis of 22-deoxy-20-hydroxyecdysone as early as 1968 [9] and a full paper was released the following year [72] but the ecdysteroid was named taxisterone when isolated from Taxus cuspidata [8]. Tax2a25THP22en(E) has been prepared from Pos2A [63,64] as an intermediate in the synthesis of gerardiasterone 20,23E(22R,23S) or the diastereomer 20,23E(22',23') with 22S,23R configuration. Pinnatasterone [24Tax(24?)] and 24-epi-pinnatasterone have been isolated from natural sources [73,74] but assignment of C24 absolute configuration remains uncertain. Both on acetylation afforded a tri and a tetraacetate. As a result, the epimers 24Tax2,3,24A and 24Tax2,3,24A(24'), as well as 24Tax2,3,24,25A and 24Tax2,3,24,25A(24') are available but a precise C24 absolute configuration assignment is still pending.

4.3.4

Other Derivatives

4.3.4.1

Rubrosterone Group

Rubrosterone isolation and two synthesis were reported in preliminary form in 1968 [77–79], separated by a mere 3 months. Full papers on structure/absolute configuration and synthesis of this C-17:C-20 bond cleaved product followed in 1969 [80,81], a key intermediate being Rub17H as shown in Scheme 4.6. 20E was used as starting material in one approach and 17β-acetoxy-5α-androstan3,6-dione in the other one. In this second instance, 5α-dihydrorubrosterone triacetate [Rub2,3,17A17H(5a)] was obtained in seven steps as a key intermediate [79], then converted to Rub17H, and the following steps were as previously. A third approach started with 3β-tosyloxyandrost-5-en-17-one, and the key intermediate was Rub2,3A(5a) [82]. Rub17H was isolated as a natural product much later (in 1990) from Caryophyllaceae [83], the compound being “identical with the product earlier synthesized” [84]. The title of the paper was “An alternative synthesis…”, and credited all three previous syntheses of rubrosterone. However, being alternative it was implying (at least) one precedent for Rub17H. The spectra and physical constants for the obtained Rub17H in this alternative synthesis “being consistent with those reported” (in [79]) settled the question. The synthesis starting material was now

4

Synthetic Ecdysteroidal Compounds

Scheme 4.6 Synthesis of rubrosterone and derivatives

117

118

J.C. Toledano

Table 4.7 Synthecdysteroids with short or no side-chain Ref. INP Rub (rubrosterone) Rub2a Rub2a17H Rub17A17H Rub2,3A Rub2,3A(5a) Rub2,3,17A17H(5a) Rub17H 11Rub

[78,79,81,82,84] [78,79,81,84] [78,79,81,84] [78,81,84] [81] [82] [79,84] [78,79,81,84]

Pos (poststerone) Pos2d3A(5a) Pos2d3TBDMS(5?) Pos2d3TBDMS20H(5?) Pos2D Pos3D Pos3D2A Pos2a Pos2a14EE Pos2a14SEM Pos2a14TMS Pos2A Pos2,3A Pos2,3A14TMS Pos7H Pos20H 11Pos

[36,85,86,97] [87] [31,96] [31,96]

Ref.

BNC2d3A22al(5a) BNC2a14EE22al BNC2a14SEM22al 20BNC2a14,20TMS21(3*F) 20BNC2,3A14,20TMS21(3*F) 20BNC14,20TMS21(3*F) 20BNC21(3*F) 20BNC22al

[16,89] [36,71] [36] [86] [86] [86] [86] [97]

20MNC22en 20MNC2a14EE22en 20MNC2a14SEM22en

[97] [36,71] [36,71]

CCL2d3TBDMS (5?) CCL2d3TBDMS24Me(5a) CCL2d3TBDMS24Me(5?,24') CCL2,3A(5a) CCL2,3A(5a,22')

[31] [31] [31] [88] [88]

2003

2005 [85] [85] [36,37,71,86] [36,71] [36,71] [37,86] [78,81] [37,86] [37,86] [58] [97] 2004

3β-mesyloxyandrost-5-en-17-one (or dehydroisoandrosterone methanesulphonate) and again Rub2,3,17A17H(5a) was obtained through a multistep sequence, and Rub17H there-from, as previously.

4.3.4.2

Poststerone Group

Poststerone played a pivotal role in ecdysteroid chemistry. First of all to prove the same tetracyclic structure and substitution as in 20E for a number of compounds. The formation of a “methylketone” on sodium metaperiodate oxidation of ponasterone A was reported in 1966 [90] and from 20E in 1967 (a byproduct in the synthesis of E was shown to be identical), a melting point being

4

Synthetic Ecdysteroidal Compounds

119

reported [91]. Improved results were obtained by previous selective acetylation to 20E2A, periodate cleavage to Pos2A and mild hydrolysis [9] and also from PoA2A [92]. Also to prove partial structure and substitution, Pos2A was in turn derived from MaC2A [93], MaA2A [94], 26PoA2,26A and 20E2A [95]. As a synthetic intermediate, Pos was prepared from 20E by direct oxidation with Jones reagent [72], later on found to produce also Pos3D (and Pos3D → Pos3D2A) [85]. Pregnenolone was used as starting material to prepare a Pos conveniently protected derivative [Pos2d3TBDMS(5a or 1:1)] [31,96]. In a multistep sequence [97] a previously described derivative (20β-benzoyloxy-5-pregnen-3β-ol) was converted to 20E through Pos20H, Pos, 20MNC22en, 20BNC22al, 20E25THP23in and 20E25THP. Pos2a was involved also as an immediate precursor of Pos (starting material Cy or 20E), or further used after 14-OH protection: Pos2a → Pos2a14SEM or Pos2a14EE (preferred) [36,71]

(4.12)

or, Pos2a → Tax2a25THP22en, directly [63,64]t

(4.13)

Pos was used to prepare 20E22D, and Pos(3*TMS) was also synthesized [27–29]. Mild 14-OH trimethylsilylation of Pos2,3A or Pos2a → Pos2,3A14TMS or Pos2a14TMS respectively [37], and on selective 7,8-double bond reduction Pos → Pos7H [58]. The first example of trifluoromethylation in the ecdysteroid series involved the preparation of several poststerone and bis-nor-cholane derivatives according to Eqs. 4.14 and 4.15 [86]: 20E → Pos → Pos2,3A → Pos2,3A14TMS → 20BNC2,3A14, 20TMS21(3*F) → 20BNC14,20TMS21(3*F) → 20BNC21(3*F) (20RS ) (4.14)

Pos → Pos2a → Pos2a14TMS → 20BNC2a14, 20TMS21(3*F) → 20BNC14,20TMS21(3*F) → 20BNC21(3*F) (20RS ).

4.3.4.3

(4.15)

Short Side-Chain Synthetic Ecdysteroids (BNC, MNC, CCL)

Sidisterone is a unique C-24 ecdysteroid and contains a side chain butenolide function (22-en-24,20-lactone). The parent hydrocarbon name is cholane (not cholestane), and it was selected here to provide a unified approach from C-22 to C-25 abbreviations: C-22 or bis-nor cholane (BNC) and C-23 or mono-nor-cholane (MNC) skeletons. Since C-25 compounds listed always display the 24,22-carbolactone function the choice was cholane-carbolactone (CCL).

120

J.C. Toledano Table 4.8 Branched side-chain synthecdysteroids Ref. MaA2d(5a) [31] Cy22D MaA2d(24') [31] Cy2a MaA2d(5a,24') [31] Cy2a20PB MaA2d3TBDMS(5a) [31] Cy20PB MaA2d3,22A(5a) [31] Cy2ibt MaA2,20,22,25d22en(5a,24') [17] Cy2,3,22A MaA2,20,22,25d3A22en(5a,24') [17] Cy2,3,22ibt MaA2,20,22,25d3Ms22en(5a,24') [17] Cy3A MaA2A [25] Cy22A 5MaB2,20a(24?) [70] 5MaB2,20a26MTPA(24?) [70] 11Pan20a 11Pan20 + 22PB 11Pan20PB2,3,11A 11Pan22PB2,3,11A 11Pan20hbz

4.3.4.4

Ref.

INP. 1992

[36,71] [36,71] [36,71] [98] [98] [98] 1995 1978

[99] [99] [99] [99] [99]

Branched Side-Chain Synthetic Ecdysteroids

Mainly, makisterone A and cyasterone derivatives have been reported.

4.4

Concluding Remarks

After this initial effort two major tasks may follow: adding new (or old but missing) information or correcting errors slipped in. Of course, the help of other ecdysonists will always be welcome as well as suggestions to improve and extend the usefulness of the present resource. Acknowledgements Financial support for ecdysteroid research since the early eighties (CAICYT, CICYT, DGICYT, Generalitat de Catalunya) is gratefully acknowledged.

References i. Lafont R, Harmatha J, Marion-Poll F, Dinan L, Wilson ID. The Ecdysone Handbook, 3rd edition, on-line, http://ecdybase.org ii. Lafont R, Wilson I (1992) The Ecdysone Handbook. The Chromatographic Society, Nottingham, UK

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1. Goodwin TW, Horn DHS, Karlson P, Koolman J, Nakanishi K, Robbins WE, Siddall JB, Takemoto T (1978) Nature 272:122 2. Lafont R, Koolman J, Rees H (1993) Insect Biochem Mol Biol 23:207 3. Karlson P (1995) Eur J Entomol 92:7 4. Lafont R, Horn DHS (1989) Phytoecdysteroids: Structures and occurrence. In: Koolman J (ed) Ecdysone: From Chemistry to Mode of Action. Georg Thieme-Verlag, Stuttgart/ New York, p 39 5. Kametani T, Tsubuki M (1989) Strategies for the synthesis of ecdysteroids. In: Koolman J (ed) Ecdysone: From Chemistry to Mode of Action. Georg Thieme-Verlag, Stuttgart, p 74 6. Horn DHS, Bergamasco R (1985) Chemistry of ecdysteroids. In: Kerkut GA, Gilbert LI (eds) Comprehensive Insect Physiology, Biochemistry and Pharmacology, vol. 7: Endocrinology, I. Pergamon Press, Oxford, p 185 7. Lafont R, Dauphin-Villemant C, Warren JT, Rees H (2005) Ecdysteroid chemistry and biochemistry. In: Gilbert LI, Iatrou K, Gill SS (eds) Comprehensive Molecular Insect Science, vol. 3: Endocrinology. Elsevier, Oxford, p 125 8. Nakano K, Nohara T, Tomimatsu T, Nishikawa M (1982) Phytochemistry 21:2749 9. Galbraith MN, Horn DHS, Middleton EJ, Hackney RJ (1968) Chem Commun:466 10. Lee YW, Lee E, Nakanishi K (1980) Tetrahedron Lett 21:4323 11. Mori H, Shibata K, Tsuneda K, Sawai M (1968) Chem Pharm Bull 16:563 12. Mori H, Shibata K, Tsuneda K, Sawai M (1971) Tetrahedron 27:1157 13. Galbraith MN, Horn DHS, Thomson JA (1975) Experientia 31:873 14. Hetru C, Nakatani Y, Luu B, Hoffmann JA (1983) Nouv J Chim 7:587 15. Haag T, Hetru C, Kappler C, Moustier AM, Hoffmann JA, Luu B (1988) Tetrahedron 44:1397 16. Haag T, Luu B, Hetru C (1988) J Chem Soc Perkin Trans 1:2353 17. Hedtmann U, Hobert K, Milkova T, Welzel P (1988) Tetrahedron 44:1941 18. Dinan L (1988) J Steroid Biochem 31:237 19. Pis J, Girault JP, Grau V, Lafont R (1995) Eur J Entomol 92:309 20. Howarth OW, Thompson MJ, Rees HH (1989) Biochem J 259:299 21. Girault JP, Blais C, Beydon P, Rolando C, Lafont R (1989) Arch Insect Biochem Physiol 10:199 22. Dolle F, Hetru C, Roussel JP, Rousseau B, Sobrio F, Luu B, Hoffmann JA (1991) Tetrahedron 47:7067 23. Pis J, Girault JP, Larchevêque M, Dauphin-Villemant C, Lafont R (1995) Steroids 60:188 24. Roussel PG, Sik V, Turner NJ, Dinan LN (1997) J Chem Soc Perkin Trans 1:2237 25. Danieli B, Lesma G, Luisetti M, Riba S (1997) Tetrahedron 53:5855 26. Suksamrarn A, Yingyongnarongkul B (1996) Tetrahedron 52:12623 27. Hedtmann U, Welzel P (1985) Tetrahedron Lett 26:2773 28. Hedtmann U, Hobert K, Klone A, Klintz R, Pongs O, Strangmanndiekmann M, Welzel P, Frelek J (1989) Angew Chem Int Ed 28:1515 29. Hedtmann U, Klintz R, Hobert K, Frelek J, Vlahov I, Welzel P (1991) Tetrahedron 47:3753 30. Changtam C, Sukcharoen O, Yingyongnarongkul B, Suksamrarn A (2006) Steroids 71:902 31. Kametani T, Tsubuki M, Higurashi K, Honda T (1986) J Org Chem 51:2932 32. Charoensuk S, Yingyongnarongkul B, Suksamrarn A (2000) Tetrahedron 56:9313 33. Kumpun S, Yingyongnarongkul B, Lafont R, Girault JP, Suksamrarn A (2007) Tetrahedron 63:1093 34. Suksamrarn A, Yingyongnarongkul B (1997) Tetrahedron 51:3145 35. Suksamrarn A, Charoensuk S, Yingyongnarongkul B (1996) Tetrahedron 52:10673 36. Guedin-Vuong D, Nakatani Y, Luu B, Ourisson G (1985) Tetrahedron Lett 26:5959 37. Odinokov VN, Savchenko RG, Nazmeeva SR, Galyautdinov IV (2002) Russian Chem Bull Int Ed 51:1963 38. Yingyongnarongkul B, Kumpun S, Chimnoi N (2005) Steroids 70:636 39. Yingyongnarongkul B, Suksamrarn A (2000) ScienceAsia 26:15 40. Suksamrarn A, Yingyongnarongkul B, Charoensuk S (1999) Tetrahedron 55:255

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41. 42. 43. 44.

Yingyongnarongkul B, Suksamrarn A (1998) Tetrahedron 54:2795 Tomás J, Camps F, Coll J, Melé E, Pascual N (1992) Tetrahedron 48:9809 Suksamrarn A, Pattanaprateep P (1995) Tetrahedron 51:10633 Suksamrarn A, Pattanaprateep P, Tanachatchairatana T, Haritakun W, Yingyongnarongkul B, Chimnoi N (2002) Insect Biochem Mol Biol 32:193 Mamadalieva NZ, Ramazanov NS, Girault JP, Lafont R, Saatov Z (2004) Chem Nat Compd 40:488 Galyautdinov IV, Nazmeeva SR, Savchenko RG, Ves’kina NA, Nedopekin DV, Fatykhov AA, Khalilov LM, Odinokov VN (2004) Russ J Org Chem 40:675 Odinokov VN, Galyautdinov IV, Nedopekin DV, Khalilov LM (2003) Russ Chem Bull Int Ed 52:232 Odinokov VN, Savchenko RG, Nazmeeva SR, Galyautdinov IV, Khalilov LM (2002) Russ Chem Bull Int Ed 51:1937 Suksamrarn A, Yingyongnarongkul B, Promrangsan N (1998) Tetrahedron 54:14565 Homvisasevongsa S, Chuaynugul A, Chimnoib N, Suksamrarn A (2004) Tetrahedron 60:3433 Roussel PG, Turner NJ, Dinan LN (1995) J Chem Soc Chem Commun:933 Singh SB, Thakur RS (1982) Tetrahedron 38:2189 Afon’kina SR, Shafikhov RV, Savchenko RG, Galyautdinov IV, Odinokov VN (2006) Russ J Org Chem 42:1234 Odinokov VN, Savchenko RG, Shafikov RV, Afon’kina SR, Khalilov LM, Kachala VV, Shashkov AS (2005) Russ J Org Chem 41:1296 Pis J, Hykl J, Budesinsky M, Harmatha J (1994) Tetrahedron 50:9679 Galbraith MN, Horn DHS (1969) Aust J Chem 22:1045 Boehm MF, Nakanishi K, Cherbas P (1991) J Chem Soc Chem Commun:52 Suksamrarn A, Tanachatchairatana T, Sirigarn C (2002) Tetrahedron 58:6033 Bourne PC, Whiting P, Dhadialla TS, Hormann RE, Girault JP, Harmatha J, Lafont R, Dinan L (2002) J Insect Sci 1:12 Harmatha J, Budeˇšínský M, Vokácˇ K (2002) Steroids 67:127 Canonica L, Danieli B, Lesma G, Palmisano G (1985) J Chem Soc Chem Commun:1321 Suksamrarn A, Ganpinyo P, Sommechai C (1994) Tetrahedron Lett 35:4445 Honda T, Takada H, Miki S, Tsubuki M (1993) Tetrahedron Lett 34:8275 Tsubuki M, Takada H, Katoh T, Miki S, Honda T (1996) Tetrahedron 52:14515 Lee SS, Nakanishi K, Cherbas P (1991) J Chem Soc Chem Commun:51 Dinan L, Bourne P, Whiting P, Tsitsekli A, Saatov Z, Dhadialla TS, Hormann RE, Lafont R, Coll J (2003) J Insect Sci 3:6(1–11) Jayasinghe L, Jayasooriya CP, Oyama K, Fujimoto Y (2002) Steroids 67:555 Kayser H, Ertl P, Eilinger P, Spindler-Barth M, Winkler T (2002) Arch Biochem Biophys 400:180 Sadikov ZT, Saatov Z (1999) Chem Nat Compd 35:440 Shigemori H, Sato Y, Kagata T, Kobayashi J (1999) J Nat Prod 62:372 Guedin-Vuong D, Nakatani Y, Ourisson G (1985) Croat Chem Acta 58:547 Galbraith MN, Horn DHS, Middleton EJ, Hackney RJ (1969) Aust J Chem 22:1517 Suksamrarn A, Sommechai C (1993) Phytochemistry 32:303 Suksamrarn A, Kumpun S, Yingyongnarongkul B (2002) J Nat Prod 65:1690 Fukuzawa A, Miyamoto M, Kumagai Y, Masamune T (1986) Phytochemistry 25:1305 Watanabe B, Nakagawa Y, Ogura T, Miyagawa H (2004) Steroids 69:483 Takemoto T, Hikino Y, Hikino H, Ogawa S, Nishimoto N (1968) Tetrahedron Lett:3053 Hikino H, Hikino Y, Takemoto T (1968) Tetrahedron Lett:4255 Hocks P, Kerb U, Wiechert R, Furlenmeier A, Fürst A (1968) Tetrahedron Lett:4281 Takemoto T, Hikino Y, Hikino H, Ogawa S, Nishimoto N (1969) Tetrahedron 25:1241 Hikino H, Hikino Y, Takemoto T (1969) Tetrahedron 25:3389 VanBever W, Kohen F, Ranade VV, Counsell RE (1970) J Chem Soc Chem Commun:758 Girault JP, Bathori M, Varga E, Szendrei K, Lafont R (1990) J Nat Prod 53:279 Cochrane JS, Hanson JR (1971) J Chem Soc C:3730

45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84.

4

Synthetic Ecdysteroidal Compounds

85. 86. 87. 88. 89. 90. 91. 92. 93. 94. 95.

Petersen QR, Cambie RC, Russell GB (1993) Aust J Chem 46:1961 Odinokov VN, Nazmeeva SR, Savchenko RG (2003) Russian J Org Chem 39:1733 Mauvais A, Hetru C, Roussel JP, Luu B (1993) Tetrahedron 49:8597 Kametani T, Tsubuki M, Nemoto H (1980) Tetrahedron Lett 21:4855 Hetru C, Nakatani Y, Luu B, Hoffmann JA (1983) New J Chem 7:587 Nakanishi K, Koreeda M, Sasaki S, Chang ML, Hsu HY (1966) Chem Commun:915 Siddall JB, Horn DHS, Middleton EJ (1967) Chem Commun:899 Moriyama H, Nakanishi K (1968) Tetrahedron Lett:1111 Galbraith MN, Horn DHS, Porter QN, Hackney RJ (1968) Chem Commun:971 Imai S, Hori M, Fujioka S, Murata E, Goto M, Nakanishi K (1968) Tetrahedron Lett:3883 Takemoto T, Hikino Y, Arihara S, Hikino H, Ogawa S, Nishimoto N (1968) Tetrahedron Lett:2475 Mauvais A, Hetru C, Luu B (1991) Tetrahedron Lett 32:5171 Mori H, Shibata K (1969) Chem Pharm Bull 17:1970 Ramazonov NSh, Syrov VN (2006) Chem Nat Compd 42:558 Vokácˇ K, Budeˇšinský M, Harmatha J, Píš J (1998) Tetrahedron 54:1657

96. 97. 98. 99.

123

Chapter 5

Ecdysteroids and Ecdysteroid Signaling Pathways During Insect Oogenesis Luc Swevers and Kostas Iatrou

Abstract During insect oogenesis, the oocyte acquires nutrients and genetic determinants to support embryonic development (previtellogenesis and vitellogenesis) and subsequently becomes surrounded by a protective eggshell (choriogenesis). In many insects, ecdysteroids are synthesized during ovarian growth which is often followed by the accumulation of ecdysteroid conjugates into the eggs. The exact role of the ecdysteroids during oogenesis remains largely unclarified although functions as paracrine or autocrine regulators to signal the progression of follicle development or the resumption of meiosis in the oocyte have been proposed. In the silkmoth, Bombyx mori, although ecdysteroids are synthesized by the ovarian follicles, progression of follicle development towards choriogenesis requires down-regulation in ecdysteroid signaling. Using the regulation of silkmoth oogenesis by 20-hydroxyecdysone as a starting point, this review discusses the physiological roles of ecdysteroids and the function of the ecdysone regulatory pathway during insect oogenesis. Keywords ecdysone • ecdysteroid • ecdysteroid conjugate • 20E • insect oogenesis • Bombyx mori • Drosophila melanogaster • Aedes aegypti • ecdysone regulatory pathway • EcR • usp • E75 • BR-C • E74 • FTZ-F1 • Broad-Complex • ecdysone receptor • ecdysone biosynthesis • maternal ecdysteroids • vitellogenesis • choriogenesis • embryogenesis • Lepidoptera • Diptera • Orthoptera • Dictyoptera, Hymenoptera

5.1

Introduction

In insects, the function of ecdysteroids has been mostly investigated in the developmental processes that regulate molting and metamorphosis. In these processes, 20-hydroxyecdysone (20E) has been shown to activate the ecdysone receptor (EcR) complex, which consists of a heterodimer of two members of the nuclear receptor L. Swevers and K. Iatrou Insect Molecular Genetics and Biotechnology, Institute of Biology, National Center for Scientific Research “Demokritos”, Aghia Paraskevi Attikis, Greece

G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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family, EcR and USP, the latter being the homolog of the vertebrate retinoid X receptor (Thomas et al., 1993; Yao et al., 1992, 1993). Binding of the hormone to the EcR complex initially results in the activation of a conserved hierarchical cascade of gene expression consisting of an interacting set of transcription factors, encoded by the so-called ecdysone-responsive early and early-late genes, including Ets domaincontaining E74, the orphan nuclear receptors E75 and HR3, and the Broad-Complex (BR-C) zinc finger proteins (Riddiford, 1993a; Thummel, 1990, 1996, 1997; Henrich and Brown, 1995). Transduction and amplification of the hormonal signal by the conserved set of early gene products subsequently results in the regulation of numerous ecdysone-responsive late genes that define the phenotypic effects of 20E in a stageand tissue-specific manner (Thummel, 2002; Bender, 2003). However, the activation of the ecdysone regulatory cascade comprises only the first half of the developmental events in which ecdysteroids are involved. Prepupal Drosophila tissues require a decline in ecdysteroid signaling in order for development to progress and to respond to the next rise in hormone titer (Richards, 1976; Woodard et al., 1994). In lepidopteran larvae, application of strong ecdysone agonists such as tebufenozide initiate larval molting (cuticle apolysis) but failure to clear the hormone results in a developmental arrest in the middle of the molt (absence of ecdysis; Retnakaran et al., 1995). These observations support the existence of a signalling cascade that is triggered by a decline in ecdysteroid titer. Genetic studies in Drosophila have shown that the orphan nuclear receptor βFTZ-F1 likely functions at the initiation of this cascade since expression of βFTZ-F1 is induced by a decline in 20E titer and βFTZ-F1 mutants show developmental defects consistent with its role as a competence factor to prepare the prepupal-pupal transition during low ecdysteroid titers (Woodard et al., 1994; Broadus et al., 1999). Besides its role in molting and metamorphosis where the molecular mechanism of its action has been studied most extensively, 20E has been implicated in the regulation of many other biological processes in insects, such as reproduction (oogenesis, vitellogenesis and spermatogenesis; Raikhel et al., 2005), embryogenesis (Kozlova and Thummel, 2003), diapause (Denlinger et al., 2005) and polyphenism (Hartfelder and Emlen, 2005). Recent studies in the silkworm, Bombyx mori, have indicated that the development of the ovary during the pupal and pharate adult stages is regulated through the ‘classical’ hierarchical cascade of gene expression mentioned above (Swevers et al., 2005). Furthermore, in the mosquito, Aedes aegypti, the regulation of vitellogenin synthesis in the fat body by ecdysteroids produced by the ovary occurs by the conserved set of ecdysone-responsive early and early-late genes (Raikhel et al., 1999; Li et al., 2000). Genetic studies also established that progression of oogenesis in Drosophila requires the function of genes implicated in the ecdysone regulatory hierarchy such as EcR, BR-C and E75 (Buszczak et al., 1999) as well as genes involved in synthesis of ecdysteroids (Freeman et al., 1999). These findings therefore suggest that the ecdysone-regulatory hierarchy is involved in the regulation of oogenesis in insects. This review will focus on the possible role of ecdysteroids to regulate the process of oogenesis in insects. Inevitably, the review will focus on the three insects for which most knowledge has accumulated (Bombyx, Aedes, Drosophila) while oogenesis in insects of other groups (mainly cockroaches, locusts and hymenopterans) will also be discussed.

5

Ecdysteroids and Ecdysteroid Signaling Pathways During Insect Oogenesis

5.2

129

The Silkmoth Paradigm: Control of Ovarian Development by Ecdysone

In the silkmoth, ovarian development is completely coupled to metamorphosis: it occurs almost exclusively during pharate adult and adult development (Tsuchida et al., 1987; Swevers and Iatrou, 2003). The dependence of ovarian development on ecdysteroids produced by prothoracic glands has been clearly demonstrated in experiments with pupae ligated between thorax and abdomen (Tsuchida et al., 1987; Swevers and Iatrou, 1999). In the isolated abdomens, the ovaries remain undeveloped; complete ovarian development is achieved following a single injection of microgram quantities of 20E. Induction of ovarian development occurs via binding of 20E to the ecdysone receptor heterodimer consisting of BmEcR and BmUSP (Swevers et al., 1995; Tzertzinis et al., 1994) which is followed by the activation of the ecdysone regulatory cascade. Silkmoth homologs of the A- and C-isoforms of the E75 nuclear receptor (BmE75A and C; Swevers et al., 2002) are induced first which is followed by the induction of the B and C-isoforms of BmHR3 (Eystathioy et al., 2001). Concomitant with the induction of BmE75 and BmHR3 occurs the decline of the BmFTZ-F1 nuclear receptor (Sun et al., 1994; Swevers and Iatrou, 2003). The expression of other “classical” ecdysone-responsive genes, such as BR-C and E74 (Bombyx homologs of BR-C have been described recently (Uhlirova et al., 2003; Nishita and Takiya, 2004; Ijiro et al., 2004) while E74 was also described recently in Manduca (Stilwell et al., 2003), has not been described yet during induction of ovarian development in Bombyx, but it is expected that they play similar roles as in the ecdysone-response in tissues in other insects. The expression of the yolk protein produced by the follicular epithelium, eggspecific protein (ESP; Sato and Yamashita, 1991), corresponds to a late event in the ecdysone regulatory cascade induced in the silkmoth ovary (Swevers and Iatrou, 2003). Interestingly, expression of the A-isoform of the BmHR3 receptor occurs concomitantly with ESP expression and BmHR3A has been proposed to function as a regulator of ESP expression (Eystathioy et al., 2001). While the early stages of oogenesis and the initiation of vitellogenesis are dependent on the presence of active ecdysteroids in the hemolymph, experiments using dibenzoyl hydrazine ecdysone agonists such as tebufenozide (Dhadialla et al., 1998) have indicated that the completion of vitellogenesis and the process of egg shell synthesis (choriogenesis) require the absence of ecdysone signaling. Similar to 20E, tebufenozide induces early and late gene expression in ecdysone-responsive target tissues but because of its persistence in tissues gene expression that is dependent on down-regulation of ecdysone signalling does not occur (Retnakaran et al., 1995). When tebufenozide is injected in silkmoth isolated abdomens, ovarian development, including vitellogenesis, is initiated but becomes subsequently arrested (Swevers and Iatrou, 1999). Gene expression analysis in arrested versus developing ovarioles established that the developmental block occurs during middle vitellogenesis and that the orphan nuclear receptor BmFTZ-F1 is the earliest factor whose induction does not occur in arrested follicles (Swevers and Iatrou,

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1999). Thus, in silkmoth ovarian follicles BmFTZ-F1 may function as a ‘competence factor’ that regulates the transition between developmental periods with high ecdysteroid titers and low ecdysteroid titers in similar fashion as was shown in genetic experiments in Drosophila (Woodard et al., 1994; Broadus et al., 1999). The regulation of ovarian development by ecdysteroids produced by the prothoracic glands during pupation in silkmoths is summarized in Fig. 5.1. The figure illustrates that ovarian follicle development occurs in two phases: an early phase dependent on high ecdysteroid titers from previtellogenesis to middle vitellogenesis and a late phase dependent on low ecdysteroid titers from middle vitellogenesis

20E

(A) (1-2 h) (1-

EcR/USP

FTZ-F1 -F1

Early

(6 h)

HR3B HR3C

(24 h) (48-72 h)

Late

VMP30 ((vitellin membrane)

HR3 E75A

FTZ-F1

EcR (B1) E75A E75C Early-Late

(B)

HR3 EcR,USP E75C GATAβ SH3 ESP

(6 h) (12 h) (18 h)

HNF-4a

Chorion genes (24 h)

HR3A ESP

Choriogenesis

Vitellogenesis

Fig. 5.1 Regulation of silkmoth (Bombyx mori) oogenesis during pharate adult and adult development by 20E produced by the prothoracic glands. Panel a: Induction of vitellogenesis by rising titers of 20E in the hemolymph. Indicated are different phases in the hormone response: early (repression of FTZ-F1, transient induction of B1-EcR and E75C, permanent induction of E75A), early-late (induction of HR3B and HR3C) and late (induction of HNF-4A, HR3A and ESP). Expression of ESP (egg-specific protein, a yolk protein precursor produced by the follicular epithelium; Sato and Yamashita, 1991) marks the initiation of vitellogenesis. Panel b: Induction of choriogenesis by declining titers of 20E in the hemolymph. At the top of the cascade is shown the nuclear receptor FTZ-F1, which plays a pivotal role in the regulation of developmental events during low titers of ecdysone (Broadus et al., 1999). As in Drosophila, induction of FTZ-F1 may be triggered by changes in the relative levels of the HR3 and E75 receptors (Swevers et al., 2002). Expression of FTZ-F1 is followed by the repression of HR3 and EcR/USP (at 6–12 h), induction of E75C, GATAβ and SH3 (at 12 h), repression of ESP (at 12–18 h) and induction of chorion gene expression (at 18–24 h). Because the vitellin membrane protein VMP30 is co-expressed with FTZF1, it was hypothesized that FTZ-F1 is a positive regulator of the expression of VMP30 (Kendirgi et al., 2002). Note that the deduction of the regulatory cascades that trigger vitellogenesis and choriogenesis in the silkmoth is based on expression patterns of mRNAs and remains to be investigated by functional analysis (See Color Plates)

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until the end of choriogenesis. It can also be noted that silkmoth ovarian development is completely dependent on ecdysteroid signalling and that juvenile hormone (JH) does not play a role in silkmoth ovarian development (Izumi et al., 1984). Besides the production of ecdysteroids by the prothoracic glands at early pupation that triggers ovarian development, the ovary itself also starts to produce ecdysteroids at day 4 after larval-pupal ecdysis (Ohnishi and Chatani, 1977). Ecdysteroids that were identified include ecdysone and 20E as well as several of their biosynthetic precursors and metabolites such as 2-deoxyecdysone, 2-deoxy20E, 2, 22-bisdeoxy-20E, 3-epi-ecdysone and 3-epi-2-deoxy-ecdysone (Legay et al., 1976; Ohnishi et al., 1989; Sonobe and Yamada, 2004). Several lines of evidence indicate that the ecdysteroids produced by the ovary have no autocrine/ paracrine or endocrine function to regulate the progression of oogenesis in the silkmoth. First, the ecdysteroids produced by the ovary are not secreted towards the hemolymph but accumulate in the eggs, mainly as ecdysteroid conjugates (C22 and C3 phosphate esters; Sonobe and Yamada, 2004). Second, as argued above, the experiments using the ecdysone agonist tebufenozide have clearly demonstrated that progression of vitellogenesis towards choriogenesis requires decline, not activation, of ecdysone signalling (Swevers and Iatrou, 1999). Third, suppression of the ecdysteroid content of ovarian follicles through the administration of the imidazole compound KK-42, a potent inhibitor of ecdysone synthesis, does not cause a disruption in ovarian follicle development (Kadono-Okuda et al., 1994). Application of KK-42 produced instead maternal defects on fertilization, embryogenesis and hatching of silkworm larvae. These data argue for a role of ovarian ecdysteroids in progression of oocyte meiotic arrest and as ‘maternal’ ecdysteroids stored in the form of inactive ecdysteroid conjugates in the egg to regulate cuticulogenesis or morphogenetic events during embryogenesis (Lanot et al., 1989). In the silkmoth, however, contradictory results were obtained regarding correlation of ecdysteroid levels with cuticle formation in the embryo (Mizuni et al., 1981; Gharib and De Reggi, 1983; Gharib et al., 1983; discussed in Sonobe and Yamada, 2004). More recently, production of free 20E from pools of maternal ecdysteroid conjugates has been shown to be responsible for the developmental difference between diapausing and non-diapausing embryos (Makka et al., 2002). Although an ‘autocrine’ or ‘paracrine’ role for ecdysteroids during ovarian follicle development is not likely in the case of the silkmoth, more recent experiments have pointed to such a role for prostaglandins. Application of non-steroidal antiinflammatory drugs such as aspirin and indomethacin that block the production of prostaglandins by the cyclooxygenase enzyme, to ovarian follicles in culture results in arrest of follicle development (Machado et al., 2007). The arrest by aspirin and indomethacin can be reversed by exogenous application of prostaglandins and cAMP. Thus, as in mammalian models, prostaglandins may act through the activation of a membrane-bound G protein-coupled receptor (GPCR) and production of cAMP. However, prostaglandin signalling is unlikely to have a developmental role since they are required at all stages of follicle development investigated,

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from middle vitellogenesis to late choriogenesis. Of note is the observation that exogenous prostaglandins or cAMP can not rescue the developmental arrest induced by tebufenozide. Prostaglandins may play a ‘homeostatic’ role in ovarian follicle development (Machado et al., 2007). Recently, several genes that encode cytochrome P450 enzymes involved in the ecdysone biosynthetic pathway were identified in Drosophila and Bombyx (Warren et al., 2004; Gilbert and Warren, 2005; Namiki et al., 2005; Ono et al., 2006). It will be of interest to determine at which stages of oogenesis the cytochrome P450 genes are expressed in the silkmoth. Furthermore, FTZ-F1, besides its established role as a ‘competence’ factor in the regulation of developmental transitions from high to low ecdysteroid titers, is also known as the insect homolog of the nuclear receptor steroidogenic factor-1 (SF-1) which plays essential roles in the differentiation of steroidogenic organs (gonads and adrenals) in mammals (Val et al., 2003). Recently, a role for βFTZ-F1 was also proposed to regulate the ecdysteroidogenic activity in the prothoracic gland through modulation of the expression levels of ecdysteroidogenic cytochrome P450 enzymes (Parvy et al., 2005). It remains to be determined if a parallel situation exists also exists in the Bombyx ovary and that the expression of BmFTZ-F1 coincides with expression of ecdysteroidogenic genes such as phantom and disembodied.

5.3

The Silkmoth Paradigm: Applicability to Other Lepidopteran Insects?

In contrast to the silkmoth, ovarian development in many other lepidopteran insects is not initiated at the pupal stage in coordination with the metamorphic processes. In fact, ovarian development in lepidopteran insects is divided in four classes according to the extent of its coupling to metamorphic processes (review by Ramaswamy et al., 1997). The first class, to which Bombyx (Bombycidae: Bombycoidea) belongs, comprises those species where oogenesis is initiated and completed during the larval and pupal stages in concert with metamorphic processes that are orchestrated by 20E. Oogenesis (vitellogenesis) in these species can be inhibited by JH (Davis et al., 1990). Other lepidopterans belonging to this class include Hyalophora cecropia (Saturniidae: Bombycoidea), Malacosoma pluviale (Lasiocampidae: Bombycoidea) and Lymantria dispar (Lymantriidae: Noctuoidea) (Ramaswamy et al., 1997). In the second class of lepidopteran insects (example: Plodia interpunctella (Pyraloidea)), dependence of ovarian development on metamorphic events is reduced. In this moth, declining levels of ecdysteroids trigger vitellogenesis during pupation while completion of follicle development beyond vitellogenesis occurs before adult eclosion and is regulated by unknown factors (Shirk et al., 1992). A further reduction on metamorphic events occurs in the third class, exemplified by Manduca sexta (Sphingidae: Bombycoidea), Diatraea grandiosella (Pyralidae: Pyraloidea) and Choristoneura fumiferana (Tortricoidea): in those lepidopterans,

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vitellogenesis in the pharate adult occurs in the absence of ecdysteroids while completion of vitellogenesis and choriogenesis is regulated in the teneral adult by JH (Nijhout and Riddiford, 1974, 1979; Satyanarayana et al., 1994; Delisle and Cusson, 1999; Ramaswamy et al., 1997). In the last (fourth) class, oogenesis takes place in the adult stage and is exclusively regulated by JH (Ramaswamy et al., 1997). In these cases, ecdysteroids can block the gonadotropic action of JH while mating stimulates the production of eggs (Satyanarayana et al., 1992). Species belonging to this class include Pieris brassicae (Pieridae: Papilionoidea), the nymphalids (Papilionoidea) Polygonia c-aureum, Nymphalis antiopa and Vanessa cardui, Danaus plexippus (Danaidae: Papilionoidea) and the noctuids (Noctuoidea) Heliothis virescens, Helicoverpa zea and Pseudaletia unipuncta (Ramaswamy et al., 1997). From the examples described above, it appears that there is no clear correlation between phylogenetic relationships and class of ovarian development. Thus, the dependence (or independence) of ovarian development from metamorphic events orchestrated by 20E must have originated many times within the order of the Lepidoptera (Ramaswamy et al., 1997). It is possible that the mechanism by which ovarian development in lepidopteran insects becomes dependent on 20E during metamorphosis involves heterochronic shifts in the expression of the ecdysone receptor EcR/USP. Thus, species belonging from class 3 to 1 are expected to show progressive shifts of expression of EcR/USP (and, consequently, dependence on regulation by 20E) to earlier stages of ovarian development or oogenesis. Such shifts in temporal expression of the ecdysone receptor have been described in gall midges (Diptera: Cecidomyiidae) where early (first larval) versus late (last larval) expression of EcR/USP directs paedogenetic or metamorphic ovarian development, respectively (Hodin and Riddiford, 2000). Also in Drosophila it was observed that alterations in the timing of expression of EcR in the ovary can uncouple the process of ovarian differentiation from tissue differentiation in the rest of the animal (Hodin and Riddiford, 1998). In class 2 lepidopterans, such as Plodia interpunctella, it can be predicted that the regulatory cascade induced by a decline in ecdysteroid signalling occurs as in B. mori, but is shifted to earlier stages of oogenesis (vitellogenesis). On the other hand, it will also be interesting to investigate the expression pattern of the genes involved in the ecdysone regulatory cascade in representatives of classes 3 and 4 of lepidopteran ovarian development. Although oogenesis in these classes is independent of ecdysteroids produced by the prothoracic glands during metamorphosis, the ovaries synthesize ecdysteroids and ecdysteroid conjugates which accumulate in the eggs implicating a role in embryonic development as discussed above (Kaplanis et al., 1973; Bollenbacher et al., 1978; see further below). Investigation of the expression pattern of ecdysone-responsive genes during oogenesis in members of these classes could clarify whether ecdysteroids produced by the ovaries would also have an autocrine/paracrine function to regulate the progression of follicle development (as has been hypothesized in other insect species, most notably Drosophila, see further below).

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If ecdysteroids play a role in the regulation of oogenesis of class 3 lepidopterans, it can be expected that ecdysone agonists influence ovarian development after adult eclosion. This is observed in the codling moth, Cydia pomonella (Tortricidae: Tortricoidea), where application of tebufenozide and methoxyfenozide to adults results in reduction in fecundity (Sun et al., 2003a). Reduction in fecundity and/ or fertility after treatment by ecdysone agonists was also observed in other lepidopteran species such as Helicoverpa zea, Platynota idaeusalis and Spodoptera exigua (Smagghe and Degheele, 1994a, b; Carpenter and Chandler, 1994; Sun et al., 2003a). It was suggested that the reduced fecundity was due to interference with normal ovarian development such as failure of completion of choriogenesis (Sun et al., 2003a, b). EcR and USP are expressed in the adult ovary of Cydia, implicating the ovary as a direct target of ecdysone agonists, and the expression of EcR and USP is modulated by the ecdysone agonists (Sun et al., 2003a, b). However, the above studies did not investigate in detail which stages of oogenesis were affected by the ecdysone agonists and further work therefore needs to be done to elucidate the exact role of 20E in the regulation of oogenesis in Cydia. In the most studied lepidopteran (besides B. mori), Manduca sexta, the main ecdysteroid found in ovaries and eggs is 26-hydroxyecdysone 26-phosphate (Thompson et al., 1984, 1985b). As also hypothesized in other insects, this conjugate could serve as precursor for the release of free ecdysteroids to regulate developmental events during embryogenesis. Peak levels of free ecdysteroids (mainly 26-hydroxyecdysone in early embryos and 20,26-hydroxyecdysone in late embryos) have been observed to coincide with the deposition of serosal and larval cuticle (Warren et al., 1986; Dorn et al., 1987) but it is not clear whether these represent the biologically active ecdysteroids (Lanot et al., 1989). In vitro experiments have shown that α-ecdysone, 20E and makisterone A promote elongation and segmentation of the embryonic germ band in Manduca although the concentrations of these ecdysteroids during Manduca embryogenesis always remained relatively low (Warren et al., 1986; Lanot et al., 1989). Interestingly, Manduca eggs also contain a non-ecdysteroid conjugate, 5-pregnen3β,20β-diol glucoside, of which the steroid moiety correspond to a C19 steroid one biosynthetic step away from pregnenolone, the precursor of steroid hormones in vertebrates (Thompson et al., 1985a). However, this compound is located in high quantities on the surface of the egg, where it probably serves a protective role as an antibacterial or antifungal reagent or as a feeding deterrent against predators (Thompson et al., 1985a; Meinwald et al., 1985). In the noctuid Spodoptera frugiperda (class 4), ecdysteroids produced by the ovary seem to have acquired a function that is normally observed in dipteran insects. While JH regulates the formation of vitellogenic follicles in this species, it was also observed that injection of 20E into decapitated female adults results in the production of vitellogenin by the fat body (but not the uptake of vitellogenin by the ovary; Sorge et al., 2000). Thus, besides stimulation of vitellogenin uptake by the ovary, JH seems to stimulate the production of ecdysteroids which are released in the hemolymph and stimulate vitellogenin synthesis in the fat body, similar to dipteran insects (see further below).

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Control of Oogenesis by the Ecdysone Regulatory Pathway in Aedes aegypti and Drosophila melanogaster

The two dipteran species from which most is known regarding the role of ecdysteroids in the regulation of oogenesis are the fruitfly, Drosophila melanogaster, and the yellow fever mosquito, Aedes aegypti. However, knowledge in both species differs regarding the processes that are affected by ecdysteroids. In Drosophila, the availability of a large array of genetic tools has allowed the analysis of the function of genes of the ecdysone regulatory hierarchy during follicle development in the ovary (Buszczak et al., 1999; Carney and Bender, 2000). In the mosquito, on the other hand, research has focused on the molecular mechanism of the regulation of the production of yolk protein precursors in the fat body by ecdysteroids produced by the ovary (Raikhel et al., 1999, 2005; Li et al., 2000). By contrast, relatively little is known regarding the role of the ecdysteroid regulatory cascade in follicle maturation in the ovary of the mosquito.

5.4.1

Aedes aegypti

In the anautogenous mosquito Aedes aegypti, vitellogenesis and follicle maturation occur synchronously following a blood meal (Raikhel and Lea, 1990). At the time of adult eclosion, each ovariole consists of the germarium and one follicle. JH is responsible for the maturation of the primary follicles in the ovary and the fat body to a resting stage at which they become competent to respond to a blood meal and initiate vitellogenesis and yolk protein precursor synthesis, respectively (Bownes, 1986; Pierceall et al., 1999). Following a blood meal, ecdysteroid synthesis, principally of α-ecdysone, is initiated in the ovary (Hagedorn, 1985), presumably in the follicular epithelium that surrounds the oocyte/nurse cell complex of the first follicle, and ecdysteroids accumulate in the hemolymph where they stimulate the first cycle of yolk protein precursor synthesis in the fat body (Deitsch et al., 1995; Zhu et al., 2000). Other functions associated with ecdysteroids during mosquito follicle maturation include the formation of the vitellin membrane envelope in the primary follicle (Lin et al., 1993) and the separation of the secondary follicle from the germarium (Beckemeyer and Lea, 1980). Stimulation of ecdysone synthesis in the ovary following a blood meal occurs through the action of neurosecretory signals from the brain and the gut, including the ovary ecdysteroidogenic hormone (OEH), a neuroparsin homolog, and insulin-like peptides (Brown et al., 1998; Riehle and Brown, 1999; Badisco et al., 2007). The action of insulin-like peptides in the mosquito ovary has been shown to involve a conserved signaling pathway that includes an insulin receptor homolog, phosphatidyl-inositol 3-kinase and protein kinase B (Graf et al., 1997; Riehle and Brown, 1999, 2003; Wu and Brown, 2006). To which extent the transduction pathways of OEH and insulin-like peptides interact is unknown at present.

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The classical ecdysteroid regulatory cascade that has been elucidated first during molting and metamorphosis in Drosophila seems to be reiteratively used during the vitellogenic cycle of the fat body in Aedes (Raikhel et al., 1999, 2003). Significant progress has been achieved in the understanding of the regulation of the transcription of the vitellogenin gene by the ecdysteroid regulatory cascade in the fat body as well as the mechanisms that regulate the competence of the fat body to respond to ecdysone and the termination of the vitellogenic response (Martín et al., 2001; Kokoza et al., 2001; Zhu et al., 2000, 2003a, b, 2006; Sun et al., 2005; review by Raikhel et al., 2005). What is less understood, however, is whether the same regulatory mechanisms are also operational in the primary follicles during the first vitellogenic cycle when they progress through previtellogenic growth, resting stage, vitellogenesis and choriogenesis concomitantly with fluctuating titers of JH and 20E. Nevertheles, although less extensively studied than the fat body, data exist with respect to the expression of ecdysone-responsive genes in the ovary of the mosquito and the possible functional significance of these will be discussed below. Although expression patterns were studied using whole ovaries, it can be inferred that these reflect primarily changes in the primary follicle as it undergoes dramatic changes during the first vitellogenic cycle. Although the mosquito ovary traditionally is not considered a target for 20E (Bownes, 1986), several genes that are historically implicated in the ecdysone response such as E75, HR3, E74 and BR-C are expressed in the ovary during the vitellogenic cycle (Pierceall et al., 1999; Kapitskaya et al., 2000; Sun et al., 2002; Chen et al., 2004). In addition, the expression of EcR mRNA increases at the initiation of vitellogenesis concomitantly with a switch in usp isoform expression (Cho et al., 1995; Kapitskaya et al., 1996, 2000; Wang et al., 2000). Besides the traditional ecdysone-responsive genes, also the B isoform of the mosquito orphan nuclear receptor HNF-4 is induced in ovarian tissue following a blood meal (Kapitskaya et al., 1998). Possible target genes for EcR/USP and the early ecdysone-responsive genes in the ovary include the components of the machinery for uptake of yolk protein precursor genes such as the vitellogenin receptor, clathrin heavy chain, and lipophorin receptor (Sappington et al., 1995; Kokoza et al., 1997; Cho and Raikhel, 2001; Cheon et al., 2001; Seo et al., 2003). In contrast to the regulation of the vitellogenin gene in the fat body, however, nothing is known whether EcR/USP and early ecdysone-responsive gene products bind to promoter elements of the target genes and how they interact with ovary-specific factors to stimulate gene expression during vitellogenesis. It can also be pointed out that the vitellogenin receptor, clathrin heavy chain and lipophorin receptor are expressed in the oocyte/nurse cell complex and that candidate target genes in the follicular epithelium remain to be identified. Finally, one isoform of the classical ecdysone-responsive genes, E74A, is likely involved in the termination of vitellogenesis in both fat body and ovary. In the ovary, its expression coincides with the synthesis of the vitelline envelope in the terminal follicle (Sun et al., 2002). In summary, the data indicate that genes historically involved in the regulation of the ecdysone response have an expression pattern in the ovary consistent with

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those of regulators of vitellogenesis. The expression pattern also follows the rise and fall of ecdysteroid in the haemolymph and therefore could indicate regulation by ecdysteroids in similar fashion as in the fat body. Recently, it was observed that the stimulation of expression of yolk protein precursor genes by 20E in the fat body requires the target-of-rapamycin (TOR) pathway that mediates nutrient (amino acid) signaling (Attardo et al., 2003; Hansen et al., 2004). Whether this pathway is also functional in the ovary and interacts with the 20E regulatory cascade remains to be investigated. While ecdysone produced by the ovary in the mosquito is secreted in the hemolymph to act as a hormone to regulate vitellogenesis, the possible production of ecdysteroid conjugates by the ovary, as observed in other insect species, that accumulate in the oocytes and are used in egg development has not received much attention. The site of ecdysteroid synthesis in the vitellogenic ovary also remains to be determined. Since many ecdysteroidogenic enzymes have been identified in Drosophila and Bombyx (Gilbert and Warren, 2005) and the genome sequence of Aedes has become available recently (Nene et al., 2007), identification of Aedes homologs should also be straightforward and allow determination of their expression during follicle maturation in the ovary. Finally, it would be interesting to correlate the appearance of mRNAs of ecdysteroidogenic enzymes with the expression of putative transcriptional regulators such as FTZ-F1 (Parvy et al., 2005). Control of vitellogenin synthesis in the fat body by ecdysteroids produced by ovarian tissue is conserved in anautogenous flies that develop eggs in batches and require a protein meal for egg development such as the house fly, Musca domestica, and the blowflies Calliphora vicina, Neobellieria bullata and Phormia regina (Adams et al., 1985; Huybrechts and De Loof, 1982; Briers and Huybrechts, 1984). Similar to the mosquito, JH is the predominant hormone after eclosion that stimulates previtellogenic follicle development in the ovary and prepares the fat body to respond to ecdysteroids to synthesize large quantities of yolk protein precursors (Adams et al., 1985). Protein meal uptake subsequently induces ovarian growth and production of ecdysteroids that stimulate vitellogenin synthesis in the fat body. Other roles for 20E produced by the ovary in flies include the control of parturition in the tsetse fly, Glossina fuscipus (Robert et al., 1986), and the production of pheromone in Musca (Adams et al., 1984). Interestingly, 20E can also induce vitellogenin synthesis in male Neobellieria, Phormia and Lucilia (Huybrechts and De Loof, 1977, 1982) and this has also been observed in Drosophila (Bownes et al., 1983).

5.4.2

Drosophila melanogaster

In species continuously laying eggs, such as Drosophila, follicle development in the ovarioles occurs asynchronously and its progression is modulated under the influence of external environmental factors such as food intake and mating (Spradling, 1993). Early physiological studies on Drosophila oogenesis focused on the regulation of vitellogenin synthesis in the fat body and vitellogenin uptake by

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the ovary (Postlethwait and Handler, 1979; Schwartz et al., 1985; Bownes, 1989; Hagedorn, 1989). More recently, however, with the availability of genetic tools and the identification of genes involved in the biosynthesis as well as the transduction of 20E, it became possible to analyze the action of 20E within the developing ovary in a manner that has hitherto not been possible in other insects. The discussion of oogenesis in Drosophila is divided in three parts. In the first part, experiments will be considered that relate to physiological and molecular effects of 20E on the process of oogenesis. The second part involves the phenotypic changes in ovarian follicle development following genetic manipulation of genes involved in the 20E biosynthetic and regulatory pathways. A final part will address the subject of the accumulation of maternal ecdysteroids in the developing oocytes of Drosophila.

5.4.2.1

Physiological Experiments

Ecdysteroids play both positive and negative roles in the regulation of oogenesis in Drosophila (Riddiford, 1993b; Soller et al., 1999). The positive effects of JH and 20E on the development of immature ovaries immediately after eclosion are probably related to their effects on the completion of metamorphosis. In isolated abdomens or decapicitated females prepared immediately after adult eclosion, JH stimulates yolk protein synthesis in both fat body and ovary while the stimulatory action of 20E is restricted to the fat body only (Jowett and Postlethwait, 1980; Postlethwait and Shirk, 1981). The action of JH on the fat body may be indirect since it stimulates ecdysteroid production in the ovary (Schwartz et al., 1989). On the other hand, the effects of 20E on yolk protein expression in the fat body also seem to be indirect and therefore may involve the classical ecdysteroid regulatory cascade as observed in the mosquito (Bownes et al., 1987, 1996). Also in diapausing female adults, which are arrested at previtellogenic stages, termination of diapause and induction of vitellogenesis is achieved after injection of 20E (Richard et al., 1998). Based on this and other observations, it was proposed that ecdysteroids produced by the ovary or other sources (Bownes, 1989) stimulate yolk protein synthesis by the fat body as well as yolk protein uptake from the ovary immediately after eclosion when follicles are at the previtellogenic stages (Richard et al., 1998, 2001). During later stages of oogenesis, alternatively, 20E has negative effects on ovarian follicle development. The critical period at which 20E exerts its negative effects is at stages 8 and 9 of follicle development which correspond to the initiation of vitellogenesis (Drummond-Barbosa and Spradling, 2001). In adverse physiological conditions that do not support oocyte maturation such as starvation, stress or absence of mating, ecdysteroid titers rise and cause nurse cell apoptosis and follicle degeneration in stage 8 and stage 9 follicles (Soller et al., 1999). In beneficial physiological conditions, on the other hand, the effects of 20E are counterbalanced by increased production of JH which stimulates initiation of vitellogenesis (Soller et al., 1999; Gruntenko et al., 2003).

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In the case of mating, its positive effects on progression of follicle development are mediated by the transfer of the sex peptide, a product of the male accessory glands, to the female (Chen et al., 1988; Kubli, 1996). The sex peptide acts on the corpora allata to stimulate the production of JH (Moshitzky et al., 1996) which counteracts the apoptotic effect of 20E on vitellogenic follicles (Soller et al., 1999). Similarly, the availability of abundant nutritional resources results in the initiation of vitellogenesis, presumably through the stimulation of the insulin pathway (Leevers, 2001), while during starvation, apoptosis of stage 8 and 9 follicles is initiated (Terashima and Bownes, 2004, 2005). Nutritional stimulation of vitellogenesis is protected by JH and counteracted by 20E (Terashima et al., 2005). Thus, a correct balance between JH and ecdysteroids is essential for the progression of vitellogenic follicle development in Drosophila (Soller et al., 1999; Gruntenko et al., 2003; Terashima et al., 2005). During the initiation of the apoptosis response by 20E or nutritional stress, switches occur in the expression of the ecdysone-responsive genes at stages 8 and 9 of follicle development (Terashima and Bownes, 2004, 2006). Increases in ecdysteroid concentrations result in increases in expression of the Z2 and Z3 isoforms of Broad-Complex in the cells of the follicular epithelium (Terashima and Bownes, 2004). The increase in Z2 and Z3 results in a decrease in E75B expression and an increase in E75A levels which induces apoptosis in the nurse cells of stage 8 and 9 follicles (Terashima and Bownes, 2006) (Fig. 5.2).

5.4.2.2

Genetic Analysis

The availability of mutants of genes involved in the ecdysone biosynthetic and regulatory pathways as well as the genetic tools to alter their expression in developing follicles has allowed an assessment of the role of ecdysteroids during oogenesis in Drosophila which hitherto has not been possible in other insects. As outlined, the discussion of genes of the ecdysone biosynthetic and regulatory pathways will be discussed separately. A third part discusses the activation of ecdysone ‘sensors’ during oogenesis in transgenic animals. (a) Genes involved in ecdysteroid biosynthesis Temperature-sensitive ecdysoneless1 (ecd1) mutants that are characterized by reduced ecdysteroid levels at the restrictive temperature show a developmental arrest at the onset of vitellogenesis during ovarian follicle development, indicating that 20E may be required for the progression of oogenesis beyond stage 8 (Audit-Lamour and Busson, 1981; Walker et al., 1987). Analysis of ovaries with non-conditional ecd-/ ecd- clones show that ecdysoneless is required in the follicle cells for appropriate follicle formation in the germarium (Gaziova et al., 2004). On the other hand, ecd-/ ecd- germline clones show developmental arrest prior to vitellogenesis due to nurse cell degeneration. Molecular characterization of ecdysoneless shows that it encodes a conserved protein with unknown function that has broad expression in both ecdysteroidogenic and non-ecdysteroidogenic tissues (Gaziova et al., 2004). In ovaries, ecd

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Fig. 5.2 Model to explain the hierarchy of ecdysone response genes regulating apoptosis of stage 8 and 9 follicles in Drosophila melanogaster. Upper Panel: Complete nutrition induces normal development of follicles during oogenesis. In this case, just the Z1 isoform of BR-C is expressed in the follicle cells at stage 8. E75B suppresses E75A expression to prevent apoptosis. Middle Panel: Injection of 20E induces apoptosis in stage 8 and 9 follicles. 20E injection results first in induction of the Z2 and Z3 isoforms of BR-C which in turn decrease E75B and increase E75A expression. While E75B is an apoptosis inhibitor, E75A is an apoptosis inducer. Lower Panel: Starvation induces apoptosis in stage 8 and 9 follicles. During starvation, ecdysone concentrations increase and the Z2 and Z3 isoforms of BR-C become expressed in the follicle cells to suppress E75B and activate E75A expression. The increase in E75A results in induction of apoptosis. JH can counteract the effects of starvation by interference with the increase in ecdysone concentration and by stimulation of the expression of E75B (Reprinted from Terashima and Bownes, (2006). E75A and E75B have opposite effects on the apoptosis/development choice of the Drosophila egg chamber. Cell Death Differ. 13, 454–464. With permission from Macmillan.) (See Color Plates)

is expressed in both nurse cells and follicle cells (Gaziova et al., 2003). However, although it was proposed that the ecdysoneless gene product is involved in the regulation of ecdysteroid biosynthesis by facilitating the translocation of sterol precursors between different subcellar compartments (Warren et al., 1996), ecd clearly affects other cellular processes besides ecdysteroidogenesis. Thus, the observed phenotype of oogenesis arrest could be caused by the involvement of the gene in other cellular processes besides ecdysteroidogenesis. Also mutations in the gene dare, which encodes the Drosophila homolog of adrenodoxin reductase, a mitochondrial protein that transports electrons to cytochrome P450 enzymes, including ecdysteroidogenic enzymes, cause developmental arrest of oogenesis at stages 8–9 (Buszczak et al., 1999; Freeman et al., 1999). Expression of dare occurs in the nurse cell complex (Freeman et al., 1999). Although it is tempting to speculate that the phenotypic effects in dare mutants are due to decreased

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ecdysteroidogenesis, the disruption of other cellular processes caused by dysfunction of other cytochrome P450 enzymes, however, can not be discounted. Recent molecular characterization of the Halloween gene family identified several members of the cytochrome P450 enzymes that are involved in 20E biosynthesis (Gilbert and Warren, 2005). These genes include disembodied (dib) that encodes the C22-hydroxylase, shadow (sad) encoding the C2-hydroxylase, shade (shd) corresponding to the C20-hydroxylase, phantom (phm) that produces the C25-hydroxylase and spook (spo) that probably catalyzes an early step in ecdysteroid synthesis (Chávez et al., 2000; Warren et al., 2002; Petryk et al., 2003; Warren et al., 2004; Namiki et al., 2005; Ono et al., 2006). For all of these genes, their expression in the ovary has been reported while for some mutant ovarian phenotypes have been described. Thus, the genes shd, phm and sad are expressed in both nurse cells and follicle cells while expression of dib and spo is restricted to the follicle cells. In all cases highest expression levels are observed in later stages of oogenesis, from stage 8 until stage 11. While it was observed that dib is not required in the germline for progression of oogenesis (Chávez et al., 2000), mutants of shd and spo, interestingly, are arrested in oogenesis at the initiation of vitellogenesis (Petryk et al., 2003; Ono et al., 2006). Because it is expected that the spo and shd gene products have very specific functions involved solely in ecdysteroid biosynthesis, these data therefore indicate a requirement for 20E production in ovarian tissue to regulate progression towards vitellogenesis. Moreover, in this case, 20E must act as a paracrine or autocrine factor since its effects are exerted within the same organ where it is produced. Other genes involved in ecdysteroidogenesis are Start1, that encodes a putative cholesterol transporter and neverland (nvd), encoding a Rieske-domain protein that probably catalyzes the conversion from cholesterol to 7-dehydro-cholesterol, the first step of ecdysteroidogenesis (Roth et al., 2004; Yoshiyama et al., 2006). As detected by in situ hybridization, both genes are expressed in the nurse cells. In the case of Start1, it is speculated that its presence in the nurse cells corresponds to its production prior to transport and storage in the oocyte as maternal mRNA (Roth et al., 2004). Transcription factors involved in the regulation of the expression of ecdysteroidogenic enzymes in the prothoracic glands include the products of the genes without children (woc) and bFTZ-F1 (Warren et al., 2001; Parvy et al., 2005). However, their expression pattern or functional analysis during oogenesis has not been described so far. (b) Genes involved in the ecdysone regulatory pathway The components of the ecdysone receptor heterodimer, EcR and usp, as well as the early gene products E75, E74 and BR-C are expressed during Drosophila oogenesis. EcR as well as USP protein can be detected from early stages in the germarium until the completion of oogenesis (Khoury-Christianson et al., 1992; Buszczak et al., 1999; Carney and Bender, 2000). High expression levels of EcR protein (B1 and B2 isoform) are observed in the border cells, a group of specialized cells that migrate from the anterior pole of the follicular epithelium through the nurse cell complex during stages 9 and 10 and constitute precursor cells of the micropyle channel (Buszczak et al., 1999; Bai et al., 2000). In situ hybridization

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also detects expression of E74 and E75, especially in the nurse cells and follicle cells of follicle stages 8 until 10 (Buszczak et al., 1999). A bimodal expression pattern is observed for BR-C protein, with an early expression pattern (stages 5–6) separated from a late pattern (stages 10–13) (Deng and Bownes, 1997; Buszczak et al., 1999; Tzolovsky et al., 1999). Changes in ecdysteroid levels such as those induced by temperature shifts in ecdysoneless1 mutants or during ovarian cultures in the presence of 20E modulate the expression of E75 and BR-C, indicating the activation of the ecdysone regulatory cascade during oogenesis (Buszczak et al., 1999). As noted above, it was observed also that administration of 20E can mimic nutritional stress in Drosophila resulting in changes in BR-C and E75 isoform expression during stages 8 and 9 followed by induction of apoptosis (Terashima and Bownes, 2004, 2006) (Fig. 5.2). However, it is clear that E75 and BR-C expression during oogenesis can be regulated by other pathways besides the involvement of 20E. Spatial patterns of E75 mRNA expression in the follicular epithelium are modulated by the epidermal growth factor receptor (EGFR) pathway (Buszczak et al., 1999). In the case of the late BR-C expression pattern, both EGFR and Decapentaplegic/transforming growth factor β (Dpp/TGFβ) pathways cooperate to restrict BR-C expression to two dorsolateral patches of follicle cells that correspond to the regions from which the dorsal appendages, two specialized eggshell structures, are derived (Deng and Bownes, 1997; Tzolovsky et al., 1999). Germline clones of EcR, E75 and E74 cause developmental arrest and degeneration of follicles (stages 6–7 for EcR; stages 8–9 for E75 and E74) (Buszczak et al., 1999; Carney and Bender, 2000). Because the similarity in phenotype with mutations that cause a defect in ecdysteroid synthesis, a model was proposed in which ecdysteroids produced at the beginning of vitellogenesis act in a paracrine or autocrine manner (i.e., within the developing follicle) to activate the ecdysone regulatory cascade and progression of oogenesis (Buszczak et al., 1999; Freeman et al., 1999). In addition, EcR (B1 and B2 isoforms) seems to be implicated in border cell migration since mutations in the taiman (tai) gene, that encodes a p160 transcriptional co-activator of EcR, cause defects in border cell migration (Bai et al., 2000; Montell, 2001). Also BR-C plays a role at the initiation of vitellogenesis to control progression of oogenesis (Huang and Orr, 1992; Terashima and Bownes, 2004, 2006). Nutritional stress or high levels of ecdysteroids stimulate expression of the Z2 and Z3 isoforms of BR-C which in turn induce apoptosis through the modulation of expression levels of E75A and E75B (Terashima and Bownes, 2004, 2006) (Fig. 5.2). In addition, BR-C regulates other functions during late oogenesis such as chorion gene amplication, chorion gene expression and dorsal appendage formation but these functions are clearly independent of 20E signalling (Huang and Orr, 1992; Deng and Bownes, 1997; Tzolovsky et al., 1999). Interestingly, also mutations in the Methoprene-tolerant (Met) gene that encodes a bHLH-PAS transcription factor that could possibly function as a JH receptor (Wilson and Ashok, 1998), cause defects in vitellogenic development and genetic interactions were observed between alleles of BR-C and Met, indicating that they may operate in the

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same genetic pathway (Wilson et al., 2006). Given the antagonism between JH and ecdysteroids to regulate progression of vitellogenesis or apoptosis, both pathways could converge on the regulation of the expression of particular BR-C isoforms to decide the developmental fate of early vitellogenic follicles (see discussion above; Terashima and Bownes, 2004, 2006). In contrast to EcR, E75, E74, and BR-C mutants, usp mutants show no defects in progression of oogenesis at the initiation of vitellogenesis (Oro et al., 1992). On the other hand, genetic studies have shown that usp is required for fertilization and chorion formation (Perrimon et al., 1985; Oro et al., 1992). The fertilization defect is likely due to the failure of border cell migration in usp mutants (Bai et al., 2000; see also above). Regarding the chorion defect, it was shown that USP binds to essential elements in chorion gene promoters (Shea et al., 1990; Khoury-Christianson et al., 1992). However, genetic analysis has shown that the usp+ function is required in the germline, and not the follicle cells (where the chorion genes are produced) to produce normal chorion (Oro et al., 1992). Thus, it was proposed that USP may not act directly as a transcriptional of the chorion genes but could function in the germline to trigger the release of a signal that acts on the follicular cells to promote chorion gene expression (Oro et al., 1992). (c) Detection of EcR/USP signalling during oogenesis If active 20E signalling occurs at particular stages during oogenesis, it should be detected by ecdysone reporters at these particular stages such as EcRE-lacZ or EcRE-GFP in transgenic animals (Kozlova and Thummel, 2003). Using these constructs, reporter gene activity is restricted to the ‘squamous’ follicle cells, that overly the nurse cells, and the border cells at stages 10–14 of oogenesis (late vitellogenesis and choriogenesis; Hackney et al., 2007). Another reporter system consisting of heat-shock promoter-Gal4-EcR or heat-shock-promoter-Gal4-USP expression constructs in combination with a Gal4-lacZ reporter shows similar active EcR/USP signalling in subsets of follicular cells at the same late stages of oogenesis. Besides high activity in the ‘squamous’ follicular cells and the border cells, intermediate to low levels of activity are observed in the centripetally migrating and main body follicle cells, respectively, using the more sensitive Gal4-based detection system. However, no reporter gene activity was observed during earlier stages, despite the broad expression of EcR and USP at these stages (see above). Interestingly, while EcR reporter activity is sensitive to exogenous ponasterone A, particularly during early stages of oogenesis, it is also negatively regulated by the EGFR-MAPK-Ras signalling pathway that defines the dorsal-anterior fate of the follicular epithelium during late oogenesis (Riechmann and Ephrussi, 2001). Expression of dominant negative EcR (Cherbas et al., 2003) in the follicular epithelium results in abnormal follicular cell migrations (including border cell migration, see above) during stage 10 and aberrant dorsal appendage formation, due to misregulation of genes that encode epithelial junction components, such as DE-cadherin, discs large and armadillo. Expression of dominant negative EcR results also in reduction of chorion gene expression and chorion gene amplification. Taken together, these data indicate a role for EcR signalling during late oogenesis

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in the follicular epithelium to regulate its morphogenetic movements at the end of vitellogenesis and the synthesis of the chorion during choriogenesis (Hackney et al., 2007). The role for EcR signalling in dorsal appendage formation, chorion gene expression and chorion gene amplification coincides with the requirement for BR-C function (Huang and Orr, 1992; Deng and Bownes, 1997; Tzolovsky et al., 1999; see above). However, surprisingly, expression of dominant negative EcR does not alter expression of BR-C (Hackney et al., 2007). As noted above, the ‘late’ functions of BR-C are considered to be ecdysteroid-independent (Deng and Bownes, 1997). In summary, EcR, usp and early genes such as E74, E75 and BR-C seem to have both ecdysteroid-dependent and ecdysteroid-independent functions during Drosophila oogenesis. Ecdysteroid-dependent functions occur earlier during oogenesis at stages 8 and 9 when concentration-dependent expression of specific isoforms of E75 and BR-C will direct the developmental decision between progression through vitellogenesis or follicle degeneration. At later stages of oogenesis, on the other hand, EcR, usp and BR-C are involved in the regulation of the morphogenetic movements that mark the end of vitellogenesis and choriogenesis as well as the synthesis of the chorion (stages 10–14). At those stages, the activity of the EcR/USP complex and BR-C may be regulated by other signalling pathways such as EGFR-MAPK-Ras or Dpp/TGFβ.

5.4.2.3

Maternal Ecdysteroids and Regulation of Embryogenesis

The ecdysteroid content in freshly oviposited Drosophila eggs is low but increases after 2 h and reaches a maximum before mid-embryogenesis (Maróy et al., 1988). Analogous to the function of ecdysteroids stored in eggs of locusts (Lanot et al., 1989; see further below), it was proposed that the rise in ecdysteroids during Drosophila embryogenesis could be achieved by the hydrolysis of maternal fatty acid ecdysteroid conjugates (Bownes et al., 1988). A key role in this process would be played by the yolk proteins which have significant sequence similarity to part of triacylglycerol lipase and were shown to bind ecdysteroid fatty acid conjugates. In this model, release of ecdysteroids at a specific period of embryogenesis would be achieved by breakdown of yolk proteins (Bownes et al., 1988). However, it was also observed that expression of the ecdysteroidogenic enzymes encoded by the genes sad (C2-hydroxylase) and dib (C22-hydroxylase) is induced in the embryonic epidermis prior to the differentiation of the prothoracic gland, indicating that de novo ecdysteroid synthesis contributes to the rise of ecdysteroid titers prior to mid-embryogenesis (Warren et al., 2002). Whatever their source, ecdysone signalling in the Drosophila embryo is involved in the regulation of the morphogenetic movements such as germ-band retraction, head-involution and dorsal closure during mid-embryogenesis as well as the deposition of the cuticle during late embryogenesis. This has been shown recently through over-expression of dominant-negative EcR during early stages of embryogenesis (Kozlova and Thummel, 2003). Similar phenotypes are observed in mutants for the genes sad, dib and spo that encode ecdysteroidogenic enzymes (Chávez et al., 2000; Warren et al., 2002).

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Ecdysone and Oogenesis in Orthoptera and Dictyoptera

Locusts and cockroaches are well known as classic physiological models of ovarian ecdysteroid production and ecdysteroid-regulated reproductive processes. At the molecular level, on the other hand, the hemimetabolous insects have lagged behind the holometabolous insects represented by the lepidopterans Manduca sexta and Bombyx mori and the dipterans Drosophila melanogaster and Aedes aegypti. Only recently, homologs of EcR and usp or the classical early ecdysteroid-responsive genes E75, BR-C and HR3 were isolated in representatives of orthopterans, dictyopterans and hemipterans (Hayward et al., 1999, 2003; Maestro et al., 2005; Cruz et al., 2006, 2007; Erezyilmaz et al., 2006). In the cockroach, Blatella germanica, the expression pattern of EcR during ovarian development was determined and functional studies established essential roles for the ecdysone regulatory pathway during oogenesis.

5.5.1

Locusts

Locusts such as Locusta migratoria and Schistocerca gregaria have served for a long time as a paradigm for the production of ovarian ecdysteroids, their storage in eggs and their reiterative use during embryonic development (Sall et al., 1983; Lanot et al., 1989; Dinan, 1997; Tawfik et al., 1999). In the classical model, ovarian development and vitellogenin synthesis in the fat body are dependent on JH secretion by the corpora allata while large amounts of ecdysteroids are synthesized by the ovary at the end of the gonadotropic cycle after the completion of vitellogenesis (Lagueux et al., 1977). The ecdysteroids are synthesized by the cells of the follicular epithelium (Glass et al., 1978; Goltzene et al., 1978) and converted mostly (>95%) to polar conjugates (Gande and Morgan, 1979; Dinan and Rees, 1981). Most of the ovarian ecdysteroids (>95%) are confined to the terminal oocytes and only low amounts accumulate in the hemolymph (Lagueux et al., 1977). Thus, in locusts, a clear role for ecdysteroids is implicated in the regulation of terminal oogenic events such as the reinitiation of meiosis and their accumulation in eggs indicates a functional role as maternal ecdysteroids involved in the regulation of embryonic events. In Locusta migratoria, titers of free (unconjugated) ecdysone and 2-deoxyecdysone peak at the posterior pole of the eggs at the times when the nuclear events of reinitiation of meiosis occur (first and second reinitation at ovulation and egg-laying, respectively; Lanot et al., 1987). Furthermore, initiation of meiosis can be stimulated by 1–100 µM ecdysone both in immature oocytes and in oocytes obtained from animals with a modified sterol profile and reduced levels of endogenous ecdysteroids (Lanot et al., 1987, 1988; Costet et al., 1987). In Locusta, the maternal conjugates of ecdysone and 2-deoxy-ecdysone are hydrolyzed during embryonic development resulting in distinct peaks in the free ecdysteroid titer (Lagueux et al., 1984; Rees and Isaac, 1984). In Locusta,

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four peaks in free ecdysteroid titer, consisting mainly of ecdysone and 2-deoxyecdysone, are observed that can be correlated to the developmental events of appearance of coelomic somites and appendage anlagen (day 3), blastokinesis (day 4), dorsal closure (day 5) and appearance of pigmentation (day 9) (Lagueux et al., 1979). The ecdysteroid peaks also correlate with the deposition of the serosal cuticle and three embryonic cuticles by the embryonic tissues. Correlations between free ecdysteroid titer and similar embryonic developmental events were also made in Schistocerca eggs (Scalia et al., 1987; Sbrenna et al., 1989; Tawfik et al., 1999). Thus, embryonic ecdysteroids in locusts likely are involved in morphogenetic and cuticular events during embryogenesis as was observed in Drosophila (Kozlova and Thummel, 2003). However, maternal ecdysteroid conjugates are believed to serve as precursors of active ecdysteroids only during the first half of embryonic development, i.e. before the differentiation of the embryonic prothoracic glands (Lagueux et al., 1979). The ecdysteroid conjugates that are deposited in the locust eggs are metabolized during embryonic development with changes occurring on both ecdysteroid genins and conjugating moieties (Sall et al., 1983). In Locusta, the main maternal ecdysteroid conjugates deposited in the egg are C22 adenosinemonophosphate (AMP) esters of 2-deoxyecdysone and C22 N6-(isopentenyl)-AMP esters of ecdysone (Tsoupras et al., 1983; Sall et al., 1983). In the course of embryonic development, the release of free ecdysteroid seems to be a minor pathway while the major pathway consists of the replacement of the initial set of polar conjugates by new types such as C22 phosphate and C3 phosphate esters in combination with conversions on the ecdysteroid moiety such as the epimerization of the 3β-hydroxyl group (Sall et al., 1983). A similar pattern of metabolic conversions during embryonic development was also observed in Schistocerca gregaria where C22 phosphates are the major conjugates that accumulate initially in the oocyte and the egg (Isaac et al., 1983; Rees and Isaac, 1984). It was proposed that the physiological role of the change in conjugate type reflects the “status” of the ecdysteroid conjugate as precursor of active ecdysteroids (C22 AMP and C22 phosphate esters) or as final inactive end product (C3 phosphate and C3 epi-3-phosphate esters) (Hoffmann et al., 1980). The deposition of maternal ecdysteroid conjugates is considered a determinant for the development of the gregarious character of the locust. Gregarious locusts accumulate larger amounts of ecdysteroids and differences are observed in the composition of the amounts of ecdysone, 20E, 2-deoxy-ecdysone and 26-hydroxyecdysone between solitary and gregarious embryos (Tawfik et al., 1999). More recently, the role for ecdysteroids acting exclusively within the ovary or as maternal ecdysteroids in Locusta was challenged following the isolation of an ecdysiotropic factor from the brain, ovary maturating parsin (OMP), that stimulates both vitellogenesis and ecdysone synthesis by the ovary (Girardie and Girardie, 1996). In these studies, it was found that part of the stimulatory effect of OMP on vitellogenin synthesis by the fat body was probably mediated by its ability to stimulate the production of ecdysone by the ovary. Further studies established that Locusta OMP appears to have two distinct roles that seem to be mediated by distinct domains of the peptide: the N-terminal domain, directly acting on fat

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body tissue, has a protective effect on the stability of vitellogenin mRNA while the ecdysteroidogenic effect on the ovary is carried out by the C-terminal domain (Girardie et al., 1998a). Thus, OMP, probably partially acting via 20E, may be necessary in conjunction with JH for complete vitellogenesis in Locusta. While the role of OMP and 20E in stimulating vitellogenesis seems to be conserved in Schistocerca (Girardie et al., 1998b), no stimulatory roles of ovarian ecdysteroids on ovarian growth or hemolymph vitellogenin titers were observed in the lubber grasshopper, Romalea microptera (Hatle et al., 2003). Thus, further investigations are necessary to establish the role of OMP-like peptides and ecdysteroids during vitellogenesis in other orthopterans. Although homologs of both EcR and RXR/USP were isolated in Locusta migratoria (Hayward et al., 1999, 2003), their expression patterns or roles during oogenesis have not been investigated in detail.

5.5.2

Crickets

Studies on other orthopterans, such as crickets, have revealed surprising differences to the pattern found in locusts with respect to ecdysteroid synthesis by the ovary (Whiting et al., 1997). In the house cricket, Acheta domesticus, ecdysteroids accumulate in the ovary as apolar instead of polar conjugates (mainly C22 fatty acyl esters of ecdysone; Whiting and Dinan, 1988a, b). In contrast to Locusta and Schistocerca, a significant part of the ecdysteroids produced by the ovary is secreted in the hemolymph and distributed throughout the body and the amount of ecdysteroid conjugates that accumulate in the eggs is 100-fold lower than in Locusta and Schistocerca (Dinan, 1997; Whiting et al., 1997). Furthermore, in crickets ecdysteroids occur both in males and females while they are confined to females in locusts. The increase in ecdysteroids in the ovary is correlated to the maturation of increasing numbers of terminal oocytes, indicating that ecdysteroid production occurs during the period of patency and vitellogenin uptake (Dinan, 1997; Whiting et al., 1997). The primary regulator of ovarian development in crickets is JH but ecdysteroids have been observed to have both inhibitory and stimulatory roles, depending on their concentration (Chudakova et al., 1982; Behrens and Hoffmann, 1983). Ecdysteroid apolar conjugates also accumulate in the oocytes and predominate in newly-laid eggs. Hydrolysis of ecdysteroid conjugates could therefore serve as a source of active ecdysteroids that regulate embryonic processes (see above).

5.5.3

Cockroaches

As in locusts and crickets, the primary regulator of ovarian development in cockroaches is JH (Engelmann and Mala, 2000; Cruz et al., 2003; Raikhel et al., 2005).

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During ovarian development, ecdysone, 20E and 2-deoxy-ecdysone, both as free ecdysteroids and as apolar conjugates, accumulate in the ovary and peak at choriogenesis (Zhu et al., 1983; Slinger and Isaac, 1988). The ecdysteroids are produced by the follicular epithelium of the synchronously developing primary follicles (Zhu et al., 1983). A significant portion of the ecdysteroids produced by the ovary accumulate in the hemolymph and an important physiological role proposed for the hemolymph ecdysteroids includes the downregulation of the JH production by the corpora allata that triggers the termination of the gonadotrophic cycle (Rankin and Stay, 1985). Autocrine paracrine roles that have been proposed for the ovarian ecdysteroids include the regulation of choriogenesis and oviposition and the programming of the developmental competence of the next generation of oocytes (Zhu et al., 1983). Experiments in vitro have shown that 20E is able to induce precocious choriogenesis in Blatella germanica (Bellés et al., 1993). As in other insects, it is attractive to consider that the ecdysteroid esters accumulate in the egg and may serve as a reservoir of active hormone to be released by hydrolytic enzymes during embryogenesis. It was observed that, during embryogenesis of Nauphoeta cinerea, the quantity of free and highly polar ecdysteroids increases between ovulation and dorsal closure, i.e. before the differentiation of the prothoracic glands (Imboden and Lanzrein, 1982; Lanzrein et al., 1985). However, it is not clear whether in this species the ecdysteroids originate from maternal sources since only minimal amounts of ecdysteroids and their conjugates exist in newly-laid eggs (Zhu et al., 1983). The cloning of the cDNA of EcR in Blatella germanica and the applicability of the RNAi technique in cockroaches has allowed the functional analysis of the role of the ecdysone regulatory pathway during oogenesis in this species (Maestro et al., 2005; Martín et al., 2006; Cruz et al., 2006, 2007). The mRNA of the A-isoform of EcR is detectable in the cells of the follicular epithelium but its expression is not regulated by 20E. RNAi-mediated inhibition of expression of EcR-A indicates a role in the proliferation and function of the cells of the follicular epithelium and normal choriogenesis (Cruz et al., 2006). On the other hand, cockroaches with reduced levels of EcR showed normal vitellin content in the ovary and normal patency of the follicular epithelium during vitellogenesis and loss of patency at the end of vitellogenesis. Also homologs of RXR (usp; two isoforms) and HR3 (three isoforms) have been cloned in Blatella and essential roles were demonstrated in the regulation of the molt (Maestro et al., 2005; Martín et al., 2006; Cruz et al., 2007). The assessment of their role in oogenesis through RNAi techniques remains to be reported.

5.6

Ecdysone and Oogenesis in Hymenoptera

Reports regarding the roles of ecdysteroids during oogenesis in other insect orders are few and in general concern the occurrence of ecdysteroids in the ovary and eggs (e.g. Hemiptera: Kaplanis et al., 1975; Coleoptera: Gelman et al., 2000; Isoptera: Delbecque et al., 1978; Phasmida: Fournier and Radallah, 1988) or physiological

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evaluation of the application of ecdysone agonist on ovarian development (e.g. Tenebrio molitor (Coleoptera): Taïbi et al., 2003). Because of its status as a beneficial insect, the role of ecdysteroids during reproduction in the honeybee, Apis mellifera, has received more attention. In addition, with the availability of its annotated genome (Honeybee Genome Sequencing Consortium, 2006), cloning of genes of the ecdysone regulatory pathway is straightforward to allow determination of their expression pattern during oogenesis. Furthermore, the technique of RNAi has been used successfully in hymenopterans (Amdam et al., 2003) and therefore can be applied for functional studies that address the role of ecdysone during oogenesis. In the honeybee, ovarian development is coupled to caste differentiation in which JH plays a crucial role while supporting roles have been hypothesized for ecdysteroids (Nijhout and Wheeler, 1982; Rachinsky et al., 1990; Robinson and Vargo, 1997). At the middle of the fifth instar larval stage, the larval ovaries become responsive to makisterone A, the principal ecdysteroid involved in moulting and metamorphosis in the honeybee (Feldlaufer et al., 1986a). Compared to workers, ecdysteroid titers in queen last instar larvae occur earlier and reach higher values, and therefore may trigger accelerated prepupal development, which includes ovarian morphogenesis (Rachinsky et al., 1990). While ovarian differentiation proceeds during metamorphosis in queen larvae and prepupae, apoptosis is induced in worker ovaries (Hepperle and Hartfelder, 2001). The higher levels of JH in queen larvae and prepupae may protect the ovary against apoptosis (Schmidt Capella and Hartfelder, 1998), as was observed in Drosophila adults (see above). Also differences in pupal ecdysteroid titers can be related to distinct modes of caste development (Pinto et al., 2002). In adults, ecdysteroid levels are higher in queens and egg-laying workers than in normal workers (Robinson et al., 1991) which probably reflects ecdysteroid synthesis by the growing ovary (Feldlaufer et al., 1986b). As in other insects, ecdysteroids may have autocrine/paracrine functions within ovarian tissue or may accumulate in the follicles/eggs as maternal determinants for embryonic development (see above). A hormonal role for ecdysteroids accumulating in the hemolymph seems unlikely because hemolymph ecdysteroid titers can not be correlated with reproductive status (Hartfelder et al., 2002). Application of 20E also can not stimulate vitellogenin synthesis in the fat body in vitro although it stimulates general protein synthesis (Engels et al., 1990). The ecdysone regulatory genes E74, E75 and Broad-Complex were cloned in Apis and their expression pattern during oogenesis was studied (Paul et al., 2005, 2006). During the adult stage, specific expression is observed in the queen ovaries, but not worker ovaries, implicating their role in the regulation of follicle maturation. In situ hybridization shows that E74 is expressed in nurse cells and oocyte during vitellogenesis while it switches expression to the cells of the follicular epithelium during later stages. The presence of E74 (as well as E75 and Broad-Complex) in follicle cells at the end of oogenesis suggest a role in the trigger of apoptosis of the follicle cells during oviposition (Paul et al., 2005, 2006). In primitively social bees and wasps, as well as the primitively eusocial bumblebee Bombus terrestris, JH acts as the main gonadotropic hormone of

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ovarian development as in other insects (Bloch et al., 2000, 2002). Ovarian growth can be correlated with the accumulation of ecdysteroids that may have autocrine/ paracrine functions or have a role as maternal determinants as discussed above. In the bumblebee, ecdysteroids have been proposed to play a secondary role to JH in the determination of social dominance (Geva et al., 2005). In the parasitoid wasp Eupelmus vuilleti, ovarian development and concomitant ecdysteroid production are dependent on host availability and therefore can be reversed if unfavourable conditions may occur (Bodin et al., 2007). Host dependence of ovarian development represents an adaptive mechanism reminiscent of food dependence of oogenesis in anautogenous flies and mosquitoes. The role of ecdysteroids produced by Eupelmus ovaries (ecdysone and 2-deoxyecdysone) remains unknown.

5.7

Conclusion

The regulation of insect oogenesis by ecdysteroids is complex and can vary considerably from species to species. Following roles for ecdysteroids have been demonstrated or proposed: 1. Production of ecdysteroids by ovarian follicles may be involved in paracrine/ autocrine regulation of follicle maturation beyond a control point determined by external factors such as food availability, environmental stress or mating. Furthermore, the concentration of ecdysteroids may determine the decision between developmental progression and follicle degeneration. 2. Secretion of ecdysteroids by ovarian tissue into the hemolymph can reflect their role as hormonal factors to regulate extra-ovarian reproductive processes such as vitellogenin synthesis by the fat body or release of gonadotropic factors from the brain. A role for ovarian ecdysteroids as hormonal factors has been demonstrated most clearly in dipteran insects. 3. Production of high levels of ecdysteroids can trigger processes that occur at the end of oogenesis such as the reinitiation of meiosis in the oocyte and the synthesis of the chorion or eggshell by the follicular epithelium. 4. Maternal ecdysteroids may accumulate as inactive ecdysteroids conjugates in the eggs and serve as a source of active ecdysteroids that regulate embryonic processes (morphogenetic movements and cuticulogenesis). 5. In some insects, such as Bombyx mori, ovarian development has become dependent of metamorphic processes regulated by ecdysteroids produced by the prothoracic glands. In such cases, the role for ovarian ecdysteroids as paracrine/autocrine or endocrine factors may have become less important than in other insects. Close inspection of the regulation of oogenesis in insects (mainly Bombyx, Aedes and Drosophila) indicates that the exact role of ecdysteroids may be different in each of these cases and that a general pathway by which ecdysteroids regulate the progression of oogenesis remains to be discovered. In the silkmoth, the process of

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oogenesis is coupled to the process of metamorphosis and therefore has become dependent on the large surge of ecdysteroid titers that occurs in the pupal stage (Ramaswamy et al., 1997; Swevers and Iatrou, 2003). The silkmoth model may therefore represent a special case that is not applicable to most insects. In the mosquito, while the role of the ecdysteroid signalling pathway to regulate vitellogenin synthesis in the fat body is well established (Raikhel et al., 2005), very little is known about the involvement of ecdysteroids to regulate oocyte development within the ovary. In the fruitfly, the regulation of oogenesis by ecdysteroids is complex as ecdysteroids can have both positive and negative effects on oocyte development (Riddiford, 1993b; Soller et al., 1999). Furthermore, it was established that the expression of genes classically implicated in the ecdysone regulatory pathway (EcR, usp, E75, BR-C) are dependent on other signalling pathways during oogenesis (Deng and Bownes, 1997; Buszczak et al., 1999; Hackney et al., 2007). Much more effort may therefore be required to clarify the roles of ecdysteroids in insects, especially given the fact that research has focused mainly on a few dipteran and lepidopteran models as opposed to the diversity of other insects that remain uninvestigated.

Note Recently, the technique of RNAi has been applied successfully to investigate the functional role of genes in many organisms, including insects (Marie et al., 1999; Amdam et al., 2003; Cruz et al., 2006). The technique, based on hemolymph injection of dsRNA fragments homologous to the target gene, promises to be an important tool to test the function of isolated genes in organisms that are not readily amenable to genetic analysis. However, it has become apparent that, for reasons that are not understood, large differences can exist among insects with respect to their sensitivity to gene silencing by injection of dsRNA (Tomoyasu et al., 2008). Regarding model insects, it became apparent that in two cases dsRNA-mediated gene silencing through simple injection in the hemolymph is very effective: the German cockroach Blatella germanica (Dictyoptera; Cruz et al., 2006) and the flour beetle Tribolium castaneum (Coleoptera; Konopova and Jindra, 2007; Tan and Palli, 2008). Because the RNAi technique is very straightforward in these insects, in comparison to other model insects, fast progress is predicted to occur in the genetic analysis of many developmental processes, including oogenesis and its regulation by ecdysone, in Blatella and Tribolium. Here, the latest results regarding the function of the ecdysone regulatory pathway during cockroach and beetle oogenesis, as assessed through RNAi, are discussed briefly. In the cockroach, RNAi-mediated knockdown of two genes implicated in the ecdysone regulatory cascade, BgE75 and BgFTZ-F1, was reported recently (ManéPadros et al., 2008; Cruz et al., 2008). Besides effects on molting and development by causing degeneration of the prothoracic gland, BgE75 deficiency also causes defects in ovarian follicle development: in the primary follicle, whose growth

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parallels the increase of ecdysteroids in the haemolymph, proliferation of the cells of the follicular epithelium was reduced resulting in reduced size of the follicles (Mané-Padros et al., 2008). Remarkably, it was also observed that BgE75 dsRNA injected last instar nymphs do not molt to adults but nevertheless initiate the adult developmental program such as vitellogenin expression in the female fat body and its uptake by the ovary. Because of the documented persistence of dsRNA-mediated knockdown in the cockroach, the latter results may suggest that BgE75 does not play a major role in the regulation of vitellogenesis in the cockroach (Mané-Padros et al., 2008). On the other hand, while knockdowns of BgFTZ-F1 through dsRNA injection also resulted in molting defects, no effects on ovarian follicle development were hitherto reported (Cruz et al., 2008). Until recently, no data existed with respect to the function of the ecdysone regulatory pathway during oogenesis in Coleoptera. During the 17th Ecdysone Workshop, the effects of dsRNA-mediated knockdown of genes involved in the ecdysone regulatory pathway on the development of the ovary in Tribolium was presented (Takaki et al., 2008). In this work, it was reported that EcR and usp are required for egg production. Interestingly, anomalies caused by EcR dsRNA in Tribolium differed from both ovarian phenotypes observed in EcR mutants in Drosophila and ovarian anomalies caused by EcR dsRNA in Blatella, which suggests that ecdysteroid signalling pathways carry out different processes during oogenesis in different insects. Because Tribolium is very sensitive to the RNAi technique, it is expected that the role of the ecdysone receptor, the genes of the ecdysone regulatory cascade as well as the genes involved in ecdysone biosynthesis during oogenesis will be revealed in detail in the near future. Acknowledgements The research on silkmoth oogenesis by the laboratory of Insect Molecular Genetics and Biotechnology is in part supported by a PENED 2003 grant (03ED 124) of the General Secretariat for Research and Technology, Ministry of Development, in Greece.

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Chapter 6

Regulation of Sciarid DNA Puffs by Ecdysone: Mechanisms and Perspectives Nadia Monesi, Juliana Aparecida Candido-Silva, Maria Luísa Paçó-Larson, and Jorge Cury de Almeida

Abstract DNA puffs, formed in the salivary gland polytene chromosomes at the end of the larval stage, are characteristic of Sciaridae. Cytological demonstrations coupled to the molecular characterization of DNA puffs revealed that they are sites of disproportional DNA synthesis and abundant transcription. The biological purpose of DNA amplification at these loci is to enable the synthesis of large amounts of proteins in short periods of time. Here, the role of ecdysone in DNA puffs formation and DNA puff genes amplification and expression is reviewed and a new model for ecdysone action on DNA puffs is proposed. Studies in transgenic Drosophila that contributed to the understanding of the mechanisms that regulate Sciaridae DNA puff gene amplification and transcription are described. Finally, the availability of antibodies raised against the EcR, will further extend the knowledge about the roles exerted by ecdysone in this unique model system. Keywords DNA puffs • Sciaridae • gene amplification • transcription • ecdysone

6.1

The Discovery of DNA Puffs

The DNA puffs, a feature of sciarid polytene chromosomes, were initially reported by Breuer and Pavan (1952), while working with R. americana (called R. angelae in their investigation). They described a structure that forms in the polytene chromosomes of the salivary glands of fourth instar larvae that were similar to the

N. Monesi() and J.A. Candido-Silva Departamento de Análises Clínicas, Toxicológicas e Bromatológicas, Faculdade de Ciências Farmacêuticas de Ribeirão Preto, Universidade de São Paulo, 14040-903, Ribeirão Preto, SP, Brazil e-mail: [email protected] M.L. Paçó-Larson and J.C. de Almedia Departamento de Biologia Celular, Molecular e Bioagentes Patogênicos, Faculdade de Medicina de Ribeirão Preto, Universidade de São Paulo, 14049-900, Ribeirão Preto, SP, Brazil G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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“Balbiani Rings” observed in Chironomidae. In 1955, the same authors observed a disproportional increase in the DNA content of these puffs by Feulgen staining when compared to the neighboring bands (Breuer and Pavan, 1955). They interpreted this phenomenon as an adaptation for increased template production in these loci, which would result in the abundant synthesis of specific salivary gland polypeptides at the end of the fourth larval instar. Experimental evidence confirming that these regions constitute sites of disproportional DNA synthesis were obtained through the observation of higher [3H]-thymidine incorporation in these puff regions (Ficq and Pavan, 1957) and with microspectrophotometric measurements of DNA (Rudkin and Corlette, 1957). DNA puffs are also loci of abundant RNA transcription. Experiments measuring [3H]-uridine incorporation revealed high levels of RNA synthesis in the DNA puff forming regions at the time the puffs are expanded (Gabrusewycz-Garcia, 1968; Pavan and Da Cunha, 1969). Pavan (1965) called these puffs “DNA puffs” in order to distinguish them from the RNA puffs that occur in salivary gland polytene chromosomes of Chironomus and Drosophila. Similar studies in other sciarids such as Sciara coprophila (Crouse and Keyl, 1968; Gabrusewycz-Garcia, 1964; Rasch, 1970; Swift, 1962), Sciara ocellaris (Perondini, 1968), Bradysia hygida (Sauaia et al., 1971), and Trichosia pubescens (Amabis, 1974) confirmed that DNA puffs are sites of developmentally regulated differential DNA replication. Formation of DNA puffs is dependent on the disproportionate synthesis of DNA (Breuer and Pavan, 1955). In this context, DNA puffs are targets of drugs that either inhibit DNA synthesis or modify the chemical properties of DNA. Three drugs, that act through distinct mechanisms, were extensively employed in B. hygida (Fig. 6.1): hydroxyurea (HU) (Sauaia et al., 1971), 2,3-dihydro-1-H-imidazo (1,2-b) pyrazole (IMPY) (Ribeiro, 1975) and the thymidine analog 5-bromodeoxyuridine (BrdUrd) (de Almeida, 1978). The age when the process of gene amplification is beginning (E3) is named the “critical time”, and the injection of either HU or IMPY at this age completely prevented the formation of the two groups of DNA puffs in B. hygida (Fig. 6.1). The inhibition of DNA puff formation occurred without affecting the pattern of RNA puffs formation and without impairing larval development (Ribeiro, 1975; Sauaia et al., 1971). In the case of BrdUrd, the injection between 18 and 10 h before E7 resulted in modifications of the chromatin structure of the DNA puff anlages and impaired the expansion of DNA puffs, but did not interfere either with the activity of the RNA puffs, or the larval development (de Almeida, 1978).

6.2

Effect of 20-Hydroxyecdysone on DNA Puff Formation

Crouse (1968) carried out the first experiments demonstrating that DNA puff formation and DNA amplification are both regulated by ecdysone. The injection of ecdysone in young larvae of S. coprophila induced DNA amplification at the DNA puff forming sites 24 h after the injection (Crouse, 1968). In Rhynchosciara angelae, experiments in which salivary glands of young larvae were implanted into the body cavity of older larvae led to extra synthesis of DNA in the salivary gland

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Fig. 6.1 B. hygida life cycle, ecdysone titers and DNA puffs expansion and regression during the end of the fourth larval instar. The sciarid B. hygida started to be cultured in 1965 (Sauaia and Alves, 1968). Top, B. hygida life cycle. In the laboratory, at 20°C, B. hygida life cycle lasts about 36 days. The embryonic stage lasts 5 days, from egg laying to hatching of first instar larvae. The

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chromosome, of both the host and the implanted gland. The implantation of salivary glands of old larvae into young larvae led to the regression of the DNA puffs in the implanted salivary gland chromosomes (Amabis and Simoes, 1971). Further experiments in Rhynchosciara demonstrated that DNA amplification and DNA puff formation are both observed 24 h after ecdysone injection (Alvarenga et al., 1991; Berendes and Lara, 1975; Fresquez, 1979; Stocker and Pavan, 1974) and is a process dependent on RNA synthesis (Alvarenga et al., 1991; Berendes and Lara, 1975). Similar results were also obtained after the injection of ecdysone in T. pubescens larvae which revealed that the effect of ecdysone on DNA synthesis and DNA puff formation are both a late response to the hormone (Amabis and Amabis, 1984a). Studies in T. pubescens demonstrated that once amplification has started, it will continue even if the hormone source is eliminated by ligature of the larvae. In this case, DNA amplification occurs in the absence of DNA puff formation, showing that higher titers of hormone are required for DNA puff formation (Amabis and Amabis, 1984b). The role of ecdysone on RNA transcription in DNA puffs was also investigated through ligature experiments in T. pubescens (Amabis and Amabis, 1984b). In these

Fig. 6.1 (continued) larval stage comprises four instars. The fourth larval instar begins on the 17th day after egg laying. At the 6th day of the fourth instar (23rd day after egg laying) the eyespots arise (age E1, 0 h) which are useful staging landmarks. The pupal molt occurs on the 27th day, metamorphosis lasts 4 days which is terminated by the emergence of the adults. Middle, results from the measurements by radioimmunoassay (RIA) of 20-hydroxyecdysone titers in the hemolymph of B. hygida larvae during the end of the fourth larval instar (reproduced from Basso et al., 2002). Mean values and standard errors represent measurements of at least four animals in each developmental stage. The small peak at stage E1 (commitment peak), corresponds to a 15-fold increase in 20-hydroxyecdysone titers and the second and broader peak at stage E7 corresponds to a 200-fold increase over basal hormone levels. A decay in the ecdysone levels was observed before the onset of pupation, after which the hormone levels start to rise again. Bottom left shows a pair of fourth instar salivary glands (reproduced from Laicine et al., 1984). The salivary glands are about 10 mm long and consist of two rows of cells and three morphologically distinct regions: S1 (anterior), S2 (granulose) and S3 (posterior). About 190 cells form each gland (Paçó-Larson, 1976; Sauaia et al., 1971), and about 45 of them are in the S1 region. All DNA puffs (bottom right) are formed in the S1 and S3 regions with the exception of DNA puff C7 which is only formed in the S1 region (reproduced from Laicine et al., 1984). The first group of DNA puffs does not form in the S2 region, and the puffs of the second group are very inconspicuous in this gland region (Sauaia et al., 1971). The eyespot E3 (48 h after E1) coincides with the beginning of the process of gene amplification and DNA puff anlage formation at several chromosomal sites in salivary gland polytene chromosomes. There are eight major DNA puff forming regions, whose expansion is finely regulated throughout development and which can be classified in two distinct groups. At stage E5, the first group of DNA puffs starts its expansion (DNA puffs C7, C5 and C4), 2 h latter the E7 pattern is attained, when these three DNA puffs are open and about 8 h latter the first group of DNA puffs has already closed. The expansion of the second group of DNA puffs, puffs A14, B3d, C6 and X4, starts about 12 h after E7, and before the pupal molt (E7 + 26 h) these puffs are already closed, with the exception of puff B3d. The DNA puff B10 is the only DNA puff active between the periods of expansion of the first and second groups of DNA puffs and its maximum size is attained at E7 + 8 h (Laicine et al., 1984)

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experiments, the ligature blocked the release of ecdysone in the posterior region of the larvae and transcriptional activity was assessed by incubation of salivary glands in medium containing 3H-uridine. In the anterior region large DNA puffs were formed which displayed a heavy 3H-uridine labeling, whereas in the posterior region DNA puffs were not formed and the labeling was uniformly distributed (Amabis and Amabis, 1984b). Together, these results constitute cytological demonstrations that ecdysone treatment induces DNA amplification, DNA puff formation and RNA transcription at DNA puff forming regions. These observations are in agreement with the ecdysone titer measurements which have been performed in fourth instar larvae of both R. americana (Stocker et al., 1984) and B. hygida (Fig. 6.1) (Basso et al., 2002).

6.3

DNA Puffs and Protein Synthesis

The expansion and regression of specific DNA puffs can be related to the developmentally regulated synthesis of groups of salivary gland polypeptides (de Almeida, 1997; Laicine et al., 1984; Winter et al., 1977). In B. hygida, the pattern of protein synthesis in the salivary gland during the fourth larval instar was carefully investigated (Laicine et al., 1984). During the first 9 days of the fourth instar, the three salivary gland regions synthesize characteristic sets of polypeptides (first period of protein synthesis). After this period of sustained protein synthesis, the S1 and S3 salivary gland regions carry out two successive, new programs of synthesis (second and third periods of protein synthesis), which last about 12 h each. The striking temporal coincidence, between the expansion of the two groups of DNA puffs (Fig. 6.1) and the second and third periods of protein synthesis, respectively, suggested that these newly synthesized proteins were the products of the DNA puffs, which were produced during short periods of time while the puffs were expanded (Laicine et al., 1984). Studies in R. americana and T. pubescens reached similar conclusions (Amabis and Janczur, 1978; de Toledo and Lara, 1978; Winter et al., 1977) and indicated a causal relationship between DNA puffs and the synthesis of specific proteins. In B. hygida, the selective inhibition of DNA puffs development in both the S1 and S3 salivary gland regions, by either HU, or IMPY or BrdUrd, is accompanied by the inhibition of the same polypeptides characteristic of the second and third periods of synthesis, without affecting those produced during the first period of synthesis (Laicine et al., 1982, 1984). These results revealed that both DNA puff formation and DNA synthesis, processes triggered at E3, are essential for the correct reprogramming of protein synthesis in the later stages (Laicine et al., 1984). These results agree with those obtained both in R. americana (Bonaldo et al., 1979; Winter et al., 1980) and T. pubescens (Amabis, 1980) and led to the suggestion that the biological purpose of DNA amplification is to enable the synthesis of large amounts of proteins in short periods of time. The process of developmentally regulated gene amplification was probably selected in order to fulfill a demand that would probably not be attained by standard gene expression mechanisms (Laicine et al., 1984).

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The pattern of DNA puff protein synthesis in B. hygida was extended through the analysis of both the patterns of global protein synthesis and the characterization of the patterns of secreted polypeptides. Radiometry measurements of the saliva of larvae injected with 3H-leucine revealed that most of the cumulative secretion of radioactive proteins occurs during the period of DNA puffs activity and reaches maximum levels of synthesis at age E7 + 6 h, at amounts that correspond to 28.5 times the amount of protein secreted at age E1 (de Almeida, 1997). The characterization of B. hygida salivary gland extracts by fluorography revealed a dynamic pattern of polypeptide synthesis in the salivary gland at the time of DNA puff formation. Correlations drawn between the time of appearance of some polypeptides and the formation of DNA puffs led to the suggestion that a 43 and a 23 kDA polypeptide constitute the products of DNA puff C4 and B10, respectively (de Almeida, 1997). These results were latter confirmed by the demonstration that polyclonal antibodies raised against either the product of DNA puff B10, the BhB10-1 protein, or against the product of DNA puff C4, the BhC4-1 protein, detect the same polypeptides as previously identified by fluorography (Fontes et al., 1999; Monesi et al., 2004). Several DNA puff genes have been cloned and searches with full length cDNAs against the public databases revealed that the DNA puffs polypeptides have no orthologs in the complete genome sequences of four other insect species (Drosophila melanogaster, Anopheles gambiae, Apis mellifera and Tribolium castaneum). Based on the similarity of the deduced amino acid sequences three DNA puff protein families have been identified: family I comprises the product of DNA puff C3 of R. americana, the II/9-1 and II/9-2 proteins of DNA puff II/9A of S. coprophila and the product of DNA puff C-4B of T. pubescens; family II comprises the BhC4-1 protein from DNA puff C4 of B. hygida, and the products of both DNA puff C8 and B2 of R. americana; family III comprises the BhB10-1 protein of DNA puff B10 of B. hygida (Penalva et al., 1997; Santelli et al., 2004).

6.4

The Role of Ecdysone in the Reprogramming of Protein Synthesis at the End of the Fourth Larval Instar

The injection of ecdysone in young larvae (E1) induced both the second and third periods of protein synthesis in the salivary gland (de Carvalho DP and de Almeida JC, 1993). These results were confirmed by the direct demonstration that the BhC4-1 protein, a member of the second period of protein synthesis, is detected in salivary gland extracts 24 h after the injection of hormone in larvae at E1, and indicated that the expression of this protein is a late response to the hormone (Basso et al., 2002) (Fig. 6.2). When salivary glands at the age E1 were cultivated in vitro in the presence of the hormone, neither the second nor the third period of protein synthesis were induced, indicating that some other factor, besides the hormone, is necessary to promote DNA puff genes expression (de Carvalho DP and de Almeida JC, 1993). These results are in agreement with those obtained in Rhynchosciara, which revealed that the complete induction of DNA puff B2

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Fig. 6.2 BhC4-1 expression and amplification are both induced by ecdysone (Reproduced from Basso et al., 2002). (a) Northern blot containing 20 µg of total RNA extracted from salivary glands of larvae injected at stage E1 with 0.4 mM ecdysone and dissected at different times after injection (1, 2, 4, 8, 16 and 24 h), 20 µg of total RNA extracted from salivary glands of larvae injected with 5% ethanol and dissected 24 h latter (Eth/24) and 20 µg of total RNA extracted from salivary glands of larvae at stage E7. The upper autoradiogram shows the result after hybridization with a BhC4-1 cDNA fragment and the bottom autoradiogram shows the result after hybridization of the same blot with a 500 bp fragment from a cDNA encoding the mitochondrial large rRNA subunit of Anopheles gambiae (Lemos et al., 1996). (b) Left hand picture, Coomassie Blue stained SDSPAGE (9%) of total salivary gland extracts of larvae injected at stage E1 with ecdysone or with 5% ethanol, and dissected at the indicated times and total salivary gland extracts of larvae at stage E7. Right hand picture, immunoblot of an identical gel after incubation with an affinity-purified anti-BhC4-1 antibody followed by detection with chemiluminescence. The 43 kDa polypeptide is the BhC4-1 product (Monesi et al., 2004). (c) Upper autoradiogram: southern blot containing 10 µg of Eco RI digested DNA extracted from salivary glands of larvae at stage E1 and E7, 10 µg of Eco RI digested DNA from salivary glands of larvae injected at stage E1 with 0.4 mM ecdysone and dissected at different times after injection (14, 16, 24 and 36 h) and 10 µg of Eco RI digested DNA from salivary glands of larvae injected at stage E1 with 5% ethanol and dissected at different times after injection (16, 24 and 36 h), after hybridization with a 1.2 kb BhC4-1 cDNA fragment which detects a 9 kb Eco RI genomic fragment. Bottom autoradiogram, the blot was stripped, followed by hybridization with a cDNA fragment encoding the elongation factor 1α (EF-1α) of R. americana (Graessmann et al., 1992)

activity in vitro was only attained when salivary glands were cultivated in the presence of both the hormone and hemolymph (Alvarenga, 1980). In older B. hygida larvae (age E7), the presence of the hormone in the culture medium was sufficient to sustain the synthesis of the polypeptides characteristic of the second period of synthesis, and inhibited the appearance of the polypeptides

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of the third period of synthesis (de Carvalho et al., 2000). Furthermore, the injection of hormone in larvae at E7 + 10 h, developmental time when the polypeptides characteristic of the third period of synthesis are induced, inhibited the synthesis of this group of polypeptides (de Carvalho et al., 2000). These results indicate that the two groups of DNA puffs of B. hygida respond to different thresholds of hormone: the expression of the genes of the first group of DNA puffs (second period of protein synthesis) depend on the presence of high titers of hormone, whereas the expression of the genes of the second group of DNA puffs (third period of protein synthesis) occurs in the presence of lower titers of hormone (de Carvalho et al., 2000). These results, together with direct measurements of 20-hydroxyecdysone concentration in the hemolymph of larvae during the end of the fourth instar (Basso et al., 2002), do not fit the model described by Lara et al., (1991) which proposed that high levels of hormone are necessary during the entire period of gene amplification and DNA puff expansion. The current model on the ecdysone action on DNA puffs, derived from studies in B. hygida, proposes that the high ecdysone titers in the hemolymph at E5 (Fig. 6.1) induce the expansion of the first group of DNA puffs and the expression of the polypeptides characteristic of the second period of protein synthesis (Basso et al., 2002; de Carvalho et al., 2000). During cocoon spinning (late E7), the levels of hormone and probably its receptor, start to decrease, which results in reduced expression levels of the amplified genes and the regression of the first group of DNA puffs. At age E7 + 12, when the hormone titers are lower, the expression of the genes of the second group of DNA puffs is induced (third period of protein synthesis) (Basso et al., 2002; de Carvalho et al., 2000). At the end of the fourth larval instar, the increase in the 20-hydroxyecdysone titers (Fig. 6.1) results in the down regulation of the genes of the second group of DNA puffs prior to the larvae to pupae transition and to the transformations typical of metamorphosis, including elimination of the salivary gland by a process of programmed cell death (Simon and de Almeida, 2004).

6.5

Mechanism of Gene Amplification in DNA Puffs

Gene amplification in DNA puff forming regions results from re-replication events triggered in specific regions of the salivary gland chromosomes. The amplified DNA accumulates in the salivary glands (Fontes et al., 1992; Paçó-Larson et al., 1992; Wu et al., 1993; Yokosawa et al., 1999) and remains bound to the DNA puff forming sites (Paçó-Larson, 1982; Rasch, 1970) until at least the end of the larval stage. Amplification levels of between 4- to 30-fold were demonstrated depending on the DNA puff analyzed (Fontes et al., 1992; Glover et al., 1982; Monesi et al., 1995; Paçó-Larson et al., 1992; Santelli et al., 2004; Wu et al., 1993). The DNA strands synthesized during amplification are arranged in parallel (Fontes et al., 1992; Paçó-Larson et al., 1992; Santelli et al., 1991; Wu et al., 1993), as opposed to the

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arrangement in tandem demonstrated for the amplicons formed as part of the drug resistance response in mammalian cells (Ma et al., 1988). Quantitative hybridization studies performed in B. hygida (Coelho et al., 1993; Monesi et al., 1995) and in R. americana (Stocker et al., 1996; Yokosawa et al., 1999) indicated the existence of an amplification gradient in the DNA puff regions similar to that described for the amplicons formed in the ovarian follicular cells of D. melanogaster (Claycomb et al., 2004; Spradling, 1981). The fragments containing the genes whose temporal pattern of expression are related to DNA puffs expansion reach higher amplification levels than fragments containing sequences expressed when the puff is absent (Coelho et al., 1993; Monesi et al., 1995). In the Drosophila chorion loci, the high frequency of replication initiation, coupled to low elongation speed, results in an amplification gradient that extends over approximately 100 kb and is centered on an origin of replication region which is repetitively triggered in the follicular cells at the end of the oogenesis (for review, Orr-Weaver, 1991). The amplification mechanism in the chorion loci was further characterized by electron microscopy studies of chromatin spreading (Osheim et al., 1988), which revealed a structure of forks within forks, named “onion skin”, and through bi-dimensional electrophoreses analysis of replication intermediates (Delidakis and Kafatos, 1989; Heck and Spradling, 1990). Quantitative hybridization results in B. hygida (Coelho et al., 1993; Monesi et al., 1995), coupled to the identification of bi-directional origins of replication which are active during the amplification period in both S. coprophila DNA puff II/9 (Liang et al., 1993) and DNA puff C3 of R. americana (Yokosawa et al., 1999), led to the proposal that the amplification in DNA puffs forming sites also occurs by an onion skin type mechanism. In these loci, replication bubbles were detected upstream of the transcription initiation site of the genes whose expression is temporally related to the respective puff expansion (Liang et al., 1993; Yokosawa et al., 1999). Similarly, the BhC4-1 gene of the B. hygida DNA puff C4 is amplified by forks that emanate from its 5′ region, although in this case the origin of replication region has not been cloned (Coelho, 1997). The characterization by bi and tri-dimensional electrophoresis of the replication zone in DNA puff II/9A of S. coprophila revealed that each DNA strand is replicated from an unique site within a ∼1 kb initiation region (ORI), located ∼2 kb upstream of the II/9A-1 gene transcription initiation site (Liang and Gerbi, 1994; Liang et al., 1993). Immediately upstream of the DNA puff II/9 ORI region lies a 80 bp fragment, which binds the origin replication complex (ORC). The binding of ORC to this 80 bp fragment is ATP dependent (Bielinsky et al., 2001). These results demonstrated that the binding of ORC to DNA fragments adjacent to the replication initiation site also occurs in metazoans (Diffley et al., 1994). The ORI region in the S. coprophila DNA puff II/9 is part of an initiation region employed during both the salivary gland endocycles and the embryonic mitotic cycles, indicating that the DNA replication machinery employed in cell proliferation and growth is also used in amplification (Lunyak et al., 2002). The results obtained in sciarids agree with those derived from the molecular characterization of mutations associated with the thin eggshell phenotype in Drosophila (Claycomb and Orr-Weaver, 2005). The demonstration that intrachromosomal amplification recruits part of the conserved

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replication initiation machinery indicates the existence of a mechanism that specifies these amplified loci for re-replication. The understanding of this mechanism is a goal that is being pursued both in Drosophila follicular cells and in sciarid DNA puffs. A ∼400 bp DNase hypersensibility region (DHS), which is more susceptible to endonuclease clevage during the amplification period than after amplification, was mapped downstream of the DNA puff II/9A ORI of S. corpophila (Urnov et al., 2002). This DHS may constitute a DNA puff amplification regulator similar to the ACEs (Amplification Control Element) formerly described in the Drosophila chorion genes loci. The ACEs are defined as elements able to induce DNA amplification when inserted in different genomic sites independently of their orientation and distance (Orr-Weaver et al., 1989; Zhang and Tower, 2004). The sequence of the ACE3 element characterized in D. melanogaster is conserved between various species of Drosophila (Swimmer et al., 1990). Heterologous systems were employed to investigate the regulation of DNA puff amplification and/or gene expression (BienzTadmor et al., 1991; Millar et al., 1985; Monesi et al., 1998). The most conclusive study was obtained by the analysis of transgenic D. melanogaster lines carrying an 18 kb fragment of the DNA puff C3 of R. americana that contained the ORI region active during amplification (Yokosawa et al., 1999) and which was flanked by su(Hw)BS elements, to minimize possible position effects (Lu and Tower, 1997). The absence of R. americana DNA puff amplification in both the salivary gland and in the follicular cells in the Drosophila transgenic lines indicated that factors involved in DNA puff C3 amplification are not conserved between the sciarids and Drosophila (Soares et al., 2003). Sequence searches performed in the NCBI non-redundant database revealed 90% identity in a 72 bp sequence present both in the S.coprophila DNA II/9A DHS sequence and in the ORI region of R. americana DNA puff C3 (Fig. 6.3), and suggested a role for this 72 bp sequence in DNA puffs amplification. The absence of identity between the DNA II/9A DHS sequence and the Drosophila ACEs is not surprising and agrees with the results derived from functional studies, which

DHS-Sci

202 AACACGAGGCGAATAAAAAGTGCATCGTTCTAAGCCGAATTTTAAAAGACAGAC---AAT 258 ||||||||| |||||||| |||||||| ||||||||||||||| |||||||||| ||| ORI-Rhy 5889 AACACGAGGTGAATAAAA-GTGCATCGGTCTAAGCCGAATTTT-AAAGACAGACTTAAAT 5946 DHS-Sci

259 ATTTACACAGTTTGC 273 ||||||||||||||| ORI-Rhy 5947 ATTTACACAGTTTGC 5961

Fig. 6.3 The R. americana DNA puff C3 ORI region share a 71–72 bp sequence displaying 90% identity with the DNase Hypersensitive Site (DHS) adjacent to the ORI of S. coprophila DNA II/9A amplicon. The alignment is the result of a Blastn of the DHS 419 pb sequence of S. coprophila against the NCBI non redundant databank (NCBI accession numbers:AY225408, R. americana and AF464926, S. coprophila)

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suggested that the factors that regulate intra-chromosomal DNA amplification during the normal course of development are not conserved between sciarids and Drosophila (Soares et al., 2003).

6.6

The Role of Ecdysone in DNA Puff Gene Amplification

In B. hygida, the selective inhibition of DNA puffs development by HU at E3 (“critical time”), suppresses the formation of DNA puff anlage (Sauaia et al., 1971) and indicates that the re-replication involved in the amplification process begins simultaneously at the DNA puffs forming regions. The fact the “critical time” defined in B. hygida coincides with the increase in ecdysone titers in the hemolymph (Fig. 6.1), agrees with the idea that the hormone is the developmental cue that triggers amplification in all DNA puffs forming sites. Accordingly, the injection of ecdysone in larvae during the pre-amplification stage, when the ecdysone levels are low, induces the amplification of B. hygida DNA puff BhC4-1 gene (Basso et al., 2002) (Fig. 6.2). Similar results were also recently described for the II/9A locus of S. coprophila both in vivo and in vitro. In in vitro experiments, the addition of transcription and protein synthesis inhibitors to the culture medium prevented the induction of amplification and suggested an indirect mechanism for the action of ecdysone in amplification (Foulk et al., 2006). Immunolocalization experiments in polytene chromosomes of T. pubescens, using a polyclonal antibody raised against the EcR of Chironomus, revealed that the EcR is bound in DNA puff-forming loci before and during DNA puffs expansion (Stocker et al., 1997). More recent experiments, using polyclonal antibodies developed against the EcR of B. hygida, revealed the presence of the receptor in all B. hygida DNA puffs when they are expanded (Fig. 6.4 and Candido-Silva et al.,

Fig. 6.4 The BhEcR colocalizes with RNA polymerase II at DNA puff forming sites. Chromosome C from Bradysia hygida larvae at age E7 labeled with antibodies anti-BhEcR (red) and anti-RNA Pol II (green). Yellow signals indicate colocalization of both antibodies. From left to right, images of the same field were captured by confocal microscopy at increasing magnifications. The bars at the bottom left of each picture correspond, from left to right, to 10 µm, 5 µm and 1 µm, respectively (See Color Plates)

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2008). Furthermore, electrophoretic mobility shift assays (EMSA) revealed that a putative ecdysone receptor binding site, localized between the S. coprophila II/9A DHS sequence and the ORC binding region, binds the Sciara EcR efficiently, perhaps suggesting that the EcR could be the elusive amplification factor (Foulk et al., 2006). At present, it is difficult to define whether the EcR/USP receptor exerts its influence by modulating amplification, transcription, or both processes. The function of the EcR in DNA puff forming sites might be broader. Remarkably, alterations in the histone acetylation in the ORC binding sites of Drosophila chorion genes interfere with chorion genes amplification (Aggarwal and Calvi, 2004). In this sense, it is possible that the EcR acts as an anchor to recruit chromatin modulator factors, which could in turn create an environment favorable for the binding of other factors that modulate the functions of DNA puffs.

6.7

The Role of Ecdysone on the Transcription of DNA Puff Genes

Sciarid DNA puffs are sites of developmentally regulated transcription (Coelho et al., 1993; DiBartolomeis and Gerbi, 1989; Fontes et al., 1992; Frydman et al., 1993; Glover et al., 1982; Monesi et al., 1995; Paçó-Larson et al., 1992; Penalva et al., 1997; Santelli et al., 1991; Santelli et al., 2004; Wu et al., 1993). In vitro experiments revealed that the presence in the culture medium of other factor(s), besides the hormone, that was or were necessary in order to fully induce DNA puff genes expression (Alvarenga, 1980; de Carvalho et al., 2000; Foulk et al., 2006). In R. americana, the expression of the B-2 mRNA in the salivary gland was induced 24 h after ecdysone injection (Alvarenga et al., 1991; Bonaldo et al., 1979; de Toledo and Lara, 1978). In B. hygida maximum levels of BhC4-1 mRNA expression are detected in the salivary gland when the ecdysteroid titers in the hemolymph are highest and the injection of ecdysone in larvae at (E1) induces BhC4-1 mRNA in the salivary gland 16 h later (Basso et al., 2002; Paçó-Larson et al., 1992) (Fig. 6.2). More recent results in S. coprophila have shown that the II/9-1 gene of DNA puff II/9A is also induced in the salivary gland 30 h after hormone injection (Foulk et al., 2006). Taken together, ecdysone injection experiments in sciarids revealed that the induction of DNA puff genes expression in the salivary gland occurs between 16–36 h after hormone injection, and constitute a late response to elevated levels of hormone in the hemolymph (Alvarenga et al., 1991; Basso et al., 2002; Foulk et al., 2006). In all cases, the complete induction of DNA puff genes expression has only been attained in in vivo experiments and depends on the injection of hormone (Alvarenga et al., 1991; Basso et al., 2002; Foulk et al., 2006). In both T. pubescens and B. hygida, most of the chromosomal regions in which the EcR is detected also correspond to regions in which RNA polymerase II is bound (Candido-Silva et al., 2008; Stocker et al., 1997). All three EcREs (ecdysone response elements) identified in the II/9-1 promoter region are able to bind protein complexes present in salivary gland nuclear extracts, which are supershifted in the

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presence of an antibody specific to DmEcR (Drosophila melanogaster ecdysone receptor) (Foulk et al., 2006). Together, the results obtained in sciarids suggests that the EcR receptor might act as a trans-activating factor in the transcriptional regulation of DNA puff genes (Candido-Silva et al., 2008; Foulk et al., 2006; Stocker et al., 1997). The characterization of DNA puff genes transcription regulatory mechanisms has been extended through functional assays in transgenic Drosophila. These studies revealed that the mechanisms that control the tissue specific developmentally regulated expression of both the II/9-1 gene of S. coprophila and the BhC4-1 gene of B. hygida are conserved in Drosophila (Bienz-Tadmor et al., 1991; Monesi et al., 1998). However, in transgenesis the DNA puff BhB10-1 of B. hygida and the C3-22 gene of R. americana are constitutively expressed at low levels throughout development (Monesi et al., 2001; Soares et al., 2003). Together, these results reveal that the conservation of DNA puff genes transcriptional regulatory mechanisms is not observed for all DNA puff genes. In transgenic Drosophila, a 718 bp fragment from the II/9-1 promoter of S. coprophila drives the expression of the CAT (chloramphenicol acetyltransferase) reporter gene in late prepupae. CAT activity is initially detected in 6 h prepupae and reaches maximum levels in 9 h prepupae (Bienz-Tadmor et al., 1991). The B. hygida BhC4-1 mRNA is initially detected during the larvae to prepupae transition in transgenesis, and reaches its highest levels of expression in 3 h prepupae (Basso et al., 2002; Monesi et al., 1998). These results indicate that the II/9-1 gene and the BhC4-1 gene are under different temporal regulation in Drosophila. While the BhC4-1 gene has been shown to be induced in response to the increase of hormone levels that trigger the larval to prepupal transition, the II/9-1 gene is most likely induced in response to the prepupal peak of hormone that promotes the prepupae to pupae transition in Drosophila. In Drosophila, it is possible to reproduce in vitro the entire sequence of RNA puffing when intermolt salivary glands are cultivated in the presence of ecdysone (Ashburner, 1972; Ashburner et al., 1974; Huet et al., 1993, 1995). In transgenesis, the induction of BhC4-1 mRNA is a late response to the hormone that occurs only after 6 h exposure of salivary glands to the hormone, which is reminiscent of the response of B. hygida larvae after hormone injection (Basso et al., 2002). The incubation of transgenic salivary glands in the presence of cycloheximide in the absence of ecdysone also resulted in the induction of BhC4-1 expression, indicating the participation of a repressor whose synthesis is necessary to maintain the gene repressed (Basso et al., 2002). A different situation has been observed for the II/9-1 gene of S. coprophila. Whereas the injection of hormone in S. coprophila induces the II/9-1 gene expression 30 h after injection (Foulk et al., 2006), in transgenesis the reporter gene expression driven by the II/9-1 promoter was rapidly induced by ecdysone (Bienz-Tadmor et al., 1991). The early response of the II/9-1 gene in transgenesis could be explained if the regulation of this gene also requires a repressor. If the tested II/9-1 promoter fragment did not include the repressor binding site, or if the repressor is not present in Drosophila, an early induction of the gene would occur (Basso et al., 2002).

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At the larvae to prepupae transition in D. melanogaster the ecdysone heterodimeric receptor EcR/USP (ultraspiracle) bound to ecdysone directly activates the expression of a small number of early response genes. The products of early genes are transcriptional regulators whose function is to induce a large set of late response genes (Thummel, 1996). As discussed above, both in B. hygida, and in transgenesis, the BhC4-1 gene is induced as a late response to the increase in ecdysone titers which precede the metamorphosis (Basso et al., 2002). The role of the Drosophila early genes, BR-C, E74 and E75 on the regulation BhC4-1 genes was recently investigated in Drosophila transformed with a chimeric BhC4-1-lacZ gene, through a loss of function approach, coupled to overexpression experiments (Basso et al., 2006). The results demonstrated that the product of the BR-C early gene, the BR-C Z3 isoform, is essential for the induction of BhC4-1-lacZ in the salivary gland and also revealed that the early gene products BR-C Z1, BR-C Z4, E75A, E74A and E74B participate to a lesser degree in the regulation of BhC41-lacZ. Since in the absence of the early gene products neither the developmental time nor the tissue specificity of BhC4-1-lacZ expression was affected, it has been suggested that the role of the early gene products is to regulate the correct levels of BhC4-1 expression in the salivary gland at the larvae to prepupae transition (Basso et al., 2006). These results constitute the first demonstration that trans-activating factors, members of the ecdysone regulatory pathway, participate in the transcriptional regulation of a DNA puff gene and suggest that this regulatory pathway is conserved between D. melanogaster and B. hygida.

6.8

Future Perspectives

The last decade has witnessed important discoveries regarding the role of ecysone in DNA puff forming sites. Ecdysone injection experiments revealed that the hormone induces DNA puffs formation, DNA amplification and transcription in sciaridae salivary glands (Basso et al., 2002; Foulk et al., 2006). Experiments in transgenesis have confirmed that the response of DNA puff genes to the hormone is a late response and have shown that the early gene products participate in the regulation of DNA puffs genes (Basso et al., 2002, 2006; BienzTadmor et al., 1991). It will be interesting to learn whether these trans-activating factors bind directly or indirectly to the BhC4-1 promoter, and to further investigate the conservation of the early gene products in sciarids and their role in DNA puff genes regulation. It is worthy of note that the early genes do not regulate either the tissue pattern or the developmental time of the DNA puff gene expression. Furthermore, previous studies from our group have indicated that the Drosophila salivary gland transcription factor, Forkhead, does not participate in the regulation of BhC4-1 gene in transgenesis. In this context, further characterization of DNA puff genes regulatory mechanisms in transgenic Drosophila will contribute to the discovery of additional tissue specific transcription factors.

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The exciting results demonstrating that the EcR is localized at DNA puff forming sites during DNA puff expansion both in T. pubescens and B. hygida (Candido-Silva et al., 2008; Stocker et al., 1997), together with EMSA experiments demonstrating that the EcR binds the II/9A locus of S. coprophila (Foulk et al., 2006) extend the characterization of the role of the EcR in sciarids. One aspect that merits further investigation is the precise role of the EcR. Is the EcR in scarids a transcription factor, an amplification factor or both? The identification of factors that interact with the sciarid EcR during the amplification process, coupled with functional assays employing RNAi, could unravel the mechanism that control DNA amplification during larval development. Furthermore, the recent cloning of the EcR and usp orthologues, both in B hygida (Valente V and de Almeida JC, unpublished results, 1999 and Candido-Silva et al., 2008) and S. coprophila (Foulk et al., 2006), provides the necessary tools to determine the role of both the receptor and the hormone in DNA puff genes transcription using cell culture based assays. In this context, the study of sciarid DNA puffs will further the knowledge of the roles exerted by the EcR receptor and its ligand, ecdysone.

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Fontes AM, Conacci ME, Monesi N, de Almeida JC, Paçó-Larson ML (1999) The DNA puff BhB10-1 gene encodes a glycine-rich protein secreted by the late stage larval salivary glands of Bradysia hygida. Gene 231: 67–75 Foulk MS, Liang C, Wu N, Blitzblau HG, Smith H, Alam D, Batra M, Gerbi SA (2006) Ecdysone induces transcription and amplification in Sciara coprophila DNA puff II/9A. Dev Biol 299: 151–163 Fresquez CL (1979) Nucleic acid synthesis in Rhynchosciara hollaenderi polytene chromosomes: I. “Dose response and temporal sequence after injection of 20-hydroxyecdysone. Insect Biochem 9: 517–523 Frydman HM, Cadavid EO, Yokosawa J, Henrique Silva F, Navarro-Cattapan LD, Santelli RV, Jacobs-Lorena M, Graessmann M, Graessmann A, Stocker AJ, et al. (1993) Molecular characterization of the DNA puff C-8 gene of Rhynchosciara americana. J Mol Biol 233: 799–803 Gabrusewycz-Garcia N (1964) Cytological and autoradiographic studies in Sciara coprophila salivary gland chromosomes. Chromosoma 15: 312–344 Gabrusewycz-Garcia N (1968) RNA metabolism of polytene chromosomes. J Cell Biol 39: 49A Glover DM, Zaha A, Stocker AJ, Santelli RV, Pueyo MT, De Toledo SM, Lara FJ (1982) Gene amplification in Rhynchosciara salivary gland chromosomes. Proc Natl Acad Sci USA 79: 2947–2951 Graessmann M, Graessmann A, Cadavid EO, Yokosawa J, Stocker AJ, Lara FJ (1992) Characterization of the elongation factor 1-alpha gene of Rhynchosciara americana. Nucleic Acids Res 20: 3780 Heck MM, Spradling AC (1990) Multiple replication origins are used during Drosophila chorion gene amplification. J Cell Biol 110: 903–914 Huet F, Ruiz C, Richards G (1993) Puffs and PCR: the in vivo dynamics of early gene expression during ecdysone responses in Drosophila. Development 118: 613–627 Huet F, Ruiz C, Richards G (1995) Sequential gene activation by ecdysone in Drosophila melanogaster: the hierarchical equivalence of early and early late genes. Development 121: 1195–1204 Laicine EM, Alves MAR, Almeida JC, Albernaz WC, Sauaia H (1982) Expressão gênica no desenvolvimento da glândula salivar de Bradysia hygida. Significado biológico dos pufes de DNA. Ciência e Cultura 34: 488–492 Laicine EM, Alves MAR, de Almeida JC, Rizzo E, Albernaz WC, Sauaia H (1984) Development of DNA puffs and patterns of polypeptide synthesis in the salivary glands of Bradysia hygida. Chromosoma 89: 280–284 Lara FJ, Stocker AJ, Amabis JM (1991) DNA sequence amplification in sciarid flies: results and perspectives. Braz J Med Biol Res 24: 233–248 Lemos FJ, Cornel AJ, Jacobs-Lorena M (1996) Trypsin and aminopeptidase gene expression is affected by age and food composition in Anopheles gambiae. Insect Biochem Mol Biol 26: 651–658 Liang C, Gerbi SA (1994) Analysis of an origin of DNA amplification in Sciara coprophila by a novel three-dimensional gel method. Mol Cell Biol 14: 1520–1529 Liang C, Spitzer JD, Smith HS, Gerbi SA (1993) Replication initiates at a confined region during DNA amplification in Sciara DNA puff II/9A. Genes Dev 7: 1072–1084 Lu L, Tower J (1997) A transcriptional insulator element, the su(Hw) binding site, protects a chromosomal DNA replication origin from position effects. Mol Cell Biol 17: 2202–2206 Lunyak VV, Ezrokhi M, Smith HS, Gerbi SA (2002) Developmental changes in the Sciara II/9A initiation zone for DNA replication. Mol Cell Biol 22: 8426–8437 Ma C, Looney JE, Leu TH, Hamlin JL (1988) Organization and genesis of dihydrofolate reductase amplicons in the genome of a methotrexate-resistant Chinese hamster ovary cell line. Mol Cell Biol 8: 2316–2327 Millar S, Hayward DC, Read CA, Browne MJ, Santelli RV, Garcia Vallejo F, Pueyo MT, Zaha A, Glover DM, Lara FJ (1985) Segments of chromosomal DNA from Rhynchosciara americana that undergo additional rounds of DNA replication in the salivary gland DNA puffs have only weak ARS activity in yeast. Gene 34: 81–86 Monesi N, Fernandez MA, Fontes AM, Basso LR, Jr., Nakanishi Y, Baron B, Buttin G, PaçóLarson ML (1995) Molecular characterization of an 18 kb segment of DNA puff C4 of Bradysia hygida (Diptera, sciaridae). Chromosoma 103: 715–724

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Monesi N, Jacobs-Lorena M, Paçó-Larson ML (1998) The DNA puff gene BhC4-1 of Bradysia hygida is specifically transcribed in early prepupal salivary glands of Drosophila melanogaster. Chromosoma 107: 559–569 Monesi N, Sousa JF, Paçó-Larson ML (2001) The DNA puff BhB10-1 gene is differentially expressed in various tissues of Bradysia hygida late larvae and constitutively transcribed in transgenic Drosophila. Braz J Med Biol Res 34: 851–859 Monesi N, Silva JA, Jr., Martins PC, Teixeira AB, Dornelas EC, Moreira JE, Paçó-Larson ML (2004) Immunocharacterization of the DNA puff BhC4-1 protein of Bradysia hygida (Diptera: Sciaridae). Insect Biochem Mol Biol 34: 531–542 Orr-Weaver TL (1991) Drosophila chorion genes: cracking the eggshell’s secrets. Bioessays 13: 97–105 Orr-Weaver TL, Johnston CG, Spradling AC (1989) The role of ACE3 in Drosophila chorion gene amplification. Embo J 8: 4153–4162 Osheim YN, Miller OL, Jr., Beyer AL (1988) Visualization of Drosophila melanogaster chorion genes undergoing amplification. Mol Cell Biol 8: 2811–2821 Paçó-Larson ML (1976) Análise quantitativa do conteúdo de DNA da glândula salivar de Bradysia hygida. M.Sc. thesis, University of São Paulo Paçó-Larson ML (1982) Citofotometria do DNA de núcleos inteiros e de segmentos cromossômicos formadores de pufes de DNA da região anterior da glândula salivar de Bradysia hygida. Ph.D. thesis, University of São Paulo Paçó-Larson ML, De Almeida JC, Edström JE, Sauaia H (1992) Cloning of a developmentally amplified gene sequence in the DNA Puff C4 of Bradysia hygida (Diptera, Sciaridae) salivary glands. Insect Biochem Mol Biol 22: 439–446 Pavan C (1965) Chromosomal differentiation. Natl Cancer Inst Monogr 18: 309–323 Pavan C, Da Cunha AB (1969) Gene amplification in ontogeny and phylogeny of animals. Genetics 61 (Suppl): 289–304 Penalva LO, Yokosawa J, Stocker AJ, Soares MA, Graessmann M, Orlando TC, Winter CE, Botella LM, Graessmann A, Lara FJ (1997) Molecular characterization of the C-3 DNA puff gene of Rhynchosciara americana. Gene 193: 163–172 Perondini ALP (1968) Estudos citológicos e autorradiográficos dos cromossomos politênicos de Sciara ocellaris. Ph.D. thesis, University of São Paulo Rasch EM (1970) Two-wavelenght cytophotometry of Sciara salivary gland chromosomes. In: Wield GL, Bahr GF (eds) Introduction to Quantitative Cytochemistry. Academic, New York Ribeiro ER (1975) Efeito do 2,3 dihidro-1H inidazo (1,2-b) pirazol (IMPY) sobre o desenvolvimento dos pufes de DNA de Bradysia hygida. M.Sc. thesis, University of São Paulo Rudkin GT, Corlette SL (1957) Disproportionate synthesis of DNA in a polytene chromosome region. Proc Natl Acad Sci USA 43: 964–968 Santelli RV, Machado-Santelli GM, Pueyo MT, Navarro-Cattapan LD, Lara FJ (1991) Replication and transcription in the course of DNA amplification of the C3 and C8 DNA puffs of Rhynchosciara americana. Mech Dev 36: 59–66 Santelli RV, Siviero F, Machado-Santelli GM, Lara FJ, Stocker AJ (2004) Molecular characterization of the B-2 DNA puff gene of Rhynchosciara americana. Chromosoma 113: 167–176 Sauaia H, Alves MAR (1968) A description of a new species of Bradysia (Diptera, Sciaridae). Pap Avul Zool 22: 85–88 Sauaia H, Laicine EM, Alves MA (1971) Hydroxyurea-induced inhibition of DNA puff development in the salivary gland chromosomes of Bradysia hygida. Chromosoma 34: 129–151 Simon CR, de Almeida JC (2004) Programmed cell death in Bradysia hygida (Diptera, Sciaridae) salivary glands presents apoptotic features. Genesis 40: 22–31 Soares MA, Monesi N, Basso LR, Jr., Stocker AJ, Paçó-Larson ML, Lara FJ (2003) Analysis of the amplification and transcription of the C3-22 gene of Rhynchosciara americana (Diptera: Sciaridae) in transgenic lines of Drosophila melanogaster. Chromosoma 112: 144–151 Spradling AC (1981) The organization and amplification of two chromosomal domains containing Drosophila chorion genes. Cell 27: 193–201 Stocker AJ, Pavan C (1974) The influence of ecdysterone on gene amplification, DNA synthesis, and puff formation in the salivary gland chromosomes of Rhynchosciara hollaenderi. Chromosoma 45: 295–319

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Stocker AJ, Troyano-Pueyo M, Pereira SD, Lara FJS (1984) Ecdysteroid titers and changes in chromosomal activity in the salivary glands of Rhynchosciara americana. Chromosoma 90: 26–38 Stocker AJ, Yokosawa J, Soares MA, Cadavid EO (1996) DNA replication and amplification during the final cycle of politeny insciarid gland chromosomes and their control by ecdysone. Ciência e Cultura 48: 306–312 Stocker AJ, Amabis JM, Gorab E, Elke C, Lezzi M (1997) Antibodies against the D-domain of a Chironomus ecdysone receptor protein react with DNA puff sites in Trichosia pubescens. Chromosoma 106: 456–464 Swift H (1962) Nucleic acids and cell morphology in Dipteran salivary glands. In: The Molecular Control of Cellular Activity. McGraw-Hill, New York Swimmer C, Fenerjian MG, Martinez-Cruzado JC, Kafatos FC (1990) Evolution of the autosomal chorion cluster in Drosophila. III. Comparison of the s18 gene in evolutionarily distant species and heterospecific control of chorion gene amplification. J Mol Biol 215: 225–235 Thummel CS (1996) Files on steroids-Drosophila metamorphosis and the mechanisms of steroid hormone action. Trends Genet 12: 306–310 Urnov FD, Liang C, Blitzblau HG, Smith HS, Gerbi SA (2002) A DNase I hypersensitive site flanks an origin of DNA replication and amplification in Sciara. Chromosoma 111: 291–303 Winter CE, de Bianchi AG, Terra WR, Lara FJS (1977) Relationships between newly synthesized proteins and DNA puff patterns in salivary glands of Rhynchosciara americana. Chromosoma 61: 193–206 Winter CE, de Bianchi AG, Terra WR, Lara FJ (1980) Protein synthesis in the salivary glands of Rhynchosciara americana. Dev Biol 75: 1–12 Wu N, Liang C, DiBartolomeis SM, Smith HS, Gerbi SA (1993) Developmental progression of DNA puffs in Sciara coprophila: amplification and transcription. Dev Biol 160: 73–84 Yokosawa J, Soares MA, Dijkwel PA, Stocker AJ, Hamlin JL, Lara FJ (1999) DNA replication during amplification of the C3 puff of Rhynchosciara americana initiates at multiple sites in a 6 kb region. Chromosoma 108: 291–301 Zhang H, Tower J (2004) Sequence requirements for function of the Drosophila chorion gene locus ACE3 replicator and ori-beta origin elements. Development 131: 2089–2099

Chapter 7

The Ecdysteroids’ Effects in the Control of Cell Proliferation and Differentiation David Siaussat, Patrick Porcheron, and Stephane Debernard

Abstract In insects, the steroid hormone 20-hydroxyecdysone (20E) plays a critical role in the control of cellular proliferation and differentiation. The cells show different responses to 20E according to the concentration to which they are exposed. The 20E at 10−7 M induces an inhibition of growth by a blockage of cells in G2/M and long term morphological transformation. The establishment of 20E responsive cell lines provided potentialities to investigate the molecular events responsible for these cellular responses as well as theirs dynamics through the cell cycle. In the Plodia interpunctella IAL-PID2 cell line, an optimal period of sensitivity of cells to 20E, in inducing G2/M arrest, was preferentially located at the transition S/G2 with a high induction of EcR, USP and HR3 mRNAs and a decrease in the expression level of B-cyclin at the end of G2 phase. On the other hand, the 20E-induced cytoskeleton rearrangement was accompanied by a redistribution of cytoplasmic microtubules which was concomitant with an increase in the β tubulin mRNA amount. The use of RNAi technique allowed to demonstrate that inhibiting the induction of EcR, USP and HR3 suppressed the 20E effects on the synthesis of B-cyclin and β tubulin then consequently prevented the arrest and the transformation o`f IAL-PID2 cells. This functional approach revealed that 20E was able to regulate the cellular differentiation and proliferation by acting on regulators of cell cycle and cytoskeleton proteins through a genomic signaling pathway involving EcR, USP and HR3. Keywords Ecdysteroid • nuclear receptor • proliferation • differentiation, 20-hydroxyecdysone • imaginal wing cells

7.1

Introduction

Through molting, insects periodically shed their rigid exoskeleton to accommodate growth during the larval life. When the larva has attained its characteristic size, the metamorphic molt(s) is initiated to produce an adult exhibiting sequential D. Siaussat (), P. Porcheron, and S. Debernard Université Pierre et Marie Curie (UPMC, Paris 6), UMR 1272A Physiologie de l’Insecte, Signalisation et Communication, 7 Quai St Bernard, 75252 Paris Cedex 05, France e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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polymorphism changes from larvae to pupae and then to the adult stage (Riddiford and Truman, 1993). This radical reorganization in body form during the metamorphosis depends on periodic changes in hemolymphatic levels both of the steroid hormone 20-hydroxyecdysone (20E) and the sesquiterpenoid juvenile hormone (JH). The 20E activates both the histolysis of larval structures and the morphogenesis of adult structures by the differentiation of diploid imaginal cells. These 20E-regulated biological responses require a hormonal action on cellular functions including cell proliferation, expansion, differentiation, programmed cell death that occurs at various times during metamorphosis in a cell- and tissue-specific manner to orchestrate this developmental transition (Yin and Thummel, 2005). Most of 20E-induced biological effects have been associated to genomic mechanisms that involve a hormonal binding to a nuclear heterodimeric complex composed of two members of the nuclear receptor superfamily: the ecdysone receptor (EcR) (Koelle et al., 1991; Schubiger et al., 1998; Bender et al., 1997) and Ultraspiracle (USP) (Yao et al., 1992; Swevers et al., 1996; Kapitskaya et al., 1996), the insect homolog of the vertebrate retinoid X receptor (RXR) (Mangelsdorf et al., 1992, 1995). Some developmental studies have shown that EcR and USP exist under isoforms that are differently expressed according to the tissue and developmental stage, thus contributing to the spatial and temporal diversity of the response to 20E (Perera et al., 1999; Talbot et al., 1993; Bender et al., 1997; Jindra et al., 1996; Schubiger et al., 1998). The activation of EcR/USP complex by 20E triggers the sequential expression of genes encoding transcription factors that ultimately regulate the activity of target genes (Henrich et al., 1999; Andres and Thummel, 1992; Thummel, 1995). Most of these genes have been first characterized in Drosophila melanogaster as transcription factors such as E75 (Segraves and Hogness, 1990), E74 (Thummel et al., 1990), E78B (Stone and Thummel, 1993), BRC (Dibello et al., 1991) and DHR3 (Koelle et al., 1992). E75 and HR3 are induced in parallel with 20E pulses throughout development (Koelle et al., 1992; Horner et al., 1995) and like other early late genes, the expression of HR3 is delayed relative to that of the E75 early gene (Horner et al., 1995; Huet et al., 1995). Genetic analysis indicated that Drosophila E75 and HR3 play a crucial role in the coordination of 20E-mediated molecular cascades leading to morphological and cellular changes that occurred at different developmental stages. Nevertheless, it yet subsists numerous gaps in the ecdysteroid signaling pathways by which the hormonal signal is transduced in order to direct the biological responses. In lepidopteran larvae, 20E is known to play a critical role in the control of cell proliferation in imaginal discs, optic lobes and epidermis (Oberlander, 1985; Kawasaki, 1995; Wieglus and Gilbert, 1978; Kato and Riddiford, 1987; Champlin and Truman, 1998). The imaginal cells show different responses to 20E according to the developmental stage and the concentration to which they are exposed (Koyama et al., 2004; Kawasaki, 1995). Through the young instars, the rate of cell proliferation of the imaginal cells is rather constant and independent of the cyclic surges of 20E. In the late fourth instar, the imaginal cells acquire the competence to respond to 20E concomitantly with their entrance into the reversible stage of pupal commitment (Fig. 7.1). At the same time, 20E induces an acceleration of cell proliferation by controlling the G2/M and G1/S transitions of cell cycle at

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Genetic reprogramming

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Arrest in G2/M

Fig. 7.1 Schematic representation of hormonal control of development of Lepidoptera imaginal discs. Through the young instars, the rate of proliferation of the imaginal cells is rather constant and independent of cyclic surges of 20E. After a genetic reprogramming of imaginal discs at the beginning of the last larval instar, the proliferation of imaginal cells becomes isometric. At the middle of the stage, the imaginal cells acquire the competence to respond to 20E concomitantly with their entrance into their reversible stage of pupal commitment in presence of 20E circulating comprised between 10−8 and 5 × 10−8 M. At the same time, 20E induces an acceleration of cell proliferation by controlling the G2/M and G1/S transitions of cell cycle. At the end of last larval instar, a high proportion of imaginal cells are blocked in G2/M phase when the hemolymphatic titer of 20E rises above 10−7 M. This arrest coincides with the entrance of imaginal cells into the pupal differentiation programme leading to the eversion and the formation of adult structures

hormonal concentrations comprised between 10−8 and 5 × 10−8 M (Koyama et al., 2004) (Fig. 7.1). At the end of last larval instar, a high proportion of imaginal cells are blocked in G2/M when the hemolymphatic titer of 20E rises above about 10−7 M (Fain and Stevens, 1982; Graves and Schubiger, 1982) (Fig. 7.1). This arrest coincides with the entrance of imaginal cells into the pupal differentiation programme leading to the formation of adult structures. Theses data revealed a 20E interaction with the cell cycle in the control of cellular growth. The establishment of various insect cell lines that have retained sensitivity to ecdysteroids provided potentialities for specifying the mode of action of these hormones in the control of cellular proliferation and differentiation processes. Kc embryonic cells from Drosophila melanogaster Diptera and IAL-PID2 cells derived from imaginal wing discs of Plodia interpunctella Lepidoptera respond to 20E treatment by an arrest of cell growth and long term spectacular modifications in the shape of cells with the formation of pseudoepithelial aggregates structures (DePasquale et al., 1994; Lin et al., 2003; Benayahu, 1997; Stevens et al., 1980; Mottier et al., 2004; Siaussat et al., 2004a, b, 2007). In Kc and IAL-PID2 cells, fluorescent flow cytometry analysis of cell cycle have shown that 20E-induced proliferative arrest resulted from a

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blockage of cells in G2/M phase (Siaussat et al., 2004a; Mottier et al., 2004; Stevens et al., 1980). This 20E-induced G2/M arrest was correlated with a high induction of Plodia interpunctella USP, EcR and HR3 (Fujiwara et al., 1995; Jindra et al., 1996). This result suggested that a quantitative increase of EcR, USP and HR3 transcripts is somehow essential in inducing G2/M arrest. It only existed some descriptive information on the control of cell cycle by ecdysteroids and there are no functional data on the 20E signaling transduction pathway responsible for G2/M arrest.

7.2

Interaction Between 20E and Cyclins in the Regulation of Cell Growth

The cell cycle is regulated by the action of a family of serine/threonine kinases known as cdks. Different cdks function at different times of the cell cycle and are positively regulated through binding to subunits called cyclins, a process that is essential for cdk activity (Buttitta et al., 2007; Meyer et al., 2000; Boylan and Gruppuso, 2005; Saab et al., 2006). More is known in mammals about the mechanisms involved in cell cycle regulation by steroid hormones. A progesterone treatment on breast cancer cells induced an arrest of cells in the G1 phase (Clarke and Sutherland, 1990; Musgrove et al., 1997). This cell cycle arrest was related to an inhibition of expression of cyclins D and E (Musgrove et al., 1998) which control the progression of cells during the G1 phase (Norbury and Nurse, 1992; Ravitz and Wenner, 1997). Genetic analysis in Drosophila melanogaster have revealed that both A- and B-type cyclins control G2/M transition (Knoblich and Lehner, 1993; Lehner and O’Farrell, 1990; Lehner et al., 1991). In IAL-PID2 cells, a decrease in the expression of B cyclin was observed as early as 8 h of exposure to 20E. This decline preceded the beginning of accumulation of cells in the G2/M phase, which occurred after 16 h of 20E treatment and could be responsible for further cellular arrest (Mottier et al., 2004). These findings brought in evidence that ecdysteroids were able to regulate the cell proliferation by acting on cell cycle regulators as cyclins.

7.3

7.3.1

Dynamic of 20E-Induced Molecular and Cellular Events in G2/M Arrest Synchronization of Insect Cells and Period of Sensitivity to 20E

The availability of synchronized insect cell model was essential to investigate events involved in the regulation of cell cycle by 20E. A protocol of cell synchronization by serum deprivation have been established from IAL-PID2 cell line (Lynn and Oberlander, 1983) and used in conjunction with studies on 20E action. Serum deprivation brought IAL-PID2 cells to a quiescence phase in which most of cells were in the

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G0/G1 stage (Hatt et al., 1997). The addition of foetal bovine serum (FBS) triggered cell cycle resumption and a majority of cells was recovered in S phase after an 8 h period following FBS supplement. Nevertheless, this synchrony was short-lived and several separate peaks of labelled mitotic cell could be detected (Hatt et al., 1997; Auzoux-Bordenave et al., 2002).Various processes of synchronization have been tested on a wide variety of insect cell types using reversible inhibitors of DNA synthesis such as thymidine, hydroxyurea, nocodazole, aphidicolin or fluorodeoxyuridine (Cress and Gerner, 1977; Hamlin and Pardee, 1976; Lynn and Hink, 1978; Tobey et al., 1990; Ananiev et al., 1977; Pittman et al., 1994; Minami et al., 1994; Gerenday et al., 1997). The chemical synchronization of the cell cycle is based on the use of appropriate drugs which have the property to be differentially toxic during the different phases of the cell cycle (Sinclair, 1967). These drugs evoke an arrest of the cell division cycle resulting in accumulation of cells at precise stage of cell cycle. The removal of drugs releases a cohort of cells that resume their cyclic activity in a good synchrony. The most effective chemical synchronization protocols were established from Plodia interpunctella and Aedes albopictus cells using hydroxyurea (Gerenday et al., 1997; Siaussat et al., 2004). The target for hydroxyurea toxicity is ribonucleotide reductase (Gerenday et al., 1997; Siaussat et al. 2004a), a rate-limiting enzyme required for DNA synthesis. A high degree of synchrony was reached when cells were exposed to two consecutive hydroxyurea treatments at 1 mM for 36 h spaced 16 h apart with 90% of cells blocked at the transition G1/S. After the removal of drug, 80% of cells were quickly recovered in S and this cohort of cells progressed through the cell cycle to enter progressively into G2 then G1 (Fig. 7.2a). Under these conditions of synchronization, a 20E treatment induced an inhibition of cell growth by an arrest of 90% of cells in G2/M phase (Siaussat et al., 2004a) (Fig. 7.2b). The efficient of this induction depended both on the time of hormonal treatment related to the position of cells in the cell cycle and the applied hormonal concentration. An optimal period of sensitivity of cells to 20E, in inducing G2/M arrest, was preferentially located at the transition S/G2 of the IAL-PID2 cell cycle in presence of 20E at 10−7 and 10−6 M (Siaussat et al., 2005) (Fig. 7.3). These concentrations were close to the hemolymphatic titer of 20E (Cherbas et al., 1989; Nijhout, 1994) and sufficient to induce in vitro the cell morphological differentiation (Milner and Sang, 1974; Milner, 1977).

7.3.2

Cell Cycle Profiles of EcR, USP, HR3, B-Cyclin Associated to G2/M Arrest

The establishment of efficient cell synchronization allowed to study throughout the cell cycle the dynamic of molecular and cellular events involved in 20E-induced G2/M arrest. From synchronized IAL-PD2 cells, the expression patterns of EcR, USP, E75, HR3 and B-cyclin were reported in correlation with this hormonal response. In the normal conditions, when cells progressed through the different phases of cell cycle, EcR, USP were constitutively expressed at low level and PHR3 mRNA was never detected (Fig. 7.4a–c). In presence of 20E, EcR, USP and HR3 mRNAs were induced

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Fig. 7.2 Protocol of synchronization by hydroxyurea in the IAL-PID2 cell line. A high degree of synchrony was reached when cells were exposed to two consecutive hydroxyurea treatments at 1 mM for 36 h spaced 16 h apart leading to a massive arrest of cells at the transition G1/S (90%). (a) Only 2 h after the removal of the drug, 70% of cells were recovered in S. This cohort of cells began to enter into G2 at 10 h then kept on progressing through the cell cycle to re-enter progressively into G1 after 18 h. (b) Under these conditions, 20E at physiological concentration induced an inhibition of cell growth by an arrest of 90% of the cells in G2/M phase. In the bubble of each figures, corresponding cytometry profiles are presented (See Color Plates)

7

The Ecdysteroids’ Effects in the Control of Cell Proliferation and Differentiation

Fig. 7.3 Window of sensitivity to 20E of IAL-PID2 cells. An optimal period of sensitivity of cells to 20E, in inducing G2/M arrest, was preferentially located at the transition S/G2 of the IAL-PID2 cell cycle in presence of 20E at 10−7 and 10−6 M. The duration of each phase of the cell cycle are indicated

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G1 (12-14h)

G2 (9-11h)

S (10-12h) Window of sensitivity to 20E

as early as the beginning of S phase and the highest inductions were observed at the S/G2 transition (Siaussat et al., 2004a) (Fig. 7.4a–c). These data were in agreement with the high levels of USP, EcR and HR3 expressions reported in Manduca sexta wing discs and epidermis in response to the rising ecdysteroid titer responsible for G2/M arrest just before the pupation (Fujiwara et al., 1995; Jindra et al., 1996). As concerns B cyclin, in untreated cells, its expression increased progressively to reach a maximum at the beginning of the G2/M phase then declined just before the re-enter in G1/G0 phase (Fig. 7.5). On the other hand, 20E induced a sharp decline in the level of B cyclin mRNA that preceded the beginning of accumulation of cells in G2/M phase (Mottier et al., 2004) (Fig. 7.5). The control of G2/M transition is associated to a high expression of B cyclin. Therefore, the inhibitory effect of 20E on B cyclin expression, at the end of G2 phase, would be in part responsible for the blocking of cells in G2/M. Taken together, these results provided evidence that the 20E-induced G2/M arrest was correlated to a high induction of EcR, USP, E75 and HR3 mRNAs at the S/G2 transition and a decrease in B cyclin mRNA level at the end of G2 phase.

7.4

7.4.1

Identification of 20E Signaling Pathway in Cell Proliferation and Differentiation RNA Interference in Insect Cells

RNA interference (RNAi) begins to be a quite well-known mechanism for posttranscriptional gene silencing observed in invertebrates as Caenorhabditis elegans (Fire et al., 1998) or Drosophila melanogaster and also plants (Kennerdell and Carthew, 1998; Misquitta and Paterson, 1999; Sharp, 1999). This evolutionarily conserved mechanism for silencing gene expression is also present in vertebrates and mammalian cells. This phenomenon is based on double-stranded RNA (dsRNA)

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a

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S

G2/M

20E Fig. 7.4 Expression profiles of EcR, USP and HR3 in synchronized IAL-PID2 cells

The Ecdysteroids’ Effects in the Control of Cell Proliferation and Differentiation

Fig. 7.5 Schematic representation of expression profile of B cyclin mRNA in synchronized IAL-PID2 cells. In absence of 20E, the amount of PcycB mRNA increased progressively to reach a maximum at the beginning of the G2/M phase then declined just before the re-enter in G1/G0 phase. In presence of 20E, a sharp decline in the level of PcycB mRNA occurred preceding the beginning of accumulation of cells in G2 phase

Relative expression (%)

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that inhibits gene expression in a sequence-specific manner by triggering mRNA degradation (Hannon, 2002). Currently, this technology has a high potentiality as a quick and powerful tool for functional studies of genes in all domains of research (Fraser et al., 2000; Herold et al., 2003). Thus, recent experiments brought to light the in vitro usefulness of RNAi for the determination of gene functions in the synchronization of cellular processes (Maiato et al., 2003; Giet and Glover, 2001; Somma et al., 2002). This RNAi technique has been reported to be effective in mammalian somatic, neuronal and embryonic cell lines, including Hela, HEK293, P19 (Harborth et al., 2001; Paddison et al., 2002). In insect cell lines, its efficacy was mainly established in Drosophila melanogaster Diptera, and, under normal conditions, the Drosophila S2, KC, BGG2-C6 cells, originally isolated from different embryonic cell types, take up external nucleic acid with a high efficiency (Clemens et al., 2000). Using a lipophilic transfection reagent, the Lipofectamine 2000, the RNAi technique has been recently extended to the 20E responsible IAL-PID2 cell line of Plodia interpunctella Lepidoptera. The rapidity, efficiency, specificity and longevity of the RNAi effects on ecdysteroid inducibility of genes in IAL-PID2 cells make this an ideal system both for identifying the role of these genes in the ecdysteroid-regulated cellular functions and aiding in the elucidation of the mechanisms that allow dsRNA to inhibit target protein synthesis (Siaussat et al., 2007). The RNAi technique was efficient toward a broad range of ecdysteroid-inducible proteins belonging to the nuclear receptors superfamily.

7.4.2

Interaction Between EcR, USP, HR3 and B Cyclin in G2/M Arrest

From the IAL-PID2 cell line, recent works were undertaken to dissect the 20E molecular cascade responsive for G2/M arrest of cells using dsRNAi method.

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The obtained results showed that inhibiting the 20E induction of EcR, USP and HR3 mRNAs prevented the decreased expression of B cyclin (Fig. 7.6) and consequently the G2/M arrest of cells (Fig. 7.7). On the other hand, a decreased induction of PHR3 mRNA was observed by blocking the activation of EcR or USP gene and applying EcR dsRNA reduced the level of USP mRNA induction (Fig. 7.8). This functional analysis revealed the participation of EcR, USP and HR3 in a 20E common signaling pathway that directs the growth in insect cells by regulating B cyclin expression. In Manduca sexta, the promoter region of the HR3 gene contains four putative ecdysone response elements (EcRE) and is activated by 20E through a binding of EcR/USP complex to EcRE (Riddiford et al., 2003; Hiruma and Riddiford, 2004). In Drosophila, it has demonstrated that 5' flanking sequences of 20E regulated target genes are composed of motif homologous to EcRE which appear to be involved in the level of the ecdysone response (Bruhat et al., 1993; Tourmente et al., 1993). To gain more precise information on the relationships between EcR, USP, HR3 and B cyclin in the 20E signaling pathway, it would be important to characterize the presence of EcRE and a HR3 response element in the promoter regions of a

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Fig. 7.6 Interaction between EcR, USP, HR3 and B cyclin in IAL-PID2 cells. At 10 h after applying 2 µg/ml EcR, USP or HR3 dsRNA combined with Lipofectamine 2000 at 0.5 µg/ml, the cells were treated with 10−7 M 20E for various exposure times. The levels of B cyclin expression in the presence of HR3 (a), USP (b) or EcR (c) were determined by northern blotting

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Fig. 7.8 Relationship between EcR, USP and HR3 in IAL-PID2 cells. At 10 h after applying 2 µg/ml EcR, USP or HR3 dsRNA combined with Lipofectamine 2000 at 0.5 µg/ml, the cells were treated with 10−7 M 20E for various exposure times. At each time, the induction level of HR3 mRNA was determined by northern blotting in the presence of EcR dsRNA (a) or USP dsRNA (b) and the induction level of USP mRNA in the presence of EcR dsRNA (c). For northern blots controls, a fragment of the cDNA encoding the RpL8 ribosomal protein of Plodia interpunctella was used

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the Plodia HR3 and B cyclin genes respectively. Some experiments performed in Drosophila and Plodia indicated that the A cyclin acts in synergy with B cyclin to control the G2/M transition and that the 20E is also able to modulate the expression of A cyclin in inducing G2/M arrest (Mottier et al., 2004). It would be interesting to determine whether 20E interacts with these two proteins through a common signaling pathway in the regulation of cell growth. In humans, target cells and organs are regulated by a complex interplay of genomic and non-genomic signaling mechanisms of steroid hormones, and the integrated action of these machineries has important functional roles in a variety of pathophysiological processes (Truss and Beato, 1993; Watters et al., 1997). The steroid-regulated cell proliferation is related to steroid effects on the initiation and promotion of cancer (Xing et al., 1997; Lokeshwar et al., 1996). It has been recently discovered that steroid hormones exert antiproliferative effects on human cancerous cell lines through multiple transduction pathways. Proposed mechanisms accounting for the cell cycle arrest include transcriptional repression of cyclins and cdks, the transcriptional activation of cdk inhibitors, inhibition of cdk activity by phosphorylation via the wee1 kinase and the activation of the extracellular signal-regulated kinases (ERKs) (Greenberg et al. 2002; Pradeep et al. 2002; Wright et al. 2003; Paruthiyil et al. 2004; Stumpff et al. 2004; Horner-Glister et al. 2005; Pradeep and Menon 2005; Zapata et al. 2005; Okamoto and Sagata 2007; Xia-Dong et al. 2007). In insects, ecdysteroids could also control the cell proliferation through several mechanisms similar to those described in humans.

7.4.3

Interaction Between EcR, USP, HR3 and b -Tubulin in Cell Morphological Differentiation

The 20E-induced G2/M arrest is followed by a morphological differentiation of cells with the formation of long cytoplasmic extensions accompanied by a redistribution of microtubules which were accumulated in the pseudopodia and arranged in a parallel direction to the longitudinal axis (Fig. 7.9b) (Sobrier et al., 1989; Judy, 1969; Courgeon, 1972; Dinan et al., 1990; Palli et al., 1995; Fretz and Spindler, 1999; Cassier et al., 1991; Porcheron et al., 1991; Mottier et al., 2004; Siaussat et al., 2004b). This cytoskeleton reorganization was concomitant with changes in the expression level of β tubulin that is composed of non-covalently bound α- and β-chains and represents the unit of all microtubules in eukaryotic cells (Montpied et al., 1988; Sobrier et al., 1986) (Fig. 7.9a). Using RNAi, it has been demonstrated that 20E-induced cell morphological transformation resulted from an increased synthesis of Plodia β tubulin and that the silencing of EcR, USP and HR3 genes blocked the 20E effects on the β tubulin expression and on the shape of IAL-PID2 cells (Siaussat et al., 2007). The inhibitory effect of RNAi on the 20E-induced responses demonstrated that EcR, USP and HR3 could also be involved in the control of morphological differentiation of insect cells through a modulation of β tubulin synthesis.

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It has been recently discovered that the human cancerous cell lines HT-1080 and LNCaP responded to androgen treatment by undergoing rapid cytoskeleton transformations that occurred in some minutes, a time lag noncompatible with the classical scheme of a nuclear receptor action (Chauhan et al., 2004; Papakonstanti et al., 2003; Kampa et al., 2002; Castoria et al., 2003). The cytoskeleton reorganization of these tumorous cells was mediated by a steroid extranuclear signaling pathway that involved an hormonal binding to membrane receptor with the sequential activation of protein kinases, GTPases and the production of lipidic second messengers such as phosphatidylinositol-3,4,5-triphosphate (IP3) (Papakonstanti et al., 2003). In insects, some studies had suggested the existence of extranuclear signaling for ecdysteroids by showing the involvement of nitric oxide in the control of neuroblast proliferation by 20E during Manduca sexta metamorphosis and by revealing the presence of a membrane ecdysteroid receptor in the silk gland of Bombyx mori and the mature nervous system of Drosophila melanogaster (Champlin and Truman, 2000; Elmogy et al., 2006; Srivastava et al., 2005).

198 Fig. 7.10 Schematic representation of control of cell proliferation and differentiation by ecdysteroïds. At the transition S/G2, the 20E binds to EcR/USP heterodimeric complex and induces a maximum induction of these two partners then HR3. This 20E signaling pathway is responsible for G2/M arrest of cells by inhibiting the expression of B cyclin and is also involved long term in the morphological differentiation of cells through an increase in the synthesis of β tubulin (See Color Plates)

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Finally, ecdysteroids are able to regulate the differentiation and proliferation cellular in insects by acting on regulators of cell cycle and cytoskeleton proteins through a common genomic signaling pathway involving EcR, USP and HR3 (Fig.7.10).

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Chapter 8

Applications of RNA Interference in Ecdysone Research Garry N. Hannan, Ronald J. Hill, Skarlatos G. Dedos, Luc Swevers, Kostas Iatrou, Anjiang Tan, Ramaseshadri Parthasarathy, Hua Bai, Zhaolin Zhang, and Subba R. Palli

Abstract RNA interference technology (RNAi) has been demonstrated to provide an effective tool in ecdysone receptor research. This chapter describes specific examples in three insect models. (1) Selective RNAi of the expression of the endogenous messenger RNA encoding EcR of Sf9 tissue culture cells and replacement with EcRs from other insects should permit cell-based screening for potential insecticides against a variety of pests. (2) The applicability of the technique of RNAi in Bombyx mori-derived Bm5 cells combined with the availability of a robust ecdysone reporter assay opens the possibility to carry out screens for the function of Bombyx genes in the primary ecdysone response pathway. (3) RNA interference (RNAi) works very well in the model insect, the red flour beetle, Tribolium castaneum. Methods for rearing beetles, synthesis of double-stranded RNA (dsRNA), injection of dsRNA RNA and evaluation of its efficacy by qRT-PCR analysis are presented. RNAi- mediated knock-down in the expression of genes such as EcR, USP, E75, HR3, HR4, HR39, HR38, SVP, FTZ-F1 and HR51 caused derailment of development from larvae to pupae and pupae to adult stages. These studies confirm the role of these nuclear receptors in 20E signal transduction. Keywords RNA interference • dsRNA • Bombyx mori • Spodoptera frugiperda • Tribolium castaneum • insect cell lines • Bm5 • Sf9 • EcR, usp • HR3 • E75 • FTZF1 • Broad • ecdysone-responsive genes • ecdysone regulatory pathway • molting, moulting • metamorphosis • nuclear receptors • transcription factors G.N. Hannan and R.J. Hill () CSIRO Molecular and Health Technologies, PO Box 184, North Ryde, New South Wales 1670, Australia S.G. Dedos, L. Swevers (), and K. Iatrou Insect Molecular Genetics and Biotechnology, Institute of Biology, National Centre for Scientific Research “Demokritos”, P. Grigoriou & Neapoleos Str., Aghia Paraskevi Attikis, 153 10 Athens, Greece A. Tan, R. Parthasarathy, H. Bai, Z. Zhang, and S.R. Palli () Department of Entomology, College of Agriculture, University of Kentucky, Lexington, KY 40546, USA e-mails: [email protected], [email protected], [email protected]

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Introduction

RNA interference (RNAi) is a phenomenon leading to specific silencing of genes as a result of destruction of their RNA transcripts via a ribonuclease whose target selectivity is guided by sequence information derived from exogenous double-stranded RNA (dsRNA). The mechanism was originally discovered in nematodes (Fire et al., 1998) and was subsequently observed in a whole host of organisms including mammals (Martinez et al., 2002), insects (Kennerdell and Carthew, 2000) and plants (Waterhouse et al., 2001). The process is generally initiated by cleavage of the introduced dsRNA to approximately 22 base-pair RNA molecules by a bidentate ribonuclease called Dicer (Bernstein et al., 2001). These short dsRNA fragments, referred to as short interfering RNA (siRNA), are taken up by RNA-induced silencing complexes (RISC) where their sequence information guides ribonuclease target selectivity (Ui-Tei et al., 2004). The injection of dsRNA was shown in the early experiments employing C. elegans to result in the selective elimination of cytoplasmic transcripts bearing the same sequence as the introduced dsRNA (Montgomery et al., 1998). Double-stranded RNA bearing sequence specific for the ecdysone receptor isoform-A of the cockroach Blatella germanica has been injected into B. germanica sixth instar nymphs or adults to allow effective analysis of the role of the EcR isoform in development (Cruz et al., 2006). However, in general the effects on gene expression mediated by injected dsRNA tend to be transient. For this reason, DNA constructs have often been employed for the in vivo expression of RNA sequences designed to fold back into hairpin double stranded structures and this strategy has been shown to result in effective RNAi, for example, in C. elegans (Tavernarakis et al., 2000) and in D. melanogaster (Kennerdell and Carthew, 2000). Lam and Thummel (2000) have created transgenic D. melanogaster lines expressing head-to-head RNA sequences from within DmEcR, designed to fold back into double stranded RNA sequences, to generate specific defects in larval moulting and metamorphosis. Recently, this approach has been generalised to the production of a genome-wide library of RNAi transgenes in some 20,000 transgenic D. melanogaster lines to permit selective disruption of 88% of the predicted protein encoding genes in the Drosophila genome (Dietzl et al., 2007). The following sections present specific examples of the application of the RNAi approach in ecdysone research.

8.2

8.2.1

Selective RNAi of an EcR Gene in a Spodoptera frugiperda (Lepidoptera) Cell Line Introduction

Tissue culture cells of the D. melanogaster embryonic cell line, Kc, on incubation with physiological concentrations of 20-hydroxyecdysone, become arrested at G2 phase of the cell cycle (Stevens et al., 1980). If incubation with hormone is continued, then after a period of several days to weeks, cell division is resumed and the cells that have resumed the cell cycle are resistant to effects of ecdysteroid (Stevens

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and O’Connor, 1982). Hormone insensitivity is associated with loss of ecdysteroid receptors; after long periods in culture in the absence of hormone the hormone receptor level returns to that in naïve control cells (Stevens and O’Connor, 1982). The lowering of the ecdysone receptor level in Kc cells also occurs on continued incubation in the presence of the environmentally-friendly insecticide, RH5849 and this was used as evidence for a common site of action for the insecticide and 20-hydroxyecdysone (Wing, 1988). Similarly, continuous incubation of D. melanogaster S2 cells with 20-hydroxyecdysone leads to the production of ecdysone-resistant cells that were used by Koelle et al. (1991) to demonstrate that transfection with constructs expressing a cloned cDNA encoding a putative DmEcR protein, in fact, conferred ecdysone responsiveness on an appropriate reporter plasmid. Sf 9 cells derived from the lepidopteran, Spodoptera frugiperda, provide robust insect cells capable of growth in vitro as monolayers or as suspension cultures. They are readily transfected with plasmid DNA and transfection of these cells with a plasmid expressing an ecdysone receptor protein from an insect pest, along with an appropriate reporter plasmid, offers an attractive cell-based system for screening chemical libraries for ligands binding to the pest’s receptor. Results from such assays would be complicated by the endogenous SfEcR protein expressed in the Sf9 cells. It might be possible to lower this background by continuous incubation of the cells with physiological levels of 20-hydroxyecdysone. However, in this account we describe an alternative strategy in which selective suppression of expression of SfEcR is effected by RNAi allowing effective replacement of the endogenous SfEcR with an EcR from the sheep blowfly, Lucilia cuprina, expressed from a transfected plasmid encoding LcEcR.

8.2.2

Experimental

8.2.2.1

PCR Cloning of a 631 Base Pair Segment from Within SfEcR

High-quality total RNA was prepared from Sf9 cultured cells by the guanidine thiocyanate procedure of Okayama et al. (1987). A 631 bp fragment encompassing most of the D-domain and a 5’-segment of the E-domain of the SfEcR cDNA (nucleotides 837 to 1,468 within the sequence SfEcR sequence, accession number AF411254) was reverse transcribed and PCR amplified from Sf9 cell total RNA using a forward primer 5’-GCCTCGGGGTACCATTATAAC-3’, and a partially degenerate reverse primer, 5-GGNAGNCC(C/T)TTNGCGAA(C/T)TC-3’; the product was blunt-end cloned into pCR2.1-TOPO (Invitrogen) to construct pTOPO8. 8.2.2.2

PCR Cloning of the SfEcR D-Domain Flanked by BamHI and Sf iI Sites and Also by NotI and Sf iI Sites

Oligonucleotides, 5’-CGCGGATCCGAGGCCCGAGTGTGTGGTG-3’ (providing a BamHI tail) and 5’-GGAAGGCCTAGATGGCCGTCTTCATCCGACTGCCAG-3’ (with an SfiI tail) were used to prime a PCR reaction from a TOPO8 template to produce a 363 bp fragment flanked by the unique restriction sites BamHI and SfiI.

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Oligonucleotides 5’-GAATGCGGCCGCTAGGCCCGAGTGTGTGGTG-3’ (with a NotI tail) and 5’GGAAGGCCATCTAGGCCGTCTTCATCCGACTGCCAG-3’ (with an SfiI tail) were used to prime a second PCR reaction also employing a TOPO8 template to produce a 362 bp fragment flanked by the unique restriction sites NotI and SfiI. The two fragments flanked by BamHI and SfiI and by NotI and SfiI sites, respectively, were individually cloned, employing these sites, into pCR2.1-TOPO and grown up. The fragments were then released from the plasmids by digestion with BamHI and SfiI on the one hand and NotI and SfiI on the other.

8.2.2.3

Construction of the RNAi Plasmid, SfEcRi, Designed to Express Head-to-Head Inverted Repeats of the SfEcR D-Domain RNA Sequence

pIE1-3 (Novagen) that had been cut with BglII and NotI, along with the SfEcR D-domain sequences flanked by BamHI + SfiI and NotI + SfiI sites, respectively, were subjected to a three-way ligation. This resulted in a construct expressing a 718-bp head-to-head concatamer of the DNA fragment derived from the D-domain of SfEcR under the control of a constitutive baculovirus ie-1 promoter in the new plasmid pSfEcRi (see Fig. 8.1).

8.2.2.4

Construction of Plasmid PLcEcR for Expression of a Modified LcEcR in Insect Cells

Plasmid pVPLcEcR (Hannan and Hill, 2001) was subjected to restriction digestion with Kpn I + XbaI. The excised VPLcEcR fragment was trimmed/filled with Klenow and ligated into pIE1-3 (Novagen) which had been subjected to restriction enzyme digestion with PmeI and treated with calf intestinal phosphatase.

8.2.2.5

Transient Assays

Sf9 insect cells were maintained as described in Graham et al. (2007). Transient transfections were conducted using DOTAP (Boehringer-Mannheim) at 15 µg/ ml, essentially as described previously (Hannan and Hill, 1997). Replicate 96 well dishes of subconfluent Sf9 cells were cotransfected with (1) pSfEcRi or unmodified pUC18 at 0.1 µg/ml, (2) pLcEcR or unmodified pUC18 at 0.1 µg/ml and (3) pEcRE/Adh/βgal (a plasmid containing a reporter gene β-galactosidase under the control of an EcRE bearing promoter, Koelle et al., 1991) at 1 µg/ml. For induction experiments, the ecdysone analogue ponasterone A (a gift from Dr Denis Horn) was added to cells at 1 µM, 6 h after transfection. For control experiments, cells were treated only with carrier ethanol. β-galactosidase activity in extracts of cells was measured 72 h after transfection as described previously (Hannan and Hill, 1997).

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Fig. 8.1 Molecular engineering of an RNAi specific for the S. frugiperda ecdysteroid receptor (SfEcR) gene. A novel fragment encompassing the SfEcR D-domain was cloned by reverse transcriptionpolymerase chain reaction employing total Sf9 cell RNA as template. Our SfEcR cDNA fragment (double headed white arrow) spans the 3’ end of the C DNA domain, all of the D domain and includes the first 23 amino acids of the E domain. This fragment was then used as template for two new PCRs employing primer sets designed to amplify the same D-domain encoding sub-segment and introduce flanking restriction sites: PCR1 produced a fragment indicated by the black ended arrow that, following restriction enzyme digestion was flanked by SfiI and BamHI compatible ends. PCR2 produced fragment bearing the same D-domain sequence (indicated by the white ended arrow) that, following restriction enzyme digestion, was flanked by SfiI and NotI compatible ends. Insect cell expression plasmid pIE1-3 was subjected to restriction enzyme digestion with BglII + NotI and treated with calf intestinal phosphatase (CIP) and used in a three-way ligation with the D-domain encoding fragments bearing the common Sfi1 end. This resulted in plasmid pSfEcRi engineered to constitutively transcribe a 718-bp head-to-head inverted-repeat concatamer of the DNA fragment derived from the D-domain of SfEcR

8.2.3

Results and Discussion

Sf9 cells in our laboratory have been found to grow rapidly in serum-free medium either as monolayers or in suspension culture and to express easily detectable levels of endogenous ecdysone receptor. These cells are also readily transfected with DNA plasmids. They are thus, following transfection with an ecdysone-dependent reporter gene, useful for as a cell-based screen for compounds that bind to lepidopteran ecdysone receptors. Furthermore, Sf9 cells offer the potential for screening

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against ecdysone receptors from a variety of pest insects following transfection with plasmids engineered to express EcRs from those pests. However, for Sf9 cells to provide a useful vehicle for screens against exogenous receptors, it is highly desirable to minimise ecdysone receptor activity originating from the endogenous Sf9 receptor. To this purpose we have developed a RNAi system targeting the relatively less conserved D domain of SfEcR. Figure 8.1 depicts construction of a plasmid to effect RNAi-based selectively knock down of the expression of SfEcR in SF9 cells whilst permitting the expression of an exogenous receptor from the sheep blowfly LcEcR. A segment of SfEcR encompassing part of the C-domain, all of the D-domain and part of the E-domain was prepared by RT-PCR from a Sf9 total RNA template. The sequence of the product shows good agreement with that reported for other lepidopteran EcRs and was later confirmed by comparison with the partial fragment of SfEcR cloned by Chen et al. (2002). Sequencing in fact confirmed that our amplified fragment spans the 3’ end of the C DNA domain, all of the D domain and includes the first 23 amino acids of the E domain, including an extra two novel nucleotides additional and 3’ to the fragment cloned by Chen et al. (2002). Two separate PCR reactions, that shared a common primer introducing a SfiI restriction site (see Fig. 8.1) amplified a 362 bp sub-segment and a 363 bp sub-segment each encompassing the SfEcR D-domain and the first 15 bp of the E-domain. The two fragments containing the same SfEcR segment flanked respectively by BamHI, NotI and common SfiI sites were three-way ligated into an appropriately cleaved pIE1-3 vector. This resulted in a head-to-head concatamer of the two copies of the SfEcR D-domain segment downstream of a powerful constitutive IE1 insect baculovirus promoter leading to transcription of an inverted repeat RNA sequence that will fold-back to give a hairpin double-stranded structure suitable for triggering RNAi. The effects in tissue culture Sf9 cells are recorded in Fig. 8.2. The ecdysteroid responsive β-galactosidase reporter plasmid, pEcRE/Adh/βgal was transfected into the cells in all wells. The endogenous Sf9 receptor allows induction of a very significant synthesis of β-galactosidase on addition of 1 µm ponasterone A. Such a response would be sufficient to allow screening of compounds against the endogenous receptor in a cell-based assay. Transfection with plasmid pSfEcRi in addition to the reporter plasmid lowers the response to the hormone by some 80%. Transfection with the L. cuprina EcR expressing plasmid, pLcEcR, and the reporter in the absence of RNAi led to a very significant increase in hormone responsiveness over that due to the endogenous receptor alone. Introduction of pSfEcRi, pLcEcR and the reporter again gave a strong hormone response indicative of expression of the LcEcR notwithstanding inhibition of endogenous SfEcR. These data, strongly indicating functional expression of the exogenous receptor while synthesis of the endogenous receptor is largely inhibited, suggest a strategy for the use of the Sf9 cells in cell-based screens of chemical libraries against a variety of pest ecdysone receptors. For such purposes stably transfected cell lines would provide a considerable advantage and may be produced by employing the plasmid vector pIE1-3-neo and selecting for stable neomycin (geneticin) resistance in the presence of geneticin at 500 µg/ml.

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Fig. 8.2 Selective silencing of SfEcR in tissue culture Sf9 cells in 96-well micro-titre plates and functional introduction of the sheep blowfly LcEcR. Sf9 cells in all wells were transfected with the reporter plasmid pEcRE/Adh/βgal; β-galactosidase induction was monitored by colorimetric analysis of enzyme activity. Cells in particular wells were also transfected with the RNAi plasmid pSfEcRi and/or the blowfly LcEcR expressing plasmid pLcEcR as indicated. One micrometer ponasterone A was added to the wells in the back row and the ethanol solvent to the wells represented in the front row as a negative control

8.2.4

Conclusions

Introduction of a β-galactosidase reporter gene, under the control of an ecdysone receptor-responsive promoter, into Sf9 cells provides a system suitable for cellbased screening of chemical libraries for S. frugiperda ecdysone receptor ligands. Selective RNAi of expression of the SfEcR allows effective functional replacement of the endogenous protein with LcEcR from the sheep blowfly and offers the possibility of cell-based screening of chemicals for potential insecticides binding to the blowfly ecdysone receptor. Selective RNAi of the expression of SfEcR and replacement with EcRs from other insects should permit cell-based screening for potential insecticides against a variety of pests in Sf9 tissue culture cells.

8.3

8.3.1

Double-Stranded RNA Mediated Inhibition of Nuclear Receptor Expression in a Bombyx mori-Derived Cell Line Introduction

Lepidopteran insect cell lines traditionally have occupied a biotechnological niche as the hosts for the baculovirus expression system (Kost et al., 2005; Acharya et al., 2002). More recently, however, with the isolation of strong insect promoters and the

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identification of powerful viral enhancers and activators, plasmid-based expression systems were developed that allowed, in combination with techniques of insertion of expression cassettes into the cells’ chromosomes, the generation of permanently transformed cell lines that express continuously a protein of pharmaceutical or agricultural importance (Douris et al., 2006). Such techniques also have opened the possibility of engineering insect cell lines as screening systems for biologically active molecules. For instance, transformed insect cell lines that continuously express a receptor protein combined with a reporter cassette have the potential to be used as screening systems for compounds that activate or antagonize the receptor. Using elements of the plasmid-based expression system, transformed silkmothderived Bm5 cell lines were generated that respond to the challenge of the steroid hormone 20-hydroxyecdysone (20E) by the induction of green fluorescence (Swevers et al., 2004). The cell lines were used successfully to screen collections of plant extracts and libraries of chemical compounds. Importantly, after screening of a large collection of dibenzoyl hydrazine compounds, a group of chemicals with known ecdysone agonist activity that are employed as insecticides, a quantitative structure activity relationship (QSAR) model of the molting hormone activity was obtained that corresponds well with the model of the ligand-binding pocket of the ecdysone receptor derived from the crystal structure of its ligand-binding domain (Wheelock et al., 2006). Moreover, ecdysone reporter systems were also generated using cell lines derived from other lepidopteran species such as the insect pest Spodoptera littoralis as well as from dipteran species (based on the Schneider 2 cell line from Drosophila melanogaster) (T. Soin, G. Smagghe, L. Swevers and K. Iatrou, 2007). The availability of reporter lines specific for different insect species or insect groups will allow the screening in high-throughput format for compounds with ecdysone mimetic activity that target specific insects or insect groups. Transformed Bm5 cell lines with incorporated 20E reporter cassette also provide an efficient reporter system to investigate the contribution of specific genes for proper function of the 20E regulatory cascade. For instance, it has been established that developmental events triggered by 20E occur through the induction of a conserved regulatory cascade of cross-regulating transcription factors such as the nuclear receptors E75 and HR3, the ets-related factor E74 and the zinc finger protein Broad-Complex (Thummel, 2002; Riddiford et al., 2003). Some of these factors have been shown to interfere with the function of the ecdysone receptor and restrict temporally the action of the hormone (White et al., 1997). Proper function of the ecdysone receptor also pre-requires its stabilization by chaperone proteins in a configuration appropriate for formation of EcR/USP heterodimers capable of binding DNA target sequences (Arbeitman and Hogness, 2000). Finally, transformed cell lines can provide a screening tool to clarify the mechanism by which dibenzoylhydrazines mount a stronger and more long-lasting response than the natural hormone 20E (Retnakaran et al., 1995; Swevers and Iatrou, 1999). To establish a role as modulator of the ecdysone response requires techniques for efficient over-expression or silencing of candidate genes in the cell lines. Regarding silencing, it has become apparent that introduction of dsRNA into cells results in

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efficient epigenetic suppression of mRNAs harbouring sequences homologous to the introduced dsRNAs. This phenomenon has been observed in several insects such as Drosophila melanogaster (Lam and Thummel, 2000), the lepidopterans Hyalophora cecropia, Bombyx mori and Spodoptera litura (Bettencourt et al., 2002; Quan et al., 2002; Rajagopal et al., 2002), Apis mellifera (Amdam et al., 2003), the mosquito Anopheles gambiae (Blandin et al., 2002; Hoa et al., 2003) and the cockroaches Periplaneta americana and Blatella germanica (Marie et al., 1999; Cruz et al., 2006). Here we report on dsRNA-mediated gene silencing of the components of the ecdysone receptor heterodimer, EcR and USP, in Bm5 cells, a Bombyx mori-derived cell line.

8.3.2

Experimental

8.3.2.1

Generation of dsRNA Fragments

Bluescript (pBS-SK+) clones encompassing a 1.9 kb EcoRI fragment of BmEcR (Swevers et al., 1995) and a 1.4 kb BamHI-XbaI fragment of BmUSP (BmCF1; Tzertzinis et al., 1994; Farrell et al., 2000) were linearized with appropriate restriction enzymes (BamHI and SalI for BmEcR; BamHI and XbaI for BmUSP) and subsequently used as templates for transcription reactions using T7 and T3 RNA polymerase (Fermentas). After digestion of the DNA template using RNase-free DNase (RQ1 DNase; Promega), transcribed RNAs were purified by phenolchloroform extraction and ethanol precipitation. Complementary single-stranded RNAs were annealed after heating for 2 min at 90°C in annealing buffer (150 mM NaCl, 1 mM EDTA, pH 8) followed by gradually cooling down to room temperature (Skeiky and Iatrou, 1991). Formation of dsRNA was checked by size comparison with DNA MW standards during native agarose gel electrophoresis.

8.3.2.2

Cell Cultures and Transfections

Bm5 cells were maintained in IPL-41 medium with 10% fetal calf serum as described previously (Johnson et al., 1992). Cells were transfected in 6-well microtiter plates (106 cells/well in 2 ml of medium) using lipofectin (Invitrogen) as described previously (Johnson et al., 1992). Transfections were carried out with different amounts of dsRNA in combination with Bluescript DNA (to a total amount of 1 µg of nucleic acid) in the presence or absence of 1.5 µg of an EcR reporter plasmid (pBmbA/ERE.cat; Swevers et al., 2004), depending on the type of assay that was employed to assess knockdown of BmEcR or BmUSP (Western blot or Chloramphenicol acetyl transferase (CAT) reporter assay). In the case of reporter assays, 20E was added to a final concentration of 1 µM at 2 days post transfection and cells were collected and processed for enzyme assays 24 h after addition of hormone.

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Western Blot Assays

Cell pellets were solubilized in ‘cracking buffer’ (Swevers et al., 1995) and processed for SDS polyacrylamide gel electrophoresis (PAGE) and Western blotting as described before (Swevers et al., 1995). To detect BmEcR protein, a monoclonal antibody against the common region of Manduca sexta EcR was used (Jindra et al., 1996) while BmUSP was detected by the monoclonal AB11 antibody (Khoury-Christianson et al., 1992). Primary antibodies and anti-mouse HRP-coupled secondary antibodies were used at 1:1,000 dilutions. Cross-reacting proteins were detected by enhanced luminescence as described by the manufacturer (Amersham). 8.3.2.4

Chloramphenicol Acetyltransferase (CAT) Assays

CAT assays using soluble protein extracted from transfected cells were carried out as described before (Johnson et al., 1992). Transfection efficiencies among different samples were assessed by dot blot hybridizations using the cat gene ORF as a probe (Johnson et al., 1992).

8.3.3

Results and Discussion

To investigate whether the technique of RNAi can be applied to Bm5 cells, a B. mori-derived cell line, long dsRNAs corresponding to the partners of the ecdysone receptor heterodimer, EcR and USP, were prepared in vitro and used in transfection experiments. Two types of assays were used to evaluate the effects of introduction of dsRNAs into the cells: (1) the inducibility of EcR target genes by 20E was measured after co-transfection with an EcR-dependent reporter plasmid; and (2) the amount of target protein was assessed in Western blot assay. As shown in Fig. 8.3a, transfection of increasing doses of EcR dsRNA results in a corresponding decline in the activation of the EcR-dependent reporter by 20E. Furthermore, in independent experiments a dose-dependent decrease in the amount of BmEcR protein was observed in Western blot assays (Fig. 8.3b). The decline was specific to BmEcR protein because upon re-probing of the membrane no decrease of BmUSP was detected. Similar results were observed for BmUSP dsRNA although the down-regulation was less than for BmEcR dsRNA (Fig. 8.4). Transfection of 1 µg of BmUSP dsRNA resulted in a 50% decrease in reporter activity compared to >90% for 1 µg of BmEcR dsRNA. The amount of BmUSP detected in Western blot showed a dose-dependent decrease at increasing concentrations of dsRNA while the amount of BmEcR remained constant (Fig. 8.4b). Our experiments show the applicability of the technique of RNAi to interfere with the response to 20E in Bm5 cells. In the case of BmEcR dsRNA, significant down-regulation of EcR activity was observed at doses as low as 0.1 µg (Fig. 8.3a). Such sensitive response was not observed for BmUSP, indicating that the efficiency of RNAi may depend on the gene that is targeted. Specific down-regulation of gene expression in our experiments, employing relatively large fragments of dsRNA (1.9 and 1.4 kb), was absolutely dependent on the pres-

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Fig. 8.3 Dose-dependent decrease in expression of BmEcR at increasing concentrations of BmEcR dsRNA. Panel A: CAT reporter assay. CAT activities are compared with that observed for the reporter construct in the absence of cotransfected dsRNA (100%). Panel B: western blot assay. The membrane was probed for the presence of BmEcR (upper) and BmUSP (lower) proteins. MW markers are indicated at the right (See Color Plates)

Fig. 8.4 Dose-dependent decrease in expression of BmUSP at increasing concentrations of BmUSP dsRNA. Panel a: CAT reporter assay. CAT activities are compared with that observed for the reporter construct in the absence of cotransfected dsRNA (100%). Panel b: western blot assay. The membrane was probed for the presence of BmEcR (upper) and BmUSP (lower) proteins. MW markers are indicated at the right (See Color Plates)

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ence of lipofectin, a cationic lipid, that is used to facilitate the uptake of nucleic acids by the cells. This requirement for lipofectin holds if dsRNA fragments of lower MW (0.4–0.7 kb) were used (S. Dedos, L. Swevers and K. Iatrou, 2002). The situation in Bm5 cells therefore contrasts to the one in Drosophila Schneider 2 cells where efficient uptake of dsRNA was observed even in the absence of transfection agents although the addition of transfection agent lowered the effective dose (Saleh et al., 2006). Down-regulation of BmEcR and BmUSP expression by dsRNA is gene-specific. No effects of BmEcR dsRNA on the abundance of BmUSP protein was observed and vice versa (Figs. 8.3b and 8.4b). Similarly, introduction of 0.1–1 µg quantities of GFP or BmGATAβ dsRNA did not interfere with the activation of the primary 20E-responsive reporter by EcR/USP (S. Dedos, L. Swevers and K. Iatrou, 2002). Thus, the technique can be used to systematically address the function of gene products in the primary ecdysone response in Bm5 cells. Using Schneider 2 cells, several high-throughput screens have been reported to predict the function of Drosophila genes in cell viability and growth (Boutros et al., 2004) as well as in specific signalling pathways (Clemens et al., 2000; DasGupta et al., 2005). A prerequisite for such screens is the availability of robust assays to detect changes in functioning of cellular pathways. Permanently transformed cell lines that respond to addition of 20E by strong fluorescence (Bm5/ERE. gfp cell lines; Swevers et al., 2004) provide such a robust assay and therefore could be employed for functional analysis of the primary response to 20E. High-throughput screens would pre-require the construction of genome-wide libraries for generation of dsRNAs, a task that, at least in principle, became feasible after the publication of the genome sequence of B. mori (Mita et al., 2004; Xia et al., 2004).

8.3.4

Conclusion

Besides their development as protein factories, lepidopteran cell lines can be engineered as efficient screening systems for biologically active substances. The applicability of the technique of RNAi in B. mori-derived cells in combination with the availability of robust reporter assays, such as the one for the primary ecdysone response, opens the possibility to carry out screens for the function of Bombyx genes in signalling pathways such as the ecdysone response.

8.4

8.4.1

RNAi Analysis of Ecdysteroid Response in the Red Flour Beetle, Tribolium castaneum Introduction

Ecdysteroids (20-hydroxyecdysone, 20E is generally the native hormone) play key roles in development, reproduction, immunity and many other aspects of insect’s life. 20E binds to a heterodimeric complex of two nuclear receptors,

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ecdysone receptor (EcR) and ultraspiracle (USP) (Koelle et al., 1991; Robertson et al., 1993; Thomas et al., 1993; Yao et al., 1993). The EcR:USP:20E complex then binds to the response elements present in the promoter regions of 20E response genes and regulate their expression (Palli et al., 2005). Interestingly, many direct targets of the 20E-EcR-USP complex are members of the nuclear receptor (NR) superfamily (King-Jones et al., 2005). In Drosophila melanogaster six NRs, Drosophila hormone receptor 3 (DHR3), Drosophila hormone receptor 4 (DHR4), Drosophila hormone receptor 39 (DHR39), E75, E78 and FTZ transcription factor 1 (FTZ-F1) were shown to play key roles in 20E action (King-Jones and Thummel, 2005). Among these, E75 is directly induced by 20E and encodes three isoforms that contain common C-terminal regions but different N-terminal ends (Segraves and Hogness, 1990). In D. melanogaster E75-null mutants that do not express all three isoforms, oogenesis was affected suggesting a role for E75 in female reproduction (Buszczak et al., 1999). E75Aspecific null mutation affects ecdyteroid titers resulting in a block in larval development (Bialecki et al., 2002). Recent studies have shown the presence of heme in the ligand binding pocket of E75 and the oxidation state of this molecule controls E75 activity (Reinking et al., 2005). In addition, gases NO and CO can activate E75 after binding to heme (Reinking et al., 2005). The orphan nuclear receptors, DHR3 and DHR4 are delayed-early genes that require 20E induced protein synthesis for their maximal levels of expression (Koelle et al., 1992; King-Jones et al., 2005). These nuclear receptors act as repressors of 20E-induced early genes and also induce the expression of ftz-f1 gene, another orphan nuclear receptor and mid-prepupal competence factor (Yamada et al., 2000). FTZ-F1 is necessary for stage specific response of 20E and mutation analyses showed that FTZ-F1 is an essential gene (Yamada et al., 2000). DHR39 is an orphan nuclear receptor closely related to FTZ-F1 and appears to be not essential because the null mutants of DHR39 are viable and fertile (Horner and Thummel, 1997). The red flour beetle, Tribolium castaneum (Herbst) (Coleoptera: Tenebrionidae), has become a worldwide pest on stored grains and products. Chemical control using toxic insecticides is the management practice currently used to manage populations of this pest. Moreover, this pest has developed resistance to some of the commonly used pesticides such as synthetic pyrethroids. Therefore, there is urgent need for development of newer insecticides that function through target sites other than the nervous system. Since four stable ecdysteroid analogs are already registered for pest management and these chemicals appear to be fairly safe to humans, there is a potential for development of insecticides based on 20E action. Very little is known about the action of 20E in this important insect pest. To develop a knowledge base on 20E action in T. castaneum, we determined the ecdysteroid titers during the final instar larval and pupal stages and utilized recently completed whole genome sequences, RNAi, quantitative real time reverse transcriptase PCR and other methods to study the function of some of the key genes involved in ecdysteroid action in T. castaneum. Some of the methods used in these studies will be described first and then the results from these studies will be summarized.

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8.4.2

Experimental

8.4.2.1

Rearing and Staging

Tribolium castaneum strain GA-1 beetles obtained from Dr. Beeman of USDAARS were reared on organic wheat flour containing 10% yeast at 300C under constant light conditions. Staging of insects was performed following methods described in our recent publications (Parthasarathy et al., 2008; Parthasarathy and Palli, 2008; Tan and Palli, 2008a,b). The final instar larvae were identified as soon as they molted using untanned white cuticle as a marker and designated as 0 h AEFL (after ecdysis into the final instar larval stage). The larvae were staged from that time onwards. The beginning of the quiescent stage was designated as 0 h and was determined based on cessation of feeding and movement. The following days in the quiescent stage were recognized by characteristic ‘C’ shaped larvae. White pupae were designated as 0 h AEPS (after ecdysis into the pupal stage) and staged thereafter. Formation of compound eyes at 48 AEPS, sclerotization of mandibles and forewings at 72 h AEPS, and pharate adults at 96 h AEPS were used as morphological characters for staging pupae. 8.4.2.2

Double-Stranded RNA Synthesis and Injection

To prepare dsRNA, gene specific primers were designed based on the sequences available in the Beetlebase. The primers contained T7 polymerase promoter sequence at their 5’ ends. The primers generally resulting in a product length from 200–400 bp were designed from sequences spanning a single exon. These primers and genomic DNA isolated from T. castaneum beetles using DNeasy genomic DNA isolation kit (Qiagen) were used as PCR template to amplify gene specific regions. The resultant PCR products were used for synthesis of dsRNA using the Ambion MEGAscript RNAi kit (Ambion, Austin, TX). The dsRNA thus prepared was treated with DNase and RNase and purified by extracting with phenol and chloroform mixture and precipitated with ethanol. The final concentration of dsRNA was adjusted to about 5 mg/ml and stored at −20°C until use. About 0.1 µl of dsRNA was injected into the larvae on the dorsal side of the first or second abdominal segments using a aspirator tube assembly (Sigma) fitted with 3.5″ glass capillary tube (Drummond) pulled by a needle puller (Model P-2000, Sutter Instruments Co.). Injected larvae were reared under standard conditions until use. Control larvae were injected with dsRNA made using Escheria coli malE gene as a template. 8.4.2.3

cDNA Synthesis and Quantitative Real-Time Reverse-Transcriptase PCR (qRT-PCR)

Total RNA was extracted from staged larvae and pupae using TRI reagent (Molecular Research Center Inc., Cincinnati, OH). cDNA was synthesized using 2 µg of DNAse1 (Ambion, Austin, TX) treated RNA and iScript cDNA synthesis kit

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(Biorad Laboratories, Hercules, CA) in a 20 µl reaction volume as per the manufacturer’s instructions. Real-time quantitative reverse-transcriptase PCR was performed using MyiQ single colour real-time PCR detection system (Biorad Laboratories). PCR reaction components were: 1 µl of cDNA, 1 µl each of forward and reverse sequence specific primers, 7 µl of H2O and 10 µl of supermix (Biorad Laboratories). PCR conditions were: 95°C for 3 min followed by 45 cycles of 95°C for 10 s, 60°C for 20 s, 72°C for 30 s. Both the PCR efficiency and R2 (correlation coefficient) values were taken into account prior to estimating the relative quantities. Relative expression levels of each gene were quantified using ribosomal protein, rp49 expression levels as an internal control. Ecdysteroid titers: An enzyme immunoassay (EIA) was used to estimate ecdysteroid titers as previously described (Kingan and Adams, 2000; Gelman et al., 2002). The ecdysteroid antiserum used in the EIA has a high affinity for ecdysone (E), 20E, makisterone A, 20,26-dihydroxyecdysone, 26-hydroxyecdysone and 3-dehydroecdysone but does not detect polar conjugates (Gelman et al., 2005). Ecdysteroid titers were determined during the final instar larval and pupal stages. Small increases in ecdysteroid levels at 60, 78, and 90 h after ecdysis into the final instar larval stage were detected during the feeding stage of final instar larval stage. The ecdysteroid levels increased soon after larvae entered the quiescent stage and reached the maximum levels by the end of the quiescent stage prior to pupation. The ecdysteroid levels increased again beginning at 42 h after ecdysis into the pupal stage and reached the maximum levels by 66 h after ecdysis into the pupal stage.

8.4.3

Results and Discussion

Ecdysone receptor and ultraspiracle: Two isoforms each of EcR and USP have been identified in T. castaneum (Tan and Palli, 2008b). Both EcR and USP showed isoform- specific developmental expression patterns in the whole body, epidermis and midgut tissues dissected from the final instar larval and pupal stages. Injection of dsRNA prepared using the common or isoform-specific region as templates in the early stages of final instar larvae caused derailment of development, resulting in the death of the injected larvae prior to pupation (Tan and Palli, 2008b). dsRNA prepared against EcR common region or EcRA-specific region caused more severe effects, and most of the treated larvae died prior to pupation (Fig. 8.5a). In contrast, EcRB dsRNA caused less severe effects and most of the treated larvae had undergone larval-pupal metamorphosis and became pupae but the pupae showed developmental defects. Monitoring the expression levels of key genes involved in 20E action in the larvae injected with EcRA, EcRB or EcR common region dsRNA showed that EcRA regulates the expression of EcRB but not vice versa suggesting that EcRA may be involved in initiation of ecdysteroid response in T. castaneum (Tan and Palli, 2008b). In larvae injected with EcRA dsRNA, lower levels of EcRA, EcRB, br, FTZ-F1, E74 and E75A mRNA compared to their levels in malE (control) dsRNA injected insects were observed. In the larvae injected with EcRB dsRNA,

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Fig. 8.5 Phenotypes observed after knock-down of EcR, TcE75, TcHR4, TcHr39 or Tcbr. dsRNA was injected at 24 h after ecdysis into the final instar larval stage and the phenotypes observed are shown in comparison with control. The larvae injected with TcEcR (b), TcE75 (b) or TcHR4 (c) dsRNA died during quiescent stage. In general, the dsRNA injected larvae were smaller and darker than the control larvae injected with malE dsRNA. About 50% of the larvae injected with TcHR39 successfully pupated but had problems with wing development (d). The larvae injected with Tcbr had problems undergoing larval-pupal ecdysis, the exuvium remained attached to the body (e) and these insects showed antennae, wings, legs and compound eyes similar to those found in adults (f). Scale bar: 1 mm (See Color Plates)

lower levels of EcRB and E74 mRNA were observed. The mRNA levels of HR3 and FTZ-F1 increased by more than twofold in larvae injected with EcRB dsRNA. In larvae injected with EcR common region dsRNA, lower levels of EcRA, EcRB, br, FTZ-F1, E74, E75A and E75B mRNA were observed. Only HR3 mRNA levels increased by 2.5-fold in insects injected with EcR common region dsRNA (Tan and Palli, 2008b). Only dsRNA prepared against USP common region but not against USPA or USPB isoform-specific region caused developmental defects during larval-pupal metamorphosis (Tan and Palli, 2008b). These data suggest that the EcR but not USP isoforms play distinct roles during larval-pupal metamorphosis in T. castaneum.

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Midgut epithelium is completely replaced during larval-pupal metamorphosis. Larval cells are eliminated through programmed cell death and intestinal stem cells proliferate and differentiate to form pupal/adult midgut epithelium. Previous studies in Aedes aegypti, Heliothis virescens and Tribolium castaeneum showed that 20E in the absence of juvenile hormone initiates midgut remodeling events. Recent RNAi studies in T. castaneum showed that both EcR and USP are required for 20E action in midgut remodeling (Parthasarathy and Palli, 2008). These studies also showed that EcRA but not EcRB plays a critical role in 20E regulation of midgut remodeling. Nuclear receptors: Six nuclear receptors, Drosophila hormone receptor 3 (DHR3), Drosophila hormone receptor 4 (DHR4), Drosophila hormone receptor 39 (DHR39), E75, E78 and FTZ-F1 were shown to play key roles in 20E action (King-Jones and Thummel, 2005). We investigated the function of these six nuclear receptors by injecting dsRNA prepared using the regions of these nuclear receptors as templates. RNAi analysis in T. castaneum showed that four of these nuclear receptors (TcE75, TcHR3, TcFTZ-F1 and TcHR4) and TcHR51 are critical for larval-pupal metamorphosis (Tan and Palli, 2008a). Additional three NRs TcHR38, TcHR39 and TcSVP are important for both larval-pupal and pupal-adult metamorphosis (Tan and Palli, 2008a). The dsRNA injected insects showed various phenotypes depending on the type of nuclear receptors injected. Some larvae injected with TcE75, TcHR3, TcFTZ-F1 and TcHR4 DNA died at various stages during the final instar larval stage mostly after entering quiescent stage prior to metamorphosis to pupal stage. The majority of larvae injected with TcE75, TcHR3, TcHR4 or TcFTZ-F1 dsRNA died during the quiescent stage (Fig. 8.5b–c). A few larvae injected with these dsRNAs died during molting. Sixty percent of the larvae injected with TcHR51 dsRNA died during the quiescent stage prior to pupation. Twelve percent of the pupae also died during pupal stage. Twenty eight percent of the larvae injected with TcHR51 dsRNA became adults. The majority of larvae injected with TcSVP dsRNA pupated but the pupae did not develop further after ecdysis. Most of the pupae remained untanned or less tanned and the wing development did not progress normally; as a result these pupae showed only small wings covering only thoracic region and died at this stage. About 68% of larvae injected with TcHR39 became pupae but the majority of these pupae showed abnormal wing development and died at this stage (Fig. 8.5d). Broad: Broad (br) is a transcription factor containing the Broad-TramtrackBric-a-brac (BTB) and zinc finger domains (Zollman et al., 1994). Drosophila melanogaster br null mutant non-pupariating fail to undergo larval-pupal transformation (Kiss et al., 1988). In both Maduca sexta and D.melanogaster br is expressed specifically during larval-pupal metamorphosis under the control of 20E and JH (Zhou et al., 1998; Zhou and Riddiford, 2001, 2002). In the hemimetabolous insect, milkweed bug, Oncopeltus fasciatus, br was shown to be necessary for the heteromorphic changes that occur during development (Erezyilmaz et al., 2006). We determined the roles of br during larval-pupal metamorphosis of Tribolium castaneum. Two major peaks of Tcbr mRNA, one peak at the end of the feeding stage prior to the larvae entering the quiescent stage and another peak during the

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quiescent stage were detected in the whole body and midgut tissue (Parthasarathy et al., 2008). RNAi experiments showed that expression of br during the final instar larval stage is essential for successful larval-pupal metamorphosis (Parthasarathy et al., 2008). Injection of Tcbr dsRNA during the final instar larval stage derailed larval-pupal metamorphosis and produced insects that showed larval, pupal and adult structures (Fig. 8.5e–f). Tcbr dsRNA injected into the final instar larvae caused reduction in the mRNA levels of genes known to be involved in 20E action (EcRA, E74 and E75B). Tcbr dsRNA injected into the final instar larvae also caused an increase in the mRNA levels of JH-response genes (JHE and Kr-h1b). Knockdown of Tcbr expression also affected 20E-mediated remodeling of midgut during larval-pupal metamorphosis. These data suggest that the expression of Tcbr during the final instar larval stage promotes the pupal program while suppressing the larval and adult programs ensuring a transitory pupal stage in holometabolous insects. RNAi studies in T. castaneum showed that Tcbr is required for midgut remodeling during larval-pupal metamorphosis. In T. castaneum midgut remodeling begins at 96 h after ecdysis into the final instar larval stage (Parthasarathy and Palli, 2008). Three peaks of ecdysteroids (60, 78 and 90 h AEFL) and one peak of Tcbr mRNA in the midgut (84 h AEFL) precede initiation of midgut remodeling (Fig. 8.6). RNAi aided knock-down in Tcbr expression levels during the final instar larval stage blocked midgut remodeling during larval-pupal metamorphosis. JH suppression of 20E-induced stem cell proliferation was not affected by Tcbr knock-down (Parthasarathy et al., 2008). This study also showed that Tcbr is required for 20E induced proliferation of intestinal stem cells. In contrast, Tcbr is not required for JH suppression of 20E induced intestinal stem cells proliferation suggesting that br is required for 20E action but not for JH suppression of 20E action during midgut remodeling. Recently, Konopov and Jindra (Konopova and Jindra, 2008) showed that T. castaneum br specifies pupal fate and operates as a temporal coordinator of hormonally regulated morphogenetic events across epidermal tissues. In another recent study on Tcbr, Suzuki et al. (2008) determined the functions of five Tcbr isoforms and showed that the functions of Tcbr isoforms are conserved within the Holometabolous insects suggesting that evolution of br isoform expression may have played an important role in the evolution of the pupa in holometabolous insects.

8.4.4

Summary and Conclusions

These initial studies on T. castaneum measured ecdysteroid titers and determined the function of some key genes involved in 20E signal transduction. In T. castaneum, three small peaks of ecdysteroids during the feeding stage of final instar larvae, a continuous rise in their levels during quiescent stage resulting in a large prepupal peak at the end of quiescent stage and a large peak at the end of pupal stage were detected (Parthasarathy et al., 2007; Fig. 8.6). RNAi knock-down studies showed that EcR but not USP isoforms play distinct roles during larval-pupal

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Ecdysteroids

Broad 0 6 12 18 24 30 36 42 48 54 60 66 72 78 84 90 96 0 12 24 36 0 6 12 18 24 30 36 42 48 54 60 66 72 Final Instar Larval Stage (h) Pupal Stage (h) Quiescent Stage (h)

Fig. 8.6 Ecdysteroid titers and mRNA levels of broad during final instar larval and pupal stages of Tribolium castaneum drawn based on the data reported by Parthasarathy et al. (2007)

metamorphosis. EcRA appears to be more important than EcRB as knock-down in EcRA levels caused severe phenotypes compared to the phenotypes observed after EcRB knock-down. Orphan nuclear receptors E75, HR3, HR4, FTZ-F1 and HR39 as well as transcription factor, broad play key roles in 20E action during larval-pupal metamorphosis as knock-down in expression of these genes caused derailment of larval-pupal metamorphosis. These studies lay a solid foundation for future studies on 20E action in this important pest insect. Acknowledgements Support for the research employing Spodoptera frugiperda from the AusIndustry R&D Start Grant Scheme is gratefully acknowledged. The research employing Bombyx mori was funded by a PENED 2003 grant (03ED124) of the General Secretariat for Research and Technology, Ministry of Development in Greece. This work on Tribolium castaneum was supported by National Science Foundation (IBN-0421856), National Institute of Health (GM070559-03), and National Research Initiative of the USDA-CSREES (2007-04636). This is contribution number 08-08-112 from the Kentucky Agricultural Experimental Station

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Chapter 9

The Function and Evolution of the Halloween Genes: The Pathway to the Arthropod Molting Hormone Lawrence I. Gilbert and Kim F. Rewitz

Abstract The Halloween genes of Drosophila melanogaster were first described in the 1980s using cytogenetic methodology. During the past several years the genes have been cloned, expressed and the gene products have been characterized as cytochrome P450 enzymes (CYPs) and four have been functionalized as mediating the final steps in the biosynthesis of the arthropod molting hormone, 20-hydroxyecdysone (20E). A fifth has now been studied in detail and shown to be required for ecdysteroidogenesis but its exact function has yet to be elucidated. Since both insects and crustaceans utilize 20E as their principal molting hormone we have examined by BLAST search the genome of Daphnia and demonstrated the existence of Halloween gene orthologs in this crustacean, indicating that these genes play an identical role in this class as they do in insects. Further examination of the data bases representing Lepidoptera, Coleoptera, Hymenoptera and other Diptera allowed the development of a phylogenetic scheme for this gene family and suggests that the Halloween genes and vertebrate steroidogenic P450s originated from common ancestors that were perhaps destined for steroidogenesis, and arose before the deuterostome-arthropod split. Keywords 20-hydroxyecdysone • steroid hydroxylases • cytochrome P450 Abbreviations 7dC: 7-dehydrocholesterol; 20E: 20-hydroxyecdysone; CYP: Cytochrome P450; Dib: Disembodied (CYP302A1); E: Ecdysone; Phm: Phantom (CYP306A1); Sad: Shadow (CYP315A1); Shd: Shade (CYP314A1); Spo: Spook (CYP307A1); Spok: Spookier (CYP307A2); Spot: Spookiest (CYP307B1); SRS: Substrate recognition site. L.I. Gilbert () Department of Biology, University of North Carolina, Chapel Hill, NC 27599-3280, USA K.F. Rewitz Department of Science, Systems and Models, Roskilde University, PO Box 260, DK-4000 Roskilde, Denmark e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Introduction: A 50 Year Voyage in Insect Endocrinology

Herein we summarize our work over the past several years using the genetics of Drosophila melanogaster to help decipher the biosynthetic scheme of the principal arthropod molting hormone 20-hydroxyecdysone (20E). However, one of us (L.I.G.) takes this opportunity to summarize how he was introduced to the fruit fly with the hope that this brief excursion will be of some value to young investigators. To put this in perspective I shall relate how I was introduced to insects some half century ago, and to research in biology in general. In 1955 I was finishing my tour of duty in the U.S. Navy by teaching ROTC at the NY Maritime College and decided to finish my Masters degree in Biology at night at New York University. My instructor in Cell Physiology, a young assistant professor, Paul R. Gross, suggested that I do a research thesis and taught me the rudiments of thinking about, and solving, biological problems. Paul, who ultimately became an internationally renowned developmental biologist, was instrumental in my decision to pursue a doctorate after my service in the navy. A lesson I learned at that time was that new ideas at times can endanger a career but that stalwart thinkers carry on with what they believe in. Paul Gross received his degree under the mentorship of Professor L.V. Heilbrunn who authored many articles and a general physiology text book noting that calcium was much more important to the physiology of the cell than was believed at the time. He was disparaged for this point of view although we now know that thousands of studies have shown that he was correct. Indeed, I now tell my students that calcium is the “secret of life”. I then applied for admission to three universities for graduate student status. To my surprise Professor Dan Mazia (University of California, Berkeley) called within a month to offer me a 4-year fellowship. His work on the sea urchin mitotic spindle was very exciting, his offer was most gratifying and I was considering it quite seriously. A week later I received a phone call from a young assistant professor at Cornell University, Howard A. Schneiderman, who asked me to visit him, and since he was only a few hours drive from my home in New York City, my wife and I went to Ithaca, New York as a courtesy. I knocked on the door of Dr. Schneiderman’s office and he said to come in; I saw this young man, only 2 years my senior, sitting at a table examining a strange brown object under the binocular microscope, and he used his fine #5 forceps to extirpate a white mass from this brown object. I introduced myself and asked what he was doing. He said that he had just removed the brain from a pupa of the American silkmoth, now called Hyalophora cecropia. I was amazed that one could keep this debrained animal alive and asked what a pupa was. This was all new to me since I had never taken a course in entomology and grew up in New York City where trees were rare and pupae virtually absent. I was so impressed with the intelligence and enthusiasm of Howard Schneiderman that I accepted a teaching assistantship at Cornell, turning down fellowships at both Princeton and Berkely – the best professional decision I ever made. After 6 months at Cornell, Howard asked me to read a paper in Nature by Carroll Williams which showed that juvenile hormone could

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be extracted from the abdomens of the male Cecropia moth. We discussed it and he asked if I would like to work on juvenile hormone for my doctoral dissertation and I promptly agreed to do so. Such was my entry into insect endocrinology and a life long friendship with Howard Schneiderman who unfortunately died at the relatively young age of 63 while Vice-President for Research at Monsanto Corporation. He was a superb mentor in addition to being a first class scientist and friend. I only hope that I have emulated these characteristics with my own students. My first professional epiphany was meeting Paul Gross and the second was working with Howard Schneiderman. After receiving my doctorate I began my academic career in 1958 as Assistant Professor of Biology at Northwestern University where I remained for 22 years before moving to the University of North Carolina in 1980. I worked on the Cecropia silkmoth for many years but had to raise these monovoltine animals on netted trees and had the opportunity only a few months each year to study the larvae, or we could purchase pupae from professional raisers and collectors, although the latter were sometimes not very dependable. While Howard Schneiderman later switched to Drosophila as his research insect and urged me to do so as well, I stubbornly stayed with saturniid pupae since they were perfect surgical models. I had become a skilled and accomplished micro-surgeon. After all, one could not pursue real endocrinology with fruit flies which seemed so small. It was only Dietrich Bodenstein who could remove the Drosophila larval ring gland and keep the larva alive. In the late 1960s I was invited by Robert Yamamoto to present a lecture on my work at North Carolina State University and he showed me the animal he was working on in his study of sensory perception, the tobacco hornworm, Manduca sexta. These large sphingid caterpillars weighed up to nine grams, had a short life cycle, about 1 month compared to H. cecropia which had one generation per year, and above all, they could be raised on an artificial diet in the laboratory. I immediately adopted this insect as my research object, brought back eggs and diet to my laboratory at Northwestern University, thanks to Bob Yamamoto, and began a colony which remains an object of my affection to this day. Our first paper using Manduca was published in 1971 (Sroka and Gilbert, 1971). It is now the most utilized “non-genetic” insect for studies of insect physiology, biochemistry and molecular biology throughout the world, due also to the early timing studies of Lynn Riddiford and Jim Truman and their continued excellent contributions over many decades. Although we have continued to use Manduca for almost 40 years, we did a number of studies on Diptera. These turned out to be quite interesting although they too were conducted on “non-genetic” organisms. For example, we showed effects of juvenile hormone (JH) extracts on the development of the flesh fly Sarcophaga bullata (Srivastava and Gilbert, 1968, 1969) and used the exquisite polytene chromosomes of the midge, Chirnomus tentans, in vitro to study the effects of 20E and JH at the chromosomal level (Lezzi and Gilbert, 1969, 1970). There were additional studies on the cell biology of dipterans, e.g. Robert et al. (1976); Roberts and Gilbert (1986) but it wasn’t until the 1980s that I decided that we must use genetics to study aspects of insect

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endocrinology as Howard Schneiderman had suggested some 15 years previously. This was elicited by my intuitive feeling that since a number of laboratories, including mine, had used classical biochemical techniques to no avail in an attempt to identify and characterize receptors for 20E and JH, that using a genetic approach may prove to be successful. I was beginning to understand then as Andreas Keller (2007) so aptly noted recently “In a century of research, many basic biological principles were discovered in Drosophila melanogaster, and it is now probably better understood than any other organism” (bold type by L.I.G). Further, the myriad studies on mammalian steroid hormone receptors had reached the molecular level and I wondered what might be different about insect hormone receptors that would be of interest to endocrinologists in general, rather than being of parochial interest only to insect biologists. So, how does a biologist in his mid-50s who is chairing and building a Department of Biology after moving south from one great university to another, learn genetics. As noted above, I did have some experience with Diptera and about that time received applications from two young Drosophila geneticists for postdoctoral positions in my laboratory, a time when I had just received a new grant and several members of my laboratory were leaving for permanent positions. I immediately accepted Vince Henrich and Tim Sliter into my research group, both of them wishing to learn insect endocrinology and utilize Drosophila to answer generally important endocrinological questions. In return, I would be their 50+ year old apprentice in my attempt to learn some genetics. This was my third professional epiphany which occurred shortly before the time that the Hogness laboratory identified the ecdysteroid receptor (EcR) and we had begun our search for it as well. Although we did not identify EcR using a mammalian consensus steroid hormone probe and a Drosophila library, we were one of three groups that almost simultaneously cloned the heterodimeric partner of EcR, ultraspiracle (Henrich et al., 1990) and in those early years contributed several other significant papers in the area of Drosophila endocrinology (Henrich et al., 1987a, b; Richard et al., 1989; Sliter et al., 1989; Dai and Gilbert, 1991; Sliter and Gilbert, 1992; Saunders et al., 1989, etc.). The lesson here is ‘that no matter how old you are, or how wedded to your experimental insect you are, you can still learn new paradigms and techniques that will help you solve what may seem to be unsolvable problems’. My laboratory’s increased interest in the use of Drosophila to solve endocrinological problems also brought us to the attention of the genetics community and we have been asked to collaborate with some of the very best Drosophila geneticists in the world. Most of these collaborations had excellent results. Among these splendid scientists were Steve Cohen, Huby Amrein, Matt Scott, Mike Stern, John Sisson, Linda Restifo, Kim McCall, Carl Thummel, Shalom Applebaum, Hermann Stellar and Mike O’Connor, the latter being a true partner in the research to be described herein. This collaboration began about 7 years ago when Mike telephoned me to find out if my laboratory would collaborate with him on a project he was working on, the Halloween gene family, that he believed was involved in ecdysone biosynthesis. I of course accepted immediately because I believed strongly that our

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biochemical and natural products chemistry expertise, as well as our acquaintance with Drosophila genetics, coupled with his molecular and genetics expertise would be the perfect scientific marriage. As you read this review, I hope that you conclude that I was correct.

9.2

The Halloween Genes and the Biosynthesis of the Arthropod Molting Hormone

For an as yet unexplained evolutionary reason, arthropods do not have the ability to synthesize cholesterol from simple precursors, but instead must rely on ingesting cholesterol found in other animals or converting phytosterols such as β-sitosterol to cholesterol via dealkylation (see Gilbert et al., 2002). Although cholesterol is a known critical constituent of cell membranes, it is also the ultimate precursor for the synthesis of ecdysone, the immediate precursor of the molting hormone, 20E (Gilbert et al., 2002). Obviously, this biochemical strategy has been quite successful since arthropods now inhabit virtually every ecological niche on earth and continue to be man’s primary competitor for the nutrients of this planet. It is now more than half a century since Peter Karlson and his colleagues elucidated the structure of ecdysone, but despite the persistent attempts of several insect biochemistry laboratories, including ours, very little was known regarding the essence of the sequence of reactions from cholesterol to 20E until very recently. The one step that was studied in some detail at the biochemical level was the hydroxylation of ecdysone at the C20 position to yield 20E, a reaction mediated by a cytochrome P450 (CYP) enzyme (Smith et al., 1979; Feyereisen and Durst, 1978). This CYP was termed the ecdysone 20-monooxygenase, or hydroxylase, both being used interchangeably, although hydroxylase will be used herein. These studies were important but were conducted on cell homogenates or subcellular fractions and therefore did not utilize pure enzyme preparations. Was there a way in which one could obtain pure enzyme for studying this reaction and perhaps for the unknown reactions comprising the biosynthetic cascade from cholesterol to 20E? As it turns out, despite numerous studies at the biochemical level and a great deal of laborious activity by a handful of laboratories over several decades, none succeeded. In large measure this was due to the extremely small quantities and lability of these enzymes in the prothoracic glands, site of the reactions leading from cholesterol to ecdysone. As will be discussed subsequently, the solution for this quandary was to use molecular genetic techniques and a “genetic” insect, Drosophila melanogaster. The first step in the biosynthetic sequence is the dehydrogenation of cholesterol (C) to 7-dehydrocholesterol (7dC; Fig. 9.1) and the importance of this sterol intermediate has been researched for several decades. In vitro radiochemical and analytical studies using lepidopteran tissue have shown that this is basically a rapid and irreversible reaction mediated by an enzyme with the characteristics of a CYP found in microsomal fractions of the prothoracic glands (see Gilbert et al., 2002).

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Fig. 9.1 Scheme of 20-hydroxyecdysone (20E) biosynthesis in Drosophila. Multiple arrows indicate an uncharacterized pathway, the Black Box. Yellow shade delineates the areas of the sterol that are involved in the transformation to the next compound (See Color Plates)

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This reaction is presumably present in all insects and likely in all arthropods, but not in a very specialized fruit fly D. pachea which must ingest 7dC and therefore lives only in an environment hosting a unique desert cactus containing 7dC. This insect is a virtual endocrinological twin of a D. melanogaster low ecdysteroid mutant (woc; without children) which also lacks the C7,8-dehydrogenase and can be rescued to the adult stage when 7dC is placed in the diet (Gilbert et al., 2002). Although the goal of our research on woc was to show that the gene product was the 7,8-dehydrogenase, it was unfortunately not, but was rather a zinc fingered transcription factor whose role has yet to be elucidated (Wismar et al., 2000; Warren et al., 2001). It may be involved in the regulation of the dehydrogenase itself or in the intracellular trafficking of 7dC from the endoplasmic reticulum to the outer membrane of the mitochondria, analogous perhaps to the trafficking of sterol intermediates between subcellular compartments in mammals. This transit is aided by proteins like StAR that have been suggested to be involved in the ratelimiting movement of C within the mitochondria of the adrenal cortex, source of the steroidogenic cortical hormones. What is the fate of 7dC in the reactions leading to 20E? It is assumed that 7dC enters what has euphemistically been labeled ‘The Black Box’ because so little is know of its contents, but we will return to that once we have considered the postblack box reactions, i.e. those mediated by the characterized Halloween genes. What are these Halloween genes? In the 1980s, some exquisite genetic studies were published by the laboratories of Nüsslein-Volhard and Wieschaus on this family of genes, several of which when mutated led to the death of the embryos prior to the time that they deposited an embryonic cuticle. Using the giant polytene chromosomes of the salivary glands of Drosophila they were able to map and identify these genes (data that were essential to our subsequent cloning of these genes). This was followed by the critical paper of Chavez et al. (2000) who concentrated on one of these genes, disembodied (dib), and showed that when mutated the result was embryonic lethality accompanied by a low ecdysteroid titer and that the gene appeared to code for a new CYP enzyme, CYP302A1. They postulated that this gene product, Dib, may be an enzyme that mediated a step in ecdysteroidogenesis. Shortly thereafter, the O’Connor and Gilbert laboratories joined forces to investigate this family of genes with the goal of elucidating the entire ecdysteroidogenic pathway using a combination of molecular genetics, functional genomics and biochemistry. Later we were joined by the laboratory of Dauphin-Villemant. How was our goal achieved?

9.2.1

Methodology

In addition to what we already knew from the studies cited above, we were able to access the Drosophila genome through the published fly database. For an example we will use the Halloween gene shade (shd) (which will be discussed in detail later) and whose position was mapped to position 70D2-E8, which according to

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the database was close to a gene coding for Cyp314a1, located at position 70E4 on the polytene chromosome (Petryk et al., 2003). Using an embryonic cDNA library we then amplified a full length cDNA coding for CYP314A1 and showed that this gene was altered in the various shd mutant alleles utilizing PCR amplification of the corresponding genomic DNA derived from the heterozygous mutant stock. The mutant lesion was identified by a specific base change at a particular locus after sequencing. The composite data showed unequivocally that CYP314A1 was a gene product of shd. This was the protocol used for all the Halloween genes after the first was cloned and characterized “the hard way”. Having cloned the gene, the next and perhaps equally important step was to investigate the exact role for each of the gene products, i.e. the functional genomics and biochemistry. The cDNA or the coding region of each gene was ligated into an expression vector and then transfected into the Drosophila S2 cell line. Similar promoter constructs that constitutively express GFP were also transfected as controls and taken through the same purification process as the cells receiving the Halloween genes. Three days later the S2 cells were incubated with known tritiated or non-labeled substrates and the resulting products analyzed first by RP-HPLC and TLC (normal phase) utilizing UV, RIA and/or radiotracer analysis. When necessary this was followed by LC-coupled electrospray ionization (ESI) mass spectrometry. For one of the genes (phantom), high field NMR was also utilized. Gene expression was monitored by in situ hybridization (Fig. 9.2), RT-PCR and Northern analysis. Additionally, both gene- and tissue-specific rescues were conducted using wild type sequences linked to either ubiquitous or tissue-specific promoters. For our studies on the tobacco hornworm, Manduca sexta, the Halloween genes were cloned and characterized. Since there is no genomic data base for this insect an alternative method was utilized. Total RNA was extracted from fifth instar larval prothoracic glands to obtain spo, phm, dib and sad, and midgut to obtain shd. cDNA was obtained by reverse transcription of total RNA. Degenerate primers, based on Drosophila and Bombyx mori Halloween gene sequences as templates, were used to amplify the Manduca genes. Gene specific primers were utilized to amplify the 5′ and 3′-ends by RACE-PCR. The gene products were expressed in S2 cells as discussed previously. Gene expression was analyzed by real time PCR (qPCR) as described previously (Rewitz et al., 2006a, b).

9.2.2

Phantom (phm)

As can be seen in Fig. 9.1, the first compound post-Black Box is the diketol and there is a great deal of indirect evidence that this Δ4 four compound is indeed a precursor (Fig. 9.1). Indeed, Blais et al. (1996) have described a reductase in the crustacean Y-organ (site of ecdysone production in the crab) that mediates the conversion of this diketol to the 5β(H)-diketol and ultimately to ecdysone. Although not yet identified in insects, we assume that it does exist and is the precursor of

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Fig. 9.2 In situ expression of the Halloween genes disembodied, shadow and phantom during late embryonic and larval stages. Shown are stage 17 embryos (a, e and i), late second instar (b, f and j), and both early (c, g and k) and late (d, h and l) third instar brain-ring gland complexes. Note the down regulation of the expression of all three genes between the late second and early third instars and their subsequent up regulation between the early and late third instars. RG, ring gland; Br, brain; VG, ventral ganglion (Data on embryonic dib expression from Chavez et al., 2000. Data on larval dib and sad expression from Warren et al., 2002. Data on phm expression from Warren et al., 2004) (See Color Plates)

the critical intermediate, the ketodiol, after reduction at C3. The ketodiol is successively hydroxylated at four positions to ultimately yield 20E (Fig. 9.1). The first hydroxylation takes place at the C25 position and past cytological investigations showed that phm was located between positions 17C5 and D2 on the X chromosome and that Cyp306a1 at position 17D1 in the database was a reasonable candidate for the 25-hydroxylase. Genomic DNA from a mutant allele was sequenced and a base alteration was identified (Warren et al., 2004) and the usual in situ hybridizations were conducted showing the presence of this gene in the ring gland (source of ecdysone in Drosophila, Fig. 9.2), as well as RIA analysis to show that this was a low ecdysteroid mutant. At the same time, our collaborators in Japan identified an ortholog in the prothoracic glands of the commercial silkworm, Bombyx mori, with 37% identity to the Dmphm using completely different technologies (Warren et al., 2004). After S2 cell transfection of both the Drosophila

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gene (Dmphm) and the Bombyx gene (Bmphm), we showed unequivocal and significant conversion of the ketodiol (2,22,25-trideoxyecdysone) to the ketotriol (2,22-dideoxyecdysone) i.e. hydroxylation at the C25 position. The gene product of phm was therefore the C25-hydroxylase in both the dipteran and lepidopteran (see also Niwa et al., 2004). As noted previously, for studies on Manduca we used Drosophila and Bombyx Halloween sequences as templates to design degenerate primers against conserved regions of phm and the other Halloween genes to obtain partial sequences for putative Manduca orthologs. With this paradigm we were ultimately able to obtain deduced amino acid sequences for Phm and the other Halloween gene products (Rewitz et al., 2006a). This allowed us to conduct qPCR analysis of phm expression in the prothoracic glands of Manduca and study developmental changes correlated with the ecdysteroid titer. This was important because each of the 220 cells in the prothoracic gland is about the same size as one ring gland, and Manduca is a better model for developmental timing than is Drosophila. During the fifth (last) larval instar phm expression is detected in the prothoracic glands and undergoes dramatic up- and down-regulation in concert with the ecdysteroid titer, increasing with the first ecdysteroid peak eliciting cellular reprogramming and later with the large ecdysteroid peak that elicits the metamorphic molting process to the pupal stage. Since the ecdysteroid titer increases about 13-fold from the day before until the day of the commitment peak, and the protein content of the gland only increases twofold (Rybczynski and Gilbert, 1994), this suggests that it is specifically the level of enzyme in ecdysteroid biosynthesis that is up-regulated rather than a general increase in gland activity. The data also suggest that phm is transcriptionally regulated, indicating that this change in gene expression is essential for the exact timing and amplitude of the ecdysteroid peak so necessary for normal growth, development and metamorphosis. The next step in the multiple hydroxylations to yield the molting hormone would be the hydroxylation at the C22 position to yield 2-deoxyecdysone (Figs. 9.1 and 9.3).

9.2.3

Disembodied (dib)

Dib was actually the first Halloween gene cloned and characterized but is discussed here because of its place in the natural sequence of reactions leading to ecdysone. Chavez et al. (2000) first discovered the importance of dib as noted previously and we continued that work (Warren et al., 2002). In situ analysis showed very clearly that dib is expressed in the prothoracic gland cells of the embryonic and larval ring glands (Fig. 9.2). [See Dai and Gilbert (1991) for a detailed study of the morphology of the ring gland.] Since the 20E titer varies significantly during post-embryonic development (Warren et al., 2006) and these changes are requisite for growth, development and metamorphosis, we examined these changes in dib expression during development by semi-quantified in situ hybridization and found that it was down regulated quite dramatically following ecdysis to the final larval stage when

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Fig. 9.3 Scheme of 20-hydroxyecdysone (20E) biosynthesis in Drosophila with gene names, gene products (enzymes) and CYP designations (See Color Plates)

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the 20E titer decreases (Fig. 9.2). This finding indicates that dib and its product would be excellent models for the study of the down regulation of an important gene. In any event, there was no doubt that since the mutant embryos contained low ecdysteroid levels, dib was important for ecdysteroidogenesis. The gene sequence revealed that it was Cyp302a1 and that Dib was a mitochondrial CYP. The gene was then transfected into S2 cells as already described and incubated in the presence of labeled ketotriol (2,22-dideoxyecdsyone). The transfected cells converted the substrate to labeled 2-deoxyecdysone with an efficiency of 82% during the 4–6 h incubation, whereas the GFP control cells showed no conversion. These data thus demonstrate that Dib is the C22-hydroxylase, and chronologically was the first characterized gene demonstrated to have a role in ecdysone biosynthesis. When qPCR of dib was examined in the Manduca prothoracic glands we found that the developmental changes were essentially the same as those noted for phm i.e. expression is well correlated with the hemolymph ecdysteroid titer (Rewitz et al., 2006a). The next challenge was to determine how 2-deoxyecdysone is converted to ecdysone.

9.2.4

Shadow (Sad)

As in the case of dib the sad mutants had low ecdysteroid levels and the in situ analyses were very similar, showing dramatic changes during late larval development (Fig. 9.2), while the sequence indicated that sad coded for another mitochondrial CYP (CYP315A1). Again, the gene was transfected into S2 cells which were subsequently incubated in the presence of the labeled ketotriol and the medium was then analyzed. The results were quite clear i.e. the ketotriol was converted to 22-deoxyecdysone, strongly indicating that sad encoded the C2-hydroxylase (Fig. 9.3). To confirm this, the S2 cells transfected with sad were incubated with 2-deoxyecdysone derived from the dib experiments and the resulting product was unequivocally identified as ecdysone. Indeed, when S2 cells were transfected with both sad and dib in the presence of the ketotriol, hydroxylation occurred at both the C2 and C22 positions yielding ecdysone. Studies with Dib and Sad showed that during the biosynthesis of ecdysone, hydroxylation at C2 (Sad) follows hydroxylation at C22 (Dib). Again, the expression of this gene in the prothoracic glands of Manduca during the last larval instar was consistent with its role in ecdysone biosynthesis as was shown for phm and dib. Although we had demonstrated the critical roles of three genes and their respective gene products in the biosynthesis of ecdysone, and although there have been scattered reports in the literature for several decades that ecdysone does have a role in insect growth and development, it is 20E that is generally accepted as the principal moiety that elicits and controls arthropod molting. We discussed previously that biochemical studies showed the presence of a 20-hydroxylase in insect tissues and the data indicated that it too was a CYP. However, there had been no

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analyses using pure enzyme and there were numerous discussions about where this CYP was located in the cell i.e. mitochondrial or ER. We turned our attention to the 20-hydroxylase using the same experimental paradigm as with the previous three Halloween genes.

9.2.5

Shade (Shd)

As per the paradigm noted previously shd was positioned at 70D2-E8 while position 70E4 exhibited Cyp314a1. Heterozygotic mutant DNA was sequenced following PCR and three alleles were analyzed. The data revealed that CYP314A1 was encoded by shd and that the mutant phenotype was typical of the mutants of the Halloween genes discussed previously i.e. low ecdysteroid content, no embryonic cuticle deposition, lethality, and very low expression of the 20E-responsive gene IMP-E1. On the other hand, in situ hybridization analysis showed that shd was expressed later in embryogenesis than the other genes and was not expressed in the ring gland, but was expressed in several other tissues (Petryk et al., 2003). These findings are completely consistent with the expected properties of a gene encoding the 20-hydroxylase, the enzyme responsible for mediating the conversion of ecdysone to 20E, a most critical enzyme found in tissues other than the prothoracic glands and whose activity is responsible for the rise in 20E titer leading to cellular reprogramming and molting in all arthropods. To prove unequivocally that Shd is the 20-hydroxylase, the gene was transfected into S2 cells and examined as with the previously described Halloween genes. We also demonstrated transgenic rescue of shd mutants using the Gal4/UAS expression system with two different promoters and demonstrated the absence of 20E in the mutant embryos, in contrast to significant quantities in the wild type controls. When the S2 cells were incubated in the presence of labeled and unlabelled ecdysone, there was significant conversion to 20E. The ESI mass spectrum was unequivocal in showing the molecular ion at 481 and the sequential loss of four molecules of water, all characteristic of 20E. Therefore, we conclude that shd encodes the 20-hydroxylase and this marked the completion of the characterization of the genes responsible for the last four steps (hydroxylations) leading to the synthesis of the arthropod molting hormone (Fig. 9.3). In a single experiment that sums up the success of our paradigm using Drosophila, we were able to transfect the S2 cells with all four genes (phm, dib, sad, shd) in the presence of ketodiol so that each provided the substrate for the next reaction, with the end product being significant quantities of 20E. Further, the Bmphm could successfully substitute for its Drosophila counterpart (Warren et al., 2004). The data also suggested that the sequential order of terminal hydroxylations is C25, C22, C2 and C20 (Fig. 9.1). The data on the chronology of Halloween gene product presence in the embryo indicates that Dib and Sad are present prior to the appearance of Shd, as would be expected since the former two CYPs are required for ecdysone biosynthesis and ecdysone is the substrate for Shd.

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The studies of Manduca shd (Msshd) were in a sense quite beautiful in that all the data on the gene product fit so very nicely with published data on the enzymatic and developmental studies of this very important enzyme. First, the Msshd was transfected into the Drosophila S2 cell line as was done for Dmshd, and the

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resulting Shd easily mediated the conversion of ecdysone to 20E, showing that in this heterologous system the Shd from both species were functionally equivalent (Rewitz et al., 2006b). The N-terminal sequence of the Msshd, like that of the fly gene, contains regions suggesting both ER and mitochondrial locations, although the studies on the fly showed that the DmShd locates to the mitochondria (see next section). The Msshd is expressed primarily in the fat body, Malphigian tubules and midgut, consistent with similar studies on Dmshd and numerous investigations in the past using biochemical assays for the 20-hydroxylase (see Rewitz et al., 2006b). When Shd was studied in the above noted tissues during the last larval instar and during early pupal-adult development there was a most remarkable correlation between gene expression and the 20-hydroxylase activity quantified biochemically by the Smith laboratory several years previously (Mitchell et al., 1999; Smith et al., 1983) (Fig. 9.4). Comparative studies with the several tissues suggest strongly that the fat body, because of its large mass, is the major contributor of Shd activity during the first (commitment) peak and that the midgut may contribute significantly to total 20-hydroxylase activity, but also utilize this enzyme within the organ to elicit the well documented changes that occur there during the metamorphic molts. During the final larval instar, the Malpighian tubules express high levels of Msshd during the peak of molting hormone, suggesting that this tissue contributes to the conversion of ecdysone to 20E that elicits molting to the pupa. Although in the case of both fat body, midgut and Malpighian tubules the ecdysteroid titer increases sharply during the first days of pupal-adult development and neither shd expression nor 20-hydroxylase activity increases in concert with this surge, the results can easily be explained by the finding two decades ago that in Manduca the hemolymph ecdysteroid titer during the first week of pupal-adult development is almost entirely ecdysone and therefore no 20-hydroxylase activity is required (Warren and Gilbert, 1986). Figure 9.4 also demonstrates that Msshd expression is down regulated very rapidly during this last larval instar, followed by rapid decreases in the ecdysteroid titer. This suggests that decreases in 20E may also elicit developmental changes as has been shown for several systems in Manduca by the Truman laboratory or that ecdysone itself may play a developmental role at times (e.g. Warren and Gilbert, 1986). Finally, the tight coupling between enzyme activity and transcription (gene expression) along with the rapid changes in hormone titer during development, indicates very precise tissue-specific transcriptional regulation. This aspect of the work is worthy of further investigation.

9.3

Subcellular Distribution of the Halloween Genes

On the basis of confocal microscopic study of the C-terminal HA-tagged proteins and analysis of the charged segment at the N-terminus of the sequence we were able to show that Dib and Sad were mitochondrial CYPs (Warren et al., 2002) and that Phm was microsomal (resides in the endoplasmic reticulum; Warren et al., 2004). The Shd protein on the other hand exhibited a location in the mitochondria but its

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predicted N-terminus contained not only the hydrophobic signal sequence typical of microsomal CYPs, but also an interior charged segment suggesting a mitochondrial targeting sequence (Petryk et al., 2003). This quandary is reminiscent of the past biochemical work on the 20-hydroxylase where the literature is replete with discussion over the location of this critical enzyme. Perhaps the Shd enzyme can reside in either subcellular organelle depending on post-translational changes (Petryk et al., 2003) as has been described for several mammalian CYPs. On the other hand, it is possible that 20-hydroxylase activity has evolved along different paths in various insect groups and that their primary sequences differ. All explanations are conjectural at this time. The most important question that arises from these studies showing that during 20E synthesis the product of one reaction, which is the substrate for the next reaction, must move within the cell from one organelle to another, is that there is virtually nothing known about this trafficking of 20E precursors either within the mitochondrion (inner to outer membrane) or from the ER to and from the mitochondrion. Further study of StAR or StAR-like proteins which function in trafficking during mammalian steroidogenesis is necessary (see Lafont et al., 2005). In our quest for the essence of the entire biosynthetic scheme from cholesterol to 20E (Fig. 9.3) we have been notably successful in taking advantage of the FlyBase in dissecting the four terminal steps in the generation of the molting hormone. We have also discussed our studies on Bombyx, and the expression of these Halloween genes in the white mouse of insect endocrinology, Manduca sexta. There is a vital piece missing from this puzzle, the black box reaction(s), which we have judiciously not referred to in the last dozen pages. That is because we do not have the ultimate answer although we have made some progress, and that brings us to spook.

9.3.1

The Black Box: Spook (Spo)

The Black Box refers to those reactions between 7dC and the diketol and ketodiol (Figs. 9.1 and 9.3). Our most recent studies suggest that one of these reactions is mediated by a CYP encoded by spook (spo) and its paralog spookier (spok), depending on species and developmental stage (Ono et al., 2006; see also Namiki et al., 2005). The Dmspo mutant embryos, like those of the other Halloween genes, exhibit low ecdysteroid titers, the same type of morphological abnormalities and fail to express a 20E-inducible gene (Chavez et al., 2000). Again, using the cytogenetic information from the 1980s, we found that a CYP gene, Cyp307a1 is located on the third chromosome at position 65D4 and deficiency mapping revealed that Dmspo was in that area. To determine if spo is indeed located at that position, two different mutant alleles were sequenced showing that both were probably null alleles since the stop codon eliminates the heme-binding domain necessary for CYP function [all of this work and that which follows on spo comes from Ono et al. (2006)]. The N-terminal sequence with its stretch of hydrophobic residues showed this CYP to be microsomal i.e. located in the ER.

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The in situ analysis of spo showed its presence in the early embryo, consistent with our study of phm, but in contrast to the other Halloween genes transcription is activated to a high level in the yolk nuclei and thence in the amnioserosa and is absent from the ring glands. Further, there is no expression of Dmspo in any tissue during the third (last) larval instar, including the prothoracic gland cells of the ring gland, and it makes its appearance once again in the follicle cells of the ovary of adult females during oogenesis! The absence of the gene from the larval ring gland was unexpected for a gene that we believed encoded a CYP essential for ecdysteroidogenesis. Even when the expression of Dmspo was analyzed using the Gal4/UAS system and transformant lines were analyzed, there was still no evidence of this gene in the ring gland at several developmental stages. Our feelings then, and now, are summed up by a quote from Winston Churchill when asked his opinion of the USSR. He answered, “a riddle wrapped in a mystery inside an enigma”. To investigate if Spo was only needed during embryogenesis, homozygous mutant embryos halfway through embryogenesis were pulsed with 20E and a significant number hatched into first instar larvae and a significant percentage of these completed development to the adult stage, although females were sterile. Ecdysone gave a similar result as did the ketotriol, although less effectively than ecdysone or 20E. On the other hand, the ketodiol and 7dC were ineffective (see Fig. 9.3). It therefore appeared that Spo functioned upstream of the ketotriol: in the Black Box? Indeed, in contrast to the other Halloween gene products it appeared that Spo is only required for embryonic and ovarian development while Shd, and probably the other Halloween genes, are required for larval development and metamorphosis to the pupa and then to the adult. As of this time the function of Spo remains conjectural although we, and likely other laboratories, continue to explore its exact role in the Black Box. It should also be mentioned that during this study we explored the possible presence of spo in the lepidopterans, Bombyx and Manduca, and demonstrated that both species have the spo gene (Ono et al., 2006). The Msspo encodes a deduced protein sequence with high homology to the BmSpo (79% identity) and both lepidopteran gene products are 49% identical and 68% similar to the DmSpo. This will be discussed in more depth in the Molecular Evolution section of this chapter. Of great interest however, is that spo of both lepidopterans is expressed in the prothoracic glands of fifth (last) instar larvae in stark contrast to the situation with Drosophila. Indeed, qPCR studies revealed that spo expression is virtually coincident with the major ecdysteroid peak in the hemolymph of these last stage larvae, suggesting strongly that Spo plays a cogent role in post-embryonic development in these Lepidoptera. Dmspo mutants die during embryogenesis as do all the Halloween gene mutants, but when they receive Bmspo they are rescued (Namiki et al., 2005), indicating that Spo from both the fly and the moth mediate the same reaction during the biosynthesis of ecdysone. Perhaps another enzyme takes over the role of Spo in the fly. By once again examining the FlyBase we found that a newly annotated gene was recently added to the data base and was located in the heterochromatin, a portion of the chromosome that was thought to be devoid of genes until recently (see Smith et al., 2007). This gene was 58% identical to Dmspo and was dubbed spookier

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(spok). This CYP, in contrast to spo, is expressed in the larval prothoracic gland cells of the ring gland where its expression cycles similarly to the other Halloween genes. Further, when spok is knocked down in first instar larvae, they do not molt, but fed 20E, ecdysone or 2-deoxyecysone, they proceed through the three larval instars. The ketodiol is also effective suggesting that Spok is required for molting and may very well be a constituent of the Black Box. The spook/spookier Cyp307a subfamily will be discussed in more detail in the section on Molecular Evolution. The possibility exists that Spo and Spok function upstream of cholesterol, perhaps in the dealkylation of the phytosterol (e.g. β-sitosterol) to cholesterol via a set of hydroxylations and epoxidations mediated by as yet uncharacterized CYPs. However, feeding experiments appear to negate this possibility, but not unequivocally (see Gilbert, 2004; Ono et al., 2006). At this time, we consider the site of Spo and Spok action to be the Black Box.

9.4

9.4.1

Other Low Ecdysteroid Mutants That May Influence Ecdysteroid Biosynthesis b FTZ-F1

Parvy et al. (2005) analyzed the possible role of βFTZ-F1 (βF) in conjunction with the Drosophila low ecdysteroid mutant woc which was discussed previously. βF is analogous to steroidogenic factor 1 in vertebrates where it controls the development of steroidogenic organs by the transcriptional regulation of crucial steroidogenic enzymes. These experiments were designed to determine if either gene might control the transcriptional regulation of phm or dib after it was demonstrated that βF is localized in the ring gland. Using clonal analysis it was shown that both Phm and Dib were present at very reduced levels in ftz transcription factor 1 (ftz-f1) mutant ring gland cells but that the woc mutation had no effect on the levels of these genes. The data indicate that βF regulates both phm and dib expression. The hypothesis offered is that βF is induced in the ring gland when the ecdysteroid titer falls, either by raising its expression rate or causing its transit from the cytoplasm to the nucleus. This is the only transcription factor that has been directly linked to ecdysteroidogenesis although several are indirectly implicated i.e. transcription factors expressed by woc, dre4 and gt.

9.4.2

Neverland (Nvd)

This gene encodes an oxygenase-like protein and is expressed in the prothoracic gland cells of the Drosophila ring gland and prothoracic glands of Bombyx larvae (Yoshiyama et al., 2006). Growth and molting are arrested in nvd mutants and the

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gene is highly conserved from prokaryotes to birds and fish (interestingly missing from plants, some of which synthesize ecdysteroids, and mammals). This gene is also found in the heterochromatin as is spok and rescue studies suggest that Nvd may play a role in the transport and/or metabolism of cholesterol or one of its derivatives. Since 7dC placed in the food rescues the nvd mutants and cholesterol does not (as in the woc mutants discussed previously), it is possible that this Nvd may play a role in the conversion of cholesterol to 7dC.

9.4.3

Molting Defective (Mld)

This gene encodes a zinc finger transcription factor that is required for ecdysone biosynthesis (Neubueser et al., 2005) as in the case of woc mutants. The mld mutants are arrested in their development in the first larval instar. These larvae have little or no ecdysteroid and their ring glands are enlarged (again as in the case of the woc mutants), perhaps due to cellular hypertrophy in response to the low ecdysteroid titer. The developmental arrest of the mld mutant larvae is partially reversed when 20E is added to the food. Since the mutation does not seem to affect the level of Halloween gene expression it was suggested that the zinc fingered transcription factor acted at another level. In the course of our study of spo and spok we also attempted to investigate their relationship to mld (Ono et al., 2006). Spok is not expressed in mld mutant larvae suggesting that Mld is a regulator of spok. We attempted, but failed to rescue the mld mutant phenotype by having the spo and spok genes expressed via the Gal4/ UAS in the prothoracic gland cells of the ring gland. This experiment suggested that Mld regulates some other aspect of ecdysone biosynthesis, perhaps via unknown residents of the Black Box.

9.5

Molecular Evolution of the Biosynthetic Enzymes

It is now clear that CYPs are essential for the biosynthesis of steroids in some nematodes, arthropods and vertebrates (see Payne and Hales, 2004; Lafont et al., 2005; Gilbert and Warren, 2005; Beckstead and Thummel, 2006). In the phylogenetic lineage leading to arthropods, a biosynthetic pathway yielding 20E evolved with the occurrence of the Halloween genes. These genes have been conserved through evolution because of the fundamental role of 20E in the development, growth and reproduction of arthropods (Rewitz et al., 2007). Although it has been suggested that ecdysteroids also play a role in nematode development, there is only limited evidence to support this (Lafont and Mathieu, 2007) and the genome of C. elegans does not contain orthologs of the insect Halloween genes (Rewitz et al., 2007). Exactly when in evolution the vertebrate sex steroids originated is still a matter of debate. Evidence supports the presence of vertebrate sex steroids in invertebrate Phyla such

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as Mollusca and Echinodermata, yet its physiological importance in invertebrates is equivocal (Köhler et al., 2007; Lafont and Mathieu, 2007). It is possible that the enzymes mediating the synthesis of sex steroids in vertebrates were present in metazoans before the radiation of arthropods. That would suggest that arthropods evolved ecdysteroids when the Halloween genes evolved but lost the ancestral steroidogenic CYP genes, and thereby, the ability to synthesize vertebrate-type sex steroids. Considering the evolution of steroid hormones, it is important to note that CYP genes were present in early metazoans. In fact, the most conserved and ancient CYP gene, CYP51, is present even in bacteria, fungi and plants (Lepesheva and Waterman, 2007). CYP51 is involved in sterol biosynthesis by catalyzing the removal of the 14α-methyl group, thus implying that sterol biosynthesis is one of the most ancient functions of CYPs (Nelson and Strobel, 1987). CYP enzymes that evolved from ancestors with such biochemical properties may easily have evolved the ability to modify cholesterol into forms that constituted the first steroid hormones. Thus, the molecular evolution of CYP genes, which are well known to undergo gene duplications and divergent evolution (see Feyereisen, 2005), has played an important role in the evolution of steroids. CYPs are heme-containing enzymes consisting of approximately 500 amino acids with well-conserved consensus motifs including WxxxR (helix-C), GxE/ DTT (helix-I), ExxR (helix-K), PxxFxPE/DRF (“PERF”) and PFxxGxRxCxG (heme-binding domain) (see Feyereisen, 2005). These motifs are important for the structure of the CYP and are conserved among CYPs. In contrast, six substrate recognition sites (SRSs) vary with substrate specificity (Gotoh, 1992), hence being important when considering the functional conservation of CYP genes. CYPs are divided into mitochondrial (Class I) and microsomal (Class II) enzymes. Several charged N-terminal amino acids target CYP proteins to the inner mitochondrial membrane whereas microsomal CYPs with mainly hydrophobic N-terminal amino acids reside in the outer membrane of the endoplasmic reticulum. Most insects have more than 80 CYPs, although the honey bee Apis mellifera has only 46 (Claudianos et al., 2006). Among these are a relatively small number of highly conserved orthologs and a more variable subset of paralogs depending on lineage-specific expansions. Approximately a dozen of these CYP genes are truly conserved as orthologs and the Halloween genes account for about half of these. Each insect genome, so far analyzed, contains single orthologs of phm, dib, sad and shd and a variable number of spo-like sequences comprising enzymes in the CYP307 family (Table 9.1). Structural conservation of orthologous CYPs suggests that they are critical to the physiology of insects, which has been demonstrated for those encoded by the Halloween genes that catalyze steps in the formation of 20E, as described previously. Because 20E controls molting and reproduction in arthropods, selective forces have imposed constraints to conserve the function of these biosynthetic enzymes (Rewitz et al., 2007). In addition to the few conserved orthologs, insects possess a variable number of CYP4 and CYP6/9 related paralogs evolving rapidly through gene duplication and diversification. These genes are believed to act as “environmental response” genes (Feyereisen, 2005) i.e. the evolution of these genes was driven by needs to adapt to changes in the environment e.g. detoxification of plant allelochemicals. Transposable elements surround some

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Table 9.1 Predicted occurrence of Halloween and spo-like genes in insect and Daphnia genomes Spo

Spok

Spot

CYP307 family

Phm

Dib

Shd

Sad

307A1

307a2

307a3

307B1

306A1

302A1

314A1

315A1

†

⊕

⊕

†

⊕

⊕

⊕

⊕

⊕

⊕

†

†

⊕

⊕

⊕

⊕

⊕ ⊕

⊕ ⊕

⊕ ⊕

⊕ ⊕

⊕ ⊕

⊕ ⊕

⊕

†

⊕

⊕

⊕

⊕

⊕

⊕

⊕

⊕

⊕

⊕

Hymenoptera Apis mellifera

†

⊕

⊕

⊕

⊕

⊕

Crustacea Daphnia pulex

⊕

⊕

⊕

⊕

⊕

Diptera Drosophilidae Subgenus Drosophila Subgenus Sophophoran Aedes Aegypti Anopheles gambiae Lepidoptera Bombyx mori Coleoptera Tribolium castaneum

⊕: Indicate extant genes. †: Indicate genes that likely have been lost during evolution. Note that non-Drosophila species have only one CYP307A ortholog, which is the closest ortholog of spok in Drosophila species (see phylogenetic analysis below).

of these CYP genes which produce a plastic genomic environment that probably allows the rapid evolution of these CYP genes (Chen and Li, 2007). In contrast, transposable elements are not found in the vicinity of the Halloween genes, reflecting stability in the genomic environment containing these genes. Evolutionary relationships between CYPs are often difficult to infer because of the multiple duplications and diversification alluded to above. However, the insect Halloween genes have highly conserved structures allowing orthologous relationships to be established easily. Orthologous Halloween sequences on average share 38.7–55.2% amino acid identity (Table 9.2). Orthologs exhibit significantly lower similarity to other CYPs in the genomes of these species e.g. the closest paralog of Phm is CYP18A1 with 29.2% amino acid identity (Rewitz et al., 2007). Although some of the orthologs are not 40% identical, they are given the same names in different species and are not cutoff by the conventional ≥40% rule of CYP family relationship (Nelson et al., 1996). Complete genome sequences are available for several insects and with the release of the Daphnia pulex genome sequence, as the first crustacean genome to be completely sequenced, it is possible to infer phylogenetic relationships between insects and crustaceans. Crustaceans synthesize 20E through a biochemically pathway similar to that of insects. This suggests strongly that orthologs of the insect Halloween genes exist in the Daphnia genome. Here we infer phylogenetic relationships of the Halloween genes in several orders of insects and between

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Table 9.2 Mean percentage amino acid identity (± standard deviation) of CYP enzymes in the Halloween familya Spo/spok Spot 307A1/2 307B1 Phm 306A1 Dib 302A1 Sad 315A1 Shd 314A1 Spo/spok 54.0 ± 8.6 36.5 ± 2.0 20.7 ± 1.4 16.7 ± 1.2 15.5 ± 1.2 16.5 ± 1.2 Spot – 55.2 ± 6.8 21.6 ± 1.3 16.8 ± 1.7 15.7 ± 1.4 16.5 ± 1.1 Phm – – 45.5 ± 8.0 18.4 ± 1.3 17.5 ± 1.3 16.9 ± 1.3 Dib – – – 48.9 ± 9.4 21.8 ± 1.8 25.9 ± 1.9 Sad – – – – 38.7 ± 8.8 20.6 ± 1.6 Shd – – – – – 43.9 ± 11.9 a Bold numbers indicate othologous genes. Sequences from Drosophila melanogaster, Drosophila pseudoobscura, Aedes aegypti, Anopheles gambiae, Bombyx mori, Manduca sexta, Tribolium castaneum, Apis mellifera and Daphnia pulex were used for comparison.

insects and Daphnia. In addition, we will review the existing literature and new data obtained recently from genome database analyses pertaining to the molecular evolution of these important steroidogenic enzymes.

9.5.1

The Steroid Hydroxylases

The final four steps in the biosynthesis of 20E are hydroxylations mediated by Phm, Dib, Sad and Shd discussed previously. These genes have been identified in a number of other insect species (Sieglaff et al., 2005; Rewitz et al., 2007; Sztal et al., 2007), although they have only been characterized functionally in Drosophila and lepidopterans (Warren et al., 2002, 2004; Niwa et al., 2004, 2005; Rewitz et al., 2006a, b). Figure 9.5 reveals that Phm, Dib, Sad and Shd orthologs are present in different insects. The obvious 1:1 orthology indicates that these genes have been conserved by selection during evolution. Each of these genes is believed to function through development as the sole gene, responsible for a particular step in the biosynthesis of 20E (see Ono et al., 2006; Rewitz et al., 2007) because: (1) arthropod genomes that are available contain one ortholog of each Halloween gene and no very close paralogs; (2) expression occurs in ecdysteroidogenic tissues during embryogenesis, larval development and in the adult ovaries; (3) 20E is not produced in homozygous embryonic lethal mutants; (4) in shd mutants ecdysone is not converted to 20E in tissues such as the fat body, midgut and Malpighian tubules which normally support this reaction and (5) ectopic expression of shd using a tissue-specific promoter that does not drive expression in these tissues rescues transgenic animals but does not restore ecdysone 20-hydroxylase activity in fat body, midgut and Malpighian tubules. Intron positions are highly conserved among orthologs and most differences can be attributed to intron loss with only a few introns being gained during the course of evolution (Fig. 9.5). Intron loss has occurred prevalently in those species with small genomes such as Drosophila species and the red flour beetle Tribolium castaneum.

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Fig. 9.5 Phylogenetic relationships of Phm (CYP306A1), Dib (CYP302A1), Sad (CYP315A1) and SHD (CYP314A1) orthologs from the water flea Daphnia pulex (Crustacea), honey bee Apis mellifera (Hymenoptera), red flour beetle Tribolium castaneum (Coleoptera), tobacco hornworm Manduca sexta and silkworm Bombyx mori (Lepidoptera), mosquitos Aedes aegypti and Anopheles gambiae, and Drosophila melanogaster and Drosophila pseudoobscura (Diptera). (Left) Neighborjoining phylogram with support on each node indicated as percentages of bootstrap values based on 1,000 replicates. (Right) Intron positions on the aligned protein sequences with bold horizontal lines indicating aligned primary sequences. Conserved introns are indicated by vertical lines. ● For phase 0 introns (splicing between codons), ▲ for phase 1 introns (splicing between the first and second nucleotide of a codon), and ■ for phase 2 introns (splicing between the second and the third nucleotide of a codon). Intron positions for the Manduca genes are not shown because they are unknown (Contains data from Rewitz et al. 2007)

In these species there has been selection for genome compaction (Rogozin et al., 2003). In the dib of mosquitoes (Aedes aegypti and Anopheles gambiae), two introns are unique, a phase 2 intron at amino acid position ~120 and a phase 1 intron at position ~575 in the alignment. These introns might have arisen in the line leading to mosquitoes. Several introns are conserved between dib, sad and shd

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indicating that these mitochondrial CYP gene products share common evolutionary origins and probably are rather ancient paralogs. This is consistent with the mitochondrial CYPs being a monophyletic group (Feyereisen, 2005). There are a few exceptions where gene structure does not exhibit conservation. In Tribolium, sad has no introns and shd has only two introns (compared to 5–10 introns in other species), none of which have counterparts in other shd orthologs. Although intron loss and gain may explain this, it seems more likely that at least Tribolium sad is a retrogene integrated into the genome from a processed mRNA. Since Tribolium only has one ortholog of sad it may be that this apparent retrogene was duplicated from the “source” gene (the ancestral gene) that was then lost during evolution. After gene duplication one of the copies is free to accumulate mutations while the original function is retained by the other paralog (see Zhang, 2003). The most likely result is loss of function (nonfunctionalization) of one of the copies as deleterious mutations accumulate. If the retrogene copy of Tribolium sad acquired a functional promoter it is possible that this gene retained activity and that the original gene was lost. Inheritable gene duplications by retrotransposition can only occur when genes are expressed in germ line cells (Zhang, 2003). Expression of the Halloween genes in adult ovaries (except D. melanogaster spok), and perhaps testes, agrees with the retropositional origin of Tribolium sad. Using Tribolium and Apis sequences as queries we searched the Daphnia genome database for orthologs of the insect Halloween genes using the translated Blast nucleotide (TBLASTN). Candidate orthologs of Phm, Dib, Sad and Shd were retrieved and analyzed for phylogenetic relationships and conservation of gene structure in order to establish phylogeny. The Daphnia genes cluster in a phylogenetic tree with their respective insect orthologs with high bootstrap support (Fig. 9.5). Conserved intron-exon structures between Daphnia and insect orthologs clearly support the phylogeny. Although several introns are conserved between dib, sad and shd, unique introns, like the first phase 1 intron in dib and shd, indicate orthologous relationships in each group. This suggests that Daphnia, like insects, has single orthologs of phm, dib, sad and shd which is consistent with crustacean 20E biosynthesis (see Lachaise et al., 1993). Microsynteny is the conserved order of genes on chromosomes in different species, and can be used to establish evolutionary relationships. In D. melanogaster, Cyp18a1 and phm are arranged tail-to-tail within 10 kb on the X chromosome (Fig. 9.6). As discussed previously, CYP18A1 is a paralog of phm (see also Claudianos et al., 2006) suggesting that CYP18A1 and phm originate from a gene duplication, a mechanism that involves unequal crossing over of chromatids in contrast to retrogenes that arise through an RNA intermediate (Zhang, 2003). In Apis, the microsyntenic linkage of phm and CYP18A1 is conserved i.e. CYP18A1 and phm are on opposite strands and adjacent to the CYP18A1-Phm cluster is CG6696 and fused. Daphnia phm is also located close to a CYP18A1 ortholog. However, CG6696 and fused are not adjacent to the CYP18A1-Phm cluster in Daphnia indicating that chromosomal rearrangements have occurred. This is not surprising considering the evolutionary distance between crustaceans and insects. Together with the sequence phylogeny, the evolutionary relationship indicated by

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Fig. 9.6 Conservation of microsynteny for phm in Drosophila melanogaster (Diptera), Apis mellifera (Hymenoptera) and Daphnia pulex (Crustacea). In these species, CYP18A1 and phm are paralogs closely linked on the same chromosome (see also Claudianos et al., 2006). Arrows indicate transcriptional orientation of genes on the genome DNA sequence (horizontal lines). The size of the chromosomal regions is shown in parenthesis but distances are not to scale (See Color Plates)

the presence of an adjacent CYP18A1 ortholog provides strong evidence that this is the Daphnia ortholog of phm (see also Rewitz et al., 2007). Although the Daphnia Halloween genes have not been functionally characterized, the orthologous relationship indicated by the phylogenetic analysis, conservation of intron-exon structure and microsynteny support the functional conservation of these steroidogenic CYP hydroxylases in arthropods ranging from crustaceans to insects.

9.5.2

Duplications in the CYP307 Family

Since it was discovered that spo is required for the production of 20E in Drosophila (Chavez et al., 2000), several enzymes have been added to the CYP307 family comprising spo-like genes (Ono et al., 2006; Rewitz et al., 2007; Sztal et al., 2007). The evolutionary history of the spo-like genes involves lineage-specific gene duplications and losses within this family. CYP307 family genes are referred to as follows: nonDrosophila species spo refers to the single intron-containing CYP307A gene and spot (spookiest) the CYP307B1 gene containing two introns; in Drosophilids, spo refers to the intronless Cyp307a1 gene (identified for its role in 20E production; Chavez et al., 2000) in D. melanogaster and other species belonging to the Sophophoran subgenus (see Fig. 9.7), spok the intron-containing Cyp307a2 gene, and Cyp307a3 the intronless gene in species belonging to the Drosophila subgenus. Although gene duplications and losses have occurred, insects and Daphnia contain at least one spo-like gene, consistent with it being required for 20E production (Namiki et al., 2005; Ono et al., 2006). The phylogeny inferred in Fig. 9.7 shows the division of two well-supported clusters, the CYP307A (spo, spok, Cyp307a3) and CYP307B (spot). Except for Apis, non-Drosophilid insects and Daphnia have one spo ortholog. Apis (Hymenoptera), Tribolium (Coleoptera) and mosquitoes (Diptera)

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Fig. 9.7 Phylogenetic relationships of spo (the single intron-containing CYP307A gene in nonDrosophilids and the intronless Cyp307a1 gene in Drosophila species of the Sophophoran subgenus), spok (Cyp307a2), Cyp307a3 and spot (CYP307B1) orthologs from the water flea Daphnia pulex (Crustacea), honey bee Apis mellifera (Hymenoptera), red flour beetle Tribolium castaneum (Coleoptera), tobacco hornworm Manduca sexta and silkworm Bombyx mori (Lepidoptera), mosquitoes Aedes aegypti and Anopheles gambiae (Diptera), and Sophophoran subgenus species Drosophila melanogaster, Drosophila pseudoobscura, Drosophila erecta, Drosophila willistoni, Drosophila subgenus species Drosophila virilis, Drosophila grimshawi and Drosophila mojavensis (Diptera). (Left) Neighbor-joining phylogram with support on each node indicated as percentages of bootstrap values based on 1,000 replicates. (Right) Intron positions on the aligned protein sequences. Symbols otherwise as in Fig. 9.5. Intron positions for the Manduca genes are not shown because they are unknown (Contains data from Rewitz et al. 2007)

have one spot gene indicating that the paralogous groups of spo/spok/Cyp307a3 and spot arise from an early gene duplication predating the split of hymenopterans (Hymenoptera is the earliest branching order of insects analyzed here according to Savard et al., 2006). Interestingly, the only Daphnia spo-like gene is in the CYP307A

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family, yet it has the two conserved introns found in insect spot-CYP307B genes and not in the insect CYP307A genes (spo, spok, Cyp307a3). Since insects are believed to have evolved from freshwater crustaceans (Glenner et al., 2006), the presence of spo but not spot in Daphnia indicates that the primordial gene was a CYP307A gene. It is therefore likely that spot occurred by a duplication of spo after the origin of insects from crustaceans. Subsequently, the insect spo ortholog may have lost one intron whereas this intron has been retained in spot orthologs. In Drosophila species, spo was the first CYP307 gene identified as being involved in 20E synthesis (Chavez et al., 2000). As discussed previously, D. melanogaster has two paralogs in the Cyp307a family, spo and spok, that likely mediate the same function at different stages of development (Ono et al., 2006). spo is intronless and spok contains one conserved phase 1 intron equivalent to the one found in the single orthologs of non-Drosophilids (Fig. 9.7). Because spo lacks introns in its coding region it is likely a retrogene copy of spok, the latter being located in the heterochromatin of D. melanogaster. Nvd is another gene that has been related to ecdysteroid biosynthesis (discussed previously in this contribution) and the product of nvd may be involved in conversion of cholesterol to 7dC, the first step in the biosynthesis of 20E (Yoshiyama et al., 2006). Spo and Spok are believed to be involved in the subsequent Black Box reaction(s) where 7dC is converted into the ketodiol (Ono et al., 2006; see previous discussion). Interestingly, these genes, that are believed to be involved in early steps in the biosynthesis of 20E, are adjacent to one another on the chromosomes in Apis and Daphnia (Fig. 9.8). In some Drosophila species the apparent linkage is conserved i.e. nvd is adjacent to spok (i.e. the intron-containing gene) which further supports the presumption that spok is the ancestral gene of Drosophilids. Recently Sztal et al. (2007) found that the intronless copy in some species in the Drosophila subgenus (i.e. D. grimshawi, D. mojavensis and D. virilis) was phylogenetically different from Dmspo and formed a separate clade (Fig. 9.7). Sequence phylogeny and different patterns of microsynteny suggest that in species such as D. virilis, belonging to the Drosophila subgenus, spo is replaced by another intronless gene in the Cyp307a subfamily, namely Cyp307a3 (Sztal et al., 2007). Cyp307a3 is more closely related to spok than spo is in several species of the Sophophoran subgenus (i.e. D. melanogaster, D. pseudoobscura, D. erecta and D. willistoni). The microsyntenic pattern of genes flanking Dmspo is conserved in the Sophophoran subgenus (Fig. 9.8). However, a different microsyntenic pattern emerges around Cyp307a3 of species in the Drosophila subgenus. This shows that spo (Cyp307a1) and Cyp307a3 are located in different genomic contexts. The most likely explanation for this is that spo and Cyp307a3 are different genes originating from two independent retrotranspositions (Sztal et al., 2007). In contrast to gene duplication by unequal crossing over, retrogenes are duplicated without regulatory elements to distant locations which prevents parallel evolution e.g. gene conversion (Gayral et al., 2007). This supports the proposition that spo and Cyp307a3 are distinct genes and that the sequence phylogeny can not be explained by gene conversion events (see Sztal et al., 2007). It is believed that the first duplication generated spo and spok which were maintained in the Sophophoran subgenus. The second more recent duplication that generated Cyp307a3 occurred in the line of

Fig. 9.8 Conservation of microsynteny for spo-like (CYP307A) genes in Drosophila species of the Sophophoran subgenus and Drosophila subgenus (see Fig. 9.7), Anopheles gambiae and Daphnia pulex. The intron-containing gene (spo in Daphnia and Anopheles and spok in Drosophilidae) is adjacent to neverland (nvd), a gene believed to be involved in the conversion of cholesterol to 7-dehydrocholesterol (7dC) which is the first step in the formation of 20E (Yoshiyama et al., 2006). Symbols otherwise as in Fig. 9.6 (Data on Drosophilids modified with permissionfrom Sztal et al. 2007) (See Color Plates)

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evolution leading to the Drosophila subgenus. After the second duplication, spo was lost in the Drosophila subgenus, leaving members of this group with only spok and Cyp307a3 (Sztal et al., 2007).

9.5.2.1

Evolutionary Scenario for spo-Like Genes

The phylogeny indicates that gene duplication and subsequent losses have occurred in different insect lineages. Figure 9.9 shows a possible evolutionary scenario for genes in the CYP307 family based on phylogenetic analysis and the presence of spo-like genes in different arthropod groups. Compared to the terminal hydroxylases, this serves as a more complicated history involving independent duplications and losses of genes. As discussed above, the spo ortholog found in Daphnia suggests that the ancestral gene was a CYP307A subfamily gene. It is likely that the ancient spo gene, containing two introns, duplicated somewhere during insect evolution to form spo and spot. Alternatively, spot may have arisen in crustaceans but was lost in Daphnia. Once genome sequences of earlier branching insects and more crustaceans become available one may be able to resolve this question. Spot is absent in Lepidoptera (Bombyx) and higher dipterans such as Drosophila; however, dipterans such as mosquitoes possess this gene. This suggests that spot has been lost independently during the evolution of lepidopterans and in dipterans in the line leading to Drosophila (Rewitz et al., 2007), which diverged from mosquitoes some 250 million years ago. Intriguingly, no spo orthologs have been found in Apis indicating that this gene, which is present in all the other arthropods analyzed so far, was lost. However, we have identified one putative spo ortholog in the wasp, Nasonia vitripennis, data base showing that spo was not lost in all hymenopterans (data not shown). A word of warning! It is of course possible that some of the apparently lost genes will be identified during continued efforts to complete present genome sequences. In the lineage leading to Drosophila species, a second duplication probably formed the intronless spo from spok, the ancestral gene. Recently, in the evolution within the Drosophila genus, Cyp307a3 originated from a third independent retroduplication in the Drosophila subgenus after the Sophophoran/Drosophila subgenus split and subsequently spo was lost in the Drosophila subgenus (Sztal et al., 2007).

9.5.3

Gene Duplications: Why?

9.5.3.1

Evolution of the Biosynthetic Enzymes by Gene Duplications

In terms of evolution, gene duplication is believed to be one of the most important processes in the development of complexity (Taylor and Raes, 2004) and this is certainly true for CYP enzymes (Feyereisen, 2005). Duplications lead to the occurrence of genes which, if not lost subsequently, have the potential to become new functional genes. Phm and Spo are both microsomal CYPs that are related to CYP2 enzymes,

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Fig. 9.9 Illustration of a possible evolutionary scenario of spo-like (CYP307) genes in arthropods. Diversification of a common ancestral CYP307A into spo-like genes by gene duplications and losses is predicted based on data from Rewitz et al. (2007), Sztal et al. (2007) and the Daphnia data presented here. Extant genes are shown in black while genes that have been lost are light gray. * spot is probably lost in Apis but not all hymenopterans since it is present in the wasp Nasonia vitripennis (K.F.R. and L.I.G. 2007 unpublished data)

and thus, these genes may have an early common ancestor. Dib, Sad and Shd are in a monophyletic group of mitochondrial CYPs (Feyereisen, 2005) and their conserved gene structure (Fig. 9.5) suggests that their origins evolved by duplications. The relaxed constraint experienced by duplicated copies allows one copy to diversify functionally to evolve a novel and often related function (neofunctionalization: see Li et al., 2005). Such related functions could be modification of steroid molecules at different positions. Duplication of biosynthetic enzymes is therefore believed to be an important process by which biological pathways evolve (Corbin et al., 2004). It is likely that the Halloween genes (at least situated in the mitochondria) may have evolved stepwise by gene duplications to form part of the ecdysteroid pathway (see Rewitz et al., 2007).

9.5.3.2

Has Expression Divergence Allowed Evolutionary Survival of Cyp307a Paralogs?

While the other Halloween genes have been retained as 1:1 orthologs, spo-like genes have duplicated to form a subset of species-specific paralogs. Expression of spo and spok in D. melanogaster and Cyp307a3 and spok in D. virilis indicate differences between the two subgenera (Fig. 9.10). In D. melanogaster, spo is

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Fig. 9.10 Differential expression of Cyp307a genes in Drosophila melanogaster and Drosophila virilis. Semi-quantitative RT-PCR analysis of D. melanogaster spo (Cyp307a1) and spok (Cyp307a2), and D. virilis spok (Cyp307a2) and Cyp307a3. Emb; embryo, FL3; feeding third instar larvae, WL3; wandering third instar larvae, AdF; adult females. Genomic DNA (gDNA) is a positive control for spo and Cyp307a3 but not for spok (since primers spanned the intron no product is expected). -ive; negative control contains no DNA. Arrows indicate differences in the expression, that is, in D. melanogaster spo is expressed in adult females whereas in D. virilis spok occupy expression at this stage (Adapted with permission from Sztal et al. 2007)

expressed early during embryogenesis in the amnioserosa and in the follicle cells of the adult ovaries, and spok is expressed in the ring gland from late embryogenesis through larval development (Ono et al., 2006; Sztal et al., 2007). In D. virilis, however, Cyp307a3 is expressed in the early embryo but not in the adult ovaries like Dmspo. Instead spok is expressed in both the ring gland and the adult ovaries in D. virilis (Sztal et al., 2007). Differences in the spatio-temporal expression suggest that: (1) the Sophophoran subgenus spo and the Drosophila subgenus Cyp307a3 are different genes; (2) even though spo-like genes duplicate and evolve, activity of at least one spo-like gene is required throughout development (i.e. in the embryo, at larval stages in the ring gland and in the adult ovaries for reproduction). Thus, the ancestral gene, before the duplication into spo, spok and Cyp307a3 would probably have occupied the expression pattern shared by spo and spok in D. melanogaster and by spok and Cyp307a3 in D. virilis. This appears to be true in lepidopterans which have only one ortholog of spo that could be considered the extant equivalent of the ancestral gene. In Bombyx and Manduca (both lepidopterans), spo is expressed in the embryo, prothoracic glands and the adult ovaries (Namiki et al., 2005; Ono et al., 2006).

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According to the duplication-degeneration-complementation (DDC) model duplicated genes are often preserved because of partitioning of the ancestral function e.g. by complementary degenerative mutations in regulatory elements that separate the expression of the duplicated genes (Force et al., 1999). If the expression of duplicated genes evolves so that the expression of the ancestral gene is shared divergently between the two copies, there will be little or no overlapping expression and both copies will be needed to fulfill the requirements of the ancestral gene. In such cases, known as subfunctionalization, separation of expression to different tissues and/or developmental stages may allow specialization of paralogs. In Drosophila, the ancestral gene spok probably duplicated to form spo, and the expression of the two genes diverged. Because spo and spok are believed to be functionally redundant paralogs involved in ecdysteroid biosynthesis (Ono et al., 2006), these stage-specific components have likely survived because of their expression divergence. Although the genes are believed to be functionally redundant, some degree of specialization may have occurred. The same process has probably conserved spok and Cyp307a3 in species belonging to the Drosophila subgenus. Spo and Cyp307a3 may have been retrotransposed into genomic contexts that favored this divergence of expression. Alternative explanations might be invoked for the molecular evolution of spolike genes e.g. a disproportional high frequency of retrogenes are copied from X chromosomal source genes to more favorable genomic environments on autosomes (Emerson et al., 2004). However, since spo-like genes in D. melanogaster and D. virilis are autosomal, movement from the X chromosome does not explain survival of retrocopies (Sztal et al., 2007). While sequence divergence might explain survival of spo, spok and Cyp307a3 paralogs, the reason, if any, that spot has been retained in the genome of some insects and not in others is conjectural. The phylogeny clearly shows the division of spot from spo/spok (Fig. 9.7). It is of interest to consider why spot and spo/spok genes have evolved this level of divergence. Perhaps it indicates elaboration of function e.g. altered activity or even new function. Another possibility is that spot, like spo and spok, has survived because of changes that have separated their expression, or perhaps it is a combination of these factors that has contributed to the survival of these genes. Unlike spo(k), spot is not represented in the available pools of expressed sequence tags (ESTs), which indicate that gene is only expressed at low levels or in few cells and perhaps only transiently during development. However, spot genes have been conserved over hundreds of millions of years indicating selective constraints to preserve its function. Therefore, whatever its function is, it must be physiologically important. It is conceivable that Spot catalyzes a reaction depending on an ecdysteroid molecule because of its structural similarity to Spo/Spok. Spot is the only CYP307 family member in Apis which is known to produce ecdysteroids (see Lafont et al., 2005). This raises the possibility that Spot, at least to some extent, is functionally redundant to Spo/Spok since these enzymes are critical for ecdysteroid biosynthesis (Namiki et al., 2005; Ono et al., 2006). Some insects unable to dealkylate dietary phytosterols produce primarily makisterone A which is a C28 ecdysteroid (see Lafont et al., 2005). Makisterone A is the primary ecdysteroid in

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Apis and being the only ortholog, Spot may be adapted to be part of the ecdysteroid synthetic pathway from dietary sterol sources other than cholesterol. It is difficult to understand why genes in the CYP307 family have had the evolutionary flexibility to duplicate when the other Halloween genes involved in 20E biosynthesis have not. Speculation is further hampered by the fact that spo-like genes have thus far eluded functional characterization. Assuming that spo and spok are functionally redundant genes involved in the possible rate-limiting Black Box reaction(s) discussed in the first part of this contribution, raises the possibility of evolutionary pressure on the regulation of ecdysteroid biosynthesis. Regulation of ecdysteroid biosynthesis is important to control the precisely timed pulses of 20E necessary for developmental transitions (Gilbert et al., 2002; Warren et al., 2006). Duplication of genes in the CYP307 family may have provided a mechanism to allow differences in regulation to evolve in different arthropod species. Once the functions of spo and other genes in the CYP307 family are elucidated it should be possible to clarify the mechanisms driving the molecular evolution of these paralogs.

9.5.4

Evolutionary Conservation of Steroidogenic CYPs

Present data indicate that CYP enzymes involved in steroid hormone biosynthesis evolved from enzymes related to the vertebrate CYP2 family and mitochondrial CYPs. In vertebrates, most CYP enzymes mediating steroid hormone biosynthesis belong to these two groups and the same applies to the insect Halloween genes. Spo and Phm are both related to vertebrate CYP2s whereas Dib, Sad and Shd are mitochondrial CYPs. In C. elegans, daf-9 codes for a CYP which is believed to produce 3-keto bile acid-like steroids important for development (Motola et al., 2006). Daf-9 is also most closely related to CYP2 enzymes. Conservation of intronexon structures supports the premise that the insect Halloween genes and vertebrate steroidogenic CYP share a common evolutionary origin (Rewitz et al., 2007). This suggests that ancestral CYP2-like and mitochondrial CYPs were destined for steroid biosynthesis early in metazoan evolution, at least before the deuterostome– arthropod split and perhaps even before the onset of nematodes. It has been established biochemically that 20E biosynthesis takes place in the Y-organs of crustaceans (Lachaise et al., 1993). The identification of orthologs of the insect Halloween genes in Daphnia should allow the functional characterization of the genes which is necessary to provide unequivocal evidence of their role in ecdysteroid biosynthesis, as was done for Drosophila. So far, orthologs of the Halloween genes have only been identified in insects and now crustaceans. The complete genome sequence of C. elegans reveals the absence of Halloween genes in this organism, although it is possible that other nematodes may have these genes. DNA sequence information on annelids is scarce and the presence of the Halloween genes, although unlikely, cannot be ruled out at present. The Halloween genes may be specific to arthropods, although complete genome sequences of more

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Fig. 9.11 Conservation profiles of ecdysteroidogenic CYP products of the Halloween genes and CYP6 proteins with the predicted substrate recognition sites (SRSs) 1–6. Note that the area encompassing SRS2 and SRS3 is shown as 2/3. n = number of aligned sequences. — indicates the position of the conserved heme-binding domain (Reproduced from Rewitz et al. 2007) (See Color Plates)

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non-arthropod invertebrates are needed before a firm decision is made. Nonarthropods that synthesize ecdysteroids may have evolved biosynthetic strategies different from that involving the Halloween genes. Another possibility is that some of the biosynthetic enzymes appeared prior to the radiation of arthropods and that only arthropods developed the whole pathway yielding 20E by the progressive evolution of the biosynthetic enzymes. Several plants are known to synthesize 20E (Lafont et al., 2005); however, no data presently indicate the occurrence of the Halloween genes in plants. This implies that plants by convergent evolution have evolved 20E biosynthesis as a defense mechanism against herbivorous insects. In arthropods, the biosynthetic enzymes in the pathway yielding 20E appear to be reasonably well conserved. Alignment of primary structures of Halloween orthologs shows that in addition to the conserved CYP motifs, present in all CYPs, sites believed to be involved in substrate recognition are highly conserved (Fig. 9.11). This indicates that purifying selection (selection against deleterious mutation) has shaped these genes (Rewitz et al., 2007). Although conservation of SRSs indicates purifying selection of all the Halloween genes, different levels of amino acid sequence conservation in orthologous groups (Table 9.2) indicates differences in the constraints that have shaped these steroidogenic enzymes. Even though evolution has allowed orthologs in the family of spo-like genes to duplicate, these genes are the most highly conserved of the CYPs encoded by the Halloween genes (an average 54% identity of orthologous proteins: see Table 9.2). In the biochemical pathway yielding 20E, conservation of biosynthetic enzymes by selection against mutations that modify their activity must have been important since molting and metamorphosis are controlled by the rate of ecdysteroid production (Gilbert et al., 2002). In Manduca, expression of spo, phm, dib and sad in the prothoracic glands undergoes developmental changes that correlate with changes in the hemolymph ecdysteroid titer (Rewitz et al., 2006a; Ono et al., 2006; Fig. 9.4). This indicates that the enzymatic steps catalyzed by Spo, Phm, Dib, and Sad to some extent are important for controlling the flux of ecdysteroid precursors through the pathway to produce ecdysone. However, if control of production is focused on a step in the Black Box involving Spo/Spok this might have been a particular target of selective constraint to conserve the activity of the rate-limiting step. Duplications of spo genes may have allowed regulation of ecdysteroid biosynthesis to be adapted to the diverse life histories of insects during evolution.

9.6

Conclusions

The use of low ecdysteroid mutants of Drosophila have provided tantalizing bits of information that may in the long run aid in understanding the mysteries of the Black Box, the last remaining piece needed to complete the ecdysteroid biosynthetic pathway puzzle. It must be noted that although we have referred throughout to the Black Box as housing a single reaction, it may just as well be thought of as a series of reactions. Only when this particular quandary has been resolved will we truly

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understand how ecdysone is synthesized, probably in all arthropods, and perhaps be able to design insect growth regulators that act at specific enzymatic steps in the scheme. Further, such knowledge would set the stage for an understanding of the regulation of 20E synthesis, the rate limiting step, and by the use of genetics we may be able to truly understand how arthropods tightly control the titer of ecdysone and 20E in their hemolymph and at target cells. Answers to these questions are requisite for a complete understanding of insect growth and development. Acknowledgements We thank all our collaborators noted in the text and the members of our laboratory, James T. Warren and Robert Rybczynski, and the laboratories of Chantal DauphinVillemant and Michael B. O’Connor. Dr. O’Connor in particular has been a motivating force in the work on the Halloween genes and a valued colleague and friend. Research in our laboratory has been funded by grant IBN130825 from the National Science Foundation.

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Parvy JP, Blais C, Bernard F, Warren JT, Petryk A, Gilbert LI, O’Connor MB, Dauphin-Villemant C (2005) A role for βFTZ-F1 in regulating ecdysteroid titers during post-embryonic development in Drosophila melanogaster. Dev Biol 282:84–94 Payne AH, Hales DB (2004) Overview of steroidogenic enzymes in the pathway from cholesterol to active steroid hormones. Endocrinol Rev 25:947–970 Petryk A, Warren JT, Marques G, Jarcho MP, Gilbert LI, Parvy J-P, Dauphin-Villemant C, O’Connor MB (2003) Shade is the Drosophila P450 enzyme that mediates the hydroxylation of ecdysone to the steroid insect molting hormone 20-hydroxyecdysone. Proc Natl Acad Sci USA 100:13773–13778 Rewitz KF, Rybczynski R, Warren JT, Gilbert LI (2006a) Identification, characterization and developmental expression of Halloween genes encoding P450 enzymes mediating ecdysone biosynthesis in the tobacco hornworm, Manduca sexta. Insect Biochem Mol Biol 36:188–199 Rewitz KF, Rybczynski R, Warren JT, Gilbert LI (2006b) Developmental expression of Manduca shade, the P450 mediating the final step in molting hormone synthesis. Mol Cell Endocrinol 247:166–174 Rewitz KF, O’Connor MB, Gilbert LI (2007) Molecular evolution of the insect Halloween family of cytochrome P450s: phylogeny, gene organization and functional conservation. Insect Biochem Mol Biol 37:741–753 Richard D, Applebaum SW, Sliter TJ, Baker FC, Schooley DA, Reuter CC, Henrich V, Gilbert LI (1989) Juvenile hormone bisepoxide biosynthesis in vitro by the ring gland of Drosophila melanogaster: a putative juvenile hormone in the higher Diptera. Proc Natl Acad Sci USA 86:1421–1425 Roberts B, Gilbert LI (1986) Ring gland and prothoracic gland sensitivity to interspecific prothoracicotropic hormone extracts. J Comp Physiol 156:767–771 Roberts B, Whitten J, Gilbert LI (1976) Patterns of incorporation of tritiated thymidine by the dorsal polytene foot-pad nuclei of Sarcophaga bullata (Sarcophagidae: Diptera). Chromosoma 54:127–140 Rogozin IB, Wolf YI, Sorokin AV, Mirkin BG, Koonin EV (2003) Remarkable interkingdom conservation of intron position and massive, lineage-specific intron loss and gain in eukaryotic evolution. Curr Biol 13:1512–1517 Rybczynski R, Gilbert LI (1994) Changes in general and specific protein synthesis that accompany ecdysteroid synthesis in stimulated prothoracic glands of Manduca sexta. Insect Biochem Mol Biol 24:175–189 Saunders D, Henrich V, Gilbert LI (1989) Induction of diapause in Drosophila melanogaster: photoperiodic regulation and impact of arrhythmic mutations on time measurement. Proc Natl Acad Sci USA 86:3748–3752 Savard J, Tautz D, Richards S, Weinstock GM, Gibbs RA, Werren JH, Tettelin H, Lercher MJ (2006) Phylogenomic analysis reveals bees and wasps (Hymenoptera) at the base of the radiation of holometabolous insects. Genome Res 16:1334–1338 Sieglaff DH, Duncan KA, Brown MR (2005) Expression of genes encoding proteins involved in ecdysteroidogenesis in the female mosquito, Aedes aegypti. Insect Biochem Mol Biol 35: 471–490 Sliter TJ, Gilbert LI (1992) Developmental arrest and ecdysone deficiency resulting from mutations at the dre4 locus of Drosophila. Genetics 130:555–568 Sliter TJ, Henrich V, Tucker RL, Gilbert LI (1989) The genetics of the Dras3-Roughenedecdysoneless chromosomal region (62B3–4 to 62D3–4) in Drosophila melanogaster: analysis of recessive lethal mutations. Genetics 123:327–336 Smith CD, Shu S, Mungall CJ, Karpen GH (2007) The release 5.1 annotation of Drosophila melanogaster heterochromatin. Science 316:1586–1591 Smith SL, Bollenbacher WE, Cooper DY, Schleyer H, Wielgus JJ, Gilbert LI (1979) Ecdysone 20-monooxygenase: characterization of an insect cytochrome p-450 dependent steroid hydroxylase. Mol Cell Endocrinol 15:111–133 Smith SL, Bollenbacher WE, Gilbert LI (1983) Ecdysone 20-monooxygenase activity during larval-pupal development of Manduca sexta. Mol Cell Endocrinol 31:227–251

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Srivastava US, Gilbert LI (1968) Juvenile hormone: effects on a higher dipteran. Science 161:61–62 Srivastava US, Gilbert LI (1969) The influence of juvenile hormone on the metamorphosis of Sarcophaga bullata. J Insect Physiol 15:177–189 Sroka P, Gilbert LI (1971) Studies on the endocrine control of post-emergence ovarian maturation in Manduca sexta. J Insect Physiol 17:2409–2419 Sztal T, Chung H, Gramzow L, Daborn PJ, Batterham P, Robin C (2007) Two independent duplications forming the Cyp307a genes in Drosophila. Insect Biochem Mol Biol 37: 1044–1053 Taylor JS, Raes J (2004) Duplication and divergence: the evolution of new genes and old ideas. Annu Rev Genet 38:615–643 Warren JT, Gilbert LI (1986) Ecdysone metabolism and distribution during the pupal-adult development of Manduca sexta. Insect Biochem 16:65–82 Warren JT, Wismar J, Subrahmanyam B, Gilbert LI (2001) Woc (without children) gene control of ecdysone biosynthesis in Drosophila melanogaster. Mol Cell Endocrinol 181:1–14 Warren JT, Petryk A, Marques G, Jarcho M, Parvy J-P, Dauphin-Villemaont C, O’Connor MB, Gilbert LI (2002) Molecular and biochemical characterization of two P450 enzymes in the ecdysteroidogenic pathway of Drosophila melanogaster. Proc Natl Acad Sci USA 99: 11043–11048 Warren JT, Petryk A, Marques G, Jarcho M, Parvy JP, Shinoda T, Itoyama K, Kobayashi J, Jarcho M, Li Y, O’Connor MB, Dauphin-Villemant C, Gilbert LI (2004) Phantom encodes the 25-hydroxylase of Drosophila melanogaster and Bombyx mori: a P450 enzyme critical in ecdysone biosynthesis. Insect Biochem Mol Biol 34:991–1010 Warren JT, Yerushalmi Y, Shimell MJ, O’Connor MB, Gilbert LI (2006) Discrete pulses of molting hormone, 20-hydroxyecdysone, during late larval development of Drosophila melanogaster: correlations with changes in gene activity. Dev Dynam 235:315–326 Wismar J, Habtemichael N, Warren JT, Dai JD, Gilbert LI, Gateff E (2000) The mutation without children(rgl) causes ecdysteroid deficiency in third-instar larvae of Drosophila melanogaster. Dev Biol 226:1–17 Yoshiyama T, Namiki T, Mita K, Kataoka H, Niwa R (2006) Neverland is an evolutionarily conserved Rieske-domain protein that is essential for ecdysone synthesis and insect growth. Development 133:565–2574 Zhang J (2003) Evolution by gene duplication: an update. Trends Ecol Evol 18:292–298

Chapter 10

Recent Studies on Prothoracic Gland Cell Growth and Ecdysteroidogenesis in the Silkworm, Bombyx mori Shi-Hong Gu and Ju-Ling Lin

Abstract The prothoracic glands of developing insects are the sources of ecdysteroids that elicit molting and metamorphosis. Using silkworms as our model system, recent studies from our laboratory have focused on the following aspects: (1) stage-specific changes in prothoracictropic hormone (PTTH) signal transduction; (2) the activation of extracellular signal-regulated kinase (ERK) by PTTH in prothoracic gland cells; (3) correlation between cell growth and ecdysteroidogenesis during development; (4) a new autocrine regulatory mechanism underlying cell growth and ecdysteroidogenesis; (5) nutrition-dependent cell growth and ecdysteroidgenesis. Our results showed that development-specific PTTH-cAMP signal transduction, which showed different patterns between the fourth and last larval instars, may play an important role in regulating changes in prothoracic gland activity. Developmentspecific changes in ERK phosphorylation may also play a role in PTTH stimulation of ecdysteroidogenesis. DNA synthesis in the prothoracic gland cells undergoes a specific developmental change: the dramatic increases in DNA synthesis during the early (for 3rd and 4th instars) or middle (for last instar) stages precede the major increase in ecdysteroidogenesis during the later stages. We have also demonstrated that a novel autocrine factor activates both DNA synthesis of the gland cells and ecdysteroidogenesis by silkworm prothoracic glands. Moreover, our results showed that upon starvation on day 3 of the last instar, larvae increased ecdysteroid production rate to enhance the rate of survive. This review summarized our recent advance in studies on silkworm prothoracic glands. Keywords Bombyx mori • prothoracic gland • PTTH signal transduction • cell growth • ecdysteroid secretion • starvation

S.-H. Gu () Department of Zoology, National Museum of Natural Science, 1 Kuan Chien Road, Taichung, Taiwan, ROC e-mail: [email protected]

G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Introduction

The prothoracic glands of developing insects are the sources of ecdysteroids that elicit molting and metamorphosis (Gilbert et al., 1996, 2002). The ecdysteroidogenesis is stimulated by the prothoracicotropic hormone, a neuropeptide produced by the brain neurosecretory cells (Bollenbacher and Granger, 1985; Gilbert et al., 2002). In Bombyx mori, the gland activity undergoes specific developmental changes during the penultimate and last larval instars (Okuda et al., 1985; Gu et al., 1996, 1997, 2000; Gu and Chow, 2005b). The prothoracic glands during the early fourth larval instar produce detectable ecdysteroid and show activation response to PTTH; however, glands during the first stages of the last larval instar cannot produce detectable ecdysteroid and show no response to PTTH (Okuda et al., 1985; Gu et al., 1996, 1997, 2000; Gu and Chow, 2005b). We previously demonstrated that such stage-specific changes in gland activity play a critical role in regulating larval molting and metamorphosis, with the inactive prothoracic glands being the most upstream developmental events that lead to very low ecdysteroid levels, inactivation of corpora allata, as well as larval-pupal transformation (Gu and Chow, 1996; Gu et al., 1996). Considerable information on regulation of silkworm prothoracic gland activity has been obtained (Okuda et al., 1985; Gu et al., 1996, 1997; Gu and Chow, 2005b). Moreover, recently we have focused on regulation of cell growth of both prothoracic glands and corpora allata (Gu and Chow, 2001, 2003, 2005a). We reported that DNA synthesis in the prothoracic gland cells of B. mori undergoes a specific developmental change during the third, fourth, and last larval instars: the dramatic increases in DNA synthesis during the early (for third and fourth instars) or middle (for last instar) stages precede the major increase in ecdysteroidogenesis during the later stages (Gu and Chow, 2001, 2005a). We have also demonstrated that a novel autocrine factor activates both DNA synthesis of the gland cells and ecdysteroidogenesis (Gu, 2006, 2007). Our results showed that when glands were incubated in the small volume of medium (10 µl), both DNA synthesis of prothoracic gland cells and ecdysteroidogenesis were dramatically increased as compared with those incubated in large volume (50 µl) (Gu, 2006, 2007). In addition, autocrine activation of ecdysteroidogenesis was also observed in the locusts Locusta migratoria and Schistocerca gregaria (Vandersmissen et al., 2007). Moreover, our study showed that the activation of DNA synthesis in gland cells during the middle stages of the last larval instar was nutrition-dependent with the starvation on day 3 inhibiting DNA synthesis. Our results showed that upon starvation on day 3, the dramatic increase in protein levels of the glands was inhibited as compared with that of control larvae. However, the glands from starved larvae secreted similar levels of ecdysteroids as those of control larvae, leading to the increased rate of ecdysteroidogenesis. The current review will focus on the results on regulatory mechanism of insect prothoracic gland cell growth and ecdysteroidogenesis obtained during the recent years in our lab.

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Results and Discussion Changes in PTTH-cAMP Signaling During Silkworm Development

It was previously demonstrated that PTTH stimulation of ecdysteroid synthesis appears to be mediated by cAMP as an intracellular second messenger in both Manduca sexta (Smith et al., 1984, 1985; Gilbert et al., 1996, 2002) and B. mori (Gu et al., 1996, 1997, 2000). In our study, we examined cAMP accumulation and ecdysteroid production in vitro throughout the fourth and last larval instars of the silkworm, B. mori. We compared stage-related ecdysteroidogenic responses of the glands to dibutyryl cAMP (dbcAMP), 1-methyl-3-isobutylxanthine (MIX), and PTTH. A comparison of cAMP accumulation to stimulation by PTTH was also conducted. Our results lend support to the notion that a cAMP-sensitive steroidogenic apparatus undergoes developmentspecific changes during both the fourth and last larval instars. For the fourth larval instar, according to the differential responsiveness of prothoracic glands to PTTH, the following three different stages were classified and changes in PTTH signal transduction were assumed. During the first stage (between days 0 and 1), the glands showed low basal and PTTH-stimulated activities in both cAMP accumulation and ecdysteroidogenesis, and PTTH release in vitro was maintained at low but detectable levels, implying that a low but sustained PTTH signal may be transduced to prothoracic gland cells. On day 1.5, when low basal ecdysteroid production of the prothoracic glands was being maintained, both the responsiveness of glands to the stimulation of PTTH and PTTH release in vitro dramatically increased, indicating greatly increased PTTH transduction. On day 3 (when the basal ecdysteroidogenesis became maximal) and afterwards, high PTTH release in vitro was maintained, but the gland showed no response to PTTH, implying that the refractoriness of gland cells to PTTH may occur at this stage. For the last larval instar, changes in PTTH signal transduction were also assumed and compared with those of the fourth larval instar. Results showed that the prothoracic glands during the first 3 days of the last instar cannot produce detectable ecdysteroid and showed no response to stimulation by PTTH or MIX. However, artificial elevation of cellular cAMP levels by in vitro dibutyryl cAMP treatment stimulated the glands to secrete detectable ecdysteroid, implying the presence of a cAMP-dependent ecdysteroidogenic apparatus during this stage. From days 3 to 8, basal gland activities fluctuated, but the glands showed activation responses to PTTH and to the chemicals that increase cellular cAMP levels. After the occurrence of the peak in basal gland activity on day 9, glands on day 10 showed no response to PTTH, implying a refractory state of the glands to PTTH stimulation. From these results, we concluded that development-specific PTTH signal transduction during the last larval instar, which shows a different pattern from that of the penultimate larval instar, may play an important role in regulating changes in prothoracic gland activity and in leading to larval-pupal metamorphosis.

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Stimulation of ERK by PTTH

ERKs belong to a member of mitogen-activated protein kinases (MAPKs). Upon extracellular stimulation, the ERKs are activated by a network of interacting proteins, which funnel the signals into a multitier kinase cascade (Lewis et al., 1998). Although the ERKs were first implicated in the regulation of proliferation and differentiation, it has been presently known that these kinases also participate in the control of various cell activities, including cellular morphology, learning and memory in the central nervous system (Widmann et al., 1999). In M. sexta, it was demonstrated that ERKs are involved in PTTH stimulation of ecdysteroidogenesis by prothoracic glands (Rybczynski et al., 2001; Rybczynski and Gilbert, 2003). However, such researches were conducted only on a single insect species. It is not clear whether or not the results obtained from M. sexta could apply to other insects. Indeed the available data indicate that some differences exist in cAMP-dependent steroidogenic competence of glands between M. sexta and B. mori (Smith et al., 1984, 1985; Gilbert et al., 1996, 2002; Gu et al., 1996, 1997; Gu and Chow, 2005b). Moreover, although the activation of glandular ERK phosphorylation by PTTH was observed by in vitro experiments (Rybczynski et al., 2001; Rybczynski and Gilbert, 2003), it is not clear whether ERK phosphorylation by PTTH could be demonstrated by in vivo PTTH stimulation. In the present study, we investigated the activation of ERK phosphorylation by PTTH in prothoracic gland cells of the silkworm, B. mori. Our results showed that PTTH stimulated ERK phosphorylation in time- and dose-dependent manners in vitro (Fig. 10.1). The ERK phosphorylation inhibitors PD98059 and U0126 blocked both basal and PTTH-stimulated ERK phosphorylation and ecdysteroidogenesis (Fig. 10.2). Our results showed that in vitro activation of glandular ERK phosphorylation by PTTH appears to be developmentally regulated with refractoriness of gland cells to PTTH occurring during the later stages of both the fourth

Fig. 10.1 Time- and dose-dependent effects of ERK phosphorylation activation by PTTH. Prothoracic glands from day 7 last instar larvae of B. mori were treated with PTTH for the indicated time points (A) or treated with the indicated concentrations of PTTH or incubated with control medium (con) for 30 min (B). Gland lysates were then prepared and subjected to immunoblotting analysis with anti-phospho-ERK antibody (Adapted from Lin and Gu, 2007)

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Fig. 10.2 Inhibitory effects of PTTH-stimulated ERK phosphorylation (A, B) and ecdysteroidogenesis (c) by PD98059 and U0126. (A) Prothoracic glands from day 7 last instar larvae were pretreated with 50 µM PD98059 or vehicle alone for 30 min, then transferred to medium containing the same dose of PD98059, with or without PTTH. (B) Prothoracic glands from day 7 last instar larvae were pretreated with 10 µM U0126 or vehicle alone for 30 min, then transferred to medium containing the same dose of U0126, with or without PTTH. The incubation was maintained for 30 min. Gland lysates were then prepared and subjected to immunoblot analysis with an anti-phospho-ERK antibody. (C) Prothoracic glands from day 7 last instar larvae were pretreated with either 50 µM PD98059, 10 µM U0126, or vehicle alone for 30 min, then transferred to medium containing the same doses of either PD98059 or U0126, with or without PTTH. The incubation was maintained for 2 h. Ecdysteroids released during the 2-h incubation were quantified. Each value is an average ± SEM (n = 8). (** p < 0.01 for PTTH + U0126 vs. PTTH;* p < 0.05 for PTTH + PD98059 vs. PTTH, Student’s t-test) (Adapted from Lin and Gu, 2007)

and last larval instars. Moreover, in vitro activation of ERK phosphorylation of prothoracic glands by PTTH was also verified by in vivo experiments: injection of PTTH into day 6 last instar larvae greatly increased activity of glandular ERK phosphorylation and ecdysteroidogenesis. From these results, it is suggested that development-specific changes in ERK phosphorylation may play a role in PTTH stimulation of ecdysteroidogenesis.

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Temporal Changes in DNA Synthesis of Prothoracic Gland Cells During Larval Development and Their Correlation with Ecdysteroidogenic Activity

DNA synthesis in prothoracic gland cells of B. mori, was studied immunocytochemically after in vivo labeling with 5-bromo-2'-deoxyuridine (BrdU), and its developmental changes during the third, fourth, and last larval instars were examined (Fig. 10.3). During the early stages of both the third and fourth larval instars, a dramatic increase in the number of DNA-synthesizing cells of the prothoracic glands was detected. However, during the latter stages of each instar, the number of DNA-synthesizing cells greatly decreased. The determination of glandular protein content showed that dramatic increases occurred during the latter stages of each larval instar. Comparison of changes in prothoracic gland cell DNA synthesis with ecdysteroidogenic activity showed that the increase in DNA synthesis precedes ecdysteroidogenesis. The cellular mechanism underlying changes in prothoracic gland cell DNA synthesis during the last two larval instars was further analyzed by determining the in vitro DNA synthesis of the glands, their responsiveness to hemolymph growth factors, and changes in the growth-promoting activity of hemolymph during development. We found that both growth factors and the responsiveness of the prothoracic gland cells to growth factors from hemolymph may play roles in regulating DNA synthesis of gland cells.

10.2.4

A New Autocrine Regulatory Mechanism Underlying Cell Growth and Ecdysteroidogenesis

Autocrine activation of DNA synthesis in prothoracic gland cells in the last instar larvae was studied using both a long-term in vitro organ culture system and immunocytochemical labeling with BrdU (Fig. 10.4). When prothoracic glands were incubated individually in a small volume of medium (10 µl/gland), the numbers of DNA-synthesizing cells per gland increased significantly, and they showed a lower stimulatory effect by hemolymph as compared with those incubated in a large volume of medium (50 µl/gland). Moreover, cultured groups of glands (six glands per group in a 50-µl drop) also resulted in much higher levels of DNA synthesis than those cultured individually in a 50-µl drop. The mechanism by which alternation of the volume of the incubation medium results in changes in the levels of DNA synthesis was further examined. When prothoracic glands were incubated in the medium (50-µl drop per gland) that was preconditioned with glands (in a 10-µl drop individually), a dramatic increase in DNA synthesis activity was also observed, indicating that prothoracic glands may release a factor that stimulates their own DNA synthesis. We further characterized the growth-promoting factor and found that the factor seems to be heat stable, and its molecular weight was estimated to be between 1,000 and 3,000 Da. Moreover, the factor also stimulated corpus allatum

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Fig. 10.3 Immunoperoxidase-stained whole mounts of silkworm prothoracic glands showing incorporation of a brief pulse of BrdU. (a) Last instar larvae which had been given a 2-h pulse of BrdU on day 5. (b) BrdU-labeled cells at higher magnification. BrdU-labeled cells are identified by the intense black precipitation of diaminobenzidine (arrow). Scale bar, 45 µm (Adapted from Gu and Chow, 2005a)

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Fig. 10.4 Immunoperoxidase-stained whole mounts of silkworm prothoracic glands showing the incorporation of BrdU. (a) Prothoracic gland from day 4 last instar larvae incubated individually in 30 µl Grace’s medium containing BrdU. (b) Prothoracic gland from day 4 last instar larvae incubated individually in 5 µl Grace’s medium containing BrdU. BrdU-labeled cells were identified by the intense black precipitation of diaminobenzidine. Scale bar, 75 µm (Adapted from Gu, 2006)

cell DNA synthesis in vitro. However, it did not stimulate cell DNA synthesis of salivary glands (Fig. 10.5). Injection of concentrated putative growth-promoting factor into day 4 last instar ligated larvae greatly increased cell DNA synthesis of the prothoracic glands, indicating the in vivo function of the present factor. To our knowledge, this is the first study to demonstrate that prothoracic glands secrete an autocrine growth factor in vitro which has a biological function in vivo. Using the same experimental approaches, we further found autocrine activation of ecdysteroidogenesis in prothoracic glands of B. mori. It was found that either decreasing the incubation volume, from 100 to 5 µl, or increasing the number of glands incubated per drop (50 µl) from one to five significantly increased ecdysteroid secretion. From the similarity between ecdysteroid secretion promoting factor and the autocrine growth factor reported in the above study, it was supposed that it may be the same molecular that has both growth-promoting and ecdysiotropic functions.

10.2.5

Effects of Starvation on Growth, Metamorphosis and Ecdysteroidogenesis of the Prothoracic Glands During Last Larval Instar

Stage-dependent effects of starvation on growth, metamorphosis and ecdysteroidogenesis of the prothoracic glands during the last larval instar of B. mori were studied. When last instar larvae were starved starting on day 1, all larvae died between days 5 and 7 of the instar. Although PTTH release from brain-corpus cardiacum-corpus allatum (BR-CC-CA) did not significantly alter upon starvation,

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Fig. 10.5 Immunoperoxidase-stained whole mounts of silkworm salivary glands (a, b), and corpora allata (c, d) showing the incorporation of BrdU. (a) Salivary gland from day 0 last instar larvae incubated in 50 µl Grace’s medium containing BrdU. (b) Salivary gland from day 0 last instar larvae incubated in 50 µl Grace’s medium containing BrdU and 10% hemolymph. (c) Corpus allatum from day 0 last instar larvae incubated in 50 µl of control medium containing BrdU. (d) Corpus allatum from day 0 last instar larvae incubated in 50 µl prothoracic gland-preconditioned medium. Each incubation was maintained for 2 days, and the number of BrdU-labeled cells was counted after a 2-day incubation. BrdU-labeled cells are identified by the intense black precipitation of diaminobenzidine. Scale bar, 75 µm (Adapted from Gu, 2006) (See Color Plates)

the deficiency in PTTH signal transduction was maintained, which leads to very low hemolymph ecdysteroids after starvation started. However, when starvation started on day 3 of the last larval instar, the major hemolymph ecdysteroid peak, preceding larval-pupal transformation, occurred 1 day earlier than that in control larvae. The protein content of the prothoracic glands in day 3-starved larvae maintained at low levels as compared to those in control larvae. The secretory activity of the prothoracic gland in day 3-starved larvae maintained at the levels similar to those of control larvae. However, the rate of ecdysteroidogenesis, expressed per microgram of glandular protein, was greatly enhanced in these starved larvae, indicating that upon starvation, larvae increased ecdysteroid production rate to enhance the rate of survival (Chen and Gu, 2006).

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Future Research

From molecular genetic analysis of Drosophila melanogaster, it has been well documented that Ras activity in the prothoracic glands is involved in ecdysteroid release: inhibition of Ras signaling in the prothoracic gland greatly attenuates the increase in ecdysteroid titer, whereas activating Ras causes a precocious increase in the titer (Caldwell et al., 2005). Moreover, it has been shown that both Raf and PI3K participate in the timing of ecdysteroid release (Caldwell et al., 2005). Our preliminary study showed that incubation of silkworm prothoracic glands with Ras farnesyltransferase inhibitor manumycin A greatly decreased PTTH-stimulated ecdysteroidogenesis, implying the involvement of Ras in silkworm ecdysteroidogenesis. However, the exact linking components from PTTH to Ras to ecdysteroidogenesis remain to be investigated. Our future studies will focus on the PTTH signal transduction mechanism involving Ras, Raf, MEK and ERK. In addition, our preliminary study showed that in addition to rapid stimulation of ecdysteroid secretion by PTTH, PTTH may also stimulate cellular growth including protein synthesis. Further study will focus on stimulation of several key components involving glandular cell growth by PTTH. Furthermore, an investigation into the correlation between nutrient and prothoracic cell growth and ecdysteroidogenesis will also be conducted. We will study nutritional control of prothoracic gland cell growth, and how nutrient regulates ecdysteroid secretion, as well as regulatory mechanism from feeding to signal transduction to specific gene expression. Acknowledgements The authors thank the National Science Council for grants NSC 93-2313B-178-002, 93-2313-B-178-003, 94-2313-B-178-001, 94-2313-B-178-002, 95-2311-B-178-001, and 95-2313-B-178-002 and the National Museum of Natural Science of the Republic of China for their financial support.

References Bollenbacher WE, Granger NA (1985) Endocrinology of the prothoracicotropic hormone. In: Kerkut GA, Gilbert LI (eds) Comprehensive Insect Physiology, Biochemistry and Pharmacology, Pergamon Press, New York, Vol. 7, pp. 109–151. Caldwell PE, Walkiewicz M, Stern M (2005) Ras activity in the Drosophila prothoracic gland regulates body size and developmental rate via ecdysone release. Curr Biol 15:1–11. Chen CH, Gu SH (2006) Stage-dependent effects of starvation on the growth, metamorphosis, and ecdysteroidogenesis by the prothoracic glands during the last larval instar of the silkworm, Bombyx mori. J Insect Physiol 52:968–974. Gilbert LI, Rybczynski R, Tobe S (1996) Endocrine cascade in insect metamorphosis. In: Gilbert LI, Tata J, Atkison P (eds) Metamorphosis: Post-Embryonic Reprogramming of Gene Expression in Amphibian and Insect Cells, Academic, San Diego, CA, pp. 59–107. Gilbert LI, Rybczynski R, Warren JT (2002) Control and biochemical nature of the ecdysteroidogenic pathway. Annu Rev Entomol 47:883–916. Gu SH (2006) Autocrine activation of DNA synthesis in prothoracic gland cells of the silkworm, Bombyx mori. J Insect Physiol 52:136–145. Gu SH (2007) Autocrine activation of ecdysteroidogenesis in prothoracic glands of the silkworm, Bombyx mori. J Insect Physiol 53:538–549.

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Gu SH, Chow YS (1996) Regulation of juvenile hormone biosynthesis by ecdysteroid levels during the early stages of the last two larval instars of Bombyx mori. J Insect Physiol 42:625–632. Gu SH, Chow YS (2001) Induction of DNA synthesis by 20-hydroxyecdysone in the prothoracic gland cells of the silkworm, Bombyx mori during the last larval instar. Gen Comp Endocrinol 124:269–276. Gu SH, Chow YS (2003) Stage-dependent effects of 20-hydroxyecdysone on DNA synthesis of corpus allatum cells in the silkworm, Bombyx mori. J Exp Zool 297A:138–146. Gu SH, Chow YS (2005a) Temporal changes of DNA synthesis in the prothoracic gland cells during larval development and their correlation with ecdysteroidogenic activity in the silkworm, Bombyx mori. J Exp Zool 303A:249–258. Gu SH, Chow YS (2005b) Analysis of ecdysteroidogenic activity of the prothoracic glands during the last larval instar of the silkworm, Bombyx mori. Arch Insect Biochem Physiol 58:17–26. Gu SH, Chow YS, Lin FJ, Wu JL, Ho RJ (1996) A deficiency in prothoracicotropic hormone transduction pathway during the early last larval instar of Bombyx mori. Mol Cell Endocrinol 120:99–105. Gu SH, Chow YS, Yin C-M (1997) Involvement of juvenile hormone in regulation of prothoracicotropic hormone transduction during the early last larval instar of Bombyx mori. Mol Cell Endocrinol 127:109–116. Gu SH, Tsia WH, Chow YS (2000) Temporal analysis of ecdysteroidogenic activity of the prothoracic glands during the fourth larval instar of the silkworm, Bombyx mori. Insect Biochem Mol Biol 30:499–505. Lewis TS, Shapiro PS, Ahn NG (1998) Signal transduction through MAP kinase cascades. Adv Cancer Res 74:49–139. Lin JL, Gu SH (2007) In vitro and in vivo stimulation of extracellular signal-regulated kinase (ERK) by prothoracicotropic hormone in prothoracic gland cells and its developmental regulation in the silkworm, Bombyx mori. J Insect Physiol 53:622–631. Okuda M, Sakurai S, Ohtaki T (1985) Activity of the prothoracic gland and its sensitivity to prothoracicotropic hormone in the penultimate and last-larval instar of Bombyx mori. J Insect Physiol 31:455–461. Rybczynski R, Bell SC, Gilbert LI (2001) Activation of an extracellular signal-regulated kinase (ERK) by the insect prothoracicotropic hormone. Mol Cell Endocrinol 184:1–11. Rybczynski R, Gilbert LI (2003) Prothoracicotropic hormone stimulated extracellular signal-regulated kinase (ERK) activity: the changing roles of Ca2 + - and cAMP-dependent mechanisms in the insect prothoracic glands during metamorphosis. Mol Cell Endocrinol 205:59–68. Smith WA, Gilbert LI, Bollenbacher WE (1984) The role of cyclic AMP in the regulation of ecdysone synthesis. Mol Cell Endocrinol 37:285–294. Smith WA, Gilbert LI, Bollenbacher WE (1985) Calcium-cyclic AMP interactions in prothoracicotropic hormone stimulation of ecdysone synthesis. Mol Cell Endocrinol 39:71–78. Vandersmissen T, De Loof A, Gu SH (2007) Both prothoracicotropic hormone and an autocrine factor are involved in control of prothoracic gland ecdysteroidogenesis in Locusta migratoria and Schistocerca gregaria. Peptides 28:44–50. Widmann C, Gibson S, Jarpe MB, Johnson GL (1999) Mitogen-activated protein kinase: conservation of a three-kinase module from yeast to human. Physiol Rev 79:143–180.

Chapter 11

Diversity in Factors Regulating Ecdysteroidogenesis in Insects Sandrien Van de Velde$, Liesbeth Badisco$, Elisabeth Marchal$, Jozef Vanden Broeck, and Guy Smagghe

Abstract Ecdysteroid synthesis in insects was long considered as an exclusive feature of the ecdysial glands in larvae, and control of molting and metamorphosis was long thought to be the only function of ecdysteroids. The ‘classical dogma’ (Delbecque et al., 1990) of insect endocrinology states that ecdysteroids are produced by the prothoracic glands (PGs) (or analogs: ventral glands and ring glands), which are under control of the neuroprotein prothoracicotropic hormone (PTTH) secreted by the insect brain. However, extensive research during the last decades revealed that ecdysteroid synthesis can be performed by several other tissues besides the PGs, and is not solely restricted to larvae. Moreover, other factors besides PTTH have been shown to control ecdysteroid biosynthesis as well. It is now well established that in adult females, the ovaries are the primary source of ecdysteroids, where they play a role in several aspects of reproduction and are incorporated into the eggs for future embryonic development. In males, it has been demonstrated for some insect species that both immature testes in late larval stages and adult testes are capable of producing ecdysteroids. However, in comparison with ovarian ecdysteroidogenesis, not much attention has been paid to research on testicular ecdysteroidogenesis, and the biological role of testicular ecdysteroids remains far less clear. In addition, oenocytes and epidermis have been proposed as alternative sites for ecdysteroid production (reviewed by Delbecque et al., 1990). Ecdysteroidogenesis in insects and multifactorial regulation of ecdysteroidogenesis thus seems far more complex than was initially stated by the classical dogma. In this chapter, we aim to present an as complete and up-to-date as possible overview of the variety of factors involved in ecdysteroidogenesis, performed by a S. Van de Velde and G. Smagghe (* ü) Laboratory of Agrozoology, Department of Crop Protection, Faculty of Bioscience Engineering, Ghent University, Coupure Links 653, B-9000 Ghent, Belgium e-mail: [email protected] L. Badisco, E. Marchal and J. Vanden Broeck Section of Animal Physiology and Neurobiology, Zoological Institute, K.U.Leuven, Naamsestraat 59, B-3000 Leuven, Belgium Equally contributed

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variety of tissues. Factors are discussed per developmental stage (juvenile/adult) and per ecdysiosynthetic tissue. Keywords Ecdysteroidogenesis • Insects • Development-reproduction

11.1

11.1.1

Factors Regulating Ecdysteroidogenesis in Juvenile Insects Factors Acting on the Larval Ecdysial Glands

It is widely accepted that the tropic hormone PTTH is the most important regulator of PG ecdysteroidogenesis. Alterations of the hemolymph ecdysteroid titer can basically be explained by alterations of PTTH levels. However, other factors – tropic and static – are also at play, responsible for the ‘fine-tuning’ of the hemolymph ecdysteroid titer. Most of them are peptides originating in the brain, but other, extracerebral factors of both peptidic and non-peptidic nature also affect PG ecdysteroidogenesis. Moreover, the ‘classic’ insect hormones juvenile hormone (JH) and 20-hydroxyecdysone (20E) are involved in the regulation of PG ecdysteroidogenesis as well. The action of JH and 20E is not exclusively stimulatory or inhibitory, but depends on the age and physiological state of the insect. A major challenge for the future is the determination of the relative importance of all these factors in ecdysteroidogenesis throughout development. A delicate balance between ecdysiotropic and ecdysiostatic factors leads to the complex secretory pattern observed in vivo.

11.1.1.1

PTTH – Prothoracicotropic Hormone

Already in 1922, Kopec´ provided evidence that the brain of the caterpillar, Lymantria dispar, secretes a factor that can control metamorphosis. Later research focussed primarily on two Lepidoptera, the silkworm, Bombyx mori and the tobacco hornworm, Manduca sexta. These experiments showed that this factor is synthesized by a pair of large neurosecretory cells (NSCs) and stored in the corpora allata, the neurohemal organ for PTTH. Release is done at particular developmental stages under control of several physiological factors, such as nutritional state, and environmental cues, such as photoperiod and time of day. The factor exerts its function via the control of ecdysteroidogenesis by the prothoracic glands. Once released into the hemolymph, it stimulates the glands to synthesize ecdysteroids (as reviewed by Gilbert et al., 1997, 2002; Gilbert, 2004; Rybczynski, 2005). Several attempts were made to characterize and purify this PTTH, but it was not until 1991 that Kataoka et al. (1991) purified and fully determined the primary structure of the PTTH of B. mori. Bombyx-PTTH was shown to be synthesized

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as a prohormone of 224 amino acids that is cleaved and results in a 109 amino acid monomer to be assembled into a glycosylated homodimeric structure with intra- and intermolecular disulfide (cysteine-cysteine) bonds (Kawakami et al., 1990; Kataoka et al., 1991). Since then PTTH sequences were derived of several other Lepidoptera: Antheraea pernyi, Samia cynthia and Hyalophora cecropia (as reviewed by Rybczynski, 2005) and more recently molecular characterization yielded putative sequences of M. sexta, Helicoverpa zea, Heliothis virescens, Helicoverpa armigera and Spodoptera exigua (Shionoya et al., 2003; Xu and Denlinger, 2003; Xu et al., 2003, 2007; Wei et al., 2005). Only modest amino acid identity and similarity was found when pair-wise comparisons were made, though some structural features such as the presence of seven cysteine residues, glycosylation site and the location of hydrophobic regions, were clearly conserved. These proteins seem to be inactive across species; the primary amino acid sequence is thus critical for bioactivity. The search for PTTH-like molecules in other insect orders still seems to be difficult and cloudy. Reports have been made on the characterization of PTTHs from the very important dipteran model organism, Drosophila melanogaster, but so far general consensus about this discovery does not seem to have been reached (Kim et al., 1997; as reviewed by Rybczynski, 2005; McBrayer et al., 2007). It should be mentioned that other smaller molecules were also shown to exhibit PTTH activity, namely bombyxin of Bombyx and small PTTH of M. sexta. These will later be discussed (Sections 11.1.1.2 and 11.1.1.3). The last decades, a lot of research has been done on the PTTH signal transductory cascade in the PGs of especially Bombyx and Manduca (as reviewed by Gilbert et al., 2002; Rybczynski, 2005). A short summary will be given here. After release into the hemolymph, PTTH binds to a receptor at the cell membrane surface of the PG. This receptor remains as yet unidentified but belongs very likely to the family of G-protein coupled receptors (GPCRs). Recently, Nagata et al. (2006) found a new receptor cDNA in the PG of B. mori during a screening targeting the PTTH receptor. First data indicate that this cDNA is a good candidate for the PTTH receptor, but confirmation requires further analysis. After interaction with its receptor, PTTH causes an influx of extracellular Ca2+. This second messenger increases cAMP generation, which indicates the participation of a Ca2+-calmodulin dependent adenylate cyclase. It has been shown that this rise in intracellular Ca2+ content activates protein kinases and a series of protein phosphorylations, of which the most well-known is the phosphorylation of protein S6 (that is known to be responsible for increased rates of translation initiation). A rather complicated signal transduction cascade has been revealed, involving protein kinase A and several additional kinases such as p70 S6 kinase and possibly a tyrosine kinase (as reviewed by Rybczynski, 2005) and an extracellular signal-regulated kinase (ERK) (Lin and Gu, 2007). Recent research also demonstrated that protein kinase C also participates in PTTH signalling (Rybczynski and Gilbert, 2006). The final signal transduction step in this peptide-regulated steroid hormone production is the rapid translation of several proteins (Keightley et al., 1990). Two of these proteins are identified as a 70 kDa heat shock protein and a β-tubulin. As in vertebrates, neuropeptides regulating steroidogenesis exert their effect via transcriptional regulation of steroidogenic enzymes (Kagawa et al., 1999; Sewer and Waterman, 2003).

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An illustration thereof was recently demonstrated by Niwa et al. (2005) in B. mori: the expression of the ‘Halloween’-gene ‘disembodied’ (Cyp302a1) was found to be upregulated by treatment with PTTH (Niwa et al., 2005). It was already known that PTTH treatment stimulates gene transcription in the PG (Keightley et al., 1990), but disembodied is the first gene found to be regulated in this way.

11.1.1.2

‘Small PTTH’: Bombyxin and Other Insulin-Like Peptides (ILPs)

First, a remark should be made on the use of the term ‘prothoracicotropic hormone’ or ‘PTTH’. It is often cited in literature that there exist at least two different PTTHs, namely ‘big PTTH’ and ‘small PTTH’ (Koolman, 1995). ‘Big PTTH’ is the true PTTH as described above (Section 11.1.1.1). ‘Small PTTH’ has been used to indicate smaller (MW < 10 kDa) neuropeptides with tropic activity on the PGs, although these molecules are structurally not related to big PTTH (except maybe Manduca small PTTH, see Section 11.1.1.3). In fact, since other factors besides big and small PTTH have been demonstrated to stimulate ecdysteroidogenesis in the PGs, the term ‘prothoracicotropic hormone’ could cover in principle any of these factors with tropic activity on the PGs. Members of the insulin superfamily are not restricted to vertebrates, but have also been identified in invertebrate species, where they have been termed ‘insulin-like peptides’ (ILPs) or ‘insulin-related peptides’ (IRPs). For a review on insect ILPs, we refer the reader to Claeys et al. (2002), and Wu and Brown (2006). Bombyxin, isolated from the brain of the silkworm Bombyx mori, was the first insulin-like molecule identified in insects (Nagasawa et al., 1984). Bombyxin was originally identified as the Bombyx small PTTH (4 K-PTTH), since it exerted ecdysiotropic activity in vivo in the related silkworm species, Samia cynthia ricini. Curiously, Bombyx small PTTH failed to activate in vivo the PGs of Bombyx itself. This result was later confirmed when small PTTH was tested for its ecdysteroidogenic effects on Bombyx PGs in vitro (Kiriishi et al., 1992). Because of this lack of homologous prothoracicotropic activity, Mizoguchi et al. (1987) proposed calling it bombyxin instead of small PTTH. However, at present, over 30 bombyxin genes have been identified from B. mori (Kondo et al., 1996; Tsuzuki et al., 1997; Yoshida et al., 1997, 1998). Thus the possibility that one of these genes might code for a true prothoracicotropic peptide cannot be ruled out (Rybczynski, 2005). Few other reports have been made on the involvement of ILPs in PG ecdysteroidogenesis. Two SBRPs, Samia cynthia ricini bombyxin-related peptides (Kimura-Kawakami et al., 1992), actually do possess prothoracicotropic activity in Samia itself (Nagata et al., 1999), but their activity appears to be 20- to 100-fold weaker than that of Bombyx bombyxin in the same Samia cynthia in vivo assay. Moreover, the SBRPs seem 200–1,000 fold less effective than the Samia ‘big PTTH’. The physiological relevance of these factors thus remains rather obscure. It is, however, worth mentioning that also the Samia genome contains many other SBRP genes (Kimura-Kawakami et al., 1992), which possibly can code for SBRPs with stronger

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activity than that of the two forms tested. Injection of anti-insulin antibodies at specific moments during the fifth instar (i.e. exactly at the time when insulin-like molecules are released from the NSCs), prevented molting in Rhodnius prolixus larvae (Sevala et al., 1992), suggesting a stimulatory role for Rhodnius ILPs in PG ecdysteroidogenesis. Whether ILPs act directly on the PGs is not known. It is however intriguing, according to the observations of Sevala et al. (1992), that the timing of the release of ILPs and of the sensitivity of molting to the injection of anti-insulin antibodies, corresponds to what is known about the timing of the release of PTTH in Rhodnius. These observations lead to the hypothesis that ILPs might have some role in the control of PTTH release, thus acting indirectly on the PGs. A direct effect can be distinguished from an indirect effect by means of in vitro PG incubation assays. Vafopoulou and Steel (1997) demonstrated that bombyxin was much less effective than Bombyx PTTH at stimulating ecdysteroidogenesis by Rhodnius PGs in vitro, while on the contrary bombyxin was the most effective in the Samia in vivo assay (Rybczynski, 2005). The combination of these results points at the possibility of an indirect control mechanism. ILPs may not only affect PG ecdysteroidogenesis, but also seem to trigger ovarian ecdysteroidogenesis. Action of ILPs on ovaries will be thoroughly discussed further in this chapter (Section 11.2.1.1.4).

11.1.1.3

Manduca Small PTTH

In 1984 Bollenbacher et al., presented evidence for the existence of two molecular forms of PTTH in M. sexta, namely ‘big’ PTTH (∼25.5 kDa) and ‘small’ PTTH (∼7 kDa). This small PTTH was shown to act as an ecdysiotropin both in vitro and in vivo and appeared to be considerably more effective at stimulating ecdysteroid synthesis in larval glands compared to pupal glands (Bollenbacher et al., 1984). Manduca small PTTH appears to exert its ecdysiotropic effect via the same second messenger cascade as big PTTH: Ca2+ influx and Ca2+-dependent cAMP accumulation (Watson et al., 1993; Hayes et al., 1995) which stimulated Gilbert et al. (2002) to stipulate that small PTTH may be a proteolytic fragment of PTTH generated during its purification. This also suggests that Manduca small PTTH is not a bombyxin homolog (belonging to the insulin peptide family and discussed previously in Section 11.1.1.2) (as reviewed by Rybczyinski, 2005). Further research is necessary to fully characterize small PTTH and determine its exact physiological role (De Loof, 2008).

11.1.1.4

Ecdysteroid Feedback

A characteristic feature of endocrine systems is feedback regulation (Koolman, 1995). Both negative and positive feedback control of PG ecdysteroidogenesis exists. Moreover, hemolymph ecdysteroids may control PG ecdysteroidogenesis both directly and indirectly. Direct control (‘short loop feedback’) involves a direct action

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of 20E or/and E on the PGs. Indirect control (‘long loop feedback’) involves feedback regulation by 20E or/and E of brain NSCs, which in turn secrete(s) factors that act on the PGs. The existence of direct feedback has been doubtlessly demonstrated, but evidence for indirect feedback is far less solid. An extensive review on ecdysteroid feedback was recently written by Sakurai (2005). Direct Feedback Regulation of PG Activity The very first evidence for the phenomenon of feedback activation was presented by Williams in 1952. The first evidence for feedback inhibition was presented by Siew and Gilbert in 1971. Both research groups performed their pioneering work on the silkworm Hyalophora cecropia. The work of Williams (1952) can be summarized as follows. A series of H. cecropia pupae were first stabilized in diapause by removing their brains. Subsequently, a chain of eight brainless diapausing pupae was established in serial parabiosis, yielding, as it were, a single elongate organism possessing continuity of hemolymph and hypodermis. Implanting a single brain into the first animal of the chain caused the pupae to undergo adult development one by one downwards the chain. In a similar experiment, a pupa still containing brain and PGs, was grafted to a chain of four pupal abdomens (thus lacking brain and PGs). Adult development was initiated only in the pupa and the two anterior-most abdomens, not in abdomens #3 and #4. The latter experiment proves that the ecdysteroids produced by one (the anterior-most) animal in the chain do not reach the necessary concentration in the posterior part of the chain to cause development. Likewise, in the former experiment, the tropic factor released by the implanted brain in the anterior-most animal in the chain does not reach the necessary concentration in the posterior part of the chain to trigger the PGs sufficiently. Therefore, it was concluded that development of each successive pupa was due to an activation cascade of the PGs without decrement, i.e. the PGs of each member of the chain being triggered by the ecdysteroids arising in the preceding pupa. Siew and Gilbert (1971) confirmed this positive feedback relationship in Samia cynthia. Injection of 20E into diapausing Samia pupae profoundly stimulated PG activity. On the contrary, injection of the same amount of 20E into developing (postdiapause) H. cecropia pupae dramatically decreased PG activity. Thus, PGs can be either stimulated or inhibited by ecdysteroids. Sakurai and Williams (1989) observed that Manduca sexta PGs having relatively high activity in feeding larvae, wandering larvae, prepupae and developing pupae, were inhibited by 20E and E. By contrast, PGs with low activity from feeding larvae, diapausing pupae and day 1 postdiapause pupae were stimulated by 20E and E. They concluded that the occurrence of either stimulation or inhibition depends on both the state of activity of the PGs and the effective level of ecdysteroids in the hemolymph. Feedback inhibition of Manduca PGs was confirmed by Song and Gilbert (1998). In vitro incubation of glands from 5-days-old fifth instars with 20E for various exposure times, revealed that 20E not only suppressed basal ecdysteroidogenesis, but also – after a longer exposure time – suppressed the gland’s responsiveness to PTTH, thus interfering

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with PTTH-stimulated ecdysteroidogenesis as well. The theory of Sakurai and Williams (1989), i.e. PGs having very low to zero secretory activity are stimulated by 20E/E, is further supported by the results of Bodnaryk (1986). Injection of 20E into 1-day-old postdiapause pupae of Mamestra configurata stimulated ecdysteroidogenesis. Furthermore, Beydon and Lafont (1983) demonstrated the negative feedback of 20E and E on the PGs of 2- and 3-days-old developing pupae of Pieris brassicae, as in Manduca developing pupae. Indirect Feedback Regulation of PG Activity As previously mentioned, there is no firm evidence for the appearance of feedback control of PTTH production and/or release by ecdysteroids (Sakurai, 2005), nor for feedback control of any other identified PG tropic or static factor. However, since receptors for ecdysteroids do exist in brain NSCs (Bidmon and Koolman, 1989), it is tempting to speculate that these receptors form the structural basis for ecdysteroid feedback action (Koolman, 1995). Various studies have also shown that 20E clearly affects cell shape and stain intensity of brain NSCs (e.g. Steel, 1975; Agui and Hiruma, 1977), suggesting a regulatory role for 20E in the production and/or release of brain neurosecretory material. However, whether the observation of an increase in stain intensity is due to upregulation of production or downregulation of release or both, can not be distinguished, making it difficult to designate this (putative) feedback effect as either positive or negative. According to Sakurai (2005), the weakest point of these studies is the lack of knowledge of the products of the NSCs in question. It has not unequivocally been proven if the NSCs responding to 20E are truly the PTTH-secreting cells. For instance, Agui and Hiruma (1977) demonstrated that in vitro incubation of Mamestra brassicae diapausing pupal brains with 20E caused the brain to release a factor which in turn could stimulate ecdysteroidogenesis in diapausing pupal PGs, but the identity of the factor is not known. Based on these observations plus additional cell staining data (Agui and Hiruma, 1977), Sakurai (2005) hypothesized that 20E presumably stimulated PTTH release at a moderate concentration while suppressing its release at a high concentration. This hypothesis is reflected in how hemolymph ecdysteroid titers and PTTH titers relate to each other. In Bombyx and Manduca larvae, the PTTH titer peaks several hours before the ecdysteroid titer peaks. The ecdysteroid peak then is accompanied by a sharp decline in PTTH titer, supporting the idea of feedback inhibition of PTTH release by high 20E concentrations (Bollenbacher et al., 1987; Mizoguchi et al., 2001, 2002). The majority of the previously mentioned cell staining studies was focused on the four pairs of large median NSCs, since these cells were at that time believed to be the PTTH-secreting cells. However, according to more recent studies (Agui et al., 1979; Mizoguchi et al., 1990; Gilbert et al., 2000), the PTTH-cells are localized in the lateral region of the brain, not in the pars intercerebralis. Interestingly, the four pairs of large median NSCs have now been identified as the production site

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for ILPs (El-Salhy et al., 1983; Ishizaki and Suzuki, 1994; Iwami et al., 1996; Van de Velde et al., 2007). 20E thus might exert its indirect feedback action via ILPs. Biochemical and Molecular Aspects of Feedback Regulation Little is known about the exact target(s) for the feedback induced by 20E in the PG cells. Sakurai (2005) postulated that there might be at least three different targets: (1) the enzymes involved in the ecdysone biosynthetic pathway, (2) the PTTH signaling pathway, and (3) the system for translocation of intermediary ecdysteroid metabolites between critical subcellular domains (such as mitochondria and endoplasmic reticulum). The role of the PG ecdysteroid receptor complex (EcR complex) in feedback regulation has been reviewed by Gilbert et al. (2002). They analyzed the EcR complex constituents in the PGs of M. sexta and how these constituents respond to 20E. In Manduca PGs, two isoforms of ultraspiracle (USP) are present, each of which can exist in a phosphorylated or unphosphorylated state. The small isoform is a 47 kDa protein (49 kDa when phosphorylated), the large isoform a 52 kDa protein (54 kDa when phosphorylated). Most important, only the 47/49 kDa isoform heterodimerizes with the EcR to form the functional EcR complex (Song and Gilbert, 1998). During Manduca development in the fifth larval and pupal stage, the two isoforms display very distinct expression patterns. In brief, when hemolymph ecdysteroid levels are low, the 47/49 kDa is expressed minimally and the 52/54 kDa isoform maximally. Whereas when hemolymph ecdysteroid levels are high, the 47/49 kDa is expressed maximally and the 52/54 kDa isoform minimally. In other words, the higher the ecdysteroid titer rises, the more functional EcR complexes are formed, thus the more the PG is susceptible for feedback by 20E. The simultaneous appearance of ecdysteroid titer peaks and expression peaks of functional receptor components is not a coincidence. Song and Gilbert (1998) provided direct evidence that the ecdysteroid titer actually regulates USP expression. The first step in 20E-regulated PG ecdysteroidogenesis thus appears to be 20E-regulated expression of a specific USP isoform. Binding of 20E to the active EcR complex in the PGs then may start a series of transcriptional and/or translational events which ultimately result in an alteration of ecdysteroid production (Gilbert et al., 2002).

11.1.1.5

JH – Juvenile Hormone

Insect growth, molting, metamorphosis and reproduction are highly dependent on two hormones, namely juvenile hormone (JH) and ecdysteroids. For a review on the juvenile hormones the reader is referred to Hartfelder, 2000; Goodman and Granger, 2005. JH apparently has two distinct effects on the PGs: it plays a role in the regulation of ecdysteroid biosynthesis or secretion and is also reported to prevent

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precocious degeneration of PGs (Dai and Gilbert, 1998). Here we will discuss its (in)direct effects on ecdysteroidogenesis in the PG. A very thorough review on this subject was recently written by Sakurai (2005). Most of the available data were collected from penultimate and last larval stages of the holometabolous lepidopterans B. mori and M. sexta. The regulation of ecdysteroidogenesis in the PG by JH is highly dependent on the developmental stage of the insect and our discussion is thus split up accordingly. Action of JH on the PG in the Last Larval Instar of Lepidoptera During the last larval instar, two periods of JH influence have been discovered. The early last instar is characterized by a dramatic decline in hemolymph JH titer and a small release of PTTH that appears to trigger a small ecdysteroid release from the PG, called the commitment peak, which commits the animal to a metamorphic molt (Sakurai et al., 1989; Nimi and Sakurai, 1997; Mizoguchi et al., 2002). Several studies have shown that during this period, the decline in JH controls PTTH release. Inhibition of the JH decline due to topical application or injection of the hormone or an analogue (JHA) caused a delay in commitment due to the inability of the PGs to synthesize or secrete ecdysone. In vitro incubation of Manduca PGs from JH(A) treated larvae showed that they were still able to respond to PTTH. Transplantation and ligation experiments indicated that the target for the JH effect is the brain-retrocerebral complex (brain-corpora cardiaca-corpora allata, Br-CC-CA), though the exact regulatory mechanism is uncertain. An effect on PTTH release/content could not be excluded (Rountree and Bollenbacher, 1986; Watson and Bollenbacher, 1988; as reviewed by Sakurai, 2005). In B. mori, similar results were reported (Dedos and Fugo, 1996). Sakurai et al. (1989) reported that JH in early fifth instar Bombyx larvae not only inhibits the secretion of PTTH but also acts directly on the PGs via an inhibition of their sensitivity to PTTH. This direct effect was later confirmed by Gu et al. (1997). In contrast to the above described inhibitory effect of JH on PTTH release, a report by Mizoguchi (2001) demonstrates that JH elevates PTTH hemolymph levels in Bombyx early last instar larvae. Further verification is necessary. A more recent study suggests that feedback inhibition by 20E (from the fourth larval instar) causes the inactivity of early last instar PGs. This feedback inhibition was further hypothesized to be maintained by JH in the early last instar (Takaki and Sakurai, 2003). During the later development of this instar (after commitment to pupation), the effect of JH on the PGs appears to change in B. mori, M. sexta and M. brassicae as demonstrated by several studies (Hiruma, 1986; Dedos and Fugo, 1996; Dai and Gilbert, 1998). Now JH seems to be necessary again. At the time of commitment and possibly as a result thereof, a reversal in hormone action takes place. It was suggested that JH stimulates ecdysteroid synthesis by the PG and can also stimulate PTTH secretion from Br-CC-CA.

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Action of JH on the PG in the Earlier Instars of Lepidoptera During these stages, JH titers are high to maintain the larval state. No commitment peak is seen but ecdysteroid levels rise while the JH titer is still significant before the molt occurs (as reviewed by Goodman and Granger, 2005). When methoprene (a JHA) was applied to fourth instar M. sexta, an in vivo effect on the ecdysteroid titer could not be observed, nor an in vitro effect on ecdysteroid synthesis in the PGs. When allatectomy was performed, no ecdysteroid peak could be observed. But the normal situation was restored after application of methoprene (Lonard et al., 1996). Allatectomy of M. brassicae penultimate instar larvae abolishes the appearance of the ecdysteroid peak necessary for the larval-larval molt. The titer remained low for several days until wandering occurred precociously (Hiruma, 1986). It was suggested that JH acts via the production or release of PTTH. PTTH levels were demonstrated to be elevated after application of JH to B. mori fourth instar larvae (Mizoguchi, 2001). Caution should be exercised when interpreting these results because the dissection of the CA as a source of JH, also implies the removal of the neurohemal organ of PTTH.

11.1.1.6

Bom-PTSP – Bombyx mori Prothoracicostatic Peptide

In 1999, Hua et al. isolated a small prothoracicostatic peptide (Bom-PTSP) from larval brains of B. mori. Its primary structure is an amidated nonapeptide with the sequence AWQDLNSAWamide. Remarkably, Bom-PTSP has the same sequence as Mas-MIP I, Manduca sexta myoinhibitory peptide I, a myoinhibitory W2W9amide previously isolated from the ventral nerve cord of M. sexta (Blackburn et al., 1995). Bom-PTSP/Mas-MIP also shows high sequence homology with a group of type B allatostatins, Grb-AST-B, isolated from the brain of adult Gryllus bimaculatus crickets (Lorenz et al., 1995). Grb-AST-B allatostatins likewise are W2W9amides, i.e. nonapeptides that contain tryptophane residues at positions 2 and 9. Interestingly, with regard to ecdysteroidogenesis, these allatostatins have the potential to inhibit ecdysteroidogenesis in G. bimaculatus ovaries in vitro (Lorenz et al., 1997, 1998). For a more profound discussion about B-type allatostatins and ovarian ecdysteroidogenesis, see Section 11.2.1.2.2 in this chapter. In addition, Bom-PTSP/Mas-MIP shows considerable sequence similarity with the highly conserved N-terminus of vertebrate galanins (Tatemoto et al., 1983; Bersani et al., 1991; Hua et al., 1999). Hua et al. (1999) demonstrated that Bom-PTSP inhibits both basal and PTTH-stimulated ecdysteroidogenesis in fifth instar Bombyx PGs in vitro, in a dose-dependent manner. Dedos et al. (2001) further characterized the peptide extensively. Their results suggest that Bom-PTSP inhibits ecdysteroidogenesis only at specific developmental stages during the fifth instar, via a mechanism that likely involves the blocking of Ca2+ influx through voltage-sensitive Ca2+ channels in the PG cells. They also demonstrated that Bom-PTSP specifically inhibits ecdysone synthesis and not release. They however remarked that minimum and maximum effective concentrations (23 and 230 nM Bom-PTSP respectively)

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for in vitro inhibition of PG ecdysteroidogenesis are rather high, which puts the physiological relevance of this factor in vivo to the question.

11.1.1.7

BMS – Bommo-Myosuppressin

A second Bombyx prothoracicostatic factor, Bommo-myosuppressin (BMS), was purified from pupal brains and identified by Yamanaka et al., in 2005. BMS is an amidated decapeptide (EDVVHSFLRFamide) with the conserved structure of insect myosuppressins (X1DVX2HX3FLRFamide), which are members of a larger peptide family, FaRPs, FMRFamide-related peptides (Nichols, 2003). BMS is identical to Manduca FLRFamide I, that previously had been isolated from M. sexta (Lu et al., 2002). Yamanaka et al. (2005) proved that BMS dose-dependently inhibited both basal and PTTH-stimulated cAMP accumulation and ecdysteroidogenesis in fifth instar Bombyx PGs in vitro. Moreover, they compared the inhibitory potential of BMS and Bom-PTSP (cf. supra), and concluded that BMS suppressed the PGs with much lower effective concentrations then Bom-PTSP. They also demonstrated that BMS and Bom-PTSP must exert their prothoracicostatic activity through different receptors. The receptor for BMS (BMSR) could be identified and functionally characterized as a GPCR that binds its ligand BMS with high affinity and that shows a high expression in the PGs. The BMS expression pattern in the brain during the fifth instar suggests a biological role mainly in the first half of the instar, i.e. in the feeding period (Yamanaka et al., 2005).

11.1.1.8

Direct Innervation of the Ecdysial Glands

For various insect orders, anatomical evidence for PG innervation is present since decades (Scharrer, 1964; Hintze-Podufal, 1970; Giebultowicz and Denlinger, 1985). The role of innervation in regulation of PG ecdysteroidogenesis however has not been well studied. The typical method for examining PG regulation involves in vitro manipulation of explanted glands, thereby removing any regulatory innervation, thus systematically ignoring the role of innervation in favor of studies of circulating factors (Truman, 2006). Benedeczky et al. (1980) examined the PG innervation in last instar Galleria mellonella. They observed transfer of neurosecretory granules from innervating neurosecretory axons to intracellular spaces inside the PG cells. The granules could be identified as peptidergic (Knowles, 1965), suggesting that a peptidic factor, delivered via direct innervation to the PG, might regulate the PG function. In last instar Periplaneta americana, the nervous activity of the PG innervating ‘prothoracic gland nerve’ originating from the prothoracic ganglion, is in remarkable coincidence with the hemolymph ecdysteroid titer (Richter and Gersch, 1983). Additional research on the P. americana PG, as reviewed by Richter and Böhm in 1997, suggests a bimodality of regulation of PG ecdysteroidogenesis. It appears that during the period prior to the major peak of ecdysteroid production, PG ecdysteroidogenesis in P. americana

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is predominantly regulated by nervous activity of the prothoracic ganglion, whereas the major ecdysteroid peak appears to be controlled predominantly by PTTH and is not subject to nervous control. A detailed map of the innervation of the ring gland (RG) in D. melanogaster was presented by Siegmund and Korge (2001). They identified a pair of neurosecretory brain neurons that directly innervate the PG region of the Drosophila RG, and speculated that these particular neurons might regulate ecdysteroidogenesis. Whether these neurons effectively secrete a signal molecule affecting ecdysteroidogenesis is presently unknown. Interestingly, Rybczynski (2005) points the possibility that a soluble PTTH, released into the hemolymph, might be completely lacking in Drosophila, and that its function might be taken over by another prothoracicotropic brain factor delivered directly to the PG via innervation and not through the circulation. The recent work by Yamanaka et al. (2006) is an important breakthrough in revealing the molecular basis of PG regulation by innervation. Next to BMS (cf. supra; Yamanaka et al., 2005), they purified and identified four additional Bombyx FaRPs (SAIDRSMIRFamide, SASFVRFamide, DPSFIRFamide and ARNHFIRLamide) which all suppress ecdysteroidogenesis in Bombyx PGs in vitro. Opposite to BMS, which was proposed to act through the circulation, these novel FaRPs are delivered to the PG’s surface via direct innervation. All four peptides are encoded by the same gene, ‘BRFa’. BRFa is predominantly expressed in the NSCs of the thoracic ganglia. Electrophysiological investigation of the BRFa neuron activity throughout the last instar, demonstrated the increased firing activity of BRFa neurons in periods with low PG activity. Intriguingly, the BRFa peptides act via the same receptor as BMS, BMSR. Since BMSR was shown to be expressed in several other tissues besides the PGs, it was suggested that Bombyx uses BMSR-mediated signaling for both general and specific inactivation, by means of circulating BMS and BRFa peptides via direct innervation respectively (Truman, 2006; Yamanaka et al., 2006).

11.1.1.9

TMOF – Trypsin Modulating Oostatic Factor

Aea-TMOF – Aedes Aegypti Trypsin Modulating Oostatic Factor Trypsin Modulating Oostatic Factor was originally discovered by Borovsky et al. (1990). It was isolated from late vitellogenic ovaries of the yellow fever mosquito Aedes aegypti. This decapeptide (YDPAPPPPPP) was named TMOF because – after secretion into the hemolymph and binding to specific receptors in the midgut – it is able to modulate (inhibit) trypsin biosynthesis by exerting a translational control on trypsin mRNA (Borovsky et al., 1994; Borovsky, 2003). This leaves the blood meal in the gut undigested. As a result, insufficient amounts of free amino acids are available for bulk synthesis of vitellogenin in the fat body. This causes the oostatic effect. Similar effects on the trypsin synthesis in the gut were reported when Aea-TMOF was administered to larvae of the tobacco budworm, Heliothis virescens and the citrus weevil, Diaprepes abbreviatus (Yan et al., 1999; Nauen et al., 2001).

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Though no ecdysiostatic effect was ever demonstrated in Aedes itself, Aea-TMOF can inhibit the ecdysteroid production in the PGs of the caterpillar, Lymantria dispar (Gelman and Borovsky, 2000). Neb-TMOF – Neobellieria Bullata Trypsin Modulating Oostatic Factor During a search for folliculostatic peptides another TMOF (NPTNLH) was found in the ovaries of the grey fleshfly, Neobellieria bullata (Bylemans et al., 1994). Its precursor is a 75 kDa matrix protein that is abundantly present in the oocytes (De Loof et al., 1995b; Bylemans et al., 1996). Although its primary structure is completely different from Aea-TMOF, Neb-TMOF also inhibits de novo trypsin biosynthesis and hence has a similar oostatic effect. But independently of these activities, Neb-TMOF also inhibits the in vitro and in vivo ecdysone biosynthesis in the larval ring glands of N. bullata and of the blowfly Calliphora vicina with a remarkable efficacy (EC50 of 5.10−9 M) and causes a delay in pupariation (Hua et al., 1994; Bylemans et al., 1995; De Loof et al., 1995a, b). Hua and Koolman (1995) showed that this activity is associated with an elevated cAMP level in the ring glands. Using a bioassay for ecdysiostatic activity, they found evidence that a TMOF-like peptide could also be synthesized by blowfly larvae (Hua and Koolman, 1995) and is located in the cerebral complex (unpublished immunohistochemical experiments). The above described pleiotropic effects of Neb-TMOF – inhibition of trypsin synthesis and ecdysteroid synthesis – are not related because injection of 20E in female flies did not influence trypsin biosynthesis (Bylemans et al., 1995). Apparently Neb-TMOF has at least two different target tissues – the digestive tract in adults and the ring glands in larvae – that may have a different receptor for TMOF (Hua and Koolman, 1995). NebTMOF does not affect ecdysone synthesis in the ovaries (Bylemans et al., 1995). It was concluded that an oostatic effect in Neobellieria can be realised in two different ways, the inhibition of protein meal digestion and the interference with ecdysteroid biosynthesis, which is necessary for yolk polypeptide synthesis (Huybrechts and De Loof, 1977; De Loof et al., 1995a, b). Although Neb-TMOF efficiently inhibits steroidogenesis in Neobellieria and Calliphora, it does not have an effect on the ring glands of Drosophila melanogaster, which shows its species specificity (Hua and Koolman, 1995). Its in vivo gonadostatic action was nevertheless demonstrated in the non-dipteran mealworm, Tenebrio molitor. The results of this study suggest that this peptide can exert its effect by antagonising JH, which plays an important gonadotropic role in T. molitor (Wasielewski and Rosin´ski, 2007). Neb-TMOF was the first peptide identified with this very potent ecdysiostatic activity and was therefore also designated as prothoracicostatic hormone (Neb-PTSH) (Bylemans et al., 1995). TMOF in Drosophila melanogaster Liu et al. (2006) recently identified TMOF like peptides and their coding genes in silico in the fruitfly D. melanogaster, yet functional confirmation still is required.

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TMOF in Locusta migratoria and Schistocerca gregaria In preliminary experiments, Bylemans (1994) showed the presence of a TMOF-like factor in an extract of 300 vitellogenic ovaries from the migratory locust, Locusta migratoria. This was named a trypsin and/or chymotrypsin modulating oostatic factor (C/TMOF) because both trypsin and chymotrypsin contribute to the digestion process. Locusta C/TMOF was partially purified and exerts ecdysone biosynthesis inhibiting activity in C. vicina (Bylemans, 1994). Purification of Schistocerca C/TMOF was tried, but was only partially successful. Further characterization is necessary (Janssen, 1997). 11.1.1.10

PG Autocrine Factors

Very recently, Gu (2007) reported on the existence of an as yet chemically unidentified ‘autocrine factor’ involved in the activation of ecdysteroidogenesis in the PGs of B. mori. In vitro incubation experiments showed that lowering the volume of incubation medium or culturing glands in group – instead of individually – caused a significant dose-dependent increase in ecdysteroid secretion by the PG from last instar larvae. A higher ecdysteroid titer was also observed when glands were incubated in PG-conditioned medium. Moreover, injection of the concentrated putative autocrine factor into last instar larvae resulted in a dose-dependent higher ecdysteroid titer in the hemolymph as compared to the one after saline injection, proving the in vivo biological function of the factor. This autocrine factor is heat stable, and has a molecular weight estimated between 1,000 and 3,000 Da. In similar incubation and injection conditions, an increase in the number of DNA-synthesizing cells in the PGs of B. mori was also observed (Gu, 2006). The fact that in B. mori dramatic increases in DNA synthesis precede ecdysteroidogenesis (Gu and Chow, 2005), and that the putative autocrine factor in both experiments has similar characteristics with respect to molecular weight, heat stability, resistance to treatment with trypsin and proteinase K and lack of retention on a C18 column, suggests that it is the same factor. Such a factor was also demonstrated in another lepidopteran, Spodoptera littoralis (Gu, 2006), and in two orthopterans, S. gregaria and L. migratoria (Vandersmissen et al., 2007). Autocrine (or paracrine) activation of ecdysteroidogenesis within the PG may thus be a general mechanism present in insects. 11.1.1.11

Ecdysiotropic Gut Peptides

In 1991, Gelman et al. claimed that the insect gut is a source of ecdysiotropic peptides. Hindgut extracts from Ostrinia nubilalis, Lymantria dispar (Gelman et al., 1991) and Manduca sexta (Gelman and Beckage, 1995) last instar larvae dose-dependently stimulated ecdysteroidogenesis in L. dispar fifth instar PGs in vitro. The ecdysiotropic potential of the Ostrinia and Lymantria extracts was also confirmed in vivo (Gelman et al., 1991). Injection of hindgut extracts into Lymantria larvae that were head-ligated before the release of PTTH, resulted in a pupal molt. The authors however only partially purified the prothoracicotropic factor from the extracts.

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They determined the molecular weight range of the factors, not the actual structure. The majority of the prothoracicotropic activity was due to the presence of low molecular weight ecdysiotropins (500–1,500 Da) (Gelman et al., 1993; Gelman and Beckage, 1995). In addition, Gelman and Bell (1995) demonstrated the presence of low molecular weight ecdysiotropins in the hemolymph of last instar O. nubilalis and L. dispar. Whether the source of these hemolymph ecdysiotropins is the hindgut, is presently unknown. It should be mentioned that, in contrast with the clear ecdysiotropic effect of Ostrinia hindgut and hemolymph extracts on Lymantria PGs, the effect of Ostrinia extracts on Ostrinia PGs was rather sporadic (Gelman and Bell, 1995). The definite in vivo role of these factors – if any – thus remains open to speculation. Furthermore, since two additional active fractions with molecular weight in the range of 3,500–4,500 Da and molecular weight of approx. 30,000 Da were derived from the Manduca hindgut extract, Gelman and Beckage (1995) postulated that this could be due to the transport of brain PTTH and small PTTH down the ventral nerve cord to a release site in the proctodaeal nerve. If this hypothesis is true, i.e. transport of brain factors via the central nervous system to the hindgut – irrespective of whether or not the mentioned molecules are truly PTTH and small PTTH – this theory could be extended to the low molecular weight ecdysiotropins as well. If so, the question rises if we can still consider these peptides as true gut peptides.

11.1.2

Factors Acting on the Larval Testes: Lymantria Testis Ecdysiotropin (LTE) and Others

It is now well accepted that also the testes, or more specifically the testis sheaths, can function as ecdysteroid source. In adult males, where the PGs are degenerated, the testes even may represent the sole source of ecdysteroids. Testes ecdysteroidogenesis during adult life and regulation thereof will be discussed further in this chapter (Section 11.2.2). However, the testes appear not only to function as ecdysteroid source during adult life, but also during late larval and pupal development. Ecdysteroidogenesis by larval testes has been examined predominantly in lepidopteran genera (Loeb et al., 1982, 1988; Shimizu et al., 1985, 1989; Gelman et al., 1989; Jarvis et al., 1994). In immature stages, the PGs are the major source of ecdysteroids, and the amount of ecdysteroids produced by the larval testes is only a small fraction of that produced by the PGs at that time. Therefore, a rather paracrine or autocrine role is suggested for testes ecdysteroids in larvae (Gilbert et al., 2002). Testes ecdysteroids are believed to play a role in the maturation of male genital tissues and in (early) spermatogenesis (as reviewed by Hagedorn, 1985). Research on factors regulating testes ecdysteroidogenesis is almost completely covered by the pioneering work of Loeb and co-workers (reviewed by Loeb et al., 2001). In 1987, Loeb et al. presented the first evidence for the existence of ecdysiotropic factors that affect the testes. Larval and pupal brain extracts from Heliothis virescens and Lymantria dispar were capable of boosting constitutive testes ecdysteroidogenesis in vitro at the end of the last larval instar, and were capable of inducing de novo synthesis

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in younger testes (Loeb et al., 1987, 1988). Brain extracts from H. virescens not only stimulated in vitro ecdysteroidogenesis in its own testes, but also in the testes of L. dispar and vice versa. However, it was not until 1997 that Wagner et al. fully determined the primary structure of an ecdysiotropic factor isolated from the brain of L. dispar pupae. This factor, designated ‘Lymantria testis ecdysiotropin’ (LTE), is a 21 amino acid peptide of sequence ISDFDEYEPLNDADNNEVLDF. Lymantria testis ecdysiotropin is very active on early last instar testes, and is biphasic in activity. Maximal ecdysiotropic activity was observed at 10−15 M and 10−10 M, and minimal activity at 10−13 M. Two concentrations of maximal activity, separated by a minimum, suggest the existence of (at least) two different types of LTE receptors in the testis sheath, the target organ for LTE (Wagner et al., 1997; Meola et al., 1998). Loeb et al. (1993, 1994) demonstrated that the major signaling pathway for TE involves Gi protein, inositol trisphosphate, diacylglycerol and protein kinase C. LTE displays no significant sequence homology with PTTH. LTE is incapable of stimulating ecdysteroidogenesis in the PGs of L. dispar and H. virescens, and PTTH likewise does not stimulate testes ecdysteroidogenesis in these two insects. LTE was isolated from the major active fraction obtained by HPLC separation of a L. dispar pupal brain extract. The other active fractions from this separation were later analyzed by Loeb et al. (1997). They purified and sequenced nine additional peptidic factors with testis ecdysiotropic activity. Five of them were analogs of LTE and are therefore considered to form an ‘LTE family’. The other four had no significant homology with LTE nor with each other. Surprisingly, Vafopoulou and Steel found LTE-like material in the larval testis sheath of an unrelated hemipteran species, R. prolixus (Vafopoulou and Steel, personal communication in Loeb et al., 2001). In a more recent publication (Vafopoulou and Steel, 2005), they immunohistochemically localized large amounts of LTEimmunoreactive material in brain NSCs, suboesophageal ganglion and testis sheath epithelium of unfed fifth instar Rhodnius larvae. Feeding of larvae resulted in rapid and massive release of this material. Moreover, several LTE-immunoreactive peptides were detected in larval brain extracts by western blot analysis. Both larval brain extracts and LTE were capable of stimulating ecdysteroidogenesis by adult testes in vitro. Unfortunately, the possible ecdysiotropic effect on larval testes was not examined in this publication. The presence of peptides related to LTE in the Rhodnius brain, however, does not necessarily explain the testis ecdysiotropic activity of the Rhodnius brain. Other, non-LTE ecdysiotropic peptides might as well be responsible for the tropic action. The ecdysiotropic effect of Rhodnius brain extracts and LTE on adult testes will be discussed in Section 11.2.2 in this chapter.

11.1.3

Alternative Ecdysteroid Sources: Larval Oenocytes and Epidermis

There exists a wealth of reports demonstrating larval ecdysteroid production in the absence of the classical source of ecdysteroids, the PGs. Significant endogenous ecdysteroid concentrations have been measured in isolated larval abdomens

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or larvae surgically deprived of PGs, and such specimens have been shown to synthesize ecdysteroids from radiolabelled cholesterol (Nakanishi et al., 1972; Hsiao et al., 1975; Studinger and Willig, 1975; Gersch and Eibisch, 1977; Rees, 1985; Delbecque et al., 1990). However, the exact nature of the ecdysteroidogenic source within such preparations is not clarified. An ovarian nature can be excluded, since ovarian ecdysteroidogenesis generally does not start before the (pharate) adult stage. Larval testes, on the contrary, have been described as a valuable ecdysteroid source (see Section 11.1.2), but since both male and female species in abovementioned experiments synthesize equal amounts of ecdysteroids, a unique contribution from the testes can be disregarded. Several authors claim that oenocytes or/and the epidermis (also designated as “hypodermis”) are responsible for this ecdysteroid synthesis outside the PGs.

11.1.3.1

Oenocytes

Oenocytes are large, differentiated epidermis cells with special secretory function. They either remain within the epidermis or migrate into the haemocoel, where they closely associate with the fat body (Redfern, 1989; Delbecque et al., 1990). Oenocytes are involved in the synthesis of cuticular lipids (Wigglesworth, 1970; Diehl, 1975; Romer, 1980). A variety of other, though less proven, functions have been attributed to oenocytes (Philogène and McFarlane, 1967; Romer, 1980; Ma and Philogène, 1985). They have been proposed as a possible alternative ecdysteroid source, mainly because of their ultrastructural resemblance with vertebrate steroid secreting cells, i.e. the presence of smooth endoplasmic reticulum (Christensen, 1965; Bjersing, 1967; Locke, 1969; Romer, 1974; Dorn and Romer, 1976). Moreover, oenocytes share structural characteristics with PG cells, i.e. the presence of plasma-membrane invaginations (Sedlak, 1985). Apart from the ultrastructural hypothesis, real evidence came from Romer et al. (1974). They actually proved that isolated abdominal oenocytes from Tenebrio molitor larvae can synthesize ecdysteroids de novo from cholesterol. The epidermis in T. molitor appears to be an active ecdysteroid source (cf. Section 11.1.3.2; Delachambre et al., 1984; Delbecque et al., 1986). Since oenocytes have an ectodermal origin, their ecdysteroidogenic capacity is not completely surprising. However, the question remains if oenocytes participate effectively in ecdysteroidogenesis also in vivo, or do so only under experimental stress (Rees, 1985). The biological target of the ecdysteroids produced by oenocytes – if any – is unknown. To our knowledge, no reports have been made on factors regulating ecdysteroidogenesis by oenocytes so far.

11.1.3.2

Epidermis

Porcheron et al. (1984, 1988) observed in vitro ecdysteroid production by cultured larval epidermis of L. migratoria. Meister et al. (1985) demonstrated the ability of Locusta cultured larval epidermis to convert the radiolabelled precursor 2,22,25-trideoxyecdysone into ecdysone – i.e. to perform the

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last three hydroxylation steps in ecdysone biosynthesis – though Locusta PGs perform this conversion 10–100 times more efficiently. Romer (1987) demonstrated ecdysteroid production by Bombyx larval epidermis explants. In T. molitor, the epidermis has been proposed as a primary ecdysteroid source by Delachambre et al. (1984) and Delbecque et al. (1986, 1988, 1990). Most interestingly, the JHA methoprene inhibits in vitro ecdysteroid production by larval and pupal epidermis of Tenebrio (Delbecque et al., 1990), pointing at JH as a direct negative factor regulating epidermal ecdysteroidogenesis in larvae. Moreover, Soltani et al. (1987, 1989) evidenced a change in ecdysteroid source at the end of larval development in Tenebrio. After treatment of Tenebrio larvae and pupae with diflubenzuron, an insect growth regulator that targets the epidermis, they observed a significant reduction of the hemolymph ecdysteroid concentration at the end of the last larval stage, and even a complete inhibition of the ecdysteroid peak in the pupal stage. Earlier larval stages were not affected. Apparently, at least in T. molitor, the epidermis gains more importance as ecdysteroid source during pupal development, when the PGs have degenerated, whereas during larval development, epidermal ecdysteroid secretion is likely inhibited by JH and PG ecdysteroidogenesis determines the overall ecdysteroid titer. In addition, several epidermal cell lines established from larval imaginal discs were shown to have the capacity to synthesize ecdysteroids in vitro (Mesnier et al., 2000). The fact that the epidermis is both source and target of ecdysteroids is intriguing, and suggests an autocrine function for epidermal ecdysteroids. Clearly, the study of additional ecdysteroid sources, such as the epidermis, and the biological importance thereof deserves further attention. Regarding regulating factors, it would be of interest to evaluate the effect of the earlier described PG regulating factors (Section 11.1.1) on epidermal ecdysteroidogenesis.

11.2

Factors Regulating Ecdysteroidogenesis in Adult Insects

11.2.1

Factors Acting on the Ovaries

11.2.1.1

Ecdysiotropins

11.2.1.1.1

JH – Juvenile Hormone

Both classic insect hormones, JH and 20E, play an important role in the physiology of insects, especially in development and reproduction. In many cases they are indissolubly linked in their functioning, since each of these hormones may be involved in regulating the biosynthesis of the other. Allatectomized G. bimaculatus last instar larvae were shown to have a reduced release of ovarian ecdysteroids, indicating that

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JH is involved in the control of their biosynthesis (Hoffmann and Gerstenlauer, 1997). In decapitated adult female mosquitoes, JH was demonstrated to restore the impaired ovarian ecdysteroidogenesis (Birnbaum et al., 1984). In addition, JH-I or analogues appeared to induce the responsiveness of mosquito ovaries to brain extracts that were shown to stimulate ovarian steroidogenesis (Shapiro and Hagedorn, 2004). Fruit flies defective in their insulin signaling pathway were shown to display both decreased JH synthesis and impaired ovarian ecdysteroidogenesis (cf. infra) (Tatar et al., 2001; Tu et al., 2002, 2005). Moreover, application of methoprene (a JH analogue) was shown to restore these defects, suggesting that they resulted from a deficiency in JH (Tatar et al., 2001). Hitherto, no information is available about positive or negative feedback mechanisms of ecdysteroids on ovarian ecdysteroidogenesis. 11.2.1.1.2

OMP – Ovary Maturating Parsin

The ovary maturating parsin (OMP) was the first insect peptide gonadotropin that was discovered. It was initially identified from CC of the migratory locust, L. migratoria, as a factor involved in ovarian maturation (Girardie et al., 1991). ‘Parsins’ are neurosecretory peptides (or small proteins) that are produced in the pars intercerebralis. OMP has hitherto only been identified in one other insect, i.e. the desert locust, S. gregaria, a species closely related to the migratory locust (Girardie et al., 1998). Moreover, studies by Girardie et al. (1996) in both L. migratoria and S. gregaria strongly suggested that OMP exerts its maturating effect through the induction of ovarian ecdysteroidogenesis. First, injections of 20E in adult female L. migratoria appeared to have the same effect as injections of the L. migratoria OMP (Lom-OMP), i.e. a precocious appearance of vitellogenins in the hemolymph, as well as a precocious maturation of the ovaries. Second, injection of 20E appeared to compensate for the effects caused by the inactivation of Lom-OMP. Third, injection of Lom-OMP in ovariectomized females did not induce the production of vitellogenin (Girardie and Girardie, 1996). Injections of OMP in S. gregaria appeared to induce a precocious occurrence of 20E in the hemolymph. The induced ecdysteroidogenesis was suggested to be involved in oocyte growth and consequent ovarian maturation (Girardie et al., 1998). 11.2.1.1.3

OEH – Ovary Ecdysteroidogenic Hormone

The ovary ecdysteroidogenic hormone (OEH) was first characterized from the yellow fever mosquito, A. aegypti, and was one of the first invertebrate gonadotropins to be discovered (Brown et al., 1998). Later, it was also identified in the African malaria mosquito, Anopheles gambiae (Riehle et al., 2002). In response to a blood meal, OEH is released from median NSCs in the brain (Lea, 1967, 1972). Subsequently, the hormone is transported to the CC, where it is stored until its release in the hemolymph. Once secreted into the hemolymph, it targets the ovaries, which are consequently stimulated to secrete ecdysteroids (Matsumoto et al., 1989). In their turn, ecdysteroids are triggering yolk protein synthesis in the fat

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body (Raikhel, 1992; Klowden, 1997). Bioinformatics searches in protein databases revealed the sequence similarity of OEH with neuroparsin A, a peptide that was previously identified from pars intercerebralis and CC extracts of L. migratoria (Girardie et al., 1989; Brown et al., 1998). Interestingly, neuroparsins also display sequence similarity with the N-terminal part of vertebrate insulin-like growth factor binding proteins (IGFBPs) (Janssen et al., 2001; Claeys et al., 2003), the protein module that possesses the hormone binding capacity. Neuroparsin-like peptides have been identified in various other arthropods (Claeys et al., 2003; Badisco et al., 2007). Although it is not known whether neuroparsin family members are directly involved in the regulation of ecdysteroidogenesis in species other than mosquitoes, this would perhaps not be surprising, since, even in vertebrates, some IGFBPs have been reported as important regulators of gonadal steroidogenesis (Monget et al., 1996; Yoshimura, 1998; Webb et al., 1999; Arraztoa et al., 2002). In addition, since they resemble the hormone binding module of IGFBPs, it does not seem unlikely that these peptides may be capable of binding endogenous ILPs. In mosquitoes, both OEH and vertebrate insulin can stimulate ecdysteroidogenesis in ovaries (Graf et al., 1997; Riehle and Brown, 1999), suggesting a parallel role for these peptides in the control of this process. Moreover, it has been demonstrated that ovarian ecdysteroidogenesis is probably accomplished by the insulin signaling pathway (Riehle and Brown, 2002). 11.2.1.1.4

ILPs – Insulin-Like Peptides

Various studies already showed that both exogenous and endogenous ILPs (most of which can also be considered as parsins, based on their major site of production in insects) can stimulate ovarian ecdysteroidogenesis in insects. In A. aegypti, both porcine and bovine insulin were shown to trigger ecdysteroid production in ovaries (Graf et al., 1997; Riehle and Brown, 1999). In the black blowfly, Phormia regina, ecdysteroid production was stimulated by bombyxin, an ILP from the silk moth, B. mori (Manière et al., 2004). Moreover, extracts of P. regina median NSCs, which are believed to be an important source of endogenous ILPs, stimulated ecdysteroid production in ovaries of this organism (Manière et al., 2004). However, one can not exclude that some effects of insulin-mediated stimulation of ecdysteroidogenesis are indirect and may for example be exerted through stimulation of JH synthesis (cf. supra). Most ILPs appear to function through a receptor tyrosine kinase (RTK). Upon binding of the ILP to the RTK, specific tyrosine residues of the latter are phosphorylated and recruit and activate the insulin receptor substrate (IRS), an intermediate adaptor protein which subsequently can initiate two signaling pathways (Yenush et al., 1996; White, 1998), i.e. the mitogen-activated protein kinase (MAPK) pathway and the phosphatidylinositol 3-OH kinase/protein kinase B (PI3K/PKB) pathway. It appears that insulin-stimulated ovarian ecdysteroidogenesis is mediated by the PI3K/PKB pathway. In D. melanogaster, mutations in the insulin receptor resulted in impaired ovarian ecdysteroidogenesis (Tu et al., 2002). In A. aegypti,

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an insulin receptor was localized in the follicle cells surrounding the oocytes (Helbling and Graf, 1998). These follicle cells appear to be the primary source of ecdysteroids in female locusts (Kappler et al., 1986), and probably also in most other adult female insects. It was shown that, in both A. aegypti and P. regina, stimulation of ecdysteroid production by insulin-like peptides is mediated by activation of an RTK, most likely the insulin receptor. Moreover, in both species, PI3K and PKB, components of the insulin/IGF (PI3K/PKB) pathway, were activated in this process. The MAPK pathway, on the other hand, did not appear to be involved in insulin-mediated ecdysteroid production (Riehle and Brown, 1999; Manière et al., 2004). In P. regina, no significant increase in ecdysteroid production was noticed upon administration of insulin during the first 2 days following adult ecdysis, but a progressive increase was noticed during the next days that stagnated around 5–6 days after the final molt (Manière et al., 2004). This is in accordance with studies in mosquitoes which showed a progressive expression of the insulin receptor in follicle cells during the previtellogenic stage (Riehle and Brown, 2002). It is likely that insulin receptor expression is still very low immediately after eclosion, since the female blowfly reproductive system is not yet fully mature at that moment. Interestingly, insulin-like peptides and IGFs were also shown to modulate ovarian steroidogenesis in mammals. These hormones are suggested to potentiate the steroidogenic response of ovaries to gonadotropins released from the pituitary (Poretsky et al., 1999). Although they do not appear to have a direct steroidogenic effect in mammals, there seems to exist an evolutionary conserved relationship between ILPs and (ovarian) steroidogenesis.

11.2.1.2 11.2.1.2.1

Ecdysiostatins Ovarian Ecdysiostatin

In higher Diptera, and perhaps in other insects, vitellogenesis is stimulated by ovarian ecdysteroids (Raikhel, 1992; Klowden, 1997), hence the maintenance of cyclic egg development includes the complementary activity of both ecdysiotropins and ecdysiostatins. Ovarian ecdysiostatin (OES) has been purified from the domestic fly, Musca domestica, as a peptide that inhibits secretion of ovarian ecdysteroids (Adams and Li, 1998). In the same organism, an ecdysteroidogenic activity was shown from head extracts (Adams et al., 1997). This was supposed to be closely related to the A. aegypti egg development neurosecretory hormone (EDNH), which has later been termed OEH (cf. Section 11.2.1.1.3). For convenience (and according to the terminology of Adams and Li, 1998), this M. domestica ecdysteroidogenin will further be referred to as OEH. Musca domestica OES hence has an activity opposite to OEH, although both factors do not seem to interfere with each other. This suggests that OES acts directly on the ovaries, independent of OEH. Moreover, ecdysteroidogenesis can be restored upon removal of OES. Furthermore, the inhibiting activity of OES in the fly is noticed during late vitellogenesis and

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post-vitellogenesis (Adams and Li, 1998), whereas OEH activity is only noticed until mid-vitellogenesis (Adams and Filipi, 1983; Adams et al., 1985). This suggests that both OES and OEH are most likely involved in the maintenance of cyclic egg development. Remarkably, OES activity was demonstrated in abdominal extracts of both male and female flies, suggesting that it might have a role in regulating ecdysteroid levels in males as well. Although it appears that OES is produced in the abdomen, the exact site of production remains hitherto unknown (Adams and Li, 1998). Regarding molecular mass and behavior in chromatographic studies, OES appears to be unrelated to any other known insect oostatic factor. 11.2.1.2.2

Ast-B or MIPs – B-type Allatostatins/Myoinhibiting Peptides

Although allatostatins were initially identified as factors inhibiting JH synthesis, they appear to have pleiotropic functions. A class of four allatostatins, which had been termed B-allatostatins (Grb-AST B1-4), was isolated from female brain extracts of the cricket G. bimaculatus (Lorenz et al., 1995). They were structurally unrelated to any other known allatostatin, but remarkably, showed sequence similarity to myoinhibiting peptides (MIPs) from the locust, L. migratoria, and the moth, M. sexta. In addition to JH inhibition, these allatostatins were also shown to inhibit ecdysteroid synthesis in vitro in G. bimaculatus ovaries, Grb-AST B1 and B2 being the most effective inhibitors (Lorenz et al., 1997, 1998). Surprisingly, Grb-AST B1 was shown to have the opposite effect in the cockroach, Blaptica dubia, where it stimulated ovarian ecdysteroidogenesis (Lorenz et al., 2004). Since ecdysteroids can influence JH synthesis and vice versa (Granger et al., 1987; Wennauer et al., 1989; Hoffmann and Gerstenlauer, 1997), some of the in vivo effects of allatostatins on ecdysteroid synthesis may result from indirect effects involving changes in JH synthesis.

11.2.1.3

Signaling Pathways

As described in the above paragraphs, the regulation of ovarian ecdysteroidogenesis involves a range of different factors, which exert their effects through various pathways. Studies in P. regina demonstrated that there are at least two different brain factors involved in the regulation of ovarian ecdysteroidogenesis, each acting through a different pathway, cAMP dependent or independent. Interestingly, cAMP dependent factors appear to be involved in early vitellogenesis, whereas cAMP independent factors are probably involved during a longer period of the vitellogenic cycle. Moreover, cAMP dependent factors appear to originate from brain areas different from the pars intercerebralis, whereas pars intercerebralis extracts show activity through a pathway independent of cAMP (Manière et al., 2000). Although the pertinent P. regina factors are hitherto unknown, ovarian ecdysteroid synthesis through a cAMP independent signaling system has been demonstrated in other insects. ILPs, produced in the pars intercerebralis, act through the insulin signaling pathway, which does not

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involve cAMP as a second messenger. OEH in mosquitoes was also shown to act through the insulin signaling pathway (Riehle and Brown, 2002). However, it can be expected that several factors involved in the control of ecdysteroidogenesis still have to be identified. Furthermore, the mode of action of a lot of these factors probably remains unknown. In summary, the preceding overview of factors that are involved in ovarian ecdysteroidogenesis is most probably incomplete. It is likely that several factors, other than the ones mentioned above, are in one way or another involved in this process. For instance, a short neuropeptide F (sNPF) like peptide was also shown to have an effect on ovarian development in locusts. Although the exact mode of action is still unknown, this effect was suggested to be exerted through a stimulation of ecdysteroid synthesis or JH synthesis or a combination of both (Cerstiaens et al., 1999).

11.2.2

Factors Acting on the Testes

The factors triggering ecdysteroid biosynthesis in the testes of adult insects are not well characterized, but there is a report on the blood-sucking bug, Rhodnius prolixus (Vafopoulou and Steel, 2005). In R. prolixus, the level of ecdysteroids in adult testes increases in vivo after a blood meal. Testes of unfed adults do not produce significant amounts of ecdysteroids in vitro. On the other hand, higher ecdysteroid levels can be induced when these testes are incubated with larval or adult brain extracts. Furthermore, LTE, which is well known as a testis ecdysiotropin in the gypsy moth, Lymantria dispar, also stimulates the ecdysteroid level in testes of unfed adults in vitro. LTE has no ecdysteroidogenic activity on Rhodnius prothoracic glands, indicating that the effect of LTE is confined to the testes and is different from PTTH. By means of immunohistochemistry using an anti-LTE antibody, LTE-like material has been localized in brain, suboesophageal ganglion and testis sheath of unfed adult bugs, but not in fed ones. Therefore, a spatially and developmentally regulated LTE-like substance may act as an ecdysiotropin for adult testes, although the details of this regulation remain to be uncovered. Acknowledgements This work was supported by the IWT-Flanders and FWO-Vlaanderen (Belgium).

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Rybczynski R (2005) Prothoracicotropic hormone. In: Gilbert LI, Iatrou K, Gill S. (ed) Comprehensive Molecular Insect Science. Elsevier, Oxford, vol. 3, pp 61–123 Rybczynski R, Gilbert LI (2006) Protein kinase C modulates ecdysteroidogenesis in the prothoracic gland of the tobacco hornworm, Manduca sexta. Mol Cell Endocrinol 251:78–87 Sakurai S (2005) Feedback regulation of prothoracic gland activity. In: Gilbert LI, Iatrou K, Gill S. (ed) Comprehensive Molecular Insect Science. Elsevier, Oxford, vol. 3, pp 409–431 Sakurai S, Williams CM (1989) Short-loop negative and positive feedback on ecdysone secretion by prothoracic gland in the tobacco hornworm, Manduca sexta. Gen Comp Endocrinol 75:204–216 Sakurai S, Okuda M, Ohtaki T (1989) Juvenile hormone inhibits ecdysone secretion and responsiveness to prothoracicotropic hormone in prothoracic glands of Bombyx mori. Gen Comp Endocrinol 75:222–230 Scharrer B (1964) The fine structure of Blattarian prothoracic glands. Z Zellforsch Mikrosk Anat 64:301–326 Sedlak BJ (1985) Structure of endocrine glands. In: Kerkut GA, Gilbert LI (ed) Comprehensive Insect Physiology, Biochemistry and Pharmacology. Pergamon Press, Oxford, vol. 7, pp 25–60 Sevala VM, Sevala VL, Loughton BG, Davey KG (1992) Insulin-like immunoreactivity and molting in Rhodnius prolixus. Gen Comp Endocrinol 86:231–238 Sewer MB, Waterman MR (2003) ACTH modulation of transcription factors responsible for steroid hydroxylase gene expression in the adrenal cortex. Microsc Res Tech 61:300–307 Shapiro JP, Hagedorn HH (2004) Juvenile hormone and the development of ovarian responsiveness to a brain hormone in the mosquito, Aedes aegypti. Gen Comp Endocrinol 46:176–183 Shimizu T, Moribayashi A, Agui N (1985) In vitro analysis of spermiogenesis and testicular ecdysteroids in the cabbage armyworm, Mamestra brassicae L. Appl Entomol Zool 20:56–61 Shimizu T, Yagi S, Agui N (1989) The relationship of testicular and hemolymph ecdysteroid titer to spermiogenesis in the common armyworm, Leucania separata. Entomol Exp Appl 50:195–198 Shionoya M, Matsubayashi H, Asahina M, Kuniyoshi H, Nagata S, Riddiford LM, Kataoka H (2003) Molecular cloning of the prothoracicotropic hormone from the tobacco hornworm, Manduca sexta. Insect Biochem Mol Biol 33:795–801 Siegmund T, Korge G (2001) Innervation of the ring gland of Drosophila melanogaster. J Comp Neurol 431:481–491 Siew YC, Gilbert LI (1971) Effects of moulting hormone and juvenile hormone on insect endocrine gland activity. J Insect Physiol 17:2095–2104 Soltani N, Quennedey A, Delbecque J-P, Delachambre J (1987) Diflubenzuron-induced alterations during in vitro development of Tenebrio molitor pupal integument. Arch Insect Biochem Physiol 5:201–209 Soltani N, Delachambre J, Delbecque J-P (1989) Stage-specific effects of diflubenzuron on ecdysteroid titers during the development of Tenebrio molitor: evidence for a change in hormonal source. Gen Comp Endocrinol 76:350–356 Song Q, Gilbert LI (1998) Alterations in ultraspiracle (USP) content and phosphorylation state accompany feedback regulation of ecdysone synthesis in the insect prothoracic gland. Insect Biochem Mol Biol 28:849–860 Steel CGH (1975) A neuroendocrine feedback mechanism in the insect moulting cycle. Nature 253:267–269 Studinger G, Willig A (1975) Biosynthesis of α- and β-ecdysone in isolated abdomens of larvae of Musca domestica. J Insect Physiol 21:1793–1798 Takaki K, Sakurai S (2003) Regulation of prothoracic gland ecdysteroidogenic activity leading to pupal metamorphosis. Insect Biochem Mol Biol 33:1189–1199 Tatar M, Kopelman A, Epstein D, Tu MP, Yin CM, Garofalo RS (2001) A mutant Drosophila insulin receptor homolog that extends life-span and impairs neuroendocrine function. Science 292:107–110 Tatemoto K, Rökaeus A, Jörnvall H, McDonald TJ, Mutt V (1983) Galanin: a novel biologically active peptide from porcine intestine. FEBS Lett 164:124–128

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Truman JW (2006) Steroid hormone secretion in insects comes of age. PNAS 103:8909–8910 Tsuzuki S, Masuta T, Furumo M, Sakurai S, Iwami M (1997) Structure and expression of bombyxin E1 gene, a novel family gene that encodes bombyxin-IV, an insect insulin-related neurosecretory peptide. Comp Biochem Physiol 117:409–416 Tu MP, Yin CM, Tatar M (2002) Impaired ovarian ecdysone synthesis of Drosophila melanogaster insulin receptor mutants. Aging Cell 1:158–160 Tu MP, Yin CM, Tatar M (2005) Mutations in insulin signaling pathway alter juvenile hormone synthesis in Drosophila melanogaster. Gen Comp Endocrinol 142:347–356 Vafopoulou X, Steel CG (1997) Ecdysteroidogenic action of Bombyx prothoracicotropic hormone and bombyxin on the prothoracic glands of Rhodnius prolixus in vitro. J Insect Physiol 43:651–656 Vafopoulou X, Steel CG (2005) Testis ecdysiotropic peptides in Rhodnius prolixus: biological activity and distribution in the nervous system and testis. J Insect Physiol 51:1227–1239 Vandersmissen T, De Loof A, Gu SH (2007) Both prothoracicotropic hormone and an autocrine factor are involved in control of prothoracic gland ecdysteroidogenesis in Locusta migratoria and Schistocerca gregaria. Peptides 28:44–50 Van de Velde S, Badisco L, Claeys I, Verleyen P, Chen X, Vanden Bosch L, Vanden Broeck J, Smagghe G (2007) Insulin-like peptides in Spodoptera littoralis (Lepidoptera): detection, localization and identification. Gen Comp Endocrinol 153:72–79 Wagner RM, Loeb MJ, Kochansky JP, Gelman DB, Lusby WR, Bell RA (1997) Identification and characterization of an ecdysiotropic peptide from brain extracts of the gypsy moth, Lymantria dispar. Arch Insect Biochem Physiol 34:175–189 Wasielewski O, Rosin´ski G (2007) Gonadoinhibitory effects of Neb-colloostatin and Neb-TMOF on ovarian development in the mealworm, Tenebrio molitor L. Arch Insect Biochem Physiol 64:131–141 Watson RD, Bollenbacher WE (1988) Juvenile hormone regulates the steroidogenic competence of Manduca sexta prothoracic glands. Mol Cell Endocrinol 57:251–259 Watson RD, Yeh WE, Muehleisen DP, Watson CJ, Bollenbacher WE (1993) Stimulation of ecdysteroidogenesis by small prothoracicotropic hormone: role of cyclic AMP. Mol Cell Endocrinol 92:221–228 Webb R, Campbell BK, Garverick HA, Gong JG, Gutierrez CG, Armstrong DG (1999) Molecular mechanisms regulating follicular recruitment and selection. J Reprod Fertil Suppl 54:33–48 Wei ZJ, Zhang QR, Kang L, Xu WH, Denlinger DL (2005) Molecular characterization and expression of prothoracicotropic hormone during development and pupal diapause in the cotton bollworm, Helicoverpa armigera. J Insect Physiol 51:691–700 Wennauer R, Kassel L, Hoffmann KH (1989) The effects of juvenile hormone, 20-hydroxyecdysone, precocene II, and ovariectomy on the activity of the corpora allata (in vitro) in adult female Gryllus bimaculatus. J Insect Physiol 35:299–304 White MF (1998) The IRS-signaling system: a network of docking proteins that mediate insulin action. Mol Cell Biochem 182:3–11 Wigglesworth VB (1970) Structural lipids in the insect cuticle and the function of the oenocytes. Tissue Cell 2:155–179 Williams CM (1952) Physiology of insect diapause. IV. The brain and prothoracic glands as an endocrine system in the Cecropia silkworm. Biol Bull 103:120–138 Wu Q, Brown MR (2006) Signalling and function of insulin-like peptides in insects. Annu Rev Entomol 51:1–24 Xu J, Su J, Shen J, Xu W (2007) Molecular characterization and developmental expression of the gene encoding the prothoracicotropic hormone in the beet armyworm, Spodoptera exigua. Sci China C Life Sci 50:466–472 Xu WH, Denlinger DL (2003) Molecular characterization of prothoracicotropic hormone and diapause hormone in Heliothis virescens during diapause, and a new role for diapause hormone. Insect Mol Biol 12:509–516 Xu WH, Rinehart JP, Denlinger DL (2003) Structural characterization and expression analysis of prothoracicotropic hormone in the cornworm, Helicoverpa zea. Peptides 24:1319–1325

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Chapter 12

20-Hydroxyecdysone, Juvenile Hormone and Biogenic Amines: Mechanisms of Interaction in Control of Drosophila Reproduction Under Normal and Stressful Conditions Nataly Gruntenko and Inga Rauschenbach Abstract Juvenile hormone (JH) and ecdysteroids play a gonadotropic role in insect reproduction. For the normal progress of oogenesis, a proper balance between JH and 20-hydroxyecdysone (20E) is of a paramount importance. An imbalance of gonadotropins (shifting the balance either to the side of JH or 20E) leads to reproductive defects: a rise in JH titre leads to oviposition arrest, a rise in 20E level, to the degradation of vitellogenic oocytes. Upon a change in the level of one of the gonadotropins as a result of a mutation, effect of a stressor or pharmacological agent, the balance is restored owing to the relative change in the titre of the other one. Mediators in the JH and 20E interrelationship are biogenic amines, dopamine and octopamine. Existence of the mechanism of gonadotropin’s – reciprocal regulation is adaptive. Keywords Drosophila • stress • reproduction • adaptability • Juvenile hormone • 20-hydroxyecdysone • dopamine • octopamine

12.1

Introduction

It has long been established that juvenile hormone (JH) and ecdysteroids (ecdysone and 20-hydroxyecdysone (20E) ) play a gonadotropic role in insect reproduction (reviewed by Koeppe et al., 1985; Bownes, 1989; Raikhel et al., 2004). According to the model generally accepted, JH, synthesized by corpus allatum (CA), stimulates ecdysteroid synthesis in the ovaries. Ecdysteroids, produced by the ovarian follicular cells, stimulate vitellogenin (Vg) synthesis in the fat body; Vg is subsequently taken up from hemolymph by ovaries. The production of both hormones is under control of a third group of insect gonadotropins, neuropeptides (reviews: Postlethwait and Shirk, 1981; Bownes, 1989; Simonet et al., 2004). Richard et al. (1998, 2001) propose that in Drosophila JH initiates only early stages of vitellogenesis in the fat body and in the ovarian follicular cells and it stimulates ecdysteroid N. Gruntenko () and I. Rauschenbach Institute of Cytology and Genetics SD RAS, Russian Academy of Sciences, Novosibirsk 630090, Russia e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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production in the ovary, while 20E plays a prominent role in the control of oogenesis by stimulating the late stages of yolk proteins (YP) production in the fat body, their transportation from hemolymph to the nurse cells and their further uptake by the oocytes. Soller et al. (1999), based on the results of experiments on the effect of exogenous JH and 20E treatment on D. melanogaster vitellogenesis, have come to the conclusion that the development of vitellogenic oocytes, including both YP production by the follicular cells and their uptake by the oocytes is promoted by JH, while 20E regulates previtellogenic stages of the oocyte development. The authors also propose that for the normal progress of oogenesis in Drosophila, a proper balance between JH and 20E is of a paramount importance (Soller et al., 1999). We obtained data (Gruntenko et al., 2003a, b, 2005a, b, 2007; Rauschenbach et al., 2004a, b, 2007; Karpova et al., 2005) which (i) support both the supposition by Soller et al. (1999) about the importance of the gonadotropin balance in the control of Drosophila oogenesis and the concept of Richard et al. (2001) regarding the prominent role of 20E in the hormonal control of the Drosophila female reproductive function, and (ii) demonstrate that in Drosophila there is a mechanism of reciprocal regulation of JH and 20E which is responsible for their proper balance. Here we present the data in short.

12.2

Imbalance of 20E and JH Leads to Reproductive Defects

To determine the effects of shifting the balance either to the side of JH or 20E on Drosophila reproduction we used two approaches: (1) studied the effect of stress, which changes sharply levels of JH and 20E (Rauschenbach et al., 1995; Hirashima et al., 2000; Gruntenko et al., 2003b), on oogenesis in wild type females of D. virilis (wt strain) and in females carrying the mutation that prevents the change in JH level under heat stress (hs strain) (Gruntenko et al., 2003b; Rauschenbach et al., 2004a); (2) estimated the influence of exogenous JH and 20E on fertility of females of wt strain (Rauschenbach et al., 2004a; Gruntenko et al., 2005a). We first described the hs mutant phenotype as a temperature-sensitive conditional larval lethal. Mortality results from the absence of a response of the JH metabolic system to heat stress. Wt larvae enduring environmental stress (heating, crowding, starvation) are able to undergo normal metamorphosis because the factor that activates the JH-degrading enzyme in the hemolymph is inhibited. As a result, JH titre decreases slowly and JH activates ecdysone synthesis in the peritracheal gland in the absence of protoracicotropic hormone, the synthesis of which is inhibited under heat stress (Rauschenbach et al., 1984, 1987). The brain factor that activates the enzymes causing JH degradation is not inhibited under heat stress in hs larvae. As a result, JH is degraded, the peritracheal gland remains inactive, and a very low level of 20E prevents mutant larvae from undergoing metamorphosis, which leads to their death (Rauschenbach et al., 1984, 1987). Wt adult females respond to heat stress (38°C) by an increase in the level of 20E and by a decrease in JH degradation level (Table 12.1). We believe that the latter is

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Table 12.1 Effect of heat stress (38°C, 3 h) on JH degradation and 20E content in 1-day-old D. virilis females of wt and hs strains (Hirashima et al., 2000; Gruntenko et al., 2003b) JH degradation 20E content (pmol/min/fly) (pg/fly) wt strain hs strain

Control

Heat stress

Control

Heat stress

13.7 ± 0.64 7.25 ± 0.30

8.12 ± 0.67 7.11 ± 0.22

10.7 ± 0.48 13.0 ± 0.47

15.6 ± 1.05 36.72 ± 0.80

indicative of an increase in JH level. Indeed, in wild type females of D. melanogaster the regulation of JH synthesis and degradation tends to be opposing: both JH titre (Bownes and Rembold, 1987; Sliter et al., 1987) and JH synthesis (Altaratz et al., 1991) in young (1-day-old) wild type D. melanogaster females are substantially higher than in mature (5–6-days-old) flies. At the same time, JH degradation in young wild type D. melanogaster females is significantly lower than in the mature ones (Gruntenko et al., 2000, 2003a). Females of the mutant strain apterous56f of D. melanogaster were shown to have dramatically decreased JH synthesis (Altaratz et al., 1991) and sharply increased JH degradation (Gruntenko et al., 2003a). Considering all the above, we have infered (Gruntenko et al., 2003a) that (i) JH synthesis and degradation are under a common control system in the adult females of Drosophila, and (ii) the factors stimulating the hormone synthesis inhibit its degradation and vice versa. This notion agrees well with the fact that an experimental increase of the JH titre in wt females of D. virilis leads to a decrease in its degradation (Rauschenbach et al., 2004a). The idea of the correlated regulation of JH synthesis and degradation in insects is also supported by the data of Renucci et al. (1990) showing that ovariectomy of Acheta domesticus females results in the simultaneous decrease of JH synthesis and increase in the activity of JH-esterase that degrades the hormone. In microarray experiments (Terashima and Bownes, 2005) treatment of D. melanogaster starved females with JH leads to a down-regulation of JH-epoxide hydrolase 3 (the main JH-hydrolizing enzyme in adults females of D. melanogaster (Khlebodarova et al., 1996) ). In hs adult females the response of the JH metabolic system to heat stress is inhibited, but it does not interfere with the response of the 20E system. Besides, hs mutants have a significantly higher level of 20E and lower JH degradation level than wt flies under normal conditions (Table 12.1). Based on the hypothesis of Soller et al. (1999) about the importance of the gonadotropin balance in the control of Drosophila oogenesis we could expect changes in the progress of oogenesis in mutant females under normal conditions. One would also predict changes in oogenesis in wt and hs females under heat stress conditions. Figure 12.1a shows the stage distribution of egg chambers in young wt and hs females under normal and heat stress (38°C) conditions (Gruntenko et al., 2003b). Under normal conditions hs mutant females have a higher proportion of early vitellogenic oocytes (stages 8–9) than wt females. The proportion of oocytes in the late stages of development (11–14) is significantly higher in wt females. Degenerating stage 8–10 egg chambers are observed in hs mutant ovaries under normal conditions. These results suggest that in hs mutant flies the transition of oocytes through stage 10 is delayed, as evidenced by the decreased proportion of late stages (11–14)

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a degenerating oocytes / total number of stage 8-14 oocytes (%)

oocytes / number of stage 8-14 oocytes (%)

45 40 35 30 25 20 15 10 5 0 8

9

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45 40

wt heat stress

35

hs control

30

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25 20 15 10 5 0

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oocyte stage

b degenerating oocytes / total number of stage 8-14 oocytes (%)

oocytes / number of stage 8-14 oocytes (%)

45 40 35 30 25 20 15 10 5 0 8

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oocyte stage

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wt control

40

wt starvation

35

hs control

30

hs starvation

25 20 15 10 5 0 8

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oocyte stage

Fig. 12.1 Effects of (a) heat (38°C, 4 h) and (b) nutrition (flies kept on pure sugar diet during 24 h) stresses on oogenesis in 3-day-old D. virilis wild type (wt) and hs mutant females. Oocyte stages were determined for 7–12 pairs of ovaries after Hoechst staining. Left diagrams show the distribution of egg chambers and right diagrams, the distribution of degenerating egg chambers under normal conditions and 12 h after stresses. Means ± SE

and the increased proportion of early stages (8–9). Together with our data showing the increased 20E level in hs mutant flies (Table 12.1), the results are consistent with the model of Soller et al. (1999) suggesting that an elevation of 20E levels in females slows down development to stage 10. Partial degeneration of stages 8–9 oocytes in hs mutant females also fits well with the conclusion of Soller et al. (1999) that elevated titres of 20E lead to the resorption of early vitellogenic oocytes. Figure 12.1a also presents the data on the effect of heat stress on the course of oogenesis in wt and hs females. In wt females there are fewer normal oocytes at

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stages 9–13; many egg chambers degenerate during stages 8–10, and mature eggs (stage 14) accumulate. The decreased percentage of early vitellogenic oocytes is caused by the increased 20E titre induced by the heat stress (Table 12.1). One can also see a delay in oocyte transition to stage 10, similar to that observed in the hs mutant under normal conditions. This developmental delay is suggested by an increase of the total number (normal and degenerating) of egg chambers at stage 9 and a decrease of the numbers at stages 10–13 as compared to the control. The accumulation of mature eggs in wt females exposed to heat stress agrees with our data showing the cessation of oviposition after heat stress, probably resulting from a decrease in JH degradation (an increase in JH titre) (Rauschenbach et al., 1996). Following heat stress hs mutant females also have a reduced proportion of early vitellogenic oocytes, stages 9–10 and an increase in the proportion of degenerating egg chambers at stages 8–9. However, unlike wt, hs mutant females show neither an accumulation of mature eggs (Fig. 12.1a) nor a cessation of oviposition (Rauschenbach et al., 1996) in response to heat stress. Thus, hs mutant females have an ecdysteroid system which responds to heat stress, while their JH system does not; in parallel they have alterations in oogenesis after heat stress at early stages of oocyte development and do not show them at late stages. We have supposed that ecdysteroids make a bigger contribution to the control of early stages of vitellogenesis under stress conditions, while JH is more important for the completion of egg maturation. To verify this supposition, we studied JH degradation and oogenesis in hs and wt females following a different kind of stress – namely starvation. We have found that starvation is accompanied by a decrease in JH degradation levels (an increase in JH levels) in females of both wt and mutant hs strains (Table 12.2). If the accumulation of mature eggs under stress is due to a JH titre increase (a decrease of the hormone degradation) we could expect that the late stages of oogenesis in hs females, which do not change under heat stress (Fig. 12.1a), would change following starvation. Figure 12.1b shows the stage distribution of egg chambers in young wt and hs females under starvation (Rauschenbach et al., 2004a). The pattern of stage distribution in the wt strain following starvation is similar to that observed following heat stress, i.e. the changes in the progress of oogenesis are similar with different types of stress. The distribution of stages in hs mutant females following starvation is similar to that observed in wt females. It is important to note that under nutritional stress there is a significant accumulation of mature eggs in hs females. The accumulation of mature eggs in wt and hs females exposed to nutrition stress agrees with our data showing an oviposition arrest under starvation in both (Rauschenbach et al., 2004a). Table 12.2 Effect of nutrition stress (starvation for 6 h) on JH degradation in young (3-day-old) D. virilis females of wt and hs strains (Rauschenbach et al., 2004a)

JH degradation (pmol/min/fly) wt strain hs strain

Control

Starvation

13.1 ± 0.45 11.5 ± 0.47

9.31 ± 0.35 7.6 ± 0.48

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Based on the above, we have inferred that (i) oocyte accumulation at stage 14 and an oviposition arrest in females under stress occurred due to the increasing JH titre; (ii) oocyte degradation at stages 8–10 occurred due to the increasing 20E titre (Rauschenbach et al., 2004a). We have confirmed this inference when studied the effects of exogenous JH and 20E on oviposition and fertility in wt females of D. virilis: 1. JH treatment (applying 2 µg of JH-III) of wt females resulted in an oviposition arrest for 1 day (Rauschenbach et al., 2004a) similar to that observed under heat stress and starvation (Rauschenbach et al., 1996, 2004a). However, unlike heat stress and starvation, it did not cause a prolonged fertility decrease when oviposition renewed (Rauschenbach et al., 2004a). 2. Different from the JH treatment, 20E treatment (24 h feeding by the hormone) of wt females did not cause an oviposition arrest. At the same time, the exogenous 20E caused a prolonged decrease of fertility: the fecundity of the 20E-treated flies was 63–74% from the control level for 4 days (Gruntenko et al., 2005a).

12.3

The Mechanisms of Maintenance of JH and 20E Balance in Drosophila Females

It has been demonstrated in vitro that biogenic amines may have a regulatory action on JH metabolism in bees, locusts, crickets, beetles and cockroaches (Lafon-Cazal and Baehr, 1988; Thompson et al., 1990; Woodring and Hoffmann, 1994; Kaatz et al., 1994; Rachinsky, 1994; Granger et al., 1996; Hirashima et al., 1999b). The effect of 20E on the metabolism of biogenic amines has also been shown in insect (Hiruma et al., 1985; Hiruma and Riddiford, 1990; Hirashima et al., 1999a; Ferdig et al., 2000; Lehman et al., 2000; Mesce, 2002; Zufelato et al., 2004). However, in the available literature we found no researches into the effects of JH on biogenic amines and the effects of biogenic amines on 20E levels in insects. To study in vivo the interplay between biogenic amines, dopamine (DA) and octopamine (OA), and gonadotropins, JH and 20E, in Drosophila we estimated JH, 20E, OA and DA levels (i) in D. melanogaster females carrying mutations that change drastically OA, DA, JH or 20E levels, and (ii) in wild type females of D. virilis and D. melanogaster treated with exogenous OA, DA, JH or 20E.

12.3.1

The Interplay Between Biogenic Amines and JH in Drosophila In Vivo

To determine whether biogenic amines affect JH metabolism in Drosophila in vivo, we studied levels of hormone degradation in octopamineless females (strain TbhnM18 of D. melanogaster carries a null mutation of the gene tyramine-b-hydroxylase, OA biosynthesis enzyme (Monastirioti et al., 1996) ) and in females with a twofold increase of the DA content (D. melanogaster strains scarlet ebony (ste) and ebony

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carry mutation which drastically decrease activity of the enzyme converting DA into N-b-alanyldopamine (Perez et al., 1997) ). It has been found that both young and mature octopamineless females have JH degradation levels much higher than those in females of the precursor strain (p845) from which the octopamineless strain was derived by P-element transposition (Monastirioti et al., 1996) and in wild type flies (Canton S). At the same time, the young females with a twofold increase of the DA content have considerably lower JH degradation levels and the mature flies have higher its levels compared to wild type (Gruntenko et al., 2000; Rauschenbach et al., 2001; Gruntenko and Rauschenbach, 2004). It should be noted that these changes in JH metabolism are most likely to be due to the mutations changing OA and DA levels rather than to other genes. This is indicated, first, by the difference in JH degradation levels between the strain TbhnM18 and its precursor strain, p845, and second, by the similar levels of hormone degradation in females of the strains ste and ebony which carry, compared to different genetic backgrounds, a mutation that results in doubling of the DA content. Based on the above, we suggested that the pattern of the regulation of JH metabolism by the biogenic amines in Drosophila includes OA inhibiting JH degradation and stimulating its synthesis (see Section 12.2) both in young and mature Drosophila females; DA inhibits hormone degradation and stimulates its synthesis in the young females, and in contrast, it stimulates JH degradation and inhibits its synthesis in mature females (Gruntenko and Rauschenbach, 2004). To verify this suggestion we studied the effects of DA and OA treatment (feeding flies with the amines) on JH degradation levels in Drosophila wild type flies (Gruntenko et al., 2005a, 2007). Figure 12.2a presents the results of measurements of the JH-hydrolyzing activity in young (2-day-old) and sexually mature (6-day-old) wt females, both OA treated and control. The increase in OA content leads in both young and mature females to a decrease of JH degradation (an increase of JH level, see Section 12.2). We obtained similar data from a study of the effect of exogenous OA on JH metabolism in

18 16 14 12 10 8 6 4 2 0

young

mature

b 20

control DA treated

18 16 14 12 10 8 6 4 2 0

c 10

DA content (ng/mg fly mass)

20

control OA treated

JH-hydrolysing activity (pmol/min/fly)

JH-hydrolysing activity (pmol/min/fly)

a

young

mature

aceton JH treated

8 6 4 2 0

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Fig. 12.2 Effects of OA (a) and DA (b) feeding on JH degradation, and effect of JH application on DA content (c) in young and mature females of D. virilis wild type. Means ± SE

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D. melanogaster (Gruntenko et al., 2007). Note, that the increase in OA content had no effect on JH degradation levels in Drosophila males (Gruntenko et al., 2007). The effect of exogenous DA on JH-hydrolyzing activity in young (2-day-old) and mature (7-day-old) wt females is shown in Fig. 12.2b. The increase in DA content leads in young females to a sharp decrease of JH degradation (an increase of JH level) and in the mature females to its increase (decrease of JH level). Ontogenetic differences in the control of JH metabolism by DA has also been established for larvae of Manduca sexta: DA stimulates hormone biosynthesis in the corpora allata in the first 2 days of the last larval stage but inhibits the corpora allata on days 3–6, at the beginning of the prepupal stage (Granger et al., 1996). It should be emphasized, that the increase in DA content, like in OA, had no effect on JH degradation levels in wt males (Gruntenko et al., 2005a). We have also shown that threefold decrease of the DA content after 3-iodo-tyrosine (an inhibitor of tyrosine hydroxylase) treatment results in a drop of JH degradation (an increase of JH level) in the mature wt females (Gruntenko et al., 2005a). As mentioned above, the apterous56f mutation leads to dramatically decreased JH levels in D. melanogaster (Altaratz et al., 1991; Gruntenko et al., 2003a). We have found that young ap56f females have DA levels twice as high as Canton S and Oregon R females (Gruntenko et al., 2003a). Although the ap gene encodes one of the LIM homeodomain transcription factors which play key roles in a variety of developmental processes (Hobert and Westphal, 2000 for review) our data suggested it was not directly involved in the regulation of DA metabolism. Really, DA content in males of the ap56f strain did not differ from the wild type (Gruntenko et al., 2003a). We presumed that the increased DA level in ap56f females could be explained by a compensatory response to a reduced JH titre: DA levels rise in young ap56f females at the beginning of the oogenesis in order to increase JH titre by stimulation of its synthesis and suppression of its degradation (see above). Support for this hypothesis was provided by our data on JH treatment of young ap56f females: the increase of JH level resulted in a decline in DA content making it closer to wild type level (Gruntenko et al., 2003a). Based on these data, we suggested that there is a feedback loop in the regulation of JH metabolism by DA (Rauschenbach et al., 2004b). The suggestion was confirmed when we studied the effects of exogenous JH on DA contents in young and mature wild type females of D. virilis (Rauschenbach et al., 2004b). Figure 12.2c depicts the results of measurements of the DA contents in young (2-day-old) and sexually mature (7-day-old) wt females after the application of JH-III dissolved in acetone and in the control treated with acetone. The increase in JH levels leads to the decrease in DA content in young females and to the rise in DA in mature females.

12.3.2

The Interplay Between JH and 20E in Drosophila In Vivo

To find out whether an experimental increase in 20E levels has any effect on JH metabolism in D. virilis, the flies were kept for 8 days on a nutrient medium with 20E.

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Levels of JH degradation were measured in 20E-treated and control young (2-day-old) and mature (7-day-old) wt females. The results are shown in Fig. 12.3a, b. 20E treatment leads to a considerable decrease of JH degradation (an increase of JH level) in young females (Fig. 12.3a). (Note, that this decrease is dose dependent (Gruntenko et al., 2005a).) In the mature females (Fig. 12.3b) too an increase in the 20E level results in a decrease of JH degradation (an increase of JH level). This testifies to the existence of a mutual control of JH and 20E in Drosophila: not only does JH stimulate ecdysteroid production (Richard et al., 1998) and increase 20E levels in vivo (Fig. 12.3c), but also 20E, in turn, is capable of regulating JH levels. Another proof of the inference that 20E affects JH metabolism are our results of JH degradation measurement in the ecdysoneless1 (ecd1) mutant of D. melanogaster: in 1- and 5-day-old ecd1 females maintained at 29°C (shifting newly emerged ecd1 adults to 29°C results in drastically reduced 20E titres (Garren et al., 1977) ) JH degradation is significantly higher (JH level lower) as compared to that

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of ecd1 females kept at the temperature 19°C at which they have the normal 20E level (Karpova et al., 2005). A question arose whether a change in the 20E level affects directly JH metabolism or is it mediated through the DA system, since we have shown that JH metabolism under normal conditions is regulated by DA (DA inhibits JH degradation in the young females and stimulates it in the mature ones (see above) ). The data in Fig. 12.3a show a steep increase of the DA level in young wt females upon 20E treatment. This increase is dose dependent (Gruntenko et al., 2005a). However, the DA content in the mature females, unlike that in the young ones, decreases upon 20E treatment (Fig. 12.3b). The influence of 20E on DA levels in Drosophila is also confirmed by our data regarding the significant decrease of the DA level in the young and increase in the mature females of the strain ecd1 kept at 29°C (Karpova et al., 2005). Taking the above into consideration, it is reasonable to infer that the effect of 20E on JH metabolism in D. virilis is mediated through the DA metabolic system. Indeed, (i) an increase in the DA content in the young females after 20E treatment and its decrease in the mature ones should result in a decrease of the JH-hydrolyzing activity (an increase in JH level) in both; (ii) if 20E had a direct effect on JH metabolism, an increased level of the latter in 20E-treated females should lead to a decrease in DA content in young females (see Section 12.3.1) and its increase in the mature ones, and we observe the reverse (Fig. 12.3a, b). To elucidate the interplay between 20E and biogenic amines, OA and DA, we measured 20E contents in 2-day-old wt females of D. virilis fed with OA and with DA precursor, L-dihydroxyphenylalanine (L-DOPA). The females fed with L-DOPA had a much higher DA content (greater by the factor of 2.5 (Rauschenbach et al., 2007) ). As mentioned above, an increase in 20E level leads in young wt D. virilis females to an increase in DA content. In that case and if there is a feedback regulation (a direct effect of DA on 20E metabolic system), an increase in DA content in young females should result in a decrease in 20E level. Data presented in Fig. 12.3d indicate that this is not the case: feeding the flies with L-DOPA results in the increase of 20E level. At the same time, a rise in JH level (a decrease of its degradation) produced in young Drosophila females by the increase in DA content (Fig. 12.2b) should lead to a rise of 20E because JH activates ecdysone synthesis in ovaries of young females (Postlethwait and Shirk, 1981; Kelley, 1994; Richard et al., 1998). Data in Fig. 12.3c correlate with this: in JH-treated wt females the 20E level is dramatically increased. Thus we infer that DA has an effect on 20E metabolism, but this effect is indirect and mediated through the JH metabolic system. The effects of a rise in OA content on 20E level in wt females of D. virilis are shown in Fig. 12.3e. The increase in OA content in 2-day-old wt females leads to the increase of 20E content. The elevation of 20E titre could be due to the OA-induced increase in the JH titre. Indeed, the increase in OA content leads in both young and mature females to a decrease of JH degradation (an increase of JH level, Fig. 12.2a). As mentioned above (see Section 12.3.1), we have also shown that both young and mature octopamineless females of D. melanogaster strain TbhnM18 have JH degradation levels much higher (JH levels much lower) than those in its precursor strain,

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p845, flies (Gruntenko et al., 2000). If OA, like DA, regulates 20E through JH metabolic system, one could expect octopamineless females to have 20E level lower than in wild type. Indeed, we have found a considerable decrease in 20E content in TbHNM18 females (Rauschenbach et al., 2007). It cannot be ruled out, however, that besides the regulation of 20E titre mediated via a JH system OA may also regulate 20E titre directly. This possibility is supported by Hirashima et al. (1999a) who show that exogenous OA affects in vitro ecdysteroid synthesis by the prothoracic gland of Bombyx mori larvae.

12.3.3

Mechanism of the Reciprocal Regulation of Gonadotropins and Biogenic Amines in Drosophila

Taking all above into consideration, we inferred (Rauschenbach et al., 2007) that a mechanism of the reciprocal regulation of biogenic amines and gonadotropins which maintains the balance of JH and 20E in the case when either of the hormone levels is changed exists in Drosophila (Fig. 12.4): (i) DA increases JH levels (inhibiting JH degradation and apparently stimulating synthesis) in young females and decreases it by stimulating degradation and inhibiting synthesis in sexually mature flies (Gruntenko et al., 2000, 2005b; Gruntenko and Rauschenbach, 2004; see Fig. 12.2). (ii) There is a feedback loop as part of this regulation – a rise in JH levels leads to the decrease in DA content in young females and to the rise in DA in mature females (Gruntenko et al., 2003b; Rauschenbach et al., 2004b; see Fig. 12.2).

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Fig. 12.4 Scheme of the reciprocal regulation of gonadotropins (JH and 20E) and biogenic amines (DA and OA) in Drosophila. Arrows in the circles point out effect directions: upward – increasing, down – decreasing

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(iii) OA leads to a rise in JH levels by inhibiting JH degradation and apparently stimulating its synthesis in young and mature females (Gruntenko et al., 2000, 2007; Rauschenbach et al., 2007; see Fig. 12.2). (iv) 20E regulates JH indirectly via DA metabolic pathways – a rise in 20E level increases DA content in young females and decreases it in mature females, thus leading to a decrease of JH degradation and hence a rise in the hormone levels in both (Gruntenko et al., 2005a; see Fig. 12.3). (v) JH regulates 20E metabolism: a rise in JH titre increases the 20E level in young females (Rauschenbach et al., 2007; see Fig. 12.3). (vi) DA and OA regulate 20E metabolism via JH metabolic pathways – a rise in DA and OA contents increases JH level in young females, thus leading to a rise in the 20E titre (Gruntenko et al., 2007; Rauschenbach et al., 2007; see Fig. 12.3). A natural question to ask is how this mechanism works? We suggest the following (Fig. 12.5a, b). I. If 20E level rises (Fig. 12.5a): In young females: 1. 20E causes an increase in DA level. 2. The increase of DA level results in a rise in JH titre.

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3. The rise in JH titre elicits a decrease of DA content, DA level stabilizes, and the gonadotropin balance sets in at a new level. In mature females: 1. 20E causes a decrease in DA level. 2. The decrease of DA level results in a rise in JH titre. 3. The rise in JH titre elicits an increase of DA content, its level stabilizes, and the gonadotropin balance sets in at a new level. II. If JH level rises (Fig. 12.5b): In young females: 1. JH causes an increase of 20E titre and a decrease of DA content. 2. The increase of 20E titre results in a rise in DA content, its level restores, and the balance of JH and 20E sets in at a new level. In mature females: 1. JH causes an increase in both 20E titre and DA content. 2. The increase in 20E titre results in a drop of DA content, its level stabilizes, and the gonadotropin balance sets in at a new level.

12.4

Conclusions

The above allows us to conclude: 1. Stress leads to dramatic changes in the reproductive function of Drosophila females, the reason being imbalance of gonadotropins which occurs owing to non-simultaneous response of their systems of metabolism to stressor. Indeed, ecdysteroid system responds to 60 min of stress exposure (38°C) with an increase in ecdysone (E) and 20-hydroxyecdysone (20E) levels (Hirashima et al., 2000); JH metabolic system responds to stress with a decrease in JH degradation (increase in JH titre) beginning after 120 min of stress exposure (Rauschenbach et al., 1995, 1996; Gruntenko et al., 2003b). 2. A rise in JH titre under stress is adaptive because it results in mature eggs accumulation and oviposition arrest, allowing the population to “wait” until the end of unfavourable conditions without a considerable decrease in potential numbers; a rise in 20E titre is adaptive under the conditions of overpopulation or reduced food supply because it leads to resorption of a part of egg chambers and a decrease of fecundity. 3. The mechanism of a reciprocal regulation of biogenic amines and gonadotropins which maintains the balance of JH and 20E in the case when either of the hormone levels is changed exists in Drosophila. The existence of such mechanism is also adaptive because it promotes restoration of the normal levels of reproduction after stress exposure.

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References Altaratz M, Applebaum ShW, Richard DS, Gilbert LI, Segal D (1991) Regulation of juvenile hormone synthethis in wild-type and apterous mutant Drosophila. Mol Cell Endocrinol 81:205–216. Bownes M (1989) The roles of juvenile hormone, ecdysone and the ovary in the control of Drosophila vitellogenesis. J Insect Physiol 35:409–413. Bownes M, Rembold H (1987) The titre of juvenile hormone during the pupal and adult stage of the life cycle of Drosophila melanogaster. Euro J Biochem 164:709–712. Ferdig MT, Taft AS, Smartt CT, Lowenberger CA, Li J, Zhang J, Christensen BM (2000) Aedes aegypti dopa decarboxylase: gene structure and regulation. Insect Mol Biol 9:231–239. Garen A, Kauvar L, Lepesant J (1977) Roles of ecdysone in Drosophila development Proc Natl Acad. Sci USA 74:5099–5103. Granger NA, Sturgis SL, Ebersohl R, Geng C, Sparks TC (1996) Dopaminergic control of corpora allata activity in the larval tobacco hornworm, Manduca sexta. Arch Insect Biochem Physiol 32:449–466. Gruntenko NE, Rauschenbach IYu (2004) Adaptive value of genes controlling the level of biogenic amines in Drosophila. Russ J Genet 40:703–709. Gruntenko NE, Wilson TG, Monastirioti M, Rauschenbach IYu (2000) Stress-reactivity and juvenile hormone degradation in Drosophila melanogaster strains having stress-related mutations. Insect Biochem Mol Biol 30:775–783. Gruntenko NE, Chentsova NA, Andreenkova EV, Bownes M, Segal D, Adonyeva NV, Rauschenbach IYu (2003a) Stress response in a juvenile hormone deficient Drosophila melanogaster mutant apterous56f. Insect Mol Biol 12:353–363. Gruntenko NE, Bownes M, Terashima J, Sukhanova M, Rauschenbach IYu (2003b) Environmental stress affects oogenesis differently in wild type and a Drosophila virilis mutant with altered juvenile hormone and 20-hydroxyecdysone levels. Insect Mol Biol 12:393–404. Gruntenko NE, Karpova EK, Adonyeva NV, Chentsova NA, Faddeeva NV, Alekseev AA, Rauschenbach IYu (2005a) Juvenile hormone, 20-hydroxyecdysone and dopamine interaction in Drosophila virilis reproduction under normal and nutritional stress conditions. J Insect Physiol 51:417–425. Gruntenko NE, Karpova EK, Alekseev AA, Chentsova NA, Saprykina ZV, Bownes M, Rauschenbach IYu (2005b) Effects of dopamine on juvenile hormone metabolism and fitness in Drosophila virilis. J Insect Physiol 51:959–968. Gruntenko NE, Karpova EK, Alekseev AA, Chentsova NA, Bogomolova EV, Bownes M, Rauschenbach IYu (2007) Effects of octopamine on reproduction, juvenile hormone metabolism, dopamine and 20-hydroxyecdysone contents in Drosophila. Arch Insect Biochem Physiol 65:85–94. Hiruma K, Riddiford LM (1990) Regulation of dopa decarboxylase gene expression in the larval epidermis of the tobacco hornworm by 20-hydroxyecdysone and juvenile hormone. Dev Biol 138:214–224. Hiruma K, Riddiford LM, Hopkins TL, Morgan TD (1985) Roles of dopa decarboxylase and phenoloxidase in the melanization of the tobacco hornworm and their control by 20hydroxyecdysone. Comp Physiol B 155:659–669. Hirashima A, Hirokado S, Ohta H, Suetsugu E, Sakaguichi M, Kuwano E, Taniguchi E, Eto M (1999a) Titres of biogenic amines and ecdysteroids: effect of octopamine on the production of ecdysteroids in the silkworm Bombyx mori. J Insect Physiol 45:843–851. Hirashima A, Suetsugu E, Hirokado S, Kuwano E, Taniguchi E, Eto M (1999b) Effect of octopamine on the activity of juvenile-hormone esterase in the silkworm Bombyx mori and the red flour beetle Tribolium freemani. Gen Comp Endocrinol 116:373–381. Hirashima A, Rauschenbach IYu, Sukhanova MJh (2000) Ecdysteroids in stress responsive and nonresponsive Drosophila virilis lines under stress conditions. Biosci Biotech Biochem 64:2657–2662. Hobert O, Westphal H (2000) Functions of LIM-homeobox genes. Trends Genet 16:75–83. Kaatz H, Eichmuller S, Kreissl S (1994) Stimulatory effect of octopamine on juvenile hormone biosynthesis in honey bees (Apis mellifera): physiological and immunocytochemical evidence. J Insect Physiol 40:856–872.

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Karpova EK, Gruntenko NE, Rauschenbach IYu (2005) The ecdysoneless1 gene regulates metabolism of the juvenile hormone and dopamine in Drosophila melanogaster. Russ J Genet 41: 1480–1486. Kelley TJ (1994) Endocrinology of vitellogenesis in Drosophila melanogaster. Perspectives in Comparative Endocrinology. Ottawa: National Research Council of Canada, pp. 282–290. Khlebodarova TM, Gruntenko NE, Grenback LG, Sukhanova MZ, Mazurov MM, Tomas BA, Hammock BD, Rauschenbach IY (1996) A comparative analysis of juvenile hormone metabolysing enzymes in two species of Drosophila during development. Insect Biochem Mol Biol 26:829–835. Koeppe JK, Fuchs M, Chen TT, Hunt LM, Kovalick GE (1985) The role of juvenile hormone in reproduction. In: Comprehensive Insect Physiology, Biochemistry and Pharmacology. Eds. Kerkut GA, Gilbert LI, Vol. 8. New York: Pergamon Press, pp. 165–203. Lafon-Cazal M, Baehr JC (1988) Octopaminergic control of corpora allata activity in an insect. Experentia 44:895–896. Lehman HK, Klukas KA, Gilchrist LS, Mesce KA (2000) Steroid regulation of octopamine expression during metamorphic development of the moth Manduca sexta. J Comp Neurol 424: 283–296. Mesce KA (2002) Metamodulation of the biogenic amines: second-order modulation by steroid hormones and amine cocktails. Brain Behav Evolut 60:339–349. Monastirioti M, Linn CE, White K (1996) Characterization of Drosophila Tyramine b-hydroxylase gene and isolation of mutant flies lacking octopamine. J Neurosci 16:3900–3911. Perez M, Castillo-Marin N, Quesada-Allue LA (1997) β-alanyl-dopamine synthase in Drosophila melanogaster and Ceratitis capitata melanic mutants. DIS 80:39–41. Postlethwait JH, Shirk PD (1981) Genetic and endocrine regulation of vitellogenesis in Drosophila. Am Zool 21:687–700. Rachinsky A (1994) Octopamine and serotonin influence on corpora allata activity in honey bee (Apis mellifera) larvae. J Insect Physiol 40:549–554. Raikhel AS, Brown MR, Belles X (2004) Hormonal control of reproductive processes. In: Comprehensive Molecular Insect Science. Eds. Gilbert LI, Iatrou K, Gill S, Vol. 3. Elsevier Pergamon, Boston pp. 433–491. Rauschenbach IYu, Gruntenko NE, Khlebodarova TM, Mazurov MM, Grenback LG, Sukhanova MJh, Shumnaja LV, Zakharov IK, Hammock BD (1996) The role of the degradation system of the juvenile hormone in the reproduction of Drosophila under stress. J Insect Physiol 42:735–742. Rauschenbach IYu, Lukashina NS, Korochkin LI (1984) Genetic of esterases in Drosophila. VIII. The gene controlling the activity of JH-esterase in D. virilis. Biochem Genet 22:5–80. Rauschenbach IYu, Lukashina NS, Maksimovsky LF, Korochkin LI (1987) Stress-like reaction of Drosophila to adverse environmental factors. J Comp Physiol 157:519–531. Rauschenbach IYu, Khlebodarova TM, Chentsova NA, Gruntenko NE, Grenback LG, Yantsen EI, Filipenko ML (1995) Metabolism of the juvenile hormone in Drosophila adults under normal conditions and heat stress. J Insect Physiol 41:179–189. Rauschenbach IYu, Gruntenko NE, Chentsova NA, Hirashima A, Sukhanova MJh, Andreenkova EV (2001) Hormones interrelationship in the control of reproductive function of Drosophila females under stress conditions is genetically determinated. Russ J Genet 37:1243–1250. Raushenbach IYu, Gruntenko NE, Bownes M, Adonieva NV, Terashima J, Karpova EK, Faddeeva NV, Chentsova NA (2004a) The role of juvenile hormone in the control of reproductive function in Drosophila virilis under nutritional stress. J Insect Physiol 50:323–330. Rauschenbach IYu, Gruntenko NE, Chentsova NA, Adonyeva NV, Karpova EK (2004b) Feedback in the regulation of juvenile hormone titre by biogenic amines in Drosophila. Dokl Biol Sci 397:324–325. Rauschenbach IYu, Chentsova NA, Alekseev AA, Gruntenko NE, Adonyeva NV, Karpova EK, Komarova TN, Vasiliev VG, Bownes M (2007) Dopamine and octopamine regulate 20hydroxyecdysone level in vivo in Drosophila. Arch Insect Biochem Physiol 65:95–102. Renucci M, Strambi C, Strambi A, Augier R, Charpin P (1990) Ovaries and regulation of juvenile hormone titer in Acheta domesticus L. (Ortoptera). Gen Comp Endocrinol 78:137–149.

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Richard DS, Watkins NL, Serafin RB, Gilbert LI (1998) Ecdysteroids regulate yolk protein uptake by Drosophila melanogaster oocytes. J Insect Physiol 44:637–644. Richard DS, Jones JM, Barbarito MR, Cerula S, Detweiler JP, Fisher SJ, Brannigan DM, Scheswohl M (2001) Vitellogenesis in diapausing and mutant Drosophila melanogaster: further evidence for the ralative roles of ecdysteroids and juvenile hormones. J Insect Physiol 47:905–913. Simonet G, Poels J, Claeys I, Van Loy T, Franssens V, De Loof A, Vanden Broeck J (2004) Neuroendocrinological and molecular aspects of Insect reproduction. J Neuroendocrinol 16:649–659. Sliter TJ, Sedlak BJ, Baker FC, Schooley DA (1987) Juvenile hormone in Drosophila. Identification and titer determination during development. Insect Biochem 17:161–165. Soller M, Bownes M, Kubli E (1999) Control of oocyte maturation in sexually mature Drosophila females. Dev Biol 208:337–351. Terashima J, Bownes M (2005) A microarray analysis of genes involved in relating egg production to nutritional intake in Drosophila melanogaster. Cell Death Differ 12:429–440. Thompson CS, Yagi K, Chen ZF, Tobe SS (1990) The effects of octopamine on juvenile hormone biosynthesis, electrophysiology, and cAMP content of the corpora allata of the cockroach Diploptera punctata. J Comp Physiol B 160:241–249. Woodring J, Hoffmann KH (1994) The effects of octopamine, dopamine and serotonin on juvenile hormone synthesis, in vitro, in the cricket, Gryllus bimaculatus. J Insect Physiol 40:797–802. Zufelato MS, Lourenco AP, Simoes ZL, Jorge JA, Bitondi MM (2004) Phenoloxidase activity in Apis mellifera honey bee pupae, and ecdysteroid-dependent expression of the prophenoloxidase mRNA. Insect Biochem Mol Biol 34:1257–1268.

Chapter 13

The Structure and Function of Ecdysone Receptors Isabelle M.L. Billas, Christopher Browning, Michael C. Lawrence, Lloyd D. Graham, Dino Moras, and Ronald J. Hill

Abstract The ligand-binding properties of recombinant ecdysone receptor EcRUSP heterodimeric ligand-binding domains (LBDs) from four insect orders are described for a range of ecdysteroids and for a dibenzoylhydrazine (DBH) insecticide (tebufenozide). Much of the order selectivity of the insecticide in the field is reproduced by the affinity of tebufenozide for the recombinant LBDs in the laboratory. Crystal structures are presented for the LBDs of ecdysone receptors from the pest insects Heliothis virescens, Bemisia tabaci and Tribolium castaneum in complex with ponasterone A, as well as of the H. virescens LBD in complex with 20-hydroxyecdysone and BYI06830 (a DBH insecticide). Comparison of ecdysteroid- and BYI06830-bound structures of the H. virescens LBD illustrates the way in which this remarkable protein can adapt its binding pocket to very different ligand chemistries. Finally, comparison of the ligand-binding pockets of H. virescens, B. tabaci and T. castaneum ecdysone receptors begins to provide insights at an atomic level of detail into the insect order selectivity of the DBH insecticides. Keywords Ecdysone receptor • ligand binding domain • 20-hydroxyecdysone • ponasterone A • tebufenozide • dibenzoylhydrazine • X-ray crystal structure

I.M.L. Billas, C. Browning and D. Moras IGBMC (Institut de Génétique et de Biologie Moléculaire et Cellulaire), Département de Biologie et de Génomique Structurales, Illkirch, F-67400 France; INSERM, U596, Illkirch, F-67400 France; CNRS, UMR7104, F-67400 Illkirch, France email: [email protected] M.C. Lawrence Walter and Eliza Hall Institute of Medical Research, 1G Royal Parade, Parkville, Victoria 3050, Australia L.D. Graham and R.J. Hill CSIRO Molecular and Health Technologies, PO Box 184, North Ryde, New South Wales 1670, Australia email: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Introduction

Insect metamorphosis has fascinated mankind since ancient times (Peck, 1970). The puffing of specific sites on the giant polytene chromosomes of Drosophila melanogaster at the end of larval life and preceding metamorphosis exemplifies the response of the genome to a rise in the titre of the moulting hormone 20-hydroxyecdysone (20E) in the hemolymph (Becker, 1962). Ashburner et al. (1973) proposed a formal model to explain control of the transcription of the vast network of genes whose activity is induced by the hormone and is manifested by puffing at over a hundred chromosomal sites. The first step in this model is the binding of 20E to a postulated receptor protein, the existence of which was suggested by the binding of tritiated hormone to D. melanogaster salivary gland chromosomal proteins (Emmerich, 1972) and by analogy to mammalian steroid receptors. The fact that the thiol-specific reagent, N-ethylmaleimide, inhibited the response to 20E was adduced as evidence strengthening the analogy, as this reagent had been shown to block the interaction of mammalian steroid hormones with their receptors (Ashburner, 1972). Ashburner’s experiments further demonstrated that pre-incubation with increasing concentrations of 20E appeared to protect the molecular target against reaction with N-ethylmaleimide. The existence of the hypothetical ‘ecdysone receptor’ was put on a firm experimental basis by rigorous hormone binding studies employing tritiated ponasterone A ([3H]-ponA), a radiolabelled phyto-ecdysteroid that binds to the receptor more tightly than 20E (Yund et al., 1978). Despite the realization that the receptor was present only in trace amounts (103 molecules per cell), there then followed heroic biochemical attempts at its isolation, which at best yielded very small amounts of enriched/purified protein (Landon et al., 1988; Luo et al., 1991). A major advance was provided by the cloning of the EcR gene (Koelle et al., 1991) and the realization that the ecdysone receptor is actually a heterodimer of the EcR protein and the ultraspiracle protein product (USP) of the usp gene (Thomas et al., 1993; Yao et al., 1993). USP is an insect orthologue of the vertebrate RXR protein. In keeping with the central role played by 20E in insect development, mutations in the EcR gene are generally lethal even before the completion of embryonic development (Bender et al., 1997). DNA sequencing and conceptual translation revealed that both EcR and USP proteins are members of the nuclear receptor family and display the characteristic domain structure of an N-terminal A/B domain (transcription activation), a C domain (DNA binding, very highly conserved), a D region (linker, shown to contain nuclear localisation signals in many nuclear receptors), an E domain (ligand-binding, moderately conserved), and in some cases a distinct F domain (Moras and Gronemeyer, 1998). Cloned DNA encoding EcRs and USPs led to the availability of recombinant EcR and USP proteins, as well as the corresponding ecdysone receptor heterodimers. There has been considerable interest in possible ligands for the USP protein in its own right, some of which may have biological significance (Billas et al., 2001; Jones et al., 2006). In addition to their natural role in coordinating and controlling the expression of networks of genes in arthropod development and reproduction, ecdysone receptors have been employed as targets for environmentally-friendly insecticides and also in the control of transgenes (e.g. therapeutic genes) in mammalian cells

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(Christopherson et al., 1992; Yang et al., 1995; Palli et al., 2003) and plant systems (Unger et al., 2002). The dibenzoylhydrazine (DBH) class of insecticides was discovered in a screen carried out by Rohm and Haas Company in the 1980s. Despite their lack of structural similarity to ecdysteroids, Wing (1988) realized that the mode of action of this new class of insecticides involved the targeting of ecdysone receptors. The DBH compounds were reported to induce premature and lethal moulting in different types of caterpillar pest larvae (Dhadialla et al., 1998). The compounds exhibit a high level of safety, presumably at least in part due to the absence of ecdysone receptors in mammals. Furthermore, they act selectively on particular taxonomic orders, making them safe for many non-target and beneficial insects. More recently, compounds of other chemistries have been found to bind to ecdysone receptors, including the tetrahydroquinoline (THQ) class of compounds (Smith et al., 2003) and the oxadiazolines (Hormann and Chortyk, 2004). The use of homology models based on other nuclear hormone receptor ligand-binding domains (LBDs) to interpret the binding of ecdysteroids and DBHs to the ecdysone receptor ligand-binding site have produced inconsistent results (Wurtz et al., 2000; Kasuya et al., 2003) and proved largely unsatisfactory. In this chapter we describe the ligand-binding properties of recombinant ecdysone receptor LBDs in free solution, both for a range of ecdysteroid compounds and for an insecticide of the DBH class of compounds, tebufenozide (RH5992). Interestingly, much of the order selectivity of the insecticides in the field is reproduced by the affinity of tebufenozide for the recombinant LBDs in the laboratory (Carmichael et al., 2005; Graham et al., 2007). We then describe the crystal structures of the LBDs of ecdysone receptors from the pest insects Heliothis virescens (Billas et al., 2003), Bemisia tabaci (Carmichael et al., 2005) and Tribolium castaneum (Iwema et al., 2007) in complex with ponasterone A (ponA), as well as in complex with 20E and BYI06830 (a DBH ligand) in the case of Heliothis virescens EcR/USP. Comparison of ecdysteroid- and BYI06830-bound structures of the H. virescens protein illustrates the way in which this remarkable protein can adapt its binding pocket to very different ligand chemistries. Finally, comparison of the ligand-binding pockets of H. virescens, B. tabaci and T. castaneum ecdysone receptors begins to provide insights at an atomic level of detail into the selectivity of the DBH insecticides.

13.2

Ligand-Binding Properties of Recombinant EcR/USP LBDs

Equilibrium binding experiments have been performed to assess the ability of various ligands to compete with [3H]-ponA for binding to recombinant ecdysone receptor LBD heterodimers (Carmichael et al., 2005; Graham et al., 2007). (The cloning of ecdysone receptors and expression of recombinant LBD heterodimers is dealt with in Chapter 19 in this book.) These experiments reveal that the binding affinity of the DBH ligand tebufenozide for recombinant heterodimers varies greatly (>105-fold) across taxonomic orders (Table 13.1) and reflects the lepidopteran-selective toxicity

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Table 13.1 Dissociation constants for ligands Species L. cuprina M. persicae LBD DE/F DE/F

B. tabaci DE

H. armigera DE/F E/F

87 1.0 0.1 n.d.b n.d. 170

240 4.8 5.3 n.d. n.d. >448,000

66 13 2.3 n.d. n.d. 3.7

20Ea Ponasterone A Muristerone A Inokosterone A Inoko A-fluor Tebufenozide

93 0.5 30 65 20–40 >132,000

290 58 10 n.d. n.d. 16

The table shows mean Ki values (nM) for ligands obtained by competitive inhibition of [3H]ponasterone A binding (Reproduced from Graham et al., 2007. With permission of Elsevier). a Chemical abbreviations: 20E, 20-hydroxyecdysone; Inoko A-fluor, inokosterone A conjugated to a fluorescent moiety (Chapter 19 of this book). b Not determined.

of this insecticide (a topic discussed further in Chapter 19 in this book). The Ki values for the ecdysteroid ligands studied vary less widely across orders, with the Ki for 20E varying least, presumably reflecting a physiological requirement for this ecdysteroid to bind effectively to most, if not all, insect ecdysone receptors. Accordingly, the titration curves for 20E (Fig. 13.1) show relatively little variation across the four species examined, whereas curves for some of the other ligands (e.g. muristerone A and tebufenozide) vary greatly from one species to the next. Such differences likely reflect sequence-dependent variations in the topography of the ligand-binding pocket at residues other than those critical for the binding of 20E, as even small changes in the position or nature of residues that contact a particular ligand could have a large impact on its binding affinity. Muristerone A binds better than ponA to the L. cuprina and H. armigera DE/F heterodimers, whereas the reverse is true with the equivalent heterodimers from M. persicae and B. tabaci. Since the binding affinities of L. cuprina and M. persicae for muristerone A (which bears C5 and C11 hydroxyl groups) differ by 300-fold, while the corresponding affinities for ponA and 20E (which do not carry C5 and C11 substituents) differ by less than twofold, it is likely that the ligand-binding pockets of these species exhibit steric and/or electrostatic variations in the immediate vicinity of the bound ecdysteroid C5 and C11 positions. Three-dimensional structures of ecdysone receptor LBDs co-crystallised with different ligands should shed light on the atomic basis for binding selectivity and the remarkable ability of ecdysone receptors to adopt a number of robust conformations, each capable of binding a distinct chemical family.

13.3

Structures of the Ecdysone Receptors in Complex with Ecdysteroids

To date, three crystal structures of ecdysone receptor LBDs in complex with the high-affinity phyto-ecdysteroid ponA have been determined (Billas et al., 2003; Carmichael et al., 2005; Iwema et al., 2007), these being from the moth

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Fig. 13.1 Competitive inhibition of [3H]-ponA binding to recombinant ecdysone receptor LBD heterodimers from four insect species. The competing ligands are 20E (chequered squares), ponA (filled circles), muristerone A (crosses), tebufenozide (filled triangles), shown with interpolated titration curves (solid lines). The curves for 20E have been labelled ‘20E’ for clarity. Ligand-free control data are also shown (open circles, dotted lines). Vertical axes show [3H]-ponA binding expressed as a percentage of that obtained without competing ligand, with s.e.m. shown as error bars (n = 2–4). Horizontal axes show the concentration of the competing ligand (nM) used with each DE/F heterodimer. For H. armigera, this axis also represents 0.3x the concentration of competing ligand used with the E/F heterodimer (the resulting horizontal displacement compensates for the poorer binding obtained with the E/F heterodimer and renders the E/F and DE/F curves coincident). Experimental details are described in Graham et al. (2007)

H. virescens (Lepidoptera), the whitefly B. tabaci (Hemiptera) and the beetle T. castaneum (Coleoptera). The structure of the H. virescens EcR/USP LBD in complex with the natural ecdysteroid hormone (20E) has recently also been reported (Browning et al., 2007). The overall heterodimeric arrangement of these EcR/USP LBDs is closely similar, with each LBD subunit exhibiting, as anticipated, the

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canonical nuclear receptor LBD fold (a three-layer anti-parallel α-helical sandwich with a small β-sheet between the fifth and sixth helices). We now discuss the main structural features observed for each subunit of the ecdysone receptor LBD as well as for the heterodimer as a whole, followed by a comparison of the ligand-binding pockets of the EcR/USP LBDs with a view to understanding the structural basis of ligand selectivity.

13.3.1

Structure of the USP Subunit

Sequence analysis of USP from various arthropods (Bonneton et al., 2003, 2006) reveals a distinct evolutionary diversification of lepidopteran and dipteran USP sequences with respect to those of other orders. In particular the lepidopteran and dipteran USP sequences are characterized by (i) a high degree of conservation of the loop (L1-3) connecting helices H1 and H3 (where we have denoted the 12 consecutive helices within the nuclear receptor LBD fold H1 through to H12 respectively) and (ii) a divergent C-terminal region encompassing the activation helix H12. USP sequences from other insect orders are closer to mammalian retinoid X receptor (RXR) sequences, with a high degree of conservation of those residues that in RXR interact with ligands. This suggests that these USPs may bind and thus be activated by RXR ligands, such as 9-cis retinoic acid. The USP crystal structures determined thus far reflect their sequence-based classification, with the H. virescens (Billas et al., 2001) and D. melanogaster (Clayton et al., 2001) USP structures (representing lepidopteran and dipteran insects, respectively) being distinct from those of B. tabaci (Carmichael et al., 2005) and T. castaneum (Iwema et al., 2007). The lepidopteran and dipteran USP structures contain a phospholipid molecule in the ligand-binding pocket, while those of non-lepidopteran/dipteran insects are found in an unliganded state. The activation helix H12 is in the so-called “antagonist” conformation (Brzozowski et al., 1997) in both types. The different liganded states correlate with structural differences in the inter-helical loops in the immediate vicinity of the pocket (Fig. 13.2a). For the phospholipid-bound USP, the loop L1-3 adopts a peculiar conformation, crossing over helix H3 and interacting with helix H3 and loop L11-12 which connects helices H11 and H12 through the residues conserved in the dipteran/lepidopteran protein. The observed placement of loop L1-3 prevents helix H12 from adopting the canonical agonist conformation and locks it in an antagonist conformation. The phospholipid molecule plays the role of a structural ligand by stabilizing the position of helix H3 that, as a consequence, is more linear in conformation than is its counterpart in human or mollusc RXR (Egea et al., 2000; De Groot et al., 2005) or in the (unliganded) TcUSP and BtUSP structures. For the unliganded USPs, the apo conformation is stabilized by intra-molecular interactions between structural elements. In fact, the regions that would otherwise correspond to helices H6 and H11 are reorganized into loops (denoted L6 and L11; Fig. 13.2b) that fold onto the

Fig. 13.2 Two types of USP LBD structures. Depicted are the LBD structures of HvUSP (in blue) and TcUSP (in brown). (a) The activation helices H12 (in red and orange for HvUSP and TcUSP, respectively) adopt an antagonist conformation. The global fold of the two types of USP structures is conserved, but significant structural differences occur, consistent with the ligand-binding status of the domain (phospholipid-bound HvUSP and ligand-free TcUSP). The phospholipid bound to HvUSP is shown in a ball representation with atoms coloured as follows: carbon atoms magenta, oxygen atoms red and phosphorus atom orange. The ligand-binding pocket is shown as a light blue semi-transparent surface. Helices H6 and H11 of HvUSP are replaced by loops in TcUSP. An important shift in the positioning of helix H3, and to a lesser degree of helix H1, can be observed. (b) Enlarged view of the region corresponding to helices H6 and H11 of HvUSP that in the case of TcUSP are reorganized into two loops (L6 and L11, respectively) that are folded onto the surface the LBD, stabilizing the apo conformation. The colour scheme of the activation helices and the bound ligand atoms follows that of Panel (a) (See Color Plates)

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LBD surface, resulting in a considerably more compact structure in this region than is the case in HvUSP and DmUSP. Residues within these loops L6 (TcUSP-V286, V288, I291) and L11 (TcUSP-L382, F383) fill in the interior of the protein, leaving no room for any potential ligand. The placement of helix H3 and the stabilization of loops L6 and L11 are independent of the N-terminal part of the receptor LBD (helix H1 and part of loop L1-3) since, in BtUSP that has lost this amino-terminal region due to proteolytic degradation, the positioning and the conformation of all the other structural elements are identical to those observed in the TcUSP LBD. The question arises as to whether endogenous ligands exist for USP. The functional and biological role of the phospholipid found in the ligand-binding pocket of dipteran and lepidopteran USP is still unclear. This contrasts with the functional role determined for the phospholipid molecule that occupies the ligand-binding pocket of vertebrate NR5 orphan receptors SF-1 and LRH-1 (Krylova et al., 2005). Some experimental data suggest that juvenile hormones represent the USP ligands (Jones and Sharp, 1997; Jones et al., 2002; Xu et al., 2002), but this issue still needs in vivo verification. On the other hand, in vitro and in vivo data have demonstrated that RXR-like USPs, such as TcUSP, are not activated by and do not bind RXR ligands, in contrast to original expectations from the high conservation of the residues in USP with those in RXR that interact with ligand (Iwema et al., 2007). Furthermore, no interaction is found between TcUSP and peptides derived from mammalian and insect co-activators, suggesting that the positioning of the activation helix H12 in the antagonist conformation is stable. The structural data are thus consistent with the functional and phylogenetic data, and together they strongly suggest that TcUSP acts as a ligand-independent and constitutively silent partner of TcEcR.

13.3.2

Structure of the Ecdysteroid-Bound EcR Subunit

The initial nuclear receptor structure used for solving the structure of the ponAbound EcR/USP using molecular replacement was that of vitamin D receptor (VDR) (Rochel et al., 2000). Subsequent comparison of the crystal structures of EcR and VDR indicates that their LBDs indeed have a highly similar structure. Both contain 12 α-helices and 3 small anti-parallel β-strands. However, they differ in the conformation of the loops that respectively link helices H2 to H3 and helices H11 to H12, as well as in the conformation of the loop that links the second and third strands of the β-sheet. In addition, significant shifts of helices H6 and H7 with respect to the remaining secondary structure elements are observed. Taken together, these differences at the atomic level likely explain the rather unsatisfactory studies of ecdysteroid docking into a VDR-derived homology model of EcR (Wurtz et al., 2000). The crystal structures of the ponA-bound EcR of the three insect species determined so far are also highly similar (see Fig. 13.3a). However, two main points of variation exist. First, the loop connecting helices H9 and H10 (L9-10) of EcR is

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Fig. 13.3 The ligand-binding domains of EcR/USP. (a) The overall structures of the EcR/USP LBD heterodimer in complex with an ecdysteroid are depicted in two orientations for Heliothis virescens (Hv) and Tribolium castaneum (Tc), insect species belonging to the orders Lepidoptera and Coleoptera, respectively. The EcR/USP heterodimerization interface is conserved between the different species. However, differences are seen between that of the lepidopteran/dipteran EcR/ USP and those of other insects. Globally, the EcR/USP LBD heterodimer is more compact and more symmetric in lepidopteran and dipteran species (exemplified here by H. virescens) as compared to that of other insects (exemplified here by T. castaneum) as explained in the text. HvEcR and HvUSP are shown as dark green and blue ribbons respectively, and TcEcR and TcUSP as light green and brown ribbons respectively). The activation helices H12 of HvEcR and TcEcR are depicted as a red and orange helical ribbon, respectively. (b) Enlarged view of the L9-10 loop region of HvEcR and TcEcR LBDs in the context of their heterodimer with the corresponding USP LBD partner. The differences both in sequence (a two amino-acid insertion) and in conformation are indicated for the loop L9-10 of lepidopteran/dipteran EcRs (exemplified here by HvEcR) as compared to that of EcRs of other insects (exemplified here by TcEcR). Residues of loop L9-10 and a few residues of loop L7-8 that interact with this element are shown in stick representation, with carbon atoms in dark green and light green for HvEcR and TcEcR respectively, oxygen atoms in red and nitrogen atoms in blue. Yellow dotted lines indicate hydrogen bonds between residues of the protein and the colour scheme of the ribbons follows that of Panel (a) (See Color Plates)

noticeably different in H. virescens compared to its counterpart in B. tabaci and T. castaneum. In fact, the protein sequence of loop L9-10 in lepidopteran and dipteran EcRs is peculiar compared to other arthropods, in that it is less basic and contains a two amino acid insertion with a concomitantly different backbone

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conformation (Fig. 13.3b). Second, for TcEcR, the C-terminal region of helix H10 is in closer proximity to the loop which connects helices H6 and H7 than is the case in the two other structures.

13.3.3

Overall Structure of the Heterodimer EcR/USP LBDs

The EcR/USP LBD heterodimer interface is comprised of an intricate network of hydrophobic and polar interactions mediated in both partners by the helices H7, H9, H10 and the loop connecting helices H8 and H9. Differences exist between the interface of HvEcR/HvUSP and those of BtEcR/BtUSP and TcEcR/ TcUSP. In H. virescens EcR/USP, the heterodimeric arrangement is more compact than for B. tabaci and T. castaneum EcR/USP due to concomitant movements of structural elements from both partners. More precisely, the region of HvEcR comprising the C-terminal part of helix H5, the C-terminal part of the first β-sheet strand and helices H6 and H7 moves towards HvUSP by 1.5–2.5 Å. At the same time, the loop (L8-9) connecting helices H8 and H9 of HvUSP is displaced by about 2 Å towards the HvEcR partner with respect to the position adopted in BtUSP and TcUSP (Fig. 13.3a). Interestingly, the interface observed for B. tabaci and T. castaneum EcR/USP is similar to the dimerization interface of vertebrate nuclear receptor heterodimers (Bourguet et al., 2000; Gampe, Jr. et al., 2000; Svensson et al., 2003; Xu et al., 2004; Shan et al., 2004; Suino et al., 2004). In these two cases, USP is rotated by ~10° from the C2 symmetry axis, resulting in an asymmetrical arrangement of the two subunits. A similar arrangement is seen for RXR in the case of vertebrate heterodimers. A consequence is the dissimilar distances separating helix H7 of one subunit and the loop L8-9 belonging to the other subunit (i.e., the distance between USP-H7 and EcR-L8-9 is smaller than the distance between EcR-H7 and USP-L8-9). For H. virescens EcR/USP, because of the structural displacement of the EcR region comprising helices H6 and H7, the asymmetry is strongly attenuated, resulting in an arrangement more reminiscent of the homodimeric vertebrate estrogen receptor (Brzozowski et al., 1997). Furthermore, two additional features present in H. virescens but absent in B. tabaci and T. castaneum EcR/USP heterodimers contribute to a larger dimerization interaction surface. The first is the insertion between helix H5 and the β-sheet in HvUSP, with the electron density for this loop suggesting possible contacts with the loop L8-9 of the EcR partner. Since loop L8-9 represents an essential element of the heterodimer interface, this helix H5-β-sheet insertion, characteristic of lepidopteran and dipteran USPs, is likely to play an additional yet unknown regulatory role. The second structural feature is the loop L9-10 of EcR which, as described above, is noticeably different in HvEcR (with a two amino acid insertion) compared to its counterparts in BtEcR and TcEcR. In the case of B. tabaci and T. castaneum EcRs, the loop L9-10 is stabilized by intra-molecular interactions (BtEcR-K366 interacting with BtEcRE324 (within helix H8) and TcEcR-R496 interacting with both TcEcR-R450

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(within loop L7-8) and TcEcR-E455 (within helix H8). On the other hand, in H. virescens EcR, the loop L9-10 is closer to the heterodimerization partner (as shown in Fig. 13.3b) and in particular to HvUSP-R404 (within helix H9).

13.3.4

Ligand-Binding Pocket of EcR

The ligand-binding pocket of EcR observed in the complexes with the steroidal hormones ponA and 20E has a J-shaped architecture that extends from helix H12 to helix H5 and the β-sheet, and is completely buried inside the receptor (Fig. 13.4). In B. tabaci EcR, the steroidal ligand occupies the major region of the pocket (Carmichael et al., 2005) with its A/B rings located in the proximity of the β-sheet and the alkyl side chain oriented towards helix H12. Additionally, for BtEcR and TcEcR, a smaller curved pocket (the ‘ancillary’ pocket) can be observed in the region between helices H1 and H8, close to the 2β- and 3β-OH groups of the steroid ligand. The opening to this intrusion into the protein atomic volume is intimately linked to the conformation of the side chains of two residues, in BtEcR these being E199 (within helix H1) and Y325 (within helix H8), and in TcEcR E330 and Y456 (located within the same respective helices). In HvEcR the entrance to the cavity corresponding to the ancillary pocket of BtEcR is closed by the different conformation of the side chains of the corresponding residues (E309 and Y437), which eclipse the volume between helices H1 and H8 (Fig. 13.5a). Note that for the 20E-bound HvEcR, a slightly different side chain conformation of E309 coincides with the presence of a water molecule in this region. For TcEcR, the side chain conformations E330 and Y456 do not occlude the entrance to the ancillary pocket, which is occupied by three water molecules (Fig. 13.5b). The reasons for the existence of the ancillary pocket volume are not clear, but the relatively loose packing of protein elements in this vicinity may reflect structural flexibility (Togashi et al., 2005) associated with accommodation of the incoming ligand. Nineteen residues are located at less than 4.5 Å from ponA, and six hydrogen bonds are formed between ponA and EcR, with identical interacting residues in all three species (E309, T343, T346, R383, A398, Y408 according to the HvEcR sequence corresponding the BtEcR residues E199, T231, T234, R271, A286 and Y296 depicted in Fig. 13.4). Where the hydrogen bonds involve side-chain atoms, the rotameric conformations are effectively the same in all three species and the contributing residues are completely conserved across insect orders. Interestingly, in the case of ponA- and 20E-bound HvEcR structures, the side chain amino group of N504 makes an electrostatic interaction with the aromatic ring of the strictly conserved W526 (located in helix H12; corresponding to BtEcR-W412 in Fig. 13.4). Non-polar interactions also take place between the ligand and residue side chains. Of the nineteen residues located at less than 4.5 Å from ponA, four are differentially conserved in Lepidoptera, i.e. strictly conserved in EcR sequences of lepidopteran insects but replaced by other residues in sequences of other arthropod orders. These are, according to HvEcR sequence, M342 (replaced by valine or isoleucine), V384

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Fig. 13.4 Opened-out view of the major and ancillary pockets of BtEcR LBD. In these Panels the pocket surface (grey) has been open out into two “shells” by slicing in a plane parallel to that of the page. Residues lying beyond the plane of slicing are shown in each Panel, with carbon atoms in orange if they highly conserved across all insect species, otherwise with carbon atoms in green. PonA, bound in the major pocket of the LBD, is shown in stick representation with carbon atoms shown in black. Oxygen atoms in both PonA and BtEcR are shown in red and nitrogen atoms in blue. Red dotted lines indicate hydrogen bonds between the protein and the ligand. For clarity, side chain or main chain groups of individual residues are omitted from the Figure where these are not involved in the formation of the pocket surface (See Color Plates)

(replaced by methionine, glycine or alanine), V395 (replaced by isoleucine), V416 (replaced by threonine, asparagine or serine). The remaining residues that make van der Waals’ interactions with the ligand are generally well conserved across species. In the crystal structures of BtEcR, TcEcR and HvEcR, these latter residues

Fig. 13.5 Residues in the region of the ancillary pocket of the EcR LBD. (a) The existence of an ancillary pocket, as observed in the BtEcR LBD, is intimately linked to the conformation of the side chains of residues (in particular HvEcR-E309/BtEcR-E199 and HvEcR-Y437/BtEcR-Y325) that fill or do not fill the volume between helices H1 and H8. HvEcR and BtEcR LBDs are depicted as dark green and yellow ribbons, respectively. Residues are shown in a stick representation, with carbon atoms in grey and yellow for HvEcR and BtEcR, respectively, oxygen atoms in red and nitrogen atoms in blue. The ligandbinding pocket of HvEcR and TcEcR are represented by a blue and yellow translucent molecular surface, respectively. In HvEcR the entrance to the cavity corresponding to the ancillary pocket of BtEcR is occluded by the side chains of HvEcR-E309 and HvEcR-Y437. (b) In the TcEcR LBD, the volume corresponding to the ancillary pocket of BtEcR is occupied by water molecules located between helices H1 and H8 and is not occluded by side chain atoms. The ligand-binding pocket was calculated without the three water molecules and is represented by a light green translucent molecular surface. The TcEcR LBD is depicted as a light green ribbon. The residues E330 and Y456 and the bound ponA ligand are shown in a stick representation with carbon atoms in green, oxygen atoms in red and nitrogen atoms in blue. The three water molecules are represented by red balls (See Color Plates)

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exhibit similar rotameric conformations and make similar area contributions to the pocket surface. In the case of HvEcR in complex with 20E, an additional hydrogen bond is formed compared to what is observed for the ponA-bound EcR protein complexes (Browning et al., 2007) – the hydroxyl group of 20E at the C25-position that is absent in ponA is hydrogen-bonded to the side chain amide group of Asn504 (located in helix H11). Remarkably, a water-mediated interaction network in this region bridges together the 20-, 22-, 25-OH groups and protein residues from helices H7, H10 and H11 (Fig. 13.6a). No water molecules were reported in the structure of HvEcR and BtEcR bound to ponA. However, re-examination of the ponA-bound HvEcR structure reveals a small peak in the difference electron density map close to Asn504, suggesting the possible existence of a water molecule at this location. In the case of TcEcR, several water molecules are observed in the electron density, and these form a tight interaction network bridging together the hydroxyl groups of the ponA alkyl chain with residues of helices H7 and H10-H11 in a similar fashion to that observed for the 20E-bound HvEcR structure (Fig. 13.6b). The differential conservation of the four above-mentioned pocket residues in Lepidoptera (M342, V384, V395 and V416 in HvEcR) have the potential to lead to a different pocket surface topography in HvEcR compared to that in BtEcR and TcEcR. We now examine each of these in turn. Inspection shows that replacement of HvEcR-V395 (located in the β-sheet) by isoleucine in BtEcR and TcEcR has a negligible effect on the topography of the pocket surface. However, the differential conservation of the remaining three residues has the following consequences. The presence of HvEcR-M342 (located in helix H3) instead of Ile230 in BtEcR and Ile361 in TcEcR results in an approximately 1 Å deeper intrusion of the pocket into the protein atomic volume of the HvEcR LBD adjacent to the C4 atom of the ponA ligand (Fig. 13.7a,b,d). Together with the presence of this more bulky residue, the polypeptide backbone in the vicinity of HvEcR-M342 is shifted approximately 1.3 Å towards the ecdysteroid C-ring as compared to its location for BtEcR and TcEcR, resulting in a noticeable change in the pocket geometry in this region. At the opposite side of the pocket, HvEcR-Val384 (in helix H5) is replaced by a methionine residue in both BtEcR (M272) and TcEcR (M403). Concomitant with this change in residue type is a change in the side-chain orientation of the neighbouring leucine residue located in helix H7 (HvEcR-L420, BtEcR-L308, TcEcR-L439). In fact, for HvEcR, the side-chain of L420 is oriented towards the side chain of V384 (in helix H5), while for BtEcR and TcEcR, the orientation of the corresponding leucine side chain is away from that of BtEcR-M272 and TcEcR-M403, respectively (Figs. 13.7a,b,d and 13.8a). The consequence of these differences is the closer proximity of the leucine to the ligand 22-OH group for HvEcR as compared to what is observed in the BtEcR and TcEcR structures (3.3 Å between ponA- or 20E-O22 and HvEcR-L420-Cδ1 as compared to 3.9 Å for the corresponding distance in BtEcR and TcEcR). Furthermore, the specific orientation of HvEcR-L420 contributes to a wider opening between helices H7 and H10, a region that is very sensitive and exhibits a large degree of adaptability in the binding of DBH compounds. The last differentially

Fig. 13.6 Stereoviews of water-mediated interaction network between the ecdysteroid and EcR residues. (a) In the case of 20E bound to the HvEcR LBD, the hydrogen-bond interaction network involves the 20-, 22-, and 25-hydroxyl groups of the 20E alkyl side-chain and three structural water molecules. (b) In the case of ponA bound to the TcEcR LBD, the interaction network involves the 20- and 22-hydroxyl groups of ponA alkyl side chain and up to five structural water molecules. The HvEcR and TcEcR LBDs are depicted as dark green and light green ribbon, respectively, with individual atoms within the LBD and ligand shown in stick representation and with carbon atoms coloured in light blue and green, respectively, oxygen atoms in red, nitrogen atoms in blue and sulphur atoms in yellow. The water molecules are represented by red balls. Hydrogen bonds are indicated by yellow and pink dotted lines in (a) and (b) respectively (See Color Plates)

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Fig. 13.7 Inter-species comparison of side-chain packing in walls of the EcR ligand-binding pocket. (a) B. tabaci EcR with ponA bound, (b) H. virescens EcR with ponA bound, (c) H. virescens EcR with BYI06830 bound and (d) T. castaneum EcR with ponA bound. The orientations of the EcR-LBDs in Panels (a–d) are identical, with the dashed line indicating a common line of in-plane register. Panel (a) shows (in rod representation) the side chains of B. tabaci residues I230, M272, T304, L308, M389, T393 and V404, whilst Panels (b) and (c) show their counterparts of these residues in H. virescens EcR (namely M342, V384, V416, L420, Q503, M507 and L518, respectively), and Panel (d) their counterparts in T. castaneum (namely I361, M403, T435, L439, Q520, M524 and L535, respectively). Atoms in these residues are coloured as follows: carbon atoms green, oxygen atoms red, nitrogen atoms blue and sulphur atoms yellow, with the position of the Cα atom of each residue being highlighted by a sphere for clarity. Also shown in each panel is the ligand-binding pocket (coloured transparent gold and calculated with a 1.2 Å probe radius), encapsulating the respective bound ligand. Ligand atoms are coloured according to the same scheme as that of the EcR side chains, except that carbon atoms are shown in black. Panels (e), (f), (g) and (h) show the atomic packing of the ligand-binding cavity wall in the vicinity of the residues highlighted to the left of the ligand in Panels (a–d) respectively, but viewed from the interior of the binding pocket and looking in the direction of the arrows. Atoms are shown in CPK representations and include both main and side chain atoms from all residues contributing to the immediate wall of the binding pocket in the direction of the view. The site of the protrusion into the protein atomic volume of the ligand-binding pocket walls of H. virescens EcR and T. castaneum EcR in the vicinity of the alkyl tail of bound ponA (see text) is indicated by an asterisk in Panels (b), (d), (f) and (h). The site of deeper protrusion into the protein atomic volume of ponA-bound HvEcR LBD in the vicinity of the C4 atom of the ligand is indicated by a solid star in Panel (b); this deeper protrusion with respect to the pocket walls of ponA-bound BtEcR LBD and TcEcR LBD is a consequence of the substitution of BtEcR I230 and TcEcR I361 by methionine in HvEcR (M342, see text) (See Color Plates)

conserved residue to be considered is HvEcR-V416 located in helix H7, and replaced by a threonine residue in BtEcR (T304) and TcEcR (T435). Since helix H7 of HvEcR is shifted (by 1.5–2.5 Å) towards the interior of the receptor compared to its counterpart in BtEcR and TcEcR, the side chain of V416 is rather close to the 22-OH group of ponA or 20E (3.3 Å between ponA- or 20E-O22 and HvEcR-V416-Cγ1) (Fig. 13.8b). On the other hand, the polar threonine residue is

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Fig. 13.8 Stereoviews of the differential conservation of the pocket residues HvEcR-V384 and HvEcR-V416 in Lepidoptera. (a) HvEcR-V384, which is replaced by methionine in non-lepidopteran EcRs, allows the specific side chain conformation of HvEcR-L420 and a closer van der Waals’ interaction with the ecdysteroid 22-hydroxyl group. (b) HvEcR-V416, which is replaced by threonine in TcEcR and BtEcR, is also closer to the ecdysteroid 22-hydroxyl group due to the concomitant shift of helix H7 towards the interior of the receptor pocket. The stereoviews depict HvEcR in blue, TcEcR in green and BtEcR in yellow. Amino acid residues of HvEcR, TcEcR and BtEcR are shown in stick representation colour with carbon atoms in light blue, green and yellow for the three species respectively and oxygen atoms in red. The ligands 20E (for HvEcR) and ponA (for TcEcR and BtEcR) are shown in a stick representation with carbon atoms in blue (20E), green (ponA in TcEcR) or wheat (ponA in BtEcR) and oxygen atoms in red. The activation helix H12 is also shown as a red ribbon (See Color Plates)

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located much further away from ponA in BtEcR and TcEcR (5.4 Å between ponAO22 and BtEcR-T304-Oγ1 and 5.3 Å between ponA-O22 and TcEcR-T435-Oγ1, respectively). In summary, the high level of conservation of residues that interact with 20E (and with its close homologue, ponA) and the preservation of side chain rotameric conformation from one species to the next, are likely to underpin the relative invariance of the competitive binding curves exhibited by the natural hormone across three taxonomic orders (Fig. 13.1). However, residues not directly involved in interactions with 20E (and with the features that it shares with ponA) are less tightly conserved, leading to local differences in pocket architecture, particularly in the vicinity of the ancillary pocket and in the vicinity of the alkyl chain of ponA. As will be seen, it is these residues that likely contribute to the species- and order-dependent differences in the binding affinity of ecdysone receptors for other ecdysteroid ligands and for ligands of other chemistries.

13.4

Structure of the Ecdysone Receptor in Complex with a Synthetic DBH Insecticide Ligand

Insecticide ligands of the dibenzoylhydrazine class of compounds (e.g. tebufenozide, Table 13.1) exhibit a high binding affinity for EcR/USP heterodimers of lepidopteran insects, while their affinity is lower for coleopteran and dipteran species and often negligible for the members of other orders studied to date. The crystal structure of the recombinant EcR/USP heterodimer from the moth H. virescens purified and crystallized in complex with a high-affinity DBH ligand (BYI06830) provides insight into the reasons behind this differential affinity. While the global heterodimeric arrangement of the EcR and USP LBDs is identical in this case to that observed for the ponA- and 20E-bound HvEcR/HvUSP LBD, the structure of BYI06830-bound EcR LBD is remarkably different to that of the ecdysteroidbound protein. The differences occur primarily in the region encompassing helix H6, helix H7, the β-sheet and the loop between helices H1 and H3 (Fig. 13.9a). The LBD accommodates the binding of the DBH ligand by opening up a cleft between helices H7 and H10, with the ligand situated at the end of the ecdysteroidbinding pocket in proximity to helices H7, H10, H11 and H12. The superimposition of the ecdysteroid- and the DBH-bound EcR LBD structures indicates that the tert-butyl group and A-ring of the DBH superimpose on the alkyl chain of the ecdysteroid. The region of the ecdysteroid-binding pocket formed by helix H1, helix H6 and the β-sheet is not occupied by the DBH ligand, with the β-sheet instead disrupted dramatically by the formation of interactions that lead to a loop structure between the residues in the erstwhile second and third β-strands. This structural rearrangement is the consequence of the inward motion of side chains of two aromatic residues (F397 and Y403), located on each side of the β-sheet, that fill in the void created by the absent ecdysteroid component in this region. Similarly, the loss of stabilizing interactions between the ligand and residues of H1 and of

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Fig. 13.9 Flexible region in the EcR LBD exploited by DBH compounds. (a) Ribbon diagram showing the superimposition of the structures of HvEcR-LBD in complex with 20E (dark green ribbons) and with BYI06830 (orange ribbons). The view is restricted to the region differing the most between the two EcR-LBDs that includes helices H2, H6, H7 and the β-sheet. 20E and BYI06830 are shown in stick representation with carbon atoms in cyan and light grey, respectively, oxygen atoms in red and nitrogen atoms in blue. (b) Superimposition of 20E and BYI06830 bound to HvEcR. Oxygen atoms are shown in red, nitrogen atoms in blue, 20E carbon atoms in cyan and BYI06830 carbon atoms in olive. The three structural water molecules observed in the structure of 20E-bound HvEcR, shown by red balls, superimpose well with the region of the B-ring of the DBH compound (See Color Plates)

the loop connecting H1 to H2 (as observed in the ponA- or 20E-bound EcR LBD structures) leads to the unwinding of that helix H2. These structural differences between the ecdysteroid-bound and the DBHbound receptor (Billas et al., 2003) are consistent with mutational data that also suggested distinct conformational forms (Kumar et al., 2002, 2004). Key residues that likely interact with ecdysteroids or DBH compounds were mutated, and the

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in vitro effects on receptor transcriptional activity upon induction with the two types of ligands were assessed using mammalian or insect cells. Mutation of the two residues that stabilize the co-activator AF2 helix by π-cation interaction, W526 and N504, results in a drastic reduction of transcriptional activity. These results agree with observations of the dominant negative phenotype of the DmEcR-W650A mutant (corresponding to HvEcR-W526A), for which neither ligand binding nor transcriptional activation are observed (Hu et al., 2003; Cherbas et al., 2003). Furthermore, the HvEcR-LBD structural data are consistent with functional studies of the lepidopteran Choristoneura fumiferana (Cf) EcR single mutants, in particular with the ecdysteroid-selective mutants R378A, F392A and A393P (R383A, F397A and A398P for HvEcR) (Kumar et al., 2002). The CfEcR-A393P mutant is particularly interesting for its critical role in discriminating between steroidal and nonsteroidal ligands. For the ponA-bound HvEcR-LBD, A398 lies in the connection between the second and third β-strands and in close contact with the ligand, while for BYI06830-bound HvEcR-LBD, the β-sheet region is not involved in ligand recognition. Mutation of A398 to proline introduces local distortions of the β-sheet backbone, which could explain the complete loss of activity for the steroidal compared to the non-steroidal ligand. It is interesting to notice that mutation of CfEcR-V411 (within helix H7, corresponding to HvEcR-V416) to phenylalanine or tyrosine completely abolishes the binding of ecdysteroids and dibenzoylhydrazine compounds (Kumar et al., 2004). Both crystal structures show the proximity of this residue to ponA and BYI06830, and its mutation to a bulkier residue can readily explain the inability of the mutant EcR to bind either type of ligand. On the other hand, the mutants CfEcR-V411F and CfEcR-V411Y, but not wild-type CfEcR, respond to the THQ class of compounds. However, the explanation for this behavior remains speculative, and may involve π-stacking interactions with the aromatic rings of phenylalanine or tyrosine, entropic effects, or the result of a different binding pocket geometry being adopted when EcR is in complex with THQ ligands.

13.5

Discussion and General Conclusions

Several crystal structures of the ecdysone receptors of various insect species in complex with ecdysteroids and, in the case of the moth H. virescens, with a DBH insecticidal compound, have been determined in the last few years. These structures reveal that all ecdysone receptors have virtually the same three-dimensional structure when complexed with ecdysteroid. However, it is known that there are two types of ecdysone receptors – those that bind and adapt to both ecdysteroids and synthetic DBH compounds (exemplified by the lepidopteran H. virescens EcR), and those that bind synthetic DBH insecticide compounds poorly or not at all (exemplified by hemipteran B. tabaci EcR). In the case of the coleopteran ecdysone receptors, such as T. castaneum EcR, the binding of DBH compounds is limited to certain examples such as halofenozide (Dhadialla et al., 1998). No crystal structure of TcEcR bound to this molecule has yet been determined.

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Both types of ecdysone receptors exhibit the same three-dimensional structure when bound to an ecdysteroid, likely reflecting the critical role of 20E, and possibly closely related hormones, across the phylum Arthropoda. However, a detailed structural analysis of the different EcR LBDs reveals significant differences at the atomic level. First, in addition to the ‘major’ pocket occupied by the ecdysteroid ligand, an ‘ancillary’ pocket in the region between helices H1 and H8 was detected for B. tabaci EcR (Fig. 13.5a). For H. virescens EcR, the entrance to this region is filled by residue side chains, while for T. castaneum EcR the space is occupied by water molecules (Fig. 13.5b). The question thus arises whether the ancillary pocket plays a role in the differential capacity of EcR to bind agonists with non-ecdysteroid chemistries (such as the DBH compounds). However, since the existence of the ancillary pocket is intimately linked to the conformation of the side chains of two highly conserved residues, it is rather unlikely that the existence/absence of an ancillary pocket plays a key role in differential receptor adaptability. Further EcR structures from other organisms in correlation with functional assays (binding studies on EcR LBDs containing point mutations, for example) would be needed to for a better understanding of this issue. The second difference concerns shape variations of regions of the ligand-binding pocket that are not in direct (e.g., electrostatic or van der Waals’) contact with the ecdysteroid. In fact, while the topography of those parts of the pocket that are within van der Waals’ distance of ponA are closely similar (Fig. 13.7), the topography of the those parts of the pocket that are not in direct contact with the ecdysteroid show significant evolutionary variation (Carmichael et al., 2005). Of particular note is the region located near the end of the alkyl side chain of ponA or 20E, in close proximity to helices H7 and H10. This region contains two of the four residues that are differentially conserved in Lepidoptera (HvV384 and HvEcR-V416). As shown in Fig. 13.7a,e, residues forming the pocket wall of BtEcR at this location are densely packed, closing the channel between helices H7 and H10. In contrast, in HvEcR, residues in this vicinity are considerably less tightly packed, with a protrusion extending into the protein interior (Figs. 13.7b,f). In particular, the side chains of HvEcR-L420 (in helix H7) and HvEcR-Q503 (in helix H10) are oriented away from each other, in contrast to the side chains of their BtEcR counterparts (L308 and M389 respectively). It is precisely within this channel between helices H7 and H10 that the 1,4-dioxan ring of BYI06830 is accommodated (Figs. 13.7c,g and 13.9b), an incursion probably facilitated by the lower atomic packing density in the protein at this point. For the 20E-bound HvEcR complex, three water molecules were identified in this region, with similar water molecules subsequently being identified in the TcEcR structure. A superimposition of the HvEcR structures bound to 20E and to BYI06830 reveals that the water molecules are located at the place of the 1,4-dioxan ring (Fig. 13.9b). More detailed analysis would be required to tease out reasons for the intermediate sensitivity of TcEcR towards DBH compounds, but preliminary inspection of this region of the ligand-binding pocket (Fig. 13.7d,h) suggests that it is intermediate in structure to that of HvEcR and that of BtEcR. The structural reorganization of the receptor LBD upon binding the DBH compound is remarkable and extreme (Fig. 13.9a). Nuclear receptor adaptation to

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various types of ligands was also observed for the human liver X receptor LXR (Färnegårdh et al., 2003) and for the estrogen receptor (Nettles et al., 2007). In these cases, the ligand-binding pockets of the receptors were shown to be flexible and capable of accommodating different types of ligands. However, unlike EcR, the different structures of the receptor bound to the various molecules are virtually identical and do not result in radically different cavities with different residues implicated in ligand recognition. In fact, for EcR, the binding of synthetic insecticide compounds to the H. virescens ecdysone receptor involves adaptation mechanisms requiring the inner-to-outer switch of two aromatic residues located in the region of the β-sheet, and the widening and restructuring of channel between helices H7 and H10. A better understanding of entropic and desolvation effects would provide additional insight into the mechanisms that underpin the structural adaptation of the EcR LBD to different types of ligand. Notice that since EcR must heterodimerize with USP for high-affinity ligand binding (Yao et al., 1993), the role of the USP partner in the binding of ecdysone agonists to EcR should be evaluated in the context of ligand-binding specificity, for example by considering EcR/USP cross-species hybrid heterodimers. Finally, we would like return to Ashburner’s experimental observations on the ecdysone puffing response of D. melanogaster polytene chromosomes over 30 years ago. The atomic structures of receptor-ecdysteroid complexes now offer a basis for the apparent protection of the ‘hypothetical ecdysone receptor’ against reaction with N-ethylmaleimide by pre-incubation with increasing concentrations of 20E. Examination of the B. tabaci ligand-binding pocket containing hormone (Fig. 13.10) reveals the close proximity of the S atom in BtEcR-Cys394 (within helix H11) to atoms C26 and C27 of the steroid (~4.0 Å centre-to-centre distance; 0.5 Å between van der Waals surfaces). Clearly, once hormone is in place in the ligand binding pocket, the N-ethyl-substituted five-membered ring of the thiol-specific reagent will experience

Fig. 13.10 The steric relationship between the alkyl side chain of ponA in the binding pocket of the B. tabaci ecdysone receptor LBD and the side chain sulphydryl group of BtEcR C394. The distance between the van der Waals’ surfaces of the terminal side chain carbon atom of the steroid and the sulphur atom of the cysteine residue C394 in the pocket wall is 0.5 Å. This cysteine residue is strictly conserved among all known EcR sequences (See Color Plates)

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considerable steric impediment to reaction with Cys-394. The residue BtEcR-C394 in helix H11 is in fact strictly conserved across all known EcR sequences. Acknowledgments We thank the Structural Biology and Genomics Platform at IGBMC for their help at various stages of this work; V. Chavant (IGBMC) for technical assistance. The work was supported in part by Bayer CropScience (Monheim, Germany), the Association pour la Recherche sur le Cancer and by the European Commission SPINE2 complexes (Contract N° LSHG-CT2006-031220) under the Integrated Programme “Quality of Life and Management of Living Resources”. We thank the fermentation group at CSIRO Molecular and Health Technologies (Parkville, Australia) for their steady production of recombinant protein over the years. Support from the AusIndustry R & D Start Grant Programme is gratefully acknowledged. Molecular graphics figures were generated using combinations of the following software packages: VOIDOO (Kleywegt and Jones, 1994a), DINO (Philippsen, 2000) and POVScript+ (Fenn et al., 2003) and PyMOL Molecular Graphics System (DeLano Scientific, San Carlos, CA; www. pymol.org). ‘LSQMAN’ option of O (Uppsala Software Factory, G.J. Kleywegt) was used for the superimposition of the different structures (Kleywegt and Jones, 1994b). Atomic coordinates were obtained from the Protein Data Bank entries 1R1K, 1R20, 1Z5X, 2NXX and 2R40.

References Ashburner, M. (1972). N-ethylmaleimide inhibition of the induction of gene activity by the hormone ecdysone. FEBS Lett. 22, 265–269. Ashburner, M., Chiara, C., Meltzer, P., and Richards, G. (1973). Temporal control of puffing activity in polytene chromosomes. Cold Spring Harb. Symp. Quant. Biol. XXXVIII, 655–662. Becker, H.J. (1962). Die Puffs der Speicheldrusenchromosomen von Drosophila melanogaster. II. Die Auslosung der Puffbildung, ihre Spezifitat und ihre Beziehung zur Funktion der Ringdruse. Chromosoma (Berlin) 13, 341–384. Bender, M., Imam, F.B., Talbot, W.S., Ganetzsky, B., and Hogness, D.S. (1997). Drosophila ecdysone receptor mutations reveal functional differences among receptor isoforms. Cell 91, 777–788. Billas, I.M.L., Moulinier, L., Rochel, N., and Billas, I.M. (2001). Crystal structure of the ligandbinding domain of the ultraspiracle protein USP, the ortholog of retinoid X receptors in insects. J. Biol. Chem. 276, 7465–7474. Billas, I.M.L., Iwema, T., Garnier, J.M., Mitschler, A., Rochel, N., and Moras, D. (2003). Structural adaptability in the ligand-binding pocket of the ecdysone hormone receptor. Nature 426, 91–96. Bonneton, F., Zelus, D., Iwema, T., Robinson-Rechavi, M., and Laudet, V. (2003). Rapid divergence of the ecdysone receptor in Diptera and Lepidoptera suggests coevolution between ECR and USP-RXR. Mol. Biol. Evol. 20, 541–553. Bonneton, F., Brunet, F.G., Kathirithamby, J., and Laudet, V. (2006). The rapid divergence of the ecdysone receptor is a synapomorphy for Mecopterida that clarifies the Strepsiptera problem. Insect Mol. Biol. 15, 351–362. Bourguet, W., Vivat, V., Wurtz, J.-M., Chambon, P., Gronemeyer, H., and Moras, D. (2000). Crystal structure of a heterodimeric complex of RAR and RXR ligand-binding domains. Mol. Cell 5, 289–298. Browning, C., Martin, E., Loch, C., Wurtz, J.M., Moras, D., Stote, R.H., Dejaegere, A.P., and Billas, I.M.L., (2007). Critical role of desolvation in the binding of 20-hydroxyecdysone to the ecdysone receptor. J. Biol. Chem. 282, 32924–32934. Brzozowski, A.M., Pike, A.C.W., Dauter, Z., Hubbard, R.E., Bonn, T., Engström, O., Öhman, L., Greene, G.L., Gustafsson, J.-A., and Carlquist, M. (1997). Molecular basis of agonism and antagonism in the oestrogen receptor. Nature 389, 753–758.

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Carmichael, J.A., Lawrence, M.C., Graham, L.D., Pilling, P.A., Epa, V.C., Noyce, L., Lovrecz, G., Winkler, D.A., Pawlak-Skrzecz, A., Eaton, R.E., Hannan, G.N., and Hill, R.J. (2005). The X-ray structure of a hemipteran ecdysone receptor ligand-binding domain: Comparison with a lepidopteran ecdysone receptor ligand-binding domain and implications for insecticide design. J. Biol. Chem. 280, 22258–22269. Cherbas, L., Hu, X., Zhimulev, I., Belyaeva, E., and Cherbas, P. (2003). EcR isoforms in Drosophila: Testing tissue-specific requirements by targeted blockade and rescue. Development 130, 271–284. Christopherson, K.S., Mark, M.R., Bajaj, V., and Godowski, P.J. (1992). Ecdysteroid-dependent regulation of genes in mammalian cells by a Drosophila ecdysone receptor and chimeric transactivators. Proc. Natl. Acad. Sci. USA 89, 6314–6318. Clayton, G.M., Peak-Chew, S.Y., Evans, R.M., and Schwabe, J.W.R. (2001). The structure of the ultraspiracle ligand-binding domain reveals a nuclear receptor locked in an inactive conformation. Proc. Natl. Acad. Sci. USA 98, 1549–1554. De Groot, A., De Rosny, E., Juillan-Binard, C., Ferrer, J.L., Laudet, V., Pierce, R.J., PebayPeyroula, E., Fontecilla-Camps, J.C., and Borel, F. (2005). Crystal structure of a novel tetrameric complex of agonist-bound ligand-binding domain of Biomphalaria glabrata Retinoid X Receptor. J. Mol. Biol. 354, 841–853. Dhadialla, T.S., Carlson, G.R., and Le, D.P. (1998). New insecticides with ecdysteroidal and juvenile hormone activity. Annu. Rev. Entomol. 43, 545–569. Egea, P.F., Mitschler, A., Rochel, N., Ruff, M., Chambon, P., and Moras, D. (2000). Crystal structure of the human RXRα ligand-binding domain bound to its natural ligand 9-cis retinoic acid. EMBO J. 19, 2592–2601. Emmerich, H. (1972). Ecdysone binding proteins in nuclei and chromatin from Drosophila salivary glands. Gen. Comp. Endocr. 19, 543–551. Färnegårdh, M., Bonn, T., Sun, S., Ljunggren, J., Ahola, H., Wilhelmsson, A., Gustafsson, J.-Å., and Carlquist, M. (2003). The three dimensional structure of the liver X receptor β reveals a flexible ligand binding pocket that can accommodate fundamentally different ligands. J. Biol. Chem. 278, 38821–38828. Fenn, T.D., Ringe, D., and Petsko, G.A. (2003). POVScript+ : A program for model and data visualization using persistence of vision ray-tracing. J. Appl. Cryst. 36, 944–947. Gampe, R.T., Jr., Montana, V.G., Lambert, M.H., Miller, A.B., Bledsoe, R.K., Milburn, M.V., Kliewer, S.A., Willson, T.M., and Xu, H.E. (2000). Asymmetry in the PPARγ/RXRα crystal structure reveals the molecular basis of heterodimerization among nuclear receptors. Mol. Cell 5, 545–555. Graham, L.D., Johnson, W.M., Pawlak-Skrzecz, A., Eaton, R.E., Bliese, M., Howell, L., Hannan, G.N., and Hill, R.J. (2007). Ligand binding by recombinant domains from insect ecdysone receptors. Insect Biochem. Mol. Biol. 37, 611–626. Hormann, R.E., and Chortyk, O. (2004). New oxadiazoline ligands for regulating expression of nuclear receptor-based inducible genes in genetic engineering. Patent Number US2004171651-A1. Hu, X., Cherbas, L., and Cherbas, P. (2003). Transcription activation by the ecdysone receptor (EcR/USP): Identification of activation functions. Mol. Endocrinol. 17, 716–731. Iwema, T., Billas, I.M.L., Beck, Y., Bonneton, F., Nierengarten, H., Chaumot, A., Richards, G., Laudet, V., and Moras, D. (2007). Structural and functional characterization of a novel type of lignad-independent RXR-USP receptor. EMBO J. 26, 3770–3782. Jones, G., and Sharp, P.A. (1997). Ultraspiracle: An invertebrate nuclear receptor for juvenile hormones. Proc. Natl. Acad. Sci. USA 94, 13499–13503. Jones, G., Wozniak, M., Chu, Y., Dhar, S., and Jones, D. (2002). Juvenile hormone III-dependent conformational changes of the nuclear receptor ultraspiracle. Insect Biochem. Mol. Biol. 32, 33–49. Jones, G., Jones, D.E., Teal, P., Sapa, A.. and Wozniak, M. (2006). The retinoid-X receptor ortholog, ultraspiracle, binds with nanomolar affinity to an endogenous morphogenetic ligand FEBS J. 273, 4983–4996.

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Chapter 14

The Multidimensional Partnership of EcR and USP Vincent C. Henrich, Joshua Beatty, Heike Ruff, Jenna Callender, Marco Grebe, and Margarethe Spindler-Barth

Abstract Cellular signaling of the insect ecdysteroids is mediated by two nuclear receptors, the ecdysone receptor (EcR) and Ultraspiracle (USP). Considerable evidence exists that each participates in a dimerization partnership that is subject to a variety of regulatory interactions and pathways in the cell. Heterologous cell culture has been used to reconstitute and study the transcriptional activity mediated by the EcR/USP heterodimer. These studies have utilized site-directed mutagenesis and other structural modifications to dissect functional features of each dimer pair. From this work it is apparent that EcR and USP transcriptional activity depends upon the isoforms tested, the type of agonist, and the presence of juvenile hormone. USP influences the ligand affinity of EcR as revealed by the effects of mutated USP, and mutations of EcR also reveal a repressive function that apparently involves an interdomain interaction and isoform-specific. EcR retains some activity and ligand affinity in the absence of USP which may prove relevant for in vivo regulation. Cross-species comparisons and pairings of EcR and USP have shown that these dimers vary considerably in their ligand responsiveness, level of transcriptional activity, and isoform-specific effects, which in turn, is likely to be useful for developing novel insecticides that are targeted at specific receptors. The results of the cell culture work have formed a foundation for testing hypotheses about isoform-specific regulation by EcR and USP, examining the role of juvenile hormone (JH) and candidate cofactors such as MET, which itself mediates JH activity, and testing the phenotypic effects of selected mutational modifications of EcR and USP in vivo. Keywords Ecdysone • cell culture • juvenile hormone • nuclear receptor

V.C. Henrich (), J. Beatty and J. Callender Center for Biotechnology, Genomics, and Health Research, University of North Carolina-Greensboro, Greensboro, NC 27402, USA e-mail: [email protected] H. Ruff and M. Spindler-Barth Institute of General Zoology and Endocrinology, University of Ulm, 89081 Ulm, Germany M. Grebe Present address: Lütticher Str. 141, 52074 Aachen, Germany G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Introduction

Ongoing reports of ecdysone receptor (EcR) and Ultraspiracle (USP) sequences from arthropods and insect species might inadvertently create the impression that the relationship between these two nuclear receptors is static, with little to understand about it that could influence developmental processes (Henrich, 2005, and references therein). At least two features of the relationship between EcR and USP may not have been noted explicitly from studies across various experimental systems. First, a growing body of evidence indicates that the functional relationship between isoforms of EcR and USP are not equivalent, and that the EcR/USP relationship depends on contextual factors. Moreover, the observed characteristics tend to be conserved in a variety of heterologous cell types and in vivo, indicating that they involve intrinsic features of EcR and USP. Secondly, while EcR and USP share easily discernible similarities across a wide range of species, there are readily observable structural and functional differences among them. This is expected: the evolutionary distance between the Chironomids and Drosophilids within the Diptera order is 200 million years, which approximates the evolutionary distance between humans and vertebrate poikotherms. In other words, the diversity between insect orders is sufficiently large to allow for considerable diversification of EcR and USP function, with potentially important consequences for biological functions (Laudet and Bonneton, 2005).

14.2

Rationale for Using Heterologous Cell Cultures to Test EcR and USP Function

Mammalian cells, such as the well-described Chinese hamster ovary (CHO) cells, have been utilized for functional studies of the ecdysteroid receptor. The experimental paradigm is straightforward: Genes encoding EcR and USP are introduced simultaneously by cotransfection and expressed under the control of a promoter that is constitutively active in the cells (Christopherson et al., 1992). The N-terminal (transactivation) domains of EcR and/or USP, which interacts with other cofactors, are often replaced with the VP16 activation domain (AD) that is active in mammalian cells. The domain replacement removes the uncertainty associated with the sequential and functional variability in the N-terminal region of EcR and USP isoforms. This domain is also variable in sequence between insect species and consequently, difficult to compare among them (Mouillet et al., 2001; Palli et al., 2003). The cotransfection also includes a transcriptional reporter carrying single or tandem copies of an ecdysone response element (EcRE), such as the canonical inverted palindrome, hsp27 EcRE (Riddihough and Pelham, 1987; Dobens et al., 1991), attached to a weak constitutive mammalian promoter (such as thymidine kinase), and a reporter gene encoding luciferase or green fluorescent protein. After transfection, the

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cells are challenged with test ligands dissolved in the cell culture medium. Later, the cells are harvested, processed, and the reporter protein activity measured, thus providing an indication of transcriptional activity mediated by the EcR/USP complex. Despite its widespread use over the years, an obvious question is: Why use a heterologous, mammalian cell culture system to study the insect ecdysteroid receptor? The answer is that mammalian cells possess no endogenous response to ecdysteroids and therefore, any response to ecdysteroids seen in these cells can be ascribed to the expression of EcR and USP. For the purposes of measuring ecdysteroid-inducible transcriptional activity via the hsp27 EcRE, for instance, both receptors are expressed in the system. Conveniently, CHO cells do not express a detectable level of the vertebrate USP homologue, RXR (Nieva et al., in press) that can form a transcriptionally active dimer with EcR in the presence of some ecdysteroids (Christopherson et al., 1992; Henrich et al., 2003). The vertebrate EcR homologue, FXR, which is highly responsive to juvenile hormone as a dimer with RXR, is also undetectable in CHO cells (Kitareewan et al., 1996). Control experiments have shown that the expression of USP alone in the cells evokes no transcriptional activity via the hsp27 EcRE. By contrast, when EcR alone is expressed in CHO cells, a basal and induced transcriptional response via the hsp27 EcRE is detectable in cell culture, albeit at a level that is comparatively low (this will be discussed later see Costantino et al., 2008). The absence of an endogenous ecdysteroid response in mammalian cells also provides the opportunity to introduce and test structurally modified forms of EcR and USP, so that the relationship between structure and individual receptor subfunctions: dimerization, DNA affinity and binding, ligand affinity and binding, and cofactor interactions, can be dissected. By contrast, insect cells usually possess an endogenous response to ecdysteroids, and typically express some combination of EcR, USP, and unidentified cofactors, so that interpretation can be confounded by endogenous basal receptor activity and the presence of undefined insect comodulators. When comparing species specific receptor attributes, the introduction of nuclear receptors from one insect species (for instance, Leptinotarsa decemlineata) into the cell line of a different insect species (such as a Drosophila S2 cell line) could be more difficult than simply using CHO cells, which require a heterologous reconstitution of the ecdysteroid-inducible transcriptional system. While the logic of assembling a transcriptional system in a heterologous cell type may address the conceptual concern, a second question arises: Is there any evidence that the system is valid for species-specific EcR/USP comparisons, structure-function study, and understanding biologically relevant mechanisms? For the case of species-specific attributes, differences in agonist responsiveness for several insect receptor complexes have been reported that conform to expectations based on in vivo work (Graham et al., 2007a, b). For structure-function analysis, the cell culture system has proven itself by verifying ligand-binding residues for 20-hydroxyecdysone (20E) and BYI06830 in the ligand-binding domain (LBD) of Heliothis zea EcR based on computational models. Site-directed mutations of the predicted binding sites yield results that confirm the model (Billas et al., 2003).

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Comparison of EcR/USP Characteristics Across Heterologous Cell Systems

Comparison of receptor ligand binding properties when expressed in different cell types (bacteria, yeast, mammalian and insect cells) reveal that EcR and USP properties tend to be independent of the cellular context (Lezzi et al., 2002; Grebe et al., 2003, 2004; Bergman et al., 2004; Przibilla et al., 2004; Beatty et al., 2006). In other words, the effects of mutagenesis and receptor alteration relate to features of the receptor partners, rather than idiosyncracies of the experimental systems. Of course, cell culture insights cannot supplant in vivo experimentation, but the system has produced several testable hypotheses for subsequent in vivo study. In the yeast system, the GAL4 activation domain (AD) has been attached to the EcR LBD, and the GAL4 DNA-binding domain (DBD) is attached to the USP LBD (Lezzi et al., 2002). The interaction of these GAL4 fusion proteins is tested by measuring lacZ reporter activity through the UAS promoter element where the GAL4 domains are brought together by the dimerization of the EcR and USP LBDs. The paradigm allows for testing the interaction of the EcR and USP LBDs under a variety of experimental conditions, without the possible complications posed by interdomain interactions with other portions of EcR and USP. Site-directed mutations have been introduced into both of the two-hybrid fusion proteins and tested for their effects (Grebe et al., 2003, 2004; Bergman et al., 2004; Przibilla et al., 2004). These included point mutations which correspond with larval lethal in vivo mutations of EcR (Bender et al., 1997). In yeast, the mutant proteins show modestly reduced basal and inducible transcriptional activity, that is, hypomorphic activity (Bergman et al., 2004) and reduced ligand affinity (Grebe et al., 2003). The partial activity seen in the two-hybrid system may explain the ability of mutants carrying these hypomorphic alleles to survive to the larval stages, since null EcR mutations cause embryonic lethality (Bender et al., 1997). Many other EcR mutations differentially affect ligand association and dissociation, revealing that these processes involve distinct mechanisms that together determine ligand affinity. As a partner for EcR, USP primarily reduces ligand dissociation and thereby increases ligand affinity of the EcR/USP complex (Grebe et al., 2004). The possibility of interdomain signalling has been inferred in the two-hybrid system from the effect of several mutations in the USP LBD which were tested in GAL4 fusion proteins for their effects on basal and induced transcriptional activity, ponasterone A binding, and DNA binding. Some of the USP LBD substitutions result in an EcR/USP two-hybrid dimer displaying reduced ponasterone A binding (Fig. 14.1) and a severe loss of transcriptional activity, which ultimately is traced to a loss of affinity for the UAS element (Przibilla et al., 2004). The effects of these mutations belie a modulatory effect of USP upon EcR ligand-binding affinity. Paradoxically, the USP mutations exert negligible effects on transcriptional activity when introduced into whole receptors and tested in the CHO cell culture system (data not shown). In other words, the relationship between the natural DBD of USP and its LBD is apparently different than the same USP LBD’s interaction with the GAL4 DBD in the yeast two-hybrid system.

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Fig. 14.1 3H-ponasterone A binding to heterodimers consisting of the ligand binding domains of D. melanogaster EcR (fused to the GAL4 activation domain) and USP (fused to the GAL4 DNAbinding domain) and expressed in yeast cells (Lezzi et al., 2002). SD < 15% based on n = 3. First column represents specific binding to ponA alone, and USP-wt represents specific binding of wild-type EcR/USP dimer (see Grebe et al., 2003 for methods)

Several EcR mutations from the yeast two-hybrid system have also been tested in full length receptors using the CHO cell system. As will be noted, these mutations bring about specific impairments of normal function which are not easily predicted or described in vivo but which are potentially important for ascertaining receptor functions. Moreover, the cross-system consistency of mutational effects indicates intrinsic receptor functions are affected. The M504A and M504R substitutions involve a residue associated with ligandbinding in the crystal structure of the EcR from Heliothis zea (Wurtz et al., 2000; Billas et al., 2003) and shared by all reported EcRs in helix 5 of the LBD. The mutation virtually abolishes ligand-binding and ligand-dependent transcription in yeast and in cell cultures, but has no discernible effect on basal transcriptional levels in yeast or mammalian cells (Grebe et al., 2003; Bergman et al., 2004; Beatty et al., 2006). EcR (M504A) retains the ability to translocate into the CHO cell nucleus in the absence of ecdysteroids at the same rate as wild-type EcR, though M504R destroys this capability. Neither M504A or M504R translocate into the nucleus at the highly elevated rate that wild-type EcR displays in the presence of ecdysteroids. The failure reveals that the elevated rate of nuclear translocation is functionally dependent on the ligand-bound EcR, and this transport is facilitated by heterodimerization with USP (Nieva et al., 2005). Two different substitutions, K497A and K497E, of a lysine residue in helix 4 of the LBD shared by all known insect EcRs. Its effects on EcR function illustrate the multifunctional properties of individual residues within the ecdysteroid receptor sequence and also reveal unexpected insights concerning receptor function. This site was originally selected because it has been associated with cofactor interactions among several

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nuclear receptors (Wurtz et al., 1995a, b). Both K497A and K497E evoke the same mutant effect in the yeast two-hybrid system: a strongly elevated level of reporter activity from the EcR/USP dimer through the UAS promoter in the absence of ligand, and a modest elevation of ligand-induced reporter gene levels (Bergman et al., 2004). The GAL4 fusion proteins encoding K497A and K497E also impose a similar effect on two distinct properties of EcR: disruption of a salt bridge between helix 4 and helix 12 and a reduction in ligand binding that is primarily associated with a severe impairment of ligand association (Grebe et al., 2004) that in turn, modestly reduces ligand-dependent dimerization. Across the salt bridge, a counterpart mutation of a residue in helix 12 only slightly decreases ligand-binding. Thus, the reduction in ligand-binding caused by K497E is not solely caused by impairment of the salt bridge. The similar effects caused by two different substitutions of K497 (A and E) further indicates that the functional change results from a loss of a function normally associated with lysine (K), rather than an unpredicted gain or change of function caused by the residue that replaces lysine. The two-hybrid results led to a series of experiments in mammalian cell cultures to examine the properties of K497E on each of the three isoforms as partners with USP. The mutant protein has a slightly reduced dimerization capability with USP, as evidenced by the effects of competition experiments with wild-type EcR for limited amounts of USP (Beatty et al., 2006). In the CHO cells, only the EcRB2 isoform shows an increase of basal activity like the K497E mutational effects seen in the yeast two-hybrid system. Basal transcriptional activity is not significantly affected by the mutation in EcRA or EcRB1. The isoform specificity reveals an interdomain interaction that resembles isoform-specific properties seen in the vertebrate androgen receptor (Berrevoets et al., 1998). It further suggests the possibility that an EcRB2-specific corepressor interaction is normally mediated at the K497 residue. The K497E mutation has further confirmed an EcR-mediated transcriptional function that occurs in the absence of USP, though basal and induced activity is reduced by about 3–4-fold of the level seen when USP is also present (Fig. 14.2). Consistent with this transcriptional effect, EcR by itself shows the capability

Fig. 14.2 Induction of luciferase reporter activity induced by YFP- D. melanogaster EcR-B1 expressed in CHO cells in the absence of D. melanogaster USP according to methods of Henrich et al. (2003). Luciferase activity is normalized on receptor concentration determined by Western blot. Black bars: No hormone. White bars: 1 µM murA. SD < 15% based on n = 3

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of binding to ecdysteroids, though ligand affinity is much higher when USP is co-expressed (Bergman et al., 2004; Grebe et al., 2004). Importantly, the direct involvement of EcR, even without USP, is illustrated by the effect of the K497E mutation, which elevates ligand-dependent activity in these experimental conditions via the hsp27 EcRE. The effect further implicates the K497 residue as a player in the interaction between the ligand-binding pocket and helix 12, since it apparently affects ligand-dependent transcriptional activity that is known to be an AF2 (ligand-dependent) function (Hu et al., 2003). In summary, both EcR and USP carry out a range of subfunctions within the context of their partnership that are altered by individual mutations. Some involve not only intermolecular signalling but also interdomain signalling. Further, there is evidence that EcR is also capable of mediating transcriptional activity in the absence of USP, though it is relatively low.

14.4

EcR, USP and Juvenile Hormone Action

Juvenile hormone (JH) molecular actions were summarized recently by Berger and Dubrovsky (2005). Despite the different modes of actions for JH proposed at the molecular level, the outcome seems to be that JH-dependent transcription could be mediated by EcR and USP (Wu et al., 2006) as well as E75 (Dubrovskaya et al., 2004), whose own expression is regulated by ecdysteroids. Together, the varied modes of action and the variety of gene targets which have been studied can be interpreted as evidence for multiple modes of JH action, particularly since seemingly contradictory conclusions do not involve the same reagents, promoters, and cell types. The recent characterization of transcriptional effects of JH incubated with dissected D. melanogaster salivary glands offers an entry point for examining gene targets of JH, and further exploring the possible role of EcR and USP for evoking changes associated with juvenile hormone (Beckstead et al., 2007). The cell culture system has also shown that insect USP and vertebrate RXR are not functionally equivalent (Billas et al., 2001; Clayton et al., 2001). RXR does not form a transcriptionally active dimer with D. melanogaster EcR in the presence of 20E (Henrich et al., 2003). Exogenously introduced USP, but not the mammalian RXR, facilitates nuclear transport of EcR in mammalian cells (Nieva et al., 2005). RXR is activated by the JH analogue, methoprene, and physically interacts with methoprene acid (Harmon et al., 1995), thus forming a basis for examining the ligand status of JH and JH analogues on RXR’s insect orthologue, USP. Based on the RXR analogy, an obvious possibility for JH action is that USP is acting as a JH receptor via its ligand binding pocket, and a computational model has been reported that allows for JH binding inside the USP ligand-binding pocket (Sasorith et al., 2002). Using a DR12 response element in insect cells, Jones et al. (2001) showed that a USP-mediated transcriptional response is detected via homodimerization, and that high affinity binding with the JH precursor, methyl farnesoate, is abolished by mutating the C472 residue in the USP LBD (Jones et al., 2006).

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CHO cells transfected with EcR and USP possess no capability for mediating a response to juvenile hormone or analogues such as methoprene and pyriproxfen via an hsp27 EcRE, and similar conclusions have been drawn from insect cells (Henrich et al., 2003; Fang et al., 2005). These experiments suggest that JH employs different mechanisms in insect cells, depending in part upon the presence of EcR and the type of promoter element involved. In CHO cells, JH exerts no effect on EcR/USP transcriptional activity but reduces the dosage of ecdysteroids necessary to achieve maximal induction by about tenfold (though it cannot act additively with ecdysteroids to evoke a supramaximal response; Beatty et al., 2006). The ability of JH and analogues to potentiate a maximal induction at otherwise submaximal ecdysteroid dosages has established a paradigm for testing known and novel compounds for their ability to affect EcR/USP activity. The analysis has been extended to the components of the mevalonate pathway, whose starting substrate is acetyl coA, which is derived from glycolysis and the Krebs cycle and which ultimately depends on the availability of nutrients (Belles et al., 2005). Several substrates along the insect mevalonate pathway ending with JH have shown the ability to potentiate the ecdysteroid response in CHO cells over the 20–100 µM range, including farnesol, farnesoic acid, farnesyl diphosphate, and methyl farnesoate (Fig. 14.3). The broad spectrum-low affinity interaction suggests a “sensing” mechanism resembling those described for the nuclear receptor PPAR,

45 0µM murA 0.1µM murA 1µM murA 0.1µM murA+ 80µM Potentiator 80µM Potentiator only

40 35 30 25 20 15 10 5

EcRB2/ USPII

EcRB1/ USPII

JHIII

Farnesoic Acid

Methyl Farnesoate

Fig. 14.3 Potentiation effects of mevalonate pathway constituents on basal and induced levels of D. melanogaster EcR/USP transcriptional activity in CHO cells following described methods (Henrich et al., 2003). Measurements of luciferase reporter activity are given as fold-induction with D. melanogaster EcRB2/USP in the absence of murA equal to 1. Standard deviations based on n = 3

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which regulates metabolism in vertebrates (Auwerx et al., 2003), the effect of glucose on LXR (Mitro et al., 2007), and the low affinity ligand interactions involving intracellular metabolites seen in other nuclear receptors such as PXR and FXR (Goodwin et al., 2003; Desvergne, 2007). Conceivably, a cellular milieu laden with mevalonate pathway substrates reduces the actual level of ecdysteroids necessary to evoke a maximal inductive response. Low ecdysteroid titer peaks generally trigger the larval-larval molts during feeding larval stages in holometabolous insects, and therefore, cross talk between low ecdysteroid titers and nutritional state would ensure an appropriate trigger for larval development. The mechanism also could account for the absence of any obvious detrimental effect when JH is absent in Drosophila, since JH may represent just one of the sensors read by EcR and USP. The mechanism for JH potentiation apparently is distinct from the high affinity interaction with methyl farnesoate. Alanine substitutions were made for each JH-binding site proposed in the Sasorith model and tested for their effects on ecdysteroid induction and JH potentiation by the EcR/USP complex in the CHO cell system. Basal and induced transcriptional activity was relatively unaffected in full-length receptors, and potentiation by JH was normal. Notably, the C472 residue associated with a high affinity interaction with methyl farnesoate affected neither JH-mediated or methyl farnesoate-mediated potentiation, nor did the mutation affect ecdysteroid inducibility (data not shown). Given the differences in behavior of USP in the presence and absence of EcR, it is conceivable that USP retains a high affinity interaction with JH as a homodimer, or in conjunction with other cofactors, whereas the EcR/USP heterodimer involves a low affinity/low specificity interaction with JH or with other cofactors that interact with JH. Differences in ligand affinity have been reported for USP’s vertebrate RXR homologue in the presence and absence of heterodimeric partners (Desvergne, 2007). The distinct mechanisms described so far may simply reveal that the functional effects of JH depend not only on USP, but also on EcR. For instance, it is plausible that JH modulates the ability of EcR and USP to recruit comodulators, such as the enhanced corepressor recruitment seen in the presence of JH (Maki et al., 2004). When viewed individually and compared together, these results further imply that JH utilizes more than one mode of action to affect transcriptional activity in the cell (Feyereisen, 1998). Another indication that JH plays multiple cellular roles is based on the gene encoding the Met-Tolerant (MET) protein. Drosophila MET protein belongs to the bHLH-PAS family and binds to JH and methoprene, though a functional or physical connection between MET and specific gene targets remains to be identified. In the absence of JH, MET dimerizes with a second bHLH-PAS factor, and is displaced by the introduction of JHIII or methoprene (Godlewski et al., 2006). Other members of the bHLH-PAS family are known to interact directly with nuclear receptors via an LXXLL motif in the PAS domain (Okino and Whitlock, 2000); MET possesses several of these motifs, though no interaction with either EcR or USP has yet been described.

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A few salient observations imply that a bHLH-PAS transcription factor may play(s) a role in mediating the low affinity potentiation described in CHO cells. The bHLH-PAS family includes the dioxin receptor (aka, aryl hydrocarbon receptor, ArHR), which is activated by polychlorinated biphenyls (PCBs). Remarkably, PCBs have been shown to potentiate the ecdysteroid response in CHO cells with about the same potency as JH, further implicating a common mode of action (Oberdorster et al., 1999). Interestingly, JH potentiation only occurs with the use of natural ecdysteroid agonists. Nonsteroidal agonists, which confer a different shape to the EcR ligandbinding pocket than natural ecdysteroids, are incapable of evoking potentiation (Beatty, unpublished, 2006). Presumably the pocket shape assumed by interaction with diacylhydrazines and other nonsteroidal agonists of EcR precludes interactions with cofactors. Whether an interaction with MET is tied to the distinct effects of ecdysteroids and nonsteroidal agonists on JH potentiation in the CHO cell culture system is an active subject of investigation.

14.5

Species Specific Aspects of EcR and USP Revealed in Cell Cultures

Species specific variation of the amino acid sequences of EcR and USP and the expression of different isoforms leads to physiological consequences like altered ligand specificity in Leptinotarsa decemlineata (Colorado potato beetle; Ogura et al., 2005) and other insects (Graham et al., 2007a, b) and changes in the dimerization efficiency (Suhr et al., 1998) and DNA binding of Bombyx mori receptors (Shirai et al., 2007). Chimeric complexes of EcR and USP derived from different species may elucidate the molecular basis for these differences, and this approach has already been applied for cell culture studies of EcR function. The evidence for interspecial differences in the USP LBD is apparent by the effects of substituting the LBD of the D. melanogaster USP with the equivalent domain of Chironomus tentans. The resulting chimeric USP rescues larval development in usp mutant flies, but suddenly fails at the onset of pupariation, meaning that the C. tentans USP LBD is unable to perform a function that is essential for metamorphosis (Henrich et al., 2000). For the three ecdysteroid receptor isoforms of D. melanogaster, deletion of the DNA-binding domain of D. melanogaster USP evokes different effects. This construct, (USP-∆DBD) forms an active dimer with EcRB1 that wholly retains its ability to mediate ecdysteroid-inducible transcriptional activity. However, while USP-∆DBD dimerizes with all three EcR isoforms, the resulting EcRA and EcRB2 complexes are only weakly active (Beatty et al., 2006). In these cases, the USP-∆DBD is dominant negative, that is, it forms a transcriptionally impaired complex with EcRA and EcRB2. USP in vivo point mutations affecting conserved residues in the DBD do not inhibit the expression of early-inducible genes, though a null mutation of USP eliminates induction. All of the USP mutations lead to premature expression of other

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genes, indicating that the USP-∆DBD normally participates in a repressive function (Schubiger and Truman, 2000; Ghbeish et al., 2001). The relationship of the cell culture results and the in vivo observations have yet to be reconciled. By contrast, the equivalent USP-∆DBD construct from Choristoneura fumiferana, forms a high activity dimer with all three D. melanogaster EcR isoforms (Henrich et al., 2003), and in fact, this heterospecies complex displays a distinctively higher activity level than the native D. melanogaster complexes using an intact USP (Fig. 14.4). The structural difference(s) that are responsible for the functional differences between the two USP sequences remains to be tested more fully. In any case, it is increasingly apparent that there is considerable divergence among the structures of insect USPs that is not obvious in sequence alignments, and that the USP proteins diverge considerably from the vertebrate RXR (Iwema et al., 2007).

65 60

0uM murA 0.1uM murA

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50

80uM JHIII

45 40 35 30 25 20 15 10 5 0 EcRB1/DmUSPII

EcRB1/DmUSPIII

EcRB1/CfUSPIII

Fig. 14.4 Effects of USP-∆DBD from two insect species on basal and induced transcriptional activity of D. melanogaster EcRB1 based on methods of Henrich et al. (2003). Fold-induction compared to transcriptional activity of EcRB2/DmUSPII in the absence of murA (equals 1). DmUSPII refers to the D. melanogaster USP in which the N-terminal domain has been replaced by VP16 AD (Beatty et al., 2006). USPIII (D. melanogaster) and CfUSP (Choristoneura fumiferana) refer to a VP16-USP (∆DBD), from which the DBD has been removed (Henrich et al., 2003 and refs. therein)

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Summary

The EcR/USP heterodimer possesses a variety of subfunctions and features which are difficult to assess in vivo and may not be apparent unless observed in isolation or through careful manipulation. The heterologous mammalian cell culture system has emerged as an effective tool to elucidate these issues. So far, the system has yielded observations that generally confirm those obtained biochemically and from yeast two-hybrid systems, indicating that these systems are dealing with actual receptor attributes. More importantly, the system has proven useful for testing computational models, structure-function studies, EcR and USP isoform comparisons within species, and interspecies receptor comparisons. New insights leading to testable hypotheses in vivo have been generated through the process about the relationship of EcR and USP isoforms, as well as the role of JH and possible cofactors. The next phase of studies will likely include many modifications and comparisons between species, with the aim of elucidating not only the evolutionarily conserved features of EcR/USP activity, but also the many nuances and variations associated with insect adaptation to specific environmental niches.

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Chapter 15

Functional Analysis of Ecdysteroid Receptor from Drosophila melanogaster “In Vitro” Anca Azoitei, Heike Ruff, Christian Tremmel, Simone Braun, and Margarethe Spindler-Barth

Abstract Ecdysone receptor (EcR) was expressed in vertebrate cells to study its functional properties in the absence of the heterodimerization partner Ultraspiracle (Usp) and to avoid interference with endogenous receptor isoforms. Comparison of different isoforms affords determination of receptor concentration, which was achieved either by determination of ligand binding sites by Scatchard analysis or by quantitative evaluation of specific Western blot signals, but not by normalization on transfection efficiency as determined by cotransfection with a constitutive reporter plasmid. Ligand- and DNA- binding, and transcriptional activity of EcR isoforms and the influence of Ultraspiracle (Usp) were described. Keywords Dimerisation • DNA • ecdysone • heterologeous expression • ligand • nuclear receptor • transcription • Ultraspiracle

15.1

Introduction

The aim of this review is to improve our understanding, how the activity of receptor proteins can be modulated to meet the special requirements in different tissues and developmental stages. We will focus on the functionality of the ecdysone receptor (EcR) at the molecular level in vitro and describe, how receptor activity is modified in an isoform specific way. We will characterize the properties of EcR isoforms and its heterodimerization partner Ultraspiracle (Usp) expressed separately in vertebrate cells (CHO-K1) with a neglectable RXR content (Nieva et al., 2008) to circumvent interactions with endogenously expressed receptor protein (Chapter 14 of this book). Next the influence of the dimerization partner Usp on EcR function was examined by coexpression of both receptors. The use of Usp variants especially Usp III, where the DNA binding domain is deleted (Beatty et al., 2006), allows to study the impact of the dimerization sites in the DNA binding and ligand

A. Azoitei, H. Ruff, Ch. Tremmel, S. Braun, and M. Spindler-Barth () Institute of General Zoology and Endocrinology, University of Ulm, 89081 Ulm, Germany e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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binding domains on ecdysone receptor function separately. The elucidation of the molecular mechanism of the interaction of EcR and Usp will certainly improve our understanding of the dual role of ultraspiracle in mediating ecdysone responses in vivo as described by Ghbeish et al. (2001).

15.2

Determination of Receptor Concentrations

For comparison of receptor functionality of EcR isoforms, it is essential that equal amounts of receptor protein complexes are used. It is common practise to normalize data on the activity of a coexpressed reporter gene coupled to a constitutive promoter. As shown in Table 15.1, determination of transfection efficiency by constitutively expressed ß-galactosidase deviates up to 40-fold from heterodimer concentrations determined by Scatchard plot. Relative EcR concentrations determined by Western blot correspond much better with Scatchard plot data (ratio of EcR concentrations determined by Scatchard plot/Western blot = 1.6 + 0.6) (Table 15.1), if sufficient amounts of Usp are present. Deletion of the C-domain of Usp (Usp III) impairs dimerization, which results in a decreased fraction of heterodimers indicated by a lower ratio: EcR-Usp (Scatchard plot)/EcR (Western blot). For comparison of the functional properties of the various heterodimeric complexes themselves, e.g. DNA binding and ligand binding studies, in vitro separately expressed Usp III had to be added to transfer EcR quantitatively into the more active heterodimers. For determination of transcriptional activity of the ecdysone receptor in transfected cells the ratio of DNA coding for EcR and Usp III has to be changed in favor of Ultraspiracle to ensure that the receptor proteins are present mainly as heterodimer. Table 15.1 Comparison of heterodimer concentrations calculated by Scatchard plot, transfection efficiency measured by cotransfection of constitutively expressed ß-galactosidase, and EcR concentration determined by Western blot EcR/UspEcR conc. EcR/Usp conc EcR/Usp ß-galacto(determined by Western (Scatchard plot) sidase activity blot, rel. units) 1.3 2.30 EcRA/UspIa EcRB1/UspI 0.95 1.54 EcRB2/UspI 1.16 6.04 EcRA/UspIIb 2.19 21.90 EcRB1/UspII 2.54 16.60 EcRB2/UspII 3.33 24.13 EcRA/UspIIIc 0.76 4.37 EcRB1/UspIII 2.74 41.52 EcRB2/UspIII 2.68 68.72 a Usp I = Vp16AB-UspCDE b Usp II = Vp16AB-UspCDE c Usp III = Vp16AB-UspDE (Beatty et al., 2006)

1.9 1.1 0.7 1.4 2.2 2.1 0.2 0.6 0.6

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Fig. 15.1 Western blot of EcR isoforms coexpressed with different Usp variants. The receptor proteins tagged at the N-terminus with YFP were detected by a GFP-specific antibody (Bioscience, Palo Alto, CA, USA)

In addition to different expression levels, the discrepancy between transfection efficiency and receptor protein concentration is certainly due to differences in the stability of the receptor proteins, which is influenced by hormone application, type and concentration of heterodimerization partners (Nieva et al., 2008), dimerization capabilities of receptor isoforms, and accessory proteins like comodulators and heatshock proteins (Cronauer and Spindler-Barth, unpublished observations, 2006). As shown in Fig. 15.1, even the coexpressed Usp variant influences EcR concentrations as determined by Western blot in an isoform specific way. For comparison of the functionality of different receptor heterodimers composed of EcR isoforms and Usp variants, we therefore normalized our data not on transfection efficiency determined by a constitutively expressed control plasmid coding for e.g. ß-galactosidase, but on receptor concentrations determined by Scatchard plot or Western blot. In addition care was taken that EcR was transferred nearly quantitatively as heterodimer, if properties of the heterodimer were studied.

15.3 Estimation of the Affinity Between EcR Isoforms and Usp As outlined above, even if the same concentrations of EcR and Usp are expressed, the concentrations of heterodimeric complexes vary according to the differences in the affinities between EcR isoforms and Usp variants. This is especially important in case of Usp III, when the dimerization interface in the C-domain is missing. Previously two hybrid assays were used to evaluate dimerization capability and differences in transactivation potency were interpreted as variations of the dimerization efficiency between EcR and Usp (Lezzi et al., 2002). Since transactivation potency depends on several additional factors like DNA binding and interaction with comodulators the conclusions are rather indirect. Comparison of Gal4 fusion proteins and full length receptors revealed also that the C-domain

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Table 15.2 Interaction of EcR isoforms with Usp variants. The concentration of the heterodimer was determined by Scatchard plot. The concentrations of EcR and Usp were determined by Western blot. The relative affinities were estimated using the law of masses EcR A Usp I = UspII >>> UspIII 1 : 1 : 25 Rel. KD(dimerization) EcR B1 UspI = UspII >> UspIII Rel. KD(dimerization) 3 : 1 : 126 EcR B2 UspI > UspII >> UspIII Rel. KD(dimerization) 1 : 10 : 150

of Gal4 transcription factor and Usp are functionally not equivalent (Chapter 14 of this book). Determination of EcR and Usp concentrations by Western blot and determination of heterodimer concentrations by Scatchard plot provide an alternative method to roughly estimate relative values for the affinity between both dimerization partners by the law of masses (Table 15.2): K D(dimerization) =

[EcR] × [Usp] [EcR/Usp]

Although no exact calculation is possible by this way, it is evident that the affinity of Usp to all isoforms of EcR is reduced dramatically if the C-domain of Usp (Usp III) is deleted. This is most pronounced for EcR-B1 and EcR-B2. The dimerization capability of EcR-A/Usp III is affected to a lesser extent, which may indicate that hormone responses, which do not afford the participation of USP – DBD (Ghbeish et al., 2001) are mediated more effectively by EcR-A.

15.4

Ligand Binding

In contrast to ecdysone receptor from most other arthropod species investigated so far EcR from Drosophila melanogaster (Grebe et al., 2004) and Leptinotarsa decemlineata (Ogura et al., 2005) bind ponasterone A specifically already in the absence of Usp. In both species ligand affinity is considerably lower in the absence of Usp and improved by heterodimerization. The ligand binding domain encompassing the C-terminal part of the D-domain and the E-domain is necessary and sufficient for binding of ponasterone A (Grebe et al., 2003; Lezzi et al., 2002). Hormone binding to EcR in the absence of Usp is associated with changes in receptor function e.g. increased nuclear localization (Nieva et al., 2007) enhanced interaction with DNA (Cronauer et al., 2007), and enhanced transcriptional activity, which underlines its physiological importance offering an additional mechanism for fine tuning of the hormonal response. In the presence of the DE-domains of Usp the affinity of ponasterone A to the heterodimeric complex is about 90-fold higher compared to DE-domains of EcR

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alone (Grebe et al., 2004). Based on mutational analysis it was concluded that the 3D-structure of the ligand binding domain of EcR is changed by dimerization with Usp (Grebe et al., 2003). Although heterodimerization is observed in the absence of hormone (Yao et al., 1993), ligand binding to EcR improves dimerization considerably. RXR can replace Usp in some instances (Henrich et al., 2003) e.g. promotes DNA binding of the heterodimeric complex and enhances transcriptional activity (Nieva et al. 2008), ligand binding is not increased by heterodimerization with RXR, which is in accordance with the high sequence identity of the DBDs in contrast to the LBDs of RXR and Usp. The KD-value of about 2 nM determined using ligand binding domains of EcR (aa 431-562) and Usp encompassing the C-terminal part of the D-domain and the E-domain (aa 224-508) (Grebe et al., 2004) is comparable to full length receptors and seems to be independent of the cellular context, since EcR/Usp expressed in bacteria (Arbeitman and Hogness, 2000; Halling et al., 1999), yeasts (Grebe et al., 2004), vertebrate cells and endogenous receptors present in insect cells (Maroy et al., 1978; Beckers et al., 1980) or insect tissue (Yund et al., 1978) bind hormone with approximately the same affinity. Only Cherbas et al. (1988) report a ten fold higher ligand affinity for EcR/Usp from insect cells. The AB-domain of EcR isoforms has no significant influence on ligand binding. Tags fused to the N-terminus of the receptors do not modify ligand affinity (Grebe et al., 2004). The fusion protein of the activation domain of VP16 and the CDE-domains of Usp, routinely used for transactivation studies to overcome the inhibitory action of the AB-domain of Usp (Henrich, 2005; Beatty et al., 2006), confers the same ligand binding capability to all EcR isoforms as wild type Usp (Azoitei, unpublished, 2006). In the absence of DNA the C-domain of Usp does not decreases the affinity of ponasterone A to the heterodimer. Interaction of the receptor complex with DNA improves ligand binding. Ponasterone A binding is enhanced significantly by the hormone response element 5x hsp 27 which is routinely used for transactivation studies, 1x hsp 27 has no significant effect.

15.5

Interaction of Ecdysone Receptor Isoforms with DNA

All three EcR isoforms are only partially localized in the nucleus in CHO cells in the absence of Usp (Gwozdz et al., 2007; see also Chapter 16 of this book). Destruction of Zn-fingers in the C-domain of EcR has no effect on intracellular localization of EcR (Cronauer et al., 2007) indicating that in the absence of hormone a considerable fraction of EcR does not bind to DNA. This is confirmed by EMSA, where only weak binding to 1x hsp 27 was found (Braun, unpublished, 2006) for all EcR isoforms in the absence of hormone and dimerization partner. In the presence of 1 µM muristerone A intracellular localization of EcR is clearly increased (Nieva et al., 2007) and DNA binding in whole cells and cell extracts is enhanced. Considerable differences in the affinity of liganded EcR isoforms to hsp 27 are not observed on EMSA gels.

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In the presence of Usp, EcR is nearly quantitatively present in the nucleus of vertebrate cells (Nieva et al., 2005; Gwozdz et al., 2007). The influence of Usp on nuclear transport is mainly due to dimerization via the ligand binding domain, since deletion of the C-domain of Usp has only a moderate effect (Cronauer et al., 2007). DNA binding of Usp fusion proteins with the AD-domain of Vp16 (Usp I and Usp II and wild type Usp are comparable. In vertebrate cells nuclear EcR/Usp interacts with DNA significantly only in the presence of hormone (Cronauer et al., 2007), which corresponds to the increased affinity of the heterodimer to hsp27 observed with muristerone A in EMSA gels. The only exception is observed with Usp III. In this case DNA binding in the presence of muristerone A is decreased for all EcR isoforms compared to basal levels (Braun, unpublished results, 2006).

15.6

Transcriptional Activity

Determination of transcriptional activity of the ecdysone receptor complex is a complicated matter due to several reasons. First, the cellular context seems to be important. For example only EcR-B1 and EcR-B2 exhibit strong transcriptional activity in Kc cells with a wild type Usp background, but not EcR-A (Hu et al., 2003), whereas basal transcriptional activity of EcR-A/Usp expressed in Hela cells is observed (Mouillet et al., 2001). Second, transcriptional activity is often normalized on expression efficiency determined by cotransfection of a constitutively expressed reporter plasmid (Beatty et al., 2006; Henrich et al., 2003). In this case transcriptional activity is the sum of transcriptional capability of the receptor complex and changes in receptor protein concentrations, which vary considerably depending on the experimental conditions. As outlined earlier, hormone binding, heterodimerization with Usp, and interaction with comodulators (Tremmel, data not shown) increase receptor stability and contribute to the enhanced transcriptional activity under these conditions. Normalization of transcriptional activity on receptor protein concentration (Western blots) reveals that basal transactivation of a reporter gene depends on the AB-domain of EcR, as reported previously (Dela Cruz 2000) and is lowest for EcR-A in vertebrate cells (Fig. 15.2a). No significant increase is observed in the presence of muristerone A, due to the low hormone binding capability in the absence of a heterodimerization partner (data not shown). Basal transcriptional activity of the heterodimeric complexes are about 3–5-fold higher, depending on the Usp variant, compared to EcR isoforms in the absence of Usp (data not shown here). The influence of the AB domain of EcR is abolished in the presence of Usp. EcR-isoform specific differences in basal (Fig. 15.2b) and hormone induced transcriptional activity (Fig. 15.2c) are observed in the heterodimer. Hormone induced increase in reporter gene activity is rather modest (about 2–4-fold), if the influence of higher receptor concentrations due to increased receptor stability in the presence of Usp and ligand is eliminated and only the transactivation potency of the receptor protein complex is determined.

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Fig. 15.2 Transcriptional activity of EcR isoforms. (a) Basal activity of EcR isoforms in the absence of a heterodimerization partner. The differences between EcR-A, EcR-B1 and -B2 are significant (p < 0.02). (b) Relative transcriptional activity of heterodimers consisting EcR isoforms and Usp II in the absence of ligand. EcR-A/Usp II = 1. The activity of the heterodimers were about 3–5-fold higher compared to EcR isoforms in the absence of a heterodimerization partner, but no isoforms specific differences were observed. (c) Increase in transcriptional activity of heterodimers consisting of ecdysone receptor isoforms and Usp II in the presence of 1 µM muristerone A. Basal activity of the corresponding heterodimers = 1

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Functional Role of Usp

Usp modulates ecdysone receptor function in multiple ways. Usp increases the stability of EcR (Nieva et al., 2008), enhances nuclear localization of EcR (Nieva et al., 2005), modifies the affinity of the ligand to EcR, affects interaction with DNA and enhances transcriptional activity of the receptor complex. The influence of the dimerization partner depends on the dimerization interface involved. For efficient nuclear transport of EcR heterodimerization in the cytoplasm via LBDs is necessary (Cronauer et al., 2007). Participation of the dimerization sites located in the C-domains of EcR and Usp seems unlikely in the absence of DNA. RXR can not replace Usp in this case (Nieva et al., 2008), which is in agreement with the fact that RXR does not enhance ligand binding to EcR/RXR. In contrast dimerization mediated by the C-domains is important for DNA binding of the receptor complex and in this case the conserved DNA binding domain of RXR can efficiently substitute Usp. These data show that not always both dimerization sites are involved equally, but depending on the circumstances different dimerization sites can be preferentially or even selectively active. Comparison of different Usp variants (Fig. 15.3a) revealed that wild type Usp increases transcriptional activity to a similar extent as Vp16 fusion proteins in CHO cells, which are usually preferred (Henrich et al., 2003), if normalized on receptor protein concentration, whereas in S2 cells no influence of the ABdomain of Usp on transcriptional activity is observed with the semisynthetic hre Eip71CD (Hu et al., 2003). Interestingly hormone induced increase of transcriptional activity is most pronounced, when the C-domain of Usp is deleted (Usp III) (Fig. 15.3b). Basal activity is rather low in this case, which shows that transcriptional activity of the heterodimer in the absence of ligand is determined mainly by DNA binding mediated by Usp. In contrast hormone induced increase is most pronounced with Usp III. As shown in Fig. 15.4 DNA binding is impaired, if the C-domain of Usp is deleted (Usp III). The influence of hormone on DNA binding of the heterodimer is rather weak for EcR-A and EcR-B1, if the same amount of receptor protein is applied as determined by ligand binding or Western blot. Independent of the EcR isoform DNA binding is diminished in the presence of hormone below basal levels in heterodimers with Usp III confirming the importance of the Usp-DBD for interaction with DNA. However DNA binding and transcriptional activity do not change in parallel. Enhanced transcriptional activity despite weak interaction with DNA determined by EMSA as shown in Fig. 15.4 was described already previously for a mutated ecdysone receptor (Beatty et al., 2006). Hormone induced stimulation of transcriptional activity is even higher with EcR/Usp III complexes despite low DNA binding, an observation also reported for mutated androgen receptor in prostate cancer cells (Haelens et al., 2007), which exemplifies the complex regulation of transcriptional activity by nuclear receptor complexes.

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Fig. 15.3 Transcriptional activity of EcR-B1/Usp complexes. Luciferase activity is normalized on EcR concentration determined by Western blot. (a) Basal activity differences between absence of Usp and presence of Usp variants (p < 0.01–0.03) and increase in activity of EcR-B1/UspIII compared to EcR-B1/UspII (p < 0.01) are significant. (b) Hormone induced increase. Differences between absence of Usp and presence of UspI, II or III are significant (p < 0.02–0.03)

This discrepancy is certainly due to modulatory effects by additional proteins like comodulators. In fact, EcR and Usp are present in high molecular weight complexes of different sizes. This is observed for DE-domains of Drosophila receptors expressed in yeasts (Greb-Markiewicz et al., 2005) and for endogenous receptors in Chironomus cells (Spindler-Barth and Spindler, 2003), but the composition of these complexes is still unknown. The influence of Hsp90 and Hsp70 on receptor stability is already demonstrated (Cronauer and Spindler-Barth, unpublished results, 2006). The requirement of heat shock proteins for DNA binding was shown previously (Arbeitman and Hogness, 2000). Coexistence of two different receptor complexes of EcR-A and EcR-B1 with Usp, which give rise to different bands on EMSA gels with the hormone response element hsp 27 confirms this hypothesis. However we have to be aware that experimental conditions for EMSAs are rather

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Fig. 15.4 Influence of muristerone A (1 µM) on binding to hsp27 as determined by EMSA (white bars) and transcriptional activity (black bars) of EcR/Usp complexes. The intensity of retarded receptor bands were quantified (Bio1D, Vilber Lourmat, France; for background substraction rolling ball mode was used) and the ratio: hormone induced/basal intensity determined. Transcriptional activity of the heterodimer was determined by induction of a reporter gene (luciferase) coupled to a promoter containing a pentameric hsp27 and normalized on receptor concentration as measured by Western blot. (a) EcR-A. The differences in transcriptional activity between EcRA/Usp I and EcR/UspIII (p < 0.05) and differences between DNA binding of EcRA/Usp I and EcRA/UspIII (p < 0.02) are significant. (b) Transcriptional activity of heterodimeric complexes with EcR-B1. Differences between EcRB1/Usp I and Usp II to EcRB1/UspIII are significant (p ≤ 0.02)

artificial compared to the physiological environment in the cells. Moreover, in EMSA the hormone response element 1x hsp 27 was applied, whereas for transactivation assays multimeric response elements were used. In the presence of muristerone A, the DNA binding domain of Usp is required for maximal EcR-A and -B2 transcriptional activity (Fig. 15.4), but is dispensable for EcR-B1 expressed in CHO cells (Beatty et al., 2006). This is in agreement with the fact that for efficient interaction of EcR-B1 with DNA in the presence of hormone Usp III is sufficient (Cronauer et al., 2007).

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Conclusions

Transcriptional activity is determined by the receptor concentration, which is the result of transcription/translation and the stability of the corresponding receptor proteins, which is different for individual receptor isoforms, and is further modified by ligand and heterodimerization partner. The concentration of the heterodimer is additionally influenced by the affinity of the dimerization partners. Normalization of transcriptional activity on receptor complex concentration allows to investigate the functional properties of the receptor complex itself. Comparison of results obtained with the same receptor proteins in vertebrate cells, which were normalized on transfection efficiency (Henrich et al., 2003; Beatty et al., 2006) with the data presented here indicate that a considerable part of the hormone induced increase of transcriptional activity is due to enhanced receptor stability. No difference in ligand binding affinity is observed for individual EcR isoforms, but a considerable influence of interaction with DNA on ligand binding is detected. On the other hand DNA binding as determined by EMSA is increased in the presence of hormone depending on the EcRE and the ecdysteroid applied (Elke et al., unpublished observations, 2002). Taken together the results show a complex mixture of multiple intramolecular interactions of various domains of both dimerization partners for diversification of the hormonal response e.g. the EcR-A isoform seems to mediate more effectively basal transcription in the absence of Usp, whereas the hormone effect is much more pronounced, if only EcR but not Usp interact with the hormone response element – a mode of action proposed already previously by Ghbeish et al. (2001).

References Arbeitman MN, Hogness DS (2000) Molecular chaperones activate the Drosophila ecdysone receptor, an RXR heterodimer. Cell 101:67–77 Beatty J, Fauth Th, Callendar JL, Spindler-Barth M, Henrich VC (2006) Functional analysis of ecdysteroid receptor isoforms in Drosophila melanogaster in a cell culture system. Insect Mol Biol 15:785–795 Beckers C, Maroy P, Dennis R, O’Connor JD, Emmerich H (1980) The uptake and release of ponasterone A by the Kc cell line of Drosophila melanogaster. Mol Cell Endocrinol 17:51–59 Cherbas P, Cherbas L, Lee SS, Nakanishi K (1988) 26-[125I]iodoponasterone A is a potent ecdysone and a sensitive radioligand for ecdysone receptors. Proc Natl Acad Sci USA 85:2096–2100 Cronauer MV, Braun S, Tremmel Ch, Krönke K-D, Spindler-Barth M (2007) Nuclear localization and DNA binding of ecdysone receptor and ultraspiracle. Arch Insect Biochem Physiol 65:125–133 Dela Cruz FE, Kirsch DR, Heinrich JN (2000) Transcriptional activity of Drosophila melanogaster ecdysone receptor isoforms and ultraspiracle in Saccharomyces cerevisiae. J Mol Endocrinol 24:183–191 Ghbeish N, Tsai CC, Schubiger M, Zhou JY, Evans RM, McKeown M (2001) The dual role of ultraspiracle, the Drosophila retinoid X receptor, in the ecdysone response. Proc Natl Acad Sci USA 98:3867–3872

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Grebe M, Przibilla S, Henrich VC, Spindler-Barth M (2003) Characterization of the ligand-binding domain of the ecdysteroid receptor from Drosophila melanogaster. Biol Chem 384:105–116 Grebe M, Fauth T, Spindler-Barth M (2004) Dynamic of ligand binding to Drosophila melanogaster ecdysteroid receptor. Insect Biochem Mol Biol 34:981–989 Greb-Markiewicz B, Fauth T, Spindler-Barth M (2005) Ligand binding is without effect on complex formation of the ligand binding domain of the ecdysone receptor (EcR). Arch Insect Biochem Physiol 59:1–11 Gwozdz T, Dutko-Gwozdz J, Nieva C, Betanska K, Orlowski M, Kowalska A, Dobrucki J, Spindler-Barth M, Spindler K-D, Ozyhar A (2007) EcR and Usp, components of the ecdysteroid nuclear receptor complex, exhibit differential distribution of molecular determinants directing subcellular trafficking. Cell Signal 19:490–503 Haelens A, Tanner T, Denayer S, Callewaert L, Claessens F (2007) The hinge region regulates DNA binding, nuclear translocation, and transactivation of the androgen receptor. Cancer Res 67:4514–4523 Halling BP, Yuhas DA, Eldridge RR, Gilbey SN, Deutsch VA, Herron JD (1999) Expression and purification of the hormone binding domain of the Drosophila ecdysone and ultraspiracle receptors. Protein Expr Purif 17:373–386 Henrich VC (2005) The ecdysteroid receptor. In: Comprehensive Molecular Insect Science (eds. L.J. Gilbert, K. Iatrou, and S.S. Gill), Oxford, Elsevier/Pergamon. Vol 3, 243–282 Henrich VC, Burns E, Yelverton DP, Christensen E, Weinberger C (2003) Juvenile hormone potentiates ecdysone receptor-dependent transcription in a mammalian cell culture system. Insect Biochem Mol Biol 33:1239–1244 Hu X, Cherbas L, Cherbas P (2003) Transcription activation by the ecdysone receptor (EcR/USP): identification of activation functions. Mol Endocrinol 17:716–731 Lezzi M, Bergman T, Henrich VC, Vogtli M, Fromel C, Grebe M, Przibilla S, Spindler-Barth M (2002) Ligand-induced heterodimerization between the ligand binding domains of the ecdysone receptor. Eur J Biochem 269:3237–3245 Maroy P, Dennis R, Beckers C, Sage BA, O’Connor JD (1978) Demonstration of an ecdysteroid receptor in a cultured cell line of Drosophila melanogaster. Proc Natl Acad Sci USA 75:6035–6038 Mouillet JF, Henrich VC, Lezzi M, Vögtli M (2001) Differential control of gene activity by isoforms A, B1 and B2 of the Drosophila ecdysone receptor. Eur J Biochem 268:1811–1819 Nieva C, Gwozdz T, Dutko-Gwozdz J, Wiedenmann J, Spindler-Barth M, Wieczorek E, Dobrucki J, Dus D, Henrich V, Ozyhar A, Spindler K-D (2005) Ultraspiracle promotes the nuclear localization of ecdysteroid receptor in mammalian cells. Biol Chem 386:463–470 Nieva C, Spindler-Barth M, Spindler K-D (2007) Influence of hormone on intracellular localization of the Drosophila melanogaster ecdysteroid receptor (EcR). Cell Signal 19:2582–2587 Nieva C, Spindler-Barth M, Azoitei A, Spindler K-D (2008) Impact of heterodimerization on intracellular localization of the ecdysteroid receptor (EcR). Arch Insect Biochem Physiol 68:40–49 Ogura T, Minakuchi C, Nakagawa Y, Smagghe G, Miyagawa H (2005) Molecular cloning, expression analysis and functional confirmation of ecdysone receptor and ultraspiracle from the Colorado potato beetle Leptinotarsa decemlineata. FEBS J 272:4114–4128 Spindler-Barth M, Spindler K-D (2003) Ecdysteroid receptors (EcR/USP). In: Encyclopedia of Hormones and Related Cell Regulators (eds. H.L. Henry and A.W. Norman), Academic Press, New-York, London, pp 466–470. Yao TP, Forman BM, Jiang Z, Cherbas L, Chen JD, McKeown M, Cherbas P, Evans RM (1993) Functional ecdysone receptor is the product of EcR and Ultraspiracle genes. Nature 366:476–479 Yund MA, King DS, Fristrom JW (1978) Ecdysteroid receptors in imaginal discs of Drosophila melanogaster. Proc Natl Acad Sci USA 75:6039–6043

Chapter 16

Intracellular Localization of the Ecdysteroid Receptor Klaus-Dieter Spindler, Katarzyna Betan´ska, Claudia Nieva, Tomasz Gwóz´dz´ , Joanna Dutko-Gwóz´dz´ , Andrzej Oz˙ yhar, and Margarethe Spindler-Barth

Abstract The components and mechanisms of nucleocytoplasmatic shuttling of the ecdysteroid receptor, especially the presence of nuclear localization (NLS) and export signals (NES) and their interaction with importins and exportins are described. The influence of hormone and heterodimerization partner, energy supply and cell cycle on the intracellular distribution of the ecdysteroid receptor are discussed. The data are compared with results from vertebrate nuclear receptors. Keywords Ecdysteroid receptor (EcR) • exportins • importins • nucleocytoplasmic shuttling • Ran-GDP-GTP-cycle • ultraspiracle (Usp)

16.1

Introduction

In this review an overview on the recent data on intracellular distribution of the ecdysteroid receptor (EcR) and its heterodimerization partner ultraspiracle (USP) will be given. In addition the data will be discussed in light of the current knowledge on mechanisms responsible for nucleoplasmic shuttling, especially of vertebrate nuclear receptors. General features of the ecdysteroid receptor and the applicability of this nuclear receptor for gene switch experiments and as a target for insecticides are described in detail in various reviews (e.g. Spindler-Barth and Spindler, 2000, 2003, 2004; Henrich, 2005; Palli et al., 2005; see also this volume).

K.-D. Spindler (), K. Betan´ska, and M. Spindler-Barth Institute of General Zoology and Endocrinology, University of Ulm, Ulm, Germany e-mail: [email protected] T. Gwóz´dz´ , J. Dutko-Gwóz´dz´ , and A. Oz˙ yhar Department of Biochemistry, Wroclaw University of Technology, Wroclaw, Poland C. Nieva Evolutive Biology Institute (CSIC), Barcelona, Spain

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The results summarized here were obtained for EcR and USP fused with fluorescent derivatives of green fluorescent protein and expressed in mammalian cell lines. This allows monitoring of nuclear transport and visualizing the action of steroid hormone receptors into living cells (Griekspoor et al., 2007) and avoids problems associated with immunocytochemistry, which quite often led to controversial results due to artefacts (Yamashita, 2001; Scheller et al., 2000). Most importantly heterologous expression allows to investigate the fate of EcR and of USP separately, whereas in insect tissue both receptors are always present simultaneously in variable amounts.

16.2

Components of the Transport Machinery

Nuclear hormone receptors shuttle continuously between cytoplasm and nucleus (Maruvada et al., 2003). A gradient distribution of the small GTPase Ran is a key determinant in the directionality of the nuclear transport (Weis, 2003). Nuclear Ran is present mainly in GTP-bound form, whereas cytoplasmatic Ran is predominantly in the GDP-bound form. The nucleotide exchange is due to cellular compartmentalization of its regulatory proteins. RanGTP is generated by a nuclear, chromatin-bound Ran guanine nucleotide exchange factor RCC1 (RanGEF). In the cytoplasm RanGTP is hydrolyzed by GTPase-activating protein RanGAP (Bischoff et al., 1994; Fig. 16.1). Shuttling between cytoplasm and nucleus occurs through nuclear pore complexes (NPCs; Fahrenkrog and Aebi, 2003), and is mediated by a family of transport factors, called karyopherins, which includes 14 members identified so far in yeast and more than 20 in human cells (Fried and Kutay, 2003). Karyopherins recognise distinct cargo proteins that contain specific import or export signals. The molecular interactions of transport proteins with their cargos are regulated by the small GTPase Ran/TC4.

Fig. 16.1 RanGDP/GTP cycle and the control of cargo binding to importins and exportins by Ran. NPC = nuclear pore complex

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The first described and best characterised “classical” NLS (nuclear localization signal)-dependent nuclear import pathway involves importin. α and β. Importin α is a 60 kDa protein composed of three functional domains: a hydrophilic amino-terminal domain that interacts with importin β (Görlich et al., 1996), a large hydrophobic NLS-binding domain consisting of ten repeats known as armadillo arm (Conti et al., 1998), and a short acidic region that binds to the cellular apoptosis susceptibility gene product (CAS). Importin β is characterised by large domains composed of 14–15 tandemly arranged HEAT motifs (Malik et al., 1997) which bind to the importin α – NLS-protein complex and guide the subsequent translocation through the NPC by direct interaction with nucleoporins (Görlich et al., 1996). Once the NLS-cargo has entered the nucleus, the import complex dissociates after RanGTP binding to importin β, and the transport vehicles are recycled into cytoplasm separately: importin β as a complex with RanGTP, whereas importin α is bound to the mediator CAS (Kutay et al., 1997) (Fig. 16.2). Participation in nuclear transport is not the only function of karyopherins. Especially importin β has been shown to be a regulator of numerous other processes together with RanGTP (for review see: Harel and Forbes, 2004; Mosammaparast and Pemberton, 2004).

Fig. 16.2 Generalized scheme of the nucleocytoplasmic transport of proteins

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The molecular mechanism of rapid protein entry into the nucleus through the nuclear pore complex (520–1,000 macromolecules per pore per second; Ribbeck and Görlich, 2001; Smith et al., 2002) is not fully understood at the moment (Rout et al., 2000) but it has been demonstrated that nucleoporins (nuclear pore complex proteins), bind with high affinity to transport receptors and mediate both import and export (Feldherr et al., 1984). For nuclear export different carriers were described: the best investigated ones are exportin-1 (CRM1, Fukuda et al., 1997), CAS (Kutay et al., 1997) and calreticulin (Holaska et al., 2001). CRM1 (chromosome region maintenance protein; also called exportin-1) has been identified in yeast and higher eukaryotes as a member of the karyopherin β family. It binds cargos with a leucine-rich NES (nuclear export signal) in a RanGTP-dependent manner (Fornerod et al., 1997; Kudo et al., 1997). The recognized signal corresponds to the consensus Ψ − X2–3 − Ψ − X2–3 − Ψ − X − Ψ (Ψ = L, I, V, F, M; Fornerod and Ohno, 2002). Exportin-1 was characterised as an export mediator by its leptomycin B (LMB) sensitivity. The drug inhibits NES-mediated protein export due to suppression of the interaction of exportin-1 with nuclear export signals (Fukuda et al., 1997; Kudo et al., 1998). Calreticulin, whose sequence is unrelated to any known transport protein, was first described as a new export receptor by Holaska et al. (2001). Cytosol depleted of CRM1 can still support nuclear export of protein kinase inhibitor (PKI) mediated by calreticulin. Black et al. (2001) have demonstrated that a calreticulin-binding region is located in the DNA binding domain between the two zinc fingers of some nuclear receptors. The same authors proposed that the DNA binding domain of nuclear receptors, which does not contain a leucine-rich NES, may function as an export signal, recognized by calreticulin. Surprisingly, calreticulin exhibits also an inhibitory effect on transcriptional activity of some nuclear receptors e.g. androgen, retinoic acid and vitamin D (Dedhar et al., 1994) and glucocorticoid receptor (Burns et al., 1994). According to their homology to vertebrate proteins three classes of Drosophila melanogaster importins α are discriminated: α1 (Giarrè et al., 2002), α2 (pendulin encoded by the oho31 locus; Török et al., 1995; Küssel and Frasch, 1995) and α3 (Dockendorff et al., 1999). All three have a conserved phylogenetic structure containing arm repeats and an importin β binding domain. Importin α1 and α3 are present at almost constant levels throughout insect development (Giarrè et al., 2002; Máthé et al., 2000). Importin α2 is expressed in early embryogenesis and a second time in the pupal stage (Küssel and Frasch, 1995). The molecular basis of action of Drosophila importins in intracellular transport is still unclear. For instance, Drosophila melanogaster homozygous null allele of importin α2 is viable but sterile. This excludes the possibility that the protein plays an important role in nuclear import. Eventually its function as transport receptor may be restricted to a few proteins (Gorjánácz et al., 2002). However, importin α2 is critically required for male and female gametogenesis. It regulates the assembly of ring canals during females’ oogenesis. Interestingly, ring canals are functionally related to nuclear pores, both structures mediate translocation. This reflects an ancestral transport mechanism, common for vertebrates and metazoans (Gorjánácz et al., 2002). Importin α3 is required for development of larval and adult tissues (Mason et al., 2003). Importin

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α3 null mutants die around the transition from first to second larval stage, whereas adults reveal severely damaged eyes due to lack of photoreceptor cells. Lippai et al. (2000) identified a Drosophila homolog of vertebrate importin β, called Ketel. The protein shares 60% identity and 78% similarity with its human relative. Ketel reveals characteristic features of importin β. It has been found to support nuclear transport of NLS-containing cargos in digitonin-permeabilized HeLa cells and like other members of the family shows cytoplasmic localization with predominant accumulation in the nuclear envelope. However, the function of Ketel protein as a nuclear receptor is controversial, since Ketel gene is not expressed in every cell type (Lippai et al., 2000).

16.3

Biochemical and Immuno-histochemical Investigations on the Localization of the Ecdysteroid Receptor

The intracellular localization of EcR and USP has been studied previously by biochemical and immuno-histochemical techniques. With biochemical techniques both in crustaceans (Kuppert et al., 1978, 1981; Spindler-Barth et al., 1981; Londershausen and Spindler, 1981; Kuppert and Spindler, 1982) and in the insects Drosophila melanogaster (Yund et al., 1978; Maroy et al., 1978; Schaltmann and Pongs, 1982) and Chironomus tentans (Turberg et al., 1988; Turberg and Spindler, 1992) a considerable portion of the ecdysteroid binding activity was found in the cytosol. With antibodies against EcR and USP these proteins have been mainly detected in the nucleus (Riddiford et al., 2000). For a Chironomus tentans cell line it was shown that EcR was also present in the cytoplasm to a considerable degree and was shifted into the nucleus by moulting hormone (Lammerding-Köppel et al., 1998). A transport of the ecdysteroid receptor into the nucleus has been demonstrated in vitro using isolated nuclei from crayfish hypodermis and receptor containing fractions (Londershausen et al., 1982) and by photoaffinity labelling in Drosophila melanogaster Kc-cells and imaginal discs (Schaltmann and Pongs, 1982). In none of these studies the molecular mechanisms of nucleocytoplasmic shuttling and the components involved – except EcR and USP – were studied in detail. In addition, EcR/USP ratios certainly influence the mechanisms of intracellular distribution of EcR and USP.

16.4

16.4.1

Recent Studies Using Fluorescent Labelled Ecdysteroid Receptor and Ultraspiracle Localization of the Ecdysteroid Receptor in the Absence of a Heterodimerization Partner

In Drosophila melanogaster three EcR isoforms, different only in AB region, are expressed (EcRA, EcRB1 and EcRB2) (Talbot et al., 1993). The isoforms exhibit

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different distribution within mammalian cells. EcRA and EcRB1 are distributed heterogeneously. Even within one batch of cells these isoforms can be localized exclusively in the nucleus in some cells, but they can also be more or less equally distributed between cytoplasm and nucleus and in some rare cases there is even exclusively cytoplasmic localization (Nieva et al., 2005; Gwóz´dz´ et al., 2007). In contrast, EcRB2 is distributed evenly within the cell and was never found exclusively within the nucleus (Gwóz´dz´ et al., 2007). The reasons for this heterogeneous localization of EcR are not yet clear. A likely explanation for the heterogeneous localization might be caused by differences in cell cycle. We therefore performed cell cycle analyses by FACS. COS cells were synchronized and then analyzed for intracellular distribution of EcR. Preliminary experiments indicate that there is a cell cycle dependent distribution: when the percentage of cells in the S phase decreases there is also a decrease of cells with cytoplasmatic localization of EcR-B1 and an inverse relation to nuclear localization.

16.4.2

Influence of Hormone on Ecdysteroid Receptor Localization

Addition of moulting hormone slightly shifts EcR of Drosophila melanogaster into the nucleus (Nieva et al., 2007), even in the absence of USP thus confirming binding of hormone in the absence of USP as shown previously (Grebe et al., 2003, 2004). Within a cell population the proportion of cells with exclusively nuclear EcR increased after hormone treatment. Single cell analysis and quantitative measurement of fluorescence in the nucleus confirmed these results. The rapid stimulation of nuclear import caused by muristerone A leads to increased nuclear localization which persists for at least 24 h (Nieva et al., in press). This increased initial nuclear import and the shift to the nucleus in the presence of muristerone A demonstrates that hormone binding to EcR in the absence of USP is associated also with changes in receptor function. The molecular mechanism of hormone action on increased EcR import is not yet clear. The preferential localisation of EcR in the nucleus could be due to hormone induced enhanced dimerization, reduced anchoring in the cytoplasm, or a quicker import in the presence of the ligand. Another likely explanation is that the nuclear export signal situated in helix 3 (Gwóz´dz´ et al., 2007) is hidden by hormone induced homodimerization, as was shown for the heterodimer EcR/USP (Betanska et al., 2007).

16.4.3

Localization of Ultraspiracle

From the first signs of expression up to at least 24 h (Nieva et al., 2005) USP is nearly exclusively in the nucleus independent whether a heterodimerization partner is present or not. Obviously heterodimerization is no prerequisite for transport of

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USP into the nucleus. This indicates that USP like EcR as shown above can enter the nucleus independently and that intracellular distribution is regulated individually for each receptor. These results further strengthen the hypothesis that EcR and USP may be involved in additional separate functional roles, which is also supported by the fact that developmental profiles of EcR and USP do not coincide (Riddiford et al., 2000) and USP homodimers are discussed as a putative juvenile hormone receptor (Jones and Sharp, 1997).

16.4.4

Localization of Ecdysteroid Receptor/Ultraspiracle

In the presence of USP, EcR is localized exclusively in the nucleus (Nieva et al., 2005; Gwóz´dz´ et al., 2007). The increased nuclear localisation of EcR in the presence of USP indicates that the heterodimerisation between the two receptor proteins occurs already in the cytoplasm and does not afford the presence of the ligand (Cronauer et al., 2007). The retinoic acid receptor RXR, the vertebrate orthologue of USP, can not replace USP concerning nuclear transport in Cos cells (Nieva et al., 2008), although other functions of the heterodimerization partner e.g. binding to hormone response elements or transactivation (Christopherson et al., 1993; Thomas et al., 1993; Yao et al., 1993; No et al., 1996; Henrich et al., 2003; Devarakonda et al., 2003) are also mediated by RXR. This is not due to insufficient amounts of RXR since overexpression of RXR never leads to an exclusively nuclear localization of EcR (Nieva et al., 2008). A reasonable explanation is that in the absence of DNA in the cytoplasm heterodimerization is only possible via the ligand binding domain. The E-domains of USP and RXR vary and can obviously not replace each other. In contrast transactivation of the EcR/RXR complex is possible (Nieva et al., 2008) due to the highly conserved DBD of RXR in comparison to USP, which allows heterodimerization in the presence of DNA in the nucleus. Because of the exclusively nuclear localization of the heterodimerEcR/USP an effect of hormone on the translocation is not possible in vertebrate cells but has been shown in the epithelial cell line from Chironomus tentans using antibodies against EcR and USP (Lammerding-Köppel et al., 1998). It remains to be clarified, whether this is due to the different methods used for studying intracellular receptor concentration, or whether the different intracellular localization is due to species specific differences.

16.4.5

Nuclear Localization (NLS) and Export Signals (NES)

Investigation of EcR deletion mutants fused to YFP as well as immunoprecipitation experiments have revealed presence of both NLS and NES activities in this receptor (Gwóz´dz´ et al., 2007). This fact is consistent with heterogeneous distribution of EcR which, as mentioned above (see Section 16.4.1), is not restricted to any specific

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compartment of the cell. AB regions of EcRA and EcRB1 isoforms are localized within the nucleus indicating presence of NLS in their sequence. In EcRA/AB a short classical NLS, similar to SV40 one (Kalderon et al., 1984), was found between amino acid residues 134–141. It is still not known which part of the AB region of EcRB1 acts as an NLS. Interestingly the amino acid sequence of this region does not contain any of so far described NLSs which means that a completely new NLS remains to be revealed in EcRB1/AB. The other possibility is that EcRB1/AB functions as a retention signal (Lixin et al., 2001) and not as the true NLS. In contrast, AB region of EcRB2 is lacking any functional NLS. Presence or lack of the NLS within AB regions seems to explain the differences in subcellular localization of full-length EcRs. The next part which is likely to be involved in nuclear import of EcR is the fragment including C (DBD) and D (CTE, hinge) regions (Gwóz´dz´ et al., 2007). The fusion proteins containing both regions were found predominantly in the nucleus, which in turn suggests presence of an NLS. However, the results obtained for isolated regions C and D are not so unambiguous. These regions were localized in the whole cell with a slight nuclear advantage in some cases. Then it is still unclear which sequence may function as an NLS in the C-D fragment of EcR. The analysis of the primary structure did not reveal any characteristic pattern similar to the classical NLS but one cannot rule out the presence of an unknown NLS. Interpretation of the above results is further complicated by the possibility of DNA binding by DBD which can cause retention of fragments containing this domain within the nucleus. The exclusively cytoplasmic localization of LBD revealed the presence of the NES function within this domain. Analysis of its primary structure showed two leucine-rich NESs within helix 1 and 3. These results are consistent with the cytoplasmic localization of full-length EcR (see Section 16.4.1). The presence of both the NLS and NES within the EcR sequence is additionally supported by immunoprecipitation experiments which showed interaction of EcR with importin α and exportin 1 (Gwóz´dz´ et al., 2007). It indicates that EcR can shuttle between the nucleus and the cytoplasm using the mechanism described in Section 16.2. In contrast to EcR, USP seems to possess only an NLS within DBD. Only this region fused to YFP was localized exclusively within the nucleus although the exact sequence which could function as an NLS remains unknown. Presence of this domain within the nucleus could be explained by its interaction with DNA as well. However, immunoprecipitation experiment revealed that USP is able to interact with importin α but not with exportin 1, which in turn indicates that USP possesses an NLS but not a NES (Gwóz´dz´ et al., 2007). These data are consistent with the exclusively nuclear localization of USP (Nieva et al., 2005).

16.4.6

Energy Requirement

Energy requirement for nuclear transport was demonstrated using inhibitors of ATP synthesis, oligomycin and sodium azide. For the sequential phosphorylation of GMP to GDP and next to GTP, ATP is required (Zalkin and Dixon, 1992).

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Therefore an absence of ATP inhibits de novo synthesis of free GTP required to produce RanGTP (Schwoebel et al., 2002). In the presence of oligomycin, the percentage of cells with nuclear localization of separately expressed YFP-EcR or YFP-USP or coexpressed EcR/USP decreases about 50%. It indicates that a subpopulation of receptor proteins shuttles between both cell compartments (Betanska et al., 2007). Muristerone A treatment does not prevent cytoplasmatic accumulation in this case and demonstrates that ligand binding to the edysteroid receptor can not compensate for the lack of energy necessary for nuclear transport. Retention of ecdysone receptor in the nucleus is also energydependent. The exportin-1 inhibitor leptomycin (Yoshida and Horinouchi, 1999) reduces nuclear export of YFP-EcR. However, if cells were incubated with oligomycin prior to leptomycin treatment, the percentage of cells with nuclear localization of YFP-EcR was significantly reduced due to inhibition of nuclear import by oligomycin (Betanska et al., 2007).

16.5

16.5.1

Comparison of Ecdysteroid Receptor/ Ultraspiracle with Vertebrate Nuclear Receptors General Importance of the Intracellular Localization of Nuclear Receptors for Regulation of Gene Expression

In addition to the intracellular localization of nuclear receptors the intranuclear distribution of steroid hormone receptors is important for transcriptional activity. DNA binding is a prerequisite for transcription of hormone-dependent genes. Cronauer et al. (2007) have reported that DNA binding of the ecdysteroid receptor increases in the presence of hormone. The impact of the ligand has been shown also for the androgen receptor. The agonist dihydrotestosterone shifts AR quantitatively into the nucleus leading to the formation of nuclear foci which are interpreted as sites of transcriptional activity (Tomura et al., 2001). In contrast, the antagonistic endocrine disruptor 1,1-dichloro-2,2-bis(p-chlorophenyl)ethylene Translocates GFP-AR only partially into the nucleus resulting in an homogenous intranuclear distribution of GFP-AR without the formation of foci within the nucleus. A similar situation has also been found for GR, ER, PR and MR (for a review: see Kumar et al., 2006) and was also seen occasionally for EcR (Fig. 16.3). The intranuclear distribution is not stable. By fluorescence recovery after photobleaching (FRAP) analyses it has been shown that a high mobility of nuclear proteins is a common feature. Liganded nuclear receptors are less mobile than unliganded, the time of immobility is in the order of 1–2 min (for a review see: Griekspoor et al., 2007). This has led to a “hit and run” model of gene activation (Nagaich et al., 2004). Interestingly the cycling time of nuclear receptors is much faster than those for the recruitment of more than 46 factors necessary to bind to an empty promoter to initiate transcription which is in the order of an hour (for review see: Griekspoor et al., 2007).

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Fig. 16.3 Formation of nuclear clusters of YFP-EcR in CHO cells. Confocal image

16.5.2

Influence of the Ligand

The intracellular localization of individual unliganded receptor proteins is summarized in Table 16.1 and can be classified into three categories: predominant nuclear localization, predominant cytoplasmic localization and more or less even distribution between the two compartments. USP falls into the first category, EcR into the third one. There is no correlation between localization and subclass of the nuclear receptor or homo- or heterodimerizing receptors. But common to all receptors is an exclusive or predominant localization in the nucleus after addition of ligand.

16.5.3

Comparison of NLS and NES

Employment of mutagenesis and fluorescent microscopy allowed to reveal that many of vertebrate nuclear receptors contain an NLS within DBD or hinge region (so-called NLS1) (Hsieh et al., 1998; Katagiri et al., 2000; Michigami et al., 1999; Pearce et al., 2002; Picard and Yamamoto, 1987; Prüfer and Barsony, 2002; Tang et al., 1997; Ylikomi et al., 1992; Zhu et al., 1998). Additionally, vertebrate steroid receptors possess a second NLS (so-called NLS2) in LBD (Picard and Yamamoto,

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Table 16.1 Intracellular localization of unliganded nuclear receptors detected with live cell imaging techniques Nuclear receptors Subcellular localization References rAR

Cytoplasmic

GR hERα

Cytoplasmic Predominantly nuclear

ERß rMR

hTRß

Predominantly nuclear Cytoplasmatic or equally distributed Predominantly cytoplasmic Cytoplasmic Exclusively nuclear Predominantly nuclear Less nuclear than PRα Predominantly nuclear Exclusively nuclear in mouse cells, evenly distributed in Xenopus oocytes Mostly nuclear

hTRß VDR

Evenly distributed Cytoplasmic

dmEcRB1

Exclusively nuclear, evenly distributed, predominant cytoplasmic Never exclusively nuclear Exclusively nuclear in COS and HeLa cells, evenly distributed in CHO cells Predominantly nuclear

hMR Pregnane XR mPPAR hPRα hPRß hRAR Trα

dmEcR B2 dmEcRA

dmUSP

Tyagi et al., 2000; Tomura et al., 2001 Htun et al., 1996 Htun et al., 1999; Stenoien et al., 2000 Maruvada et al., 2003 Htun et al., 1996 Fejes-Toth et al., 1988; Sartorato et al., 2004 Squires et al., 2004 Feige et al., 2005 Lim et al., 1999 Lim et al., 1999 Maruvada et al., 2003 Bunn et al., 2001

Baumann et al., 2001; Maruvada et al., 2003 Zhu et al., 1998 Racz and Barsony, 1999; Michigami et al., 1999 Nieva et al., 2005; Gwóz´dz´ et al., 2007 Gwóz´dz´ et al., 2007 Gwóz´dz´ et al., 2007

Nieva et al., 2005

1987; Saporita et al., 2003; Walther et al., 2005; Ylikomi et al., 1992). Most of the identified NLSs are basic signals similar to classical mono or bipartite NLSs. Therefore, it is generally believed that nuclear import of vertebrate nuclear receptors is governed by importin α/importin β complex. On the other hand, in transport of some nuclear receptors other importins and NLSs, different than classical ones, are involved. For instance in GR, NLS1 is recognized by importin α/importin β complex but NLS2 is bound by importin 7 (Freedman and Yamamoto, 2004). Recently, a third NLS (so-called NLS0), rich in serine/threonine residues, has been found in AB region of MR (Walther et al., 2005). EcR only partly matches the characteristics of the vertebrate counterparts. Although it seems that EcR contains an NLS activity within DBD-hinge region

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(Gwóz´dz´ et al., 2007), no similarity to the classical NLSs was found. As mentioned above, the characteristic feature of the vertebrate steroid nuclear receptors is the presence of an additional NLS in LBD. Surprisingly, neither the analysis of the primary structure nor the experimental analysis of the distribution of the YFP-tagged EcR fragments revealed any potential NLS within LBD of EcR. However, the second NLS activity was found within AB regions of EcRA and EcRB1 (Gwóz´dz´ et al., 2007). Notably in contrast to above mentioned serine/threonine-rich NLS motif identified within AB region of MR (Walther et al., 2005), the signal within EcRA/AB sequence is the classical SV40-like NLS. Comparison of the primary structures of currently known EcRs suggests that the presence of such an NLS in AB region is not accidental. At least three phylogenetically distinct EcRs from insects (Ceratitis capitata and Locusta migratoria) and from tick (Amblyomma americanum), possess sequences homological to the NLS localized within the AB region of the EcRA isoform (Gwóz´dz´ et al., 2007). In USP a clear NLS activity could be observed solely within the core-DBD and no impact on USP distribution has neither the hinge region nor LBD (Gwóz´dz´ et al., 2007). In comparison to import, export of nuclear receptors is not well characterized. In nuclear receptors leucine-rich NES was only found in case of an orphan nuclear receptor NGFI-B (Katagiri et al., 2000), and its heterodimerizing partner RXRα (Cao et al., 2004) so far and usage of LMB helped to confirm that these receptors are exported by CRM1. However results obtained for other receptors like GR and TRα are inconclusive. There are data suggesting that export of these receptors is dependent on CRM1 (Maruvada et al., 2003; Savory et al., 1999) and independent of CRM1 (Black et al., 2001; Liu and DeFranco, 2000) at the same time. On the basis of amino acid sequences it seems that vertebrate steroid nuclear receptors do not contain leucine-NES at all which suggests existence of the other export pathway. Indeed, Black et al. and Holaska et al. have identified calreticulin (CRT) as a transport factor for some proteins including some nuclear receptors (Black et al., 2001; Holaska et al., 2001). CRT recognizes an amino acid sequence between Zn-modules in DBD of nuclear receptors like AR, ER, LXR, PR, RAR, RevErb, RXR, TR and VDR which suggests that this CRT-export pathway is commonly used for export of nuclear receptors. Additionally, in case of AR, ERα and MR, there was found other NES different from leucine-rich NES within LBD but it is not known whether these NESs are also recognized by CRT (Saporita et al., 2003). However, the data discussed above achieved for EcR and indicating presence of at least two leucine-rich NESs within the LBD are in marked contrast to suggestions that steroid nuclear receptors are devoid of a leucine-rich NES. Furthermore, it has been found using immunoprecipitation experiments (Gwóz´dz´ et al., 2007) that such NES activities within LBD of EcR are dependent on exportin 1/CRM1 (Betanska et al., 2007). Amino acid sequence alignment of the EcRLBDs from various invertebrates shows that the sequences identified as the NESs in D. melanogaster EcRLBD within helix 1 and 3 are highly conserved in other species (Gwóz´dz´ et al., 2007) and makes EcR a unique receptor in the steroid receptor family. The investigation of distribution of particular regions of USP and sequence analysis indicate that USP is lacking any NES activity. It is already clear that nuclear export process

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is highly complex and receptor-specific (Shank and Paschal, 2005). Comparison of three dimensional structures of LBDs from EcR, RXRα, AR and NGFI-B shows various patterns of NESs in each case (Gwóz´dz´ et al., 2007). Although the physiological reason for this remarkable diversity is yet unclear, one can speculate that it would represent the way how the different nuclear recepors, having the same overall LBD fold (Weatherman et al., 1999), individualize the actual NES fingerprint of this domain recognized by the nuclear export machinery. Obviously, the NES fingerprint for a particular receptor could be diverse and related to its molecular, and in consequence physiological, state defined by conformation of the LBD, which is evoked by the cognate ligand and/or proteins interacting with LBD (Gwóz´dz´ et al., 2007).

16.5.4

Energy Requirement

For all nuclear receptors nuclear import needs energy. Nuclear export is regulated differently for individual receptors. In the case of PR (Guiochon-Mantel et al., 1991), TRß (Baumann et al., 2001), ER, RAR (Maruvada et al., 2003), and AR (Whitaker et al., 2004), export into cytoplasm occurs through passive diffusion. This is in agreement with the fact, that as observed for USP, no leucine-rich NES could be identified for PR (Tyagi et al., 1998), TR (Bunn et al., 2001) and AR (Saporita et al., 2003) so far. Other molecular mechanisms that are involved in nuclear transport must be taken into account. For instance, prior to export, GR is being released from association with the nuclear matrix, and this step is energydependent (Tang and DeFranco, 1996). Increased nuclear export of ER and RXR in the presence of sodium azide is prevented by the cognate ligand. In contrast nuclear export of TRß upon ATP depletion was unaffected by ligand binding (Baumann et al., 2001).

16.6 16.6.1

Open Questions and Perspectives Insect Tissue

So far our studies were performed in a heterologous system, which offers some advantages as outlined above (see also Chapter 14 in this book) but the question remains, whether the results obtained in vertebrate cells reflect also the situation in insects. The presence of importins (Erdelyi et al., 1997; Dockendorff et al., 1999; Lippai et al., 2000) and the Ran/CT4-system has been demonstrated in Drosophila melanogaster and it has been shown that these components of the nuclear transport machinery are essential for early embryogenesis, oogenesis and spermatogenesis (Epps and Tanda, 1998; Mathe et al., 2000; Mason et al., 2002). Therefore, it is apparent that the cellular machinery responsible for nuclear receptor localization

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exists also in Drosophila and may play an important role for dictating a variety of developmental events associated with ecdysteroid-induced transcriptional activity. But till now it is unclear which import and export carriers are used in insects for nucleocytoplasmic shuttling of the nuclear receptors. Since a pronounced substrate specificity of carrier proteins has been shown in vertebrate systems, if more than one protein was offered (Köhler et al., 1999) one can not exclude the possibility that EcR and USP use different carriers.

16.6.2

Role of Receptor Phosphorylation

Phosphorylation of nuclear receptors is of great importance for their activity. This is not only true for the overall transcriptional activity but also for some specific effects like protein-protein interaction and thus recruitment of cofactors, stability of the nuclear receptors, DNA binding and also for intracellular localization (for a review see: Weigel and Moore, 2007). EcR and USP are phosphorylated (Rauch et al., 1998; Song and Gilbert, 1998; Nicolaï et al., 2000; Song et al., 2003) but so far it is unclear, whether the phosphorylation status influences intracellular localization of these two nuclear receptors.

16.6.3

Role of SUMOylation

Small ubiqiuitin related modifier SUMO-1 and its homologs can be conjugated to various proteins in a cascade resembling ubiquitination. But in contrast to ubiquitin SUMO can be removed by an isopeptidase and is thus no degradation signal but instead regulates mainly protein-protein interactions, prevents some targets from ubiquitin-dependent degradation and influences nucleocytoplasmic transport and intranuclear targeting (for a review see: Pichler and Melchior, 2002). Up to now no studies were performed on insect EcR or USP.

16.6.4

Importance of Heat Shock Proteins

The importance of heat shock proteins for ecdysone receptor stability (Cronauer and Spindler-Barth, 2007) and receptor function e.g. DNA binding was demonstrated previously (Arbeitman and Hogness, 2000). It is generally assumed that after hormone binding to a nuclear receptor heat shock proteins dissociate from this complex (transformation), thus triggering nuclear entry of the nuclear receptor. This dogma is not supported by experimental data in case of the glucocorticoid and the mineralocorticoid receptor where it has been shown that the heterocomplex of receptor/hormone and heat shock proteins is required for the

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movement of the receptors to the nucleus and within the nucleus to transcription regulatory sites and for retention (Tago et al., 2004; Pratt et al., 2006; Pilipuk et al., 2007). Whether this is valid for all other nuclear receptors including ecdysteroid receptor and ultraspiracle has not been investigated so far.

16.6.5

Additional Regulators of Nucleocytoplasmic Shuttling

Agonistic or antagonistic effects of nonsteroidal hormones can be explained, at least to a certain degree, by interaction with the nucleocytoplasmic shuttling. The peptide hormone relaxin is a glucocorticoid agonist and induces translocation (Dschietzig et al., 2004) whereas melatonin as an antagonist inhibits (Presman et al., 2006) import of the glucocorticoid receptor into the nucleus. Whether cross talk with other hormonal systems is involved in the regulation of nucleocytoplasmic shuttling of other steroid hormone receptors like EcR is unknown. Acknowledgements We gratefully acknowledge the financial support of our work by the Deutsche Forschungsgemeinschaft (Graduate college 1041; K.-D. S., K. B., C. N., M. S.-B.) and the Polish Ministry of Education and Science (T. G., J. D.-G. and A. O.) and Mr. M. Burret (Ulm) for the drawings.

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Chapter 17

Genomic and Nongenomic Actions of 20-Hydroxyecdysone in Programmed Cell Death Masatoshi Iga and Sho Sakurai

Abstract 20-hydroxyecdysone (20E) induces programmed cell death (PCD) in the anterior silk glands of the silkworm Bombyx mori. When glands from gut-purged larvae are incubated with 20E, the expression of early genes required for PCD occurs within 8 h. However, continuous 20E stimulation is required until 42 h to complete PCD, suggesting the involvement of an additional nongenomic mechanism. The nongenomic pathway of 20E signaling was examined using a combination of a protein synthesis inhibitor (cycloheximide), 20E, membrane-permeable second messenger analogs, protein kinases inhibitors, and a caspase 3 inhibitor. The results showed that 20E signaling initiates with ligand binding to the putative membrane ecdysone receptor, increased intracellular Ca2+, the activation of PKC and caspase 3-like protease, and finally DNA and nuclear fragmentation. Keywords Calcium signaling • caspase 3 • 20-hydroxyecdysone • membrane ecdysone receptor • nongenomic action • programmed cell death • protein kinase C

17.1

Introduction

Steroid hormones are membrane-permeable. After penetrating cell membranes, steroids bind to nuclear receptor proteins that are present in the cytoplasm and/or the nucleus, and act as ligand-dependent transcription factors (Lösel et al., 2003). Thus, the genomic action of steroids begins with gene transcription, and their hormonal effects become overt after a series of covert effects including RNA splicing,

M. Iga and S. Sakurai () Division of Life Sciences, Graduate School of Natural Science and Technology, Kanazawa University, Kanazawa 920–1192, Japan e-mail: [email protected] M. Iga Present address: Laboratory of Agrozoology, Faculty of Bioscience Engineering, Ghent University, Coupure links 653, B-9000 Ghent, Belgium G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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protein translation, posttranslational modifications, and maturation into functional proteins. Several hours or days may pass before visible effects occur, for example, changes in cellular morphology, biochemical metabolism, and secretory activity. In addition to the genomic action of steroids, cells often exhibit quick responses to steroids, which are associated with changes in intracellular second messenger levels (Lösel and Wehling, 2003; Lösel et al., 2003). Progesterone induces oocyte maturation in the African clawed frog Xenopus laevis (Wasserman et al., 1980; Baulieu et al., 1985) by decreasing cyclic AMP (cAMP) levels (Sadler and Maller, 1981) and stimulates the acrosome reaction in human sperm (Meizel and Turner, 1991; Turner and Meizel, 1995). In the chicken Gallus gallus, estrogen elicits an increase in intracellular Ca2+ levels in granulosa cells (Morley et al., 1992), and androgen stimulation increases intracellular Ca2+ within seconds in human granulosa cells during luteinization (Machelon et al., 1998). These rapid responses suggest that a nongenomic mechanism must exist in addition to the well characterized genomic mechanism, but this issue has not been investigated until recently.

17.2

Prediction of the Membrane Steroid Hormone Receptor

The nongenomic mechanism is most likely mediated through a membrane receptor. Three such receptors have been identified: the progestin receptor in the spotted seatrout Cynoscion nebulosus (Zhu et al., 2003a, b), the estrogen membrane receptor in the mouse (Filardo et al., 2002), and the dopamine/ecdysome receptor in the fruit fly Drosophila melanogaster (Srivastava et al., 2005). Note, however, that the latter is not considered to be a true steroid receptor. Progestin is a steroid hormone that induces meiotic maturation in the oocytes of C. nebulosus. It is associated with a rapid decrease in intracellular cAMP levels, which suggests the presence of a nongenomic mechanism (Thomas et al., 2002). The progestin membrane receptor is a seven-transmembrane G protein-coupled receptor (GPCR), and an inhibitory G protein (Gi/o) is activated upon progestin binding, which decreases intracellular cAMP level within minutes (Zhu et al., 2003b). Estrogen causes the rapid elevation of cAMP levels through the activation of adenylyl cyclase (Szego and Davis, 1967; Filardo et al., 2002), which indicates the presence of a GPCR coupled to Gs (Thomas et al., 2005). This GPCR has been identified as GPR30 (Filardo, 2002), which is distinct from the classical estrogen nuclear receptors ERα and ERβ (Carmeci et al., 1997). It is suggested that GPR30 is not localized to the plasma membrane, but rather to the membrane of endoplasmic reticulum (ER), where it is coupled with calcium channels (Revankar et al., 2005). Thus, estrogen binding to GPR30 would elicit a rapid increase in intracellular calcium levels that would trigger further rapid responses. However, later studies showed that GPR30 is present at the plasma membrane (Filardo et al., 2007), and thus the mode of estrogen action is still inconclusive. Another example may be observed in the D. melanogaster dopamine/ecdysome receptor. This GPCR has been identified as a membrane receptor for dopamine that also binds 20E (Srivastava et al., 2005) at a separate binding site (Evans, this

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volume); however, treatment with 20E alone is capable of increasing intracellular cAMP. Although the role of 20E in receptor activation is not yet known, it is an example of a membrane GPCR that binds two ligands. Although we have only three examples of membrane steroid receptors, evidence exists that downstream signal transduction pathways may vary with different hormones, animal species, and tissues. These pathways include the activation of protein phosphorylating enzymes by second messengers such as cAMP, Ca2+, and diacylglycerol, and the classical mitogen-activated protein kinase (MAPK) cascade (Filardo et al., 2002; Lösel and Wehling, 2003; Lösel et al., 2003).

17.3

Nongenomic Action of Ecdysteroid in Insects

Ecdysteroid is a unique steroid hormone with respect to its properties as a polyhydroxy sterol. Ecdysone secreted from the prothoracic glands is converted to a biologically active ecdysteroid, 20E. 20E is water-soluble, and other ecdysteroids such as ponasterone A and ecdysone are also more hydrophilic than mammalian steroids. 20E elicits its genomic actions (Kroeger, 1963) by binding to a heterodimeric receptor complex that consists of the nuclear ecdysone receptor (EcR) and its partner protein Ultraspiracle (USP), a mammalian retinoid X receptor analog, which acts as a transcription factor (Yao et al., 1992; Huet et al., 1993). Rapid responses to 20E have been reported in insects and crustaceans; these responses are involved in the nongenomic action of ecdysteroids. In the hypodermis of the crayfish Orconectes limosus, a 20E transport system is mediated by proteins that specifically bind ecdysone (Daig and Spindler, 1983a, b), and a Na+/K+ pump is involved in 20E uptake (Spindler and Grossmann, 1987). In the midge Chironomus thummi, puff formation is induced by changes in the Na+/K+ ratio (Kroeger, 1963). In Drosophila salivary glands, 20E induces nuclear swelling (Wünsch et al., 1993) and increases the intracellular pH through the activation of Na+/H+ exchange across the plasma membrane (Schneider et al., 1996). In the saturnid moth Hyalophora gloveri, 20E increases K+ conductivity (Schneider et al., 1996), activates adenylyl cyclase, and increases intracellular cAMP levels in wing disk epidermal cells (Applebaum and Gilbert, 1972). In the flesh fly Sarcophaga peregrina, 20E promotes the incorporation of storage proteins into fat body cells through the phosphorylation of intracellular proteins (Ueno et al., 1983; Ueno and Natori, 1984).

17.4

Programmed Cell Death of Anterior Silk Glands Induced by 20E

At the time of pupal metamorphosis, larval-specific tissues degenerate and are removed from the pupal bodies after pupation (Buszczak and Segraves, 2000). This process of elimination is carried out via programmed cell death (PCD;

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Lockshin and Williams, 1965) induced by 20E. In recent years, remarkable progress has been made regarding the molecular mechanisms of apoptosis in cultured and immunocompetent mammalian cells. In contrast, very little information is available regarding the molecular mechanism of PCD during insect development and metamorphosis. Silk gland, a larval-specific tissue, undergoes PCD in response to the pre-metamorphic peak of hemolymphatic ecdysteroids, and PCD is complete shortly after pupation (Chinzei, 1975). PCD in the anterior silk glands of the silkworm Bombyx mori begins with membrane blebbing, followed by the formation of intracellular spaces, cell shrinkage, nuclear condensation, DNA fragmentation, nuclear fragmentation, and finally apoptotic body formation (Fig. 17.1; Terashima et al., 2000; Kakei et al., 2005). Thus, PCD in the anterior silk glands involves the same fundamental changes observed in mammalian apoptosis. Generally, apoptosis in cultured mammalian and insect cells is completed quickly, in less than 10 or several hours, respectively. Such rapid progression makes it somewhat difficult to follow the temporal sequence of events or to determine what factors are involved in each step. PCD in the anterior silk glands proceeds at a substantially slower pace, requiring approximately 120 h after the initial 20E stimulus to complete

0

8

Time after exposure to 20E (h) 42 48 72 96

18 24

Transcription Translation

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d e

Requirement of 20E Nuclear condensation Cell shrinkage

a b

DNA fragmentation

c

Nuclear fragmentation Apoptotic body formation

Fig. 17.1 In vitro progression of 20E-induced PCD in anterior silk glands. Anterior silk glands from B. Mori obtained at the time of gut purge were cultured with 1 µM 20E and monitored until the formation of apoptotic bodies. (a–e) Anterior silk glands as observed under a light microscope; (a'–e') Nuclear shape was examined in each sample via DAPI staining. (a) Freshly dissected gland showing the transparent, hexagonal, epithelial cells. (b) Gland becomes opaque by 48 h. (c) Cells become round-shaped as a result of cell shrinkage. (d) Cell Shrinkage is intensified and cell boundries become irregular. (e) Cells complete PCD with the formation of apoptotic bodies. (a') A highly branched nucleus. (b') Nuclear condensation occurs 48 h after 20E challenge. (c') Nuclear condensation is intensified. (d'), Nuclear fragmentation occurs at 96 h. (e') Finely fragmented nucleus. Bar, 100 µm (a–e), 20 µm (a’–e’). See Terashima et al. (2000) for details

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apoptotic body formation (Fig. 17.1e). This lengthy process makes it easy to study individual events that occur at the cellular and nuclear levels. Because the anterior silk glands are composed of a simple epithelium consisting of only large, flat, deformed hexagonal cells, they are easy to manipulate in vitro and suitable for observing the changes in cellular and nuclear morphologies. The nuclei are not spherical, but instead ramify into thin branches distributed throughout the cell (Fig. 17.1á), and thus the morphological changes that accompany PCD, such as nuclear condensation (Fig. 17.1b΄) and fragmentation (Fig. 17.1d΄), are clearly observable. We took advantage of these features to outline the nongenomic effects of 20E and its signal transduction pathway during PCD in the anterior silk glands of B. mori.

17.5

Genomic Action of 20E in 20E-Induced PCD

In the anterior silk glands of B. mori, completion of PCD requires both genomic and nongenomic effects of 20E. In 20E-induced PCD in vitro, all gene expression required for PCD is completed within the first 18 h of culture, during which most of the early genes exhibit expression profiles that are characteristic of individual isoforms (Sekimoto et al., 2006, 2007). The temporal profiles of gene expression during the prepupal period indicate that the EcR-A and Usp-2 isoforms are involved; neither EcR-B1 nor usp-1 increased in response to 20E stimulation in the anterior silk glands of just gut purged larvae (Sekimoto et al., 2006). Among the early gene isoforms examined so far, E74A, E75A and B, BHR-3, and BR-C z2, z3, and z4 exhibited individually distinctive expression profiles. However, few data are available regarding the hierarchical control of these genes in B. mori compared to that available in Drosophila (Jiang et al., 2000). Recently, RNAi against these early genes has been used in the cockroach Blattella germanica to show that EcR-A is closely involved in PCD of the prothoracic glands (Cruz et al., 2006). However, little information is available regarding cross talk between the genomic and nongenomic actions of 20E.

17.6

Presence of the Membrane Receptor for 20E

When the anterior silk glands of B. mori larvae were cultured with 1 µM 20E, PCD was complete in approximately 120 h. As de novo gene expression is a prerequisite for PCD, to determine the time frame of PCD-requisite gene expression stimulated by 20E, α-amanitin, a transcription inhibitor, was added at various time points after 20E challenge. Cell death was completely inhibited by the addition of α-amanitin at 0 or 6 h after 20E challenge, but no effect was observed when the drug was added at 8 h or later (Terashima et al., 2000). These results indicate that

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all gene expression required for PCD was finished within 8 h of the 20E stimulus. However, when anterior silk glands were transferred to 20E-free culture medium at various time points after 20E challenge, we found that 20E stimulus was necessary until 42 h of the initial challenge to complete PCD (Terashima et al., 2000). This raises the question of what occurred between 8 and 42 h. At the very least, the effects of 20E may not be accompanied by gene expression or mediated by the classical nuclear ecdysone receptor. To detect the presence of a membrane receptor for 20E, we measured ecdysone-binding activity in the membrane fraction prepared from anterior silk glands. We identified a protein that exhibited a binding constant (Kd) of 17 nM for ponasterone A, a phytoecdysteroid that is physiologically more active than 20E (Elmogy et al., 2004). In addition, the membrane fraction exhibited only weak binding activity for nonsteroidal ecdysone agonists such as RH-2485, -5849, and -5992, even though these agonists show higher binding affinity for the nuclear ecdysone receptor complex than 20E (Minakuchi et al., 2003). This was also the case for the dopamine/ecdysome receptor (Srivastava et al., 2005). In addition, Western blot analysis of the membrane fraction and nuclear extracts using anti-Bombyx EcR-A antiserum did not reveal any immunoreactive bands in the membrane fraction, indicating that this membrane protein is distinct from EcR. The solubilization of these binding sites showed that the putative membrane EcR (mEcR) may be an integral plasma membrane protein (Elmogy et al., 2004). These results support the existence of a membrane ecdysone receptor that is distinct from the nuclear EcR.

17.7

Separation of the Genomic and Nongenomic Actions of 20E

Cycloheximide (CHX), a translation inhibitor, induces apoptosis in Drosophila S2 cells (Fraser et al., 1997). In the anterior silk glands, a high concentration of CHX (2 mM) induces type B cell death (Fig. 17.2; Iga et al., 2007). This type of cell death is distinct from 20E-induced PCD; although it involves nuclear and DNA fragmentation, it is not accompanied by cell shrinkage, nuclear condensation, or apoptotic body formation. A much lower concentration of CHX (0.2 mM) did not induce any changes associated with PCD, but inhibited protein synthesis to the same degree as 2 mM CHX (Iga et al., 2007). To determine the genomic action of 20E, we observed the progress of PCD in the anterior silk gland after the simultaneous addition of 0.2 mM CHX and 20E. Although little change occurred in cell morphology as observed under a light microscope, DAPI staining showed that nuclei became fragmented after nuclear condensation, and DNA electrophoresis revealed the ladder pattern characteristic of PCD. This type of cell death, induced by 20E and 0.2 mM CHX, is known as type C cell death (Fig. 17.2; Iga et al., 2007). Type B and C cell death suggest that the

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Fig. 17.2 Comparison of morphological changes that occur during cell death induced by 2 mM CHX (type B cell death) or 0.2 mM CHX and 1 µM 20E (type C cell death). (a–f) Anterior silk glands as observed under a light microscope; (a'–f') DAPI staining corresponding to individual light microscopic views. (a and d) Freshly dissected glands. (b) Glands at 96 h. (c and f) Glands at 120 h. (e) Glands at 48 h. showing nuclear condensation. Note that in type B cell death, nuclear fragmentation occurs without nuclear condensation (b'), whereas nuclear condensation does occur in type C cell death (é). In both types of cell death, cell shrinkage does not occur, although the cells do not maintain their original morphology. Bar, 100 µm (a–f), 20 µm (a’–f’)

molecular mechanism that induces changes in cell morphology, such as cell shrinkage and apoptotic body formation, is different from the mechanism that mediates nuclear condensation, nuclear fragmentation, and DNA fragmentation (Fig. 17.3). Changes in cell morphology require 20E-induced de novo gene expression and protein synthesis, which is likely mediated via the genomic pathway using EcR/USP. However, gene expression and protein synthesis are not required for the pathways that result in nuclear condensation and nuclear and DNA fragmentation. These results clearly indicate the presence of a signal transduction pathway that is activated by the binding of 20E to a putative mEcR (Fig. 17.4).

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1 µM 20E + 0.2 mM CHX

1 µM 20E

Nuclear condensation

Cell shrinkage Nongenomic DNA fragmentation Genomic Nuclear fragmentation

Apoptotic body formation

Type B cell death

Type A cell death

Type C cell death

Fig. 17.3 Flowchart of PCD progression in normal 20E-induced PCD (type A) versus type B and C cell death. The genomic action of 20E primarily governs cell shrinkage and apoptotic body formation, whereas a nongenomic mechanism controls nuclear condensation, DNA fragmentation, and nuclear fragmentation

20E Membrane receptor Cytosol Nucleus Nuclear receptor

Ca2+

EcR USP

Early gene Effector gene

Cell shrinkage Apoptotic body formation Genomic

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Interaction? PKC

Caspase 3

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Fig. 17.4 Schematic representation of putative 20E signaling in anterior silk glands during 20Einduced PCD

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17.8

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Signaling Cascade Downstream of 20E

Results obtained via the time-lapse addition of α-amanitin and CHX, and 20E withdrawal experiments support the existence of a nongenomic mechanism for 20E. If this is the case, second messengers and subsequent protein kinase activation may also be involved in the pathway. To examine this issue, anterior silk glands were incubated with 20E for 18 h. Our previous experiments showed that the addition of CHX after 18 h did not affect the progression of PCD. Then, 20E was replaced with one of the following membrane-permeable second messenger analogs: dbcAMP, dbcGMP, calcium ionophore, or phorbol ester (Iga et al., 2007). Under these conditions, only the calcium ionophore mimicked the effects of 20E, although cell death stopped short at nuclear and DNA fragmentation. These results indicate that Ca2+ is a potential second messenger in 20E signaling. To confirm these results, we examined the possible involvement of protein kinase(s) using various protein kinase inhibitors. Inhibitors against mitogen-activated protein kinase kinase (MEK), protein kinase A (PKA), and c-Jun N-terminal kinase (JNK) did not affect 20E-induced PCD. In contrast, an inhibitor of protein kinase C (PKC) blocked DNA and nuclear fragmentation, which appeared in type C cell death, suggesting that the Ca2+-PKC pathway participates in the nongenomic action of 20E. In addition, the PKC inhibitor inhibited changes in cell morphology, although it did not inhibit nuclear condensation. These changes in cell morphology do not appear in type B or C cell death (Fig. 17.2), indicating that this effect was the result of the genomic action of 20E. Furthermore, these results provide evidence of cross talk between the nongenomic mechanism occurring downstream of PKC and the genomic action of 20E (Fig. 17.4). The addition of the PKC inhibitor at 24 h inhibited both DNA and nuclear fragmentation, but its addition at 48 h elicited no inhibitory effects on the progression of cell death, indicating that active PKC is required for at least 24 h after 20E stimulation to complete PCD. This is consistent with our previous observation that 20E stimulation is required for approximately 40 h to complete PCD (Fig. 17.1).

17.9

Activation of Caspase 3-Like Protease by 20E

Unlike apoptosis in mammalian cells, the cell death signal cascade in insects is not well-known, with the exception of PCD in the salivary glands of D. melanogaster (Jiang et al., 2000; Yin and Thummel, 2005). Incubating salivary glands from white puparia with 20E induces PCD. The death genes hid, reaper, and grim are induced during early gene expression, and are thought to function in the signal cascade leading to caspase 3 activation. Caspases are crucial for PCD and highly conserved among a wide variety of organisms, from the nematode Caenorhabditis elegans to mammals (Twomey and McCarthy, 2005). Caspase 3 is a key factor for DNA and nuclear fragmentation, and thus inhibitors developed against human caspase 3 are

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widely used in the study of cell death (Nezis et al., 2006). Accordingly, we examined the effects of a caspase 3 inhibitor on 20E-induced PCD in B. mori anterior silk glands. Incubation with this caspase 3 inhibitor blocked nuclear and DNA fragmentation, but not nuclear condensation. When caspase 3-like activity was measured using an artificial substrate, enzyme activity appeared 72 h after the initial 20E challenge and peaked at 96 h (Iga et al., 2007). The proteolytically activated form of Bombyx caspase 3 is detectable via Western blotting using an antibody raised against the active human caspase 3 (Cullen and McCall, 2004). Using this antibody, Western blotting revealed an immunoreacitve band that appeared at 72 h and peaked at 96 h (T. Yasanga, Y. Kaneko and S. Sakurai, unpublished data, 2006). These results coincided with the timing of both DNA and nuclear fragmentation (Fig. 17.1), suggesting that a caspase 3-like protease participates in PCD in B. mori anterior silk glands. As observed in type C cell death, the simultaneous presence of 20E and 0.2 mM CHX induced nuclear condensation and DNA fragmentation (Fig. 17.3), whereas the caspase 3 inhibitor did not prevent nuclear condensation. These results indicate that nuclear condensation in Bombyx anterior silk glands is independent of a caspase 3-like protease pathway and is regulated independently of DNA fragmentation. This is also the case in Drosophila salivary glands (Nezis et al., 2006), although its relationship with the genomic effect of 20E remains to be elucidated.

17.10

Conclusions

As summarized in Fig. 17.4, two pathways result in changes to cell morphology. The first requires gene expression via EcR/USP (genomic action), whereas the second pathway, which involves nuclear condensation and nuclear and DNA fragmentation (nongenomic action) does not. We also demonstrated the involvement of PKC and caspase 3-like protease in the signal transduction pathway downstream of the putative mEcR. These pathways do not act independently. Because gene expression is prerequisite for 20E-induced PCD, communication may occur between genomic and nongenomic processes. Likewise, because the PKC inhibitor affected cell morphology in addition to inhibiting DNA and nuclear fragmentation, communication likely takes place between nongenomic and genomic processes. Cross talk between genomic and nongenomic processes has not yet been reported in other animal or cultured cells, probably because of the difficulty in tracking sequential cellular events during PCD. A single steroid hormone may act on both the classical nuclear receptor and a putative membrane receptor, which induces PCD via the integration of EcRmediated genomic effects and mEcR nongenomic effects. Cross talk, as observed between genomic and nongenomic processes in 20E-induced PCD, may commonly occur in response to steroids in animal cells. The isolation and identification of putative plasma membrane steroid receptors are crucial to clarify this issue.

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Wünsch S., Schneider S., Schwab A., and Oberleithner H. (1993) 20-OH-ecdysone swells nuclear volume by alkalinization in salivary glands of Drosophila melanogaster. Cell Tissue Res. 274: 145–151. Yao T.P., Segraves W.A. Oro A.E., McKeown M., and Evans R.M. (1992) Drosophila ultraspiracle modulates ecdysone receptor function via heterodimer formation. Cell 71: 63–72. Yin V.P., and Thummel C.S. (2005) Mechanisms of steroid-triggered programmed cell death in Drosophila. Semin. Cell Dev. Biol. 16: 237–243. Zhu Y., Bond J., and Thomas P. (2003a) Identification, classification, and partial characterization of genes in humans and other vertebrates homologous to a fish membrane progestin receptor. Proc. Natl. Acad. Sci. USA 100: 2237–2242. Zhu Y., Rice C.D., Pang Y., Pace M., and Thomas P. (2003b) Cloning, expression, and characterization of a membrane progestin receptor and evidence it is an intermediary in meiotic maturation of fish oocytes. Proc. Natl. Acad. Sci. USA 100: 2231–2236.

Chapter 18

Rapid, Non-Genomic Responses to Ecdysteroids and Catecholamines Mediated by a Novel Drosophila G-Protein-Coupled Receptor Peter D. Evans, Deepak P. Srivastava, and Vincenzina Reale

Abstract Classically, steroid hormones have been thought to mediate their actions by binding to intracellular proteins that migrate to the nucleus and induce changes in gene expression. However, it is now becoming clear that steroids may also induce rapid actions through the activation of various second messenger pathways by cell surface receptors. Considerable controversy exists over the mechanisms underlying these non-genomic effects. Thus, some effects may be attributable to the allosteric actions of steroids on ligand-gated ion channels. Other effects may be mediated via the activation of G-protein coupled second messenger pathways, including in a few cases the direct activation of specific G-protein coupled receptors (GPCRs). This chapter reviews the evidence for the non-genomic actions of ecdysteroids and parallels are drawn with the non-genomic actions of vertebrate steroids in a number of systems. Possible molecular mechanisms for non-genomic actions of steroids are discussed. These include the possible actions of steroids on nuclear receptors that migrate to the cell surface and also their actions on cell surface receptors including, GPCRs. Recent evidence for the actions of ecdysteroids on insect cell surface receptors, including a recently cloned Drosophila GPCR, DmDopEcR, is reviewed. The direction of future research in the area of non-genomic actions of ecdysteroids is considered. Keywords Non-genomic actions of steroids • ecdysteroids • G-protein coupled receptors • dopamine • Drosophila

P.D. Evans() and V. Reale The Inositide Laboratory, The Babraham Institute, The Babraham Research Campus, Babraham, Cambridge, CB22 3AT, UK D.P. Srivastava Present address: Department of Physiology, Northwestern University, Feinberg School of Medicine, 303 E. Chicago Ave., Ward 7-174, Chicago, Il 60611, USA e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Introduction

Classically, steroid hormones, including ecdysteroids, have been thought to mediate their actions by binding to intracellular protein receptors that migrate to the nucleus and induce changes in gene expression (Fig. 18.1). Indeed the classical work on the action of ecdysteroids on the puffing patterns of insect salivary gland giant polytene chromosomes provided the first insights into the molecular mechanisms underlying the steroid control of gene expression (Clever and Karlson, 1960; Ashburner et al., 1974). Since that time the insect model system Drosophila melanogaster has been the focus of many genetic studies to define the molecular actions of steroid hormones (Koelle et al., 1991; Thomas et al., 1993; Baker et al., 2003). Such studies have

Steroid c

a b

Nuclear receptor at plasma membrane Nuclear receptor

GPCR

cyclic AMP Calcium MAPK

5 4 TMTM 3 TM TM 6 TMTM 1 2 TM 7

α GDP

γ β

mRNA

Protein Synthesis Fig. 18.1 Steroid hormone signaling. (a) Conventionally steroid hormones are thought to be lipophylic hormones and to be able to enter the cell easily. They then bind with a range of ‘nuclear’ receptors which transport them into the nucleus where the complex binds to the DNA and initiates changes in gene expression. Non-genomic actions of steroids are very rapid and mediate changes in second messenger levels without any changes in gene expression or protein synthesis. (b) It has been postulated that these rapid non-genomic effects could be mediated via steroid interactions with the nuclear binding proteins that migrate to become closely associated with the plasma membrane. Once activated the complex is then proposed to bring about rapid changes in second messenger levels. (c) It has also been proposed that steroids may interact with specific seven transmembrane-spanning GPCRs to activate second messenger pathways via the activated G-proteins (See Color Plates)

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shown that 20-Hydroxyecdysone (20E) binds to the ecdysone nuclear receptor (EcR), which in turn heterodimerises with the ultraspiracle receptor (USP). In the nucleus this complex binds to ecdysone response elements on the promoters of target genes, thereby altering their expression. These effects take hours to days to manifest and are known as genomic actions. In Drosophila, the major ecdysteroids produced are Ecdysone (E), 20E and Makisterone A (MaA) (Fig. 18.2). Ecdysone is the main ecdysteroid produced and secreted by the ring gland (Riddiford and Truman, 1978; Truman, 1988). It is then metabolised in the peripheral tissues into 20E which is believed to be the more biologically active form of the hormone in most tissues. However, recent evidence has shown that E, and not 20E, may be responsible for some insect ecdysteroid responses. For example, it was shown that E was as effective as 20E in promoting proliferation of neuroblasts in the early development of Manduca sexta (tobacco hornworm) (Champlin and Truman, 1998a, b). Evidence that the Drosophila ring gland also produces 20-deoxymakisterone A, which in turn is metabolised into MaA in fat body, has come from gas chromatography mass spectrometry studies (Redfern, 1984), and it has been suggested to function as an additional steroid hormone in Drosophila (Cottam and Milner, 1997). During development ecdysteroids control many of the processes involved in growth and maturation. Time specific increases and decreases in haemolymph edysteroid titres are accompanied by distinctive metamorphic processes and have been shown to regulate a vast number of biological processes in insects, including morphogenesis, proliferation, programmed cell death, cuticle synthesis and the timing of development (see Riddiford, 1993; Truman, 1996; Thummel and Chory, 2002). Thus, these

OH

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OH HO

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OH HO H

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OH HO

H

O Makisterone A

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N N O H RH - 0345, halofenozide

Fig. 18.2 Structures of ecdysteroids together with those of the bisacylhydrazine insecticides, RH-5849 and RH-0345. E and 20-deoxymakisterone A are released from the Drosophila ring gland and are converted into 20E and MaA in the peripheral tissues (Modified from Srivastava et al., 2005)

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hormones can also function as triggers and coordinators of many aspects of insect development during metamorphosis.

18.2

Rapid, Non-genomic Actions of Steroids

It is now clear that, in addition to their genomic actions on the control of gene expression, a wide range of vertebrate steroids also display rapid non-genomic actions that have time courses of seconds or minutes (Lösel and Wehling, 2003; Lösel et al., 2003). However, the molecular mechanisms underlying these rapid, non-genomic actions of steroids remain highly controversial and appear to show much hormone, cell and tissue specificity (Fig. 18.1). Some effects have been proposed to be due to the non-specific effects of steroids on the fluidity of lipids in the plasma membrane, whilst others have been proposed to be due to the allosteric effects of steroids on the activity of ligand-gated ion channels. Examples include the action of corticosterone on inhibiting acetylcholine induced currents through nicotinic receptors in PC12 cells (Shi et al., 2001), the effects of progesterone on the GABA-A receptor (Harrison and Simmonds, 1984; see Akk et al., 2005) and the actions of pregnenolone on NMDA receptors (Harrison and Simmonds, 1984; Bowlby, 1993). However, steroids have also been shown to rapidly activate G-protein coupled second messenger pathways that can change the activity of adenylyl or guanylyl cyclase, modulate the activity of ion channels, increase mitogen-activated protein kinase (MAP kinase) or phosphoinositide 3-kinase (PI3 kinase) activity or change intracellular calcium levels (Cato et al., 2002; Lösel and Wehling, 2003). Moreover, many of these effects have been suggested to be mediated by specific receptors expressed at the cell surface (Lösel and Wehling, 2003; Toran-Allerand et al., 2002; Thomas et al., 2007). A number of studies have suggested that these rapid effects are mediated via nuclear receptors expressed at the cell surface (Pietras and Szego, 1977; Razandi et al., 1999; Wade et al., 2001) (Fig. 18.1). Substantial evidence supports the idea that the nuclear estrogen receptor α (ERα) and the nuclear estrogen receptor β (ERβ) may show cell membrane localization in a range of estrogen sensitive cells (Razandi et al., 2004; Pedram et al., 2006). It has recently been suggested that the human nuclear ERα, ERβ, progesterone and androgen receptors may express a sequence highly homologous to the F(X6)LL motif found in many heptahelical G-protein coupled receptors (GPCRs) which is responsible for their export from the endoplasmic reticulum and their plasma membrane localization (Pedram et al., 2007). This localization sequence is suggested to mediate the palmitoylation of the nuclear receptors which facilitates their association with caveolin-1 and their subsequent membrane localization and steroid signalling. In contrast to the above, a number of recent studies have suggested that in some cases the rapid, non-genomic actions of steroids may be mediated by binding to and functional activation of specific GPCRs (Fig. 18.1). These include the rat oxytocin receptor, which was suggested to bind progesterone (Grazzini et al., 1998), and

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a novel progesterone-binding family of GPCRs that has been cloned from sea trout, mouse and humans, which have also been shown to regulate MAP kinase activity and suggested to play a role in meiotic maturation of oocytes (Zhu et al., 2003a, b; Thomas et al., 2007). However, the best characterized steroid activated GPCR is the orphan receptor GPR30. This receptor has been shown to mediate the estrogen-induced activation of extracellular signal-related kinase 1 (ERK1) and ERK2 via the transactivation of the epidermal growth factor through the release of heparin-bound epidermal growth factor in SKBR3 breast cancer cells that lack ERα and ERβ (Filardo et al., 2000; Thomas et al., 2005; Filardo and Thomas, 2005). In addition, GPR30 has been shown to mediate 17 β-estradiol regulation of c-fos in breast cancer cells (Maggiolini et al., 2004). GPR30 has been suggested to function at the plasma membrane in both SKBR3 cells and in transfected HEK-293 cells (Filardo et al., 2007). However, this suggestion has been challenged by Revankar et al. (2005, 2007) who suggest that GPR30 may mediate rapid cell signalling as a transmembrane intracellular estrogen receptor since their experimental evidence suggests it is predominantly expressed in the endoplasmic reticulum. They further suggest that there is insufficient GPR30 expressed on the cell surface to initiate signalling in response to impermeable ligands in the cell lines that they examined. A functional intracellular localization of a nuclearized Type 1 Sphingosine 1-phosphate GPCRs has also been reported recently (Liao et al., 2007). Two studies have reported rapid, non-genomic actions of estrogen and xenoestrogens in both pancreatic α- and β-cells (Nadal et al., 2000; Ropero et al., 2002). These effects were associated with a membrane bound receptor that was unrelated to the intracellular estrogen receptors. Interestingly, both studies showed that the catecholamines, noradrenaline, dopamine and adrenaline were able to displace estradiol-peroxidase (E-HRP) binding in these cells (Nadal et al., 2000; Ropero et al., 2002). However, when a number of common α- or β-adrenergic antagonists were also tested, no displacement of E-HRP was found, thus demonstrating that this catecholamine pharmacology is different to either classical α- or β-adrenergic pharmacology. This receptor pharmacology has also been described previously in both vertebrate blood vessels (Hirst and Neild, 1980; Hirst et al., 1982; Benham and Tsien, 1988) and in nervous tissue (Yawo, 1996, 1999). It is characterised by an equal activation by noradrenaline, dopamine and adrenaline, whilst displaying a unique pharmacology unlike that of α- or β-adrenergic receptors. These receptors have been termed ‘γ-adrenergic receptors’.

18.3

Non-Genomic Actions of Ecdysteroids

Non-genomic actions of ecdysteroids, with different time courses have also been described in a range of insect and crustacean preparations (see Tomaschko, 1999; Thummel and Chory, 2002; Schlattner et al., 2006 for references) but the molecular mechanisms underlying these effects are not well understood. In many cases these actions are very rapid, taking place within seconds or minutes, and do not require

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protein synthesis or gene transcription. Thus, ecdysteroids have been shown to rapidly modulate electrical activity in insect neurosecretory cells in the moth Manduca sexta (Ruegg et al., 1982) and to rapidly depress synaptic efficacy at crayfish (Cooper and Ruffner, 1998) and at Drosophila third instar larval (Ruffner et al., 1999; Li et al., 2001) neuromuscular junctions. Specific receptors that respond to either E or 20E have also been described in crustacean, insect and tick sensory systems (see Tomaschko, 1999) and hypothesised to under lie either pheromonal roles or to act as feeding deterrents when released from plants expressing high levels of ecdysteroids. Ecdysteroids control many aspects of the metamorphosis of the insect nervous system (see Truman, 1988, 1996) including neuronal maturation (Booker and Truman, 1987), remodelling (Truman and Reiss, 1998; Brown et al., 2006) and cell death (Truman and Schwartz, 1984). Rapid actions of ecdysteroids have been described at various stages of insect development, including the control of cell proliferation during optic lobe neurogenesis in Manduca (Champlin and Truman, 1998a). In Manduca sexta where eye development is thought to be similar to that in Drosophila (Champlin and Truman, 1998b), proliferation of cells in the optic anlagen that give rise to the neural processing regions of the optic lobe, become sensitive to the presence of ecdysteroids at the wandering stage of larval development. Proliferation of the cells can be turned on and off rapidly by repeated removal or addition of permissive levels of ecdysteroids. These effects are very rapid (less than 1 h) and are unusual since they are likely to be mediated by E rather than 20E, since the former is the predominant steroid in Manduca haemolymph at this stage of development. In addition, Champlin and Truman (2000) have provided evidence that this ecdysteroid signal can be sharpened by an ecdysteroid induced reduction in nitric oxide (NO) which removes a NO based inhibition of cell proliferation. The effect of the reduction of NO levels is very rapid, occurring within 15 min, again suggesting a non-genomic action for the E. During the development of the eye imaginal disc in the Drosophila third instar larva, a wave of retinal specification moves from the posterior margin of the disc and expands anteriorly (Wolf and Ready, 1993). Associated with the anterior boundary of retinal patterning are a synchronized cell arrest in G1, a relaxation of heterochromatic gene silencing and a coordinated apical-basal contraction of cells that produces the indentation of the ‘morphogenic furrow’ (Ready et al., 1976; Brennan et al., 2001). It is now clear that the progression of the morphogenic furrow across the disc is dependent upon the presence of ecdysteroids but not on the presence of the nuclear edysteroid receptor, EcR (Brennan et al., 1998, 2001). Equally, progress of the morphogenic furrow in embryonic eye development of the retina of the locust, Schistocerca americana, was not blocked by the nuclear EcR antagonist cucurbitacin B (Dong et al., 2003). Further, in the tobacco hornworm, Manduca sexta E, as well as 20E, is active at promoting the progression of the furrow and the effects are rapidly reversible (Champlin and Truman, 1998a, b). A wide range of non-genomic effects of ecdysteroids on ion transport, cyclic AMP production, GABAA receptor activity and the phosphorylation of insect storage proteins have been described (see Tomaschko, 1999). Many of these

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seem to be involved in the lysis of larval tissues during metamorphosis. Thus, the insect haemolymph protein, HP19, appears to mediate the non-genomic effect of ecdysteroids on acid phosphatase activity in fat body from the rice moth, Corcyra cephalonica (Arif et al., 2004). Here, although a minimum of 4 h was required in cultured fat bodies for the stimulation of the enzyme activity by 20E, it was unaffected by transcriptional or translational inhibitors and further in vitro studies with fat body homogenates showed a rapid stimulation of the enzyme activity within seconds to 1 min. A further non-genomic action of 20E has been described in the steroid-induced programmed cell death of the silkworm silk glands. Here 20E appears to mediate a non-genomic action on nuclear condensation, and DNA and nuclear fragmentation (Iga et al., 2007). However, the exact time course of action of these effects is not clear since the preparation requires a pre-exposure to 20E for 8 h before these effects can be seen. It has been suggested that these nongenomic effects might be initiated by binding to a membrane receptor resulting in intracellular calcium increases and the activation of protein kinase C. Indeed, the phosphorylation of the receptor for the storage protein, hexamerin has been suggested to be catalyzed partly by a tyrosine kinase which is activated by 20E through a non-genomic action (Arif et al., 2003). The presence of a potential membrane bound E receptor in the anterior silk glands of the silkworm, Bombyx mori has been suggested from binding studies with [3H]-ponasterone, a plant ecdysteroid (Elmogy et al., 2004). This, shows different steroid binding properties from the nuclear EcR but further experiments are required to determine if it is responsible for the effects described by Iga et al. (2007) (see above) or for the rapid 20E mediated increases in cyclic AMP levels described in the anterior silk glands (unpublished observations quoted in Elmogy et al., 2004). However, it should be noted that a cloned Drosophila GPCR has recently been shown to bind and to be activated by ecdysteroids in heterologous expression systems (see below). In a parallel with the suggestions that some of the rapid non-genomic actions of vertebrate steroids could be mediated via nuclear receptors trafficking close to the plasma membrane of sensitive cells (see above), it has been proposed that the nuclear EcR receptor may also do this (Schlattner et al., 2006). This suggestion was based on immunocytochemical evidence that indicated that EcR could be localised close to the plasma membrane in both lateral and median protocerebral neurosecretory cells in the last instar larvae of the bloodsucking insect, Rhodnius prolixus. A conformational compatibility model for EcR was proposed to explain the differential coupling of the receptor to different genomic and non-genomic pathways depending on the conformation of the receptor induced by different agonists and different scaffolding or binding proteins. However, the validation of this theory will require evidence that the antibody used is specifically interacting only with EcR and not with other proteins in these cells. In this respect it is interesting to note that antibodies against the C-terminus of the nuclear estrogen receptor ER-α, also appear to cross react with the distinct membrane bound estrogen receptor ER-X (Toran-Allerand et al., 2002). It will also be of interest to see if the ultraspiracle receptor, which dimerises with EcR to form the functional nuclear

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ecdysteroid receptor, is also associated with EcR when it is expressed close to the plasma membrane.

18.4

Identification and Characterization of an Orphan Drosophila GPCR That Can Mediate Rapid, Non-Genomic Actions of Ecdysteroids

We have recently cloned and characterized an orphan GPCR from Drosophila that binds and responds to both dopamine and to ecdysteroids (Srivastava et al., 2005). This receptor may well represent the insect equivalent of the vertebrate ‘γ-adrenergic receptor’, described above, which has not yet been cloned. We found this receptor (CG18314; DmDopEcR) as part of an investigation into the deorphanization of a range of putative Drosophila aminergic receptors identified by a bioinformatic analysis of the genome (Brody and Cravchik, 2000). Three of these receptors (CG6919, CG6989 and CG7078) turned out to encode a new family of β-adrenergic-like octopamine receptors (Maqueira et al., 2005; Evans and Maqueira, 2005). CG18314 also shared many sequence characteristics with vertebrate β-adrenergic receptors but was named DmDopEcR since the receptor could be activated preferentially by the biogenic amine dopamine (see below). Heterologous expression of the DmDopEcR receptor in CHO cells indicated that of the catecholamines tested unusually only dopamine could significantly, but modestly, couple the receptor to a stimulation of cyclic AMP levels in both a concentration dependent and saturable manner (Srivastava et al., 2005). To investigate if this modest degree of stimulation might be due to the poor expression of an insect receptor in a vertebrate cell line, we further characterized the expression of the receptor using the baculovirus expression system in the insect Sf9 cell line. In this latter expression system the dopamine response was both more efficient and showed a higher affinity than when expressed in CHO cells. However, its agonist profile was essentially the same as when expressed in CHO cells. In addition, in both systems studied, a range of vertebrate and insect synthetic receptor agonists and antagonists revealed that pharmacologically DmDopEcR could not easily be classified as similar to any particular vertebrate, or invertebrate, dopaminergic or adrenergic receptor subtype. Another common characteristic of GPCRs is their ability to interact with more than one second messenger system. Thus, we investigated if the activation of DmDopEcR could lead to changes in other second messenger pathways (Srivastava et al., 2005). We were unable to demonstrate any receptor mediated elevations in intracellular Ca2+ levels or in the activation of the MAP kinase pathway as measured by changes in the phosphorylation of ERK1 or ERK2. However, dopamine induced a receptor mediated, time-dependent and concentration-dependent activation of the PI3 kinase pathway via an increased phosphorylation of Akt, suggesting that one possible effect of dopamine through this receptor could be to influence cell survival (Scanga et al., 2000).

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As the response to dopamine in both the CHO and Sf9 cells lines was modest, it suggested the possibility that this may not be the only or the most effective, endogenous ligand for DmDopEcR. The unique pharmacology displayed by DmDopEcR, with its ability to bind both adrenergic and dopaminergic ligands, showed some similarity to an observed pharmacological response in vertebrate tissues mediated by the so-called ‘γ-adrenergic receptor’ (see above). In addition, in pancreatic cells the rapid, non-genomic actions of estrogen appear to be mediated via a cell surface receptor that also shows ‘γ-adrenergic pharmacology’ (Nadal et al., 2000; Ropero et al., 2002). Thus, we investigated the ability of DmDopEcR to bind the insect steroids (ecdysteroids) E, 20E, MaA and the plant analogue of E, ponasterone A (PoA) (Srivastava et al., 2005). Initial binding experiments using [3H] PoA with membranes isolated from Sf9 cells expressing DmDopEcR showed specific, high affinity, saturable binding (Bmax = 0.32 ± 0.04 pmol/mg protein; KD = 10.4 ± 0.38 nM). No specific binding was observed in membranes made from wild type control cells showing that the specific binding was due to the expression of the DmDopEcR receptor. Further studies indicated that the rank order of ecdysteroid binding potency for DmDopEcR (PoA > E > 20E >> MaA) was different from that of the nuclear EcR expressed in Sf9 cells (PoA > 20E > MaA > E) (Nakagawa et al., 2000). Further, the bisacylhydrazine insecticides, RH-5849 and RH-0345 (halofenozide) (Fig. 18.2), two synthetic compounds which have previously been shown to be potent agonists of the nuclear EcR’s from Lepidoptera and Drosophila (Dhadialla et al., 1998; Nakagawa et al., 2000) were unable to displace [3H] PoA binding to DmDopEcR, up to a concentration of 100 µM. This further demonstrates that [3H] PoA is binding to DmDopEcR and not to the endogenous Sf9 nuclear EcR, somehow expressed at the cell surface. It also demonstrates that E has a higher binding affinity for DmDopEcR than 20E which is generally regarded as the classical biologically active ecdysteroid in most insect systems with E acting as its non-functional synthetic precursor. However, E has been shown to be equally, or even more potent, than 20E at mediating a number of possible non-genomic effects of insect ecdysteroids (see above and Schlattner et al., 2006). Interestingly, the putative membrane ecdysone receptor from the anterior silk glands of the silkworm, Bombyx mori, also showed a similar pharmacology with PoA showing a slightly higher affinity for the receptor than 20E and the bisacylhydrazine insecticides, RH-2485 (Methoxyfenozide), RH-5992 (Tubefenozide) and RH-5849 showing a much reduced affinity compared to their actions on the nuclear EcR (Elmogy et al., 2004). However, in this preparation the bisacylhydrazine insecticides did show significant binding at concentrations above 10 µM. It is not clear if this difference between the Drosophila DmDopEcR and the Bombyx binding site are due to species differences in homologous receptors or due to the fact that they represent two totally different receptors that are capable of binding ecdysteroids. Dopamine’s ability to elevate cyclic AMP levels via the activation of DmDopEcR raises the question of the relationship between the binding sites for dopamine and ecdysteroids on DmDopEcR. To further investigate the pharmacological profile of this receptor, we used a range of natural and synthetic catecholamine agonists and antagonists to test for their ability to block specific [3H] PoA binding to

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DmDopEcR in isolated membranes from DmDopEcR expressing Sf9 cells (Srivastava et al., 2005). We found that the DA receptor antagonists flupenthixol and spiperone, as well as the adrenergic antagonist prazosin, were able to block specific binding in a dose-dependent manner (Rank order: flupenthixol = prazosin >> spiperone). The same compounds also inhibited the dopamine mediated increases in cyclic AMP levels through this receptor (Rank order: spiperone > flupenthixol > prazosin). This indicates that dopamine and PoA may share an overlapping binding site on DmDopEcR but the different rank order of potency for the compounds suggests that the receptor may exhibit a form of ‘agonist-specific coupling’ (see below). However, dopamine was not able to block the binding of [3H] PoA to the receptor at concentrations up to 100 µM, despite its ability to bind to DmDopEcR to induce changes in cyclic AMP levels. This suggests that the receptor may have a much higher affinity for steroids than for catecholamines. Although dopamine and the ecdysteroids both have ring structures with two hydroxyl groups, dopamine may not be able to effectively displace the ecdysteroids from their binding site if the steroid molecules, because of their larger size, can bind to additional amino acid residues in the receptor not accessible to dopamine. These ideas are supported by the observed importance of the positioning of the hydroxyl groups on the cholesterol-like carbon side chain of the various ecdysteroid molecules at position 17 of the steroid ring, for their ability to displace [3H] PoA from the receptor. The ideas are also compatible with recent probabilistic models of GPCR behaviour (Kenakin, 2004). To determine whether DmDopEcR might be capable of mediating any of the rapid, non-genomic signaling actions of ecdysteroids, we determined if ecdysteroids could in anyway modulate the activity of second messenger pathways through this receptor (Srivastava et al., 2005). Although we could find no evidence for direct ecdysteroid modulation of cyclic AMP levels or [Ca2+]I levels, we were able to show that ecdysteroids were able to inhibit the dopamine-mediated stimulation of cyclic AMP levels in a dose-dependent manner in both Sf9 and CHO cells expressing DmDopEcR. E, 20E and PoA all inhibited the dopamine mediated stimulation of cyclic AMP levels with great efficiency. The order of potency seen for the inhibition of the dopamine response as determined by the IC50s was: PoA > E > 20E (IC50: PoA, 12.1 ± 0.2 pM; E, 0.15 ± 0.04 nM; 20E, 1.5 ± 0.4 nM) which mirrors the order of potency described with [3H] PoA binding. No cross-reactivity with ecdysteroids was demonstrated with the endogenous octopamine receptors in Sf9 cells, which increase cyclic AMP levels (Nasman et al., 2002), or with a number of other Drosophila aminergic GPCRs expressed in either Sf9, CHO cells or in Xenopus oocytes, including the octopamine/tyramine receptor which decreases cyclic AMP levels (Robb et al., 1994), or the DopR99B (DAMB) Dopamine D1-like receptor, which increases cyclic AMP levels (Reale et al., 1997). This suggests that the actions of ecdysteroids on DmDopEcR are likely to be receptor specific and not secondary on second messenger pathways. Furthermore, the inhibitory effects of the ecdysteroids on the dopamine response were also reproducible in CHO cells expressing DmDopEcR, implying that the effects were also not cell specific. A number of the non-genomic effects of vertebrate steroids appear to be mediated via the activation of the MAP kinase pathway (Cato et al., 2002; Lösel and

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Wehling, 2003). In addition, dopamine can activate the PI3 kinase pathway through activation of DmDopEcR as mentioned above. Thus, we examined whether ecdysteroid activation of DmDopEcR could activate either of these pathways (Srivastava et al., 2005). In CHO cells expressing DmDopEcR ecdysone was not able to activate the PI3 kinase pathway as determined by changes in Akt phosphorylation. However, it was able to activate the MAP kinase pathway as determined by increases in ERK1/2 phosphorylation in both a time-dependent and dose-dependent manner; maximal activation occurred 15 min after exposure to E and the threshold concentration for activation occurred between 0.1 and 1.0 µM which is below the maximum levels of ecdysteroid reported from insect haemolymph. In addition, this response was found to be rapid and not dependent on either gene transcription or protein synthesis. Preliminary studies suggest that the ecdysteroid activation of MAP kinase via DmDopEcR may involve the transactivation of a membrane bound receptor tyrosine kinase, such as the Drosophila equivalent of the epidermal-derived growth factor receptor or the platelet-derived growth factor receptor (DP Srivastava and PD Evans, unpublished, 2008). This would parallel the mechanism of the nongenomic actions of estrogen via GPR30 (Filardo et al., 2000; Thomas et al., 2005; Filardo and Thomas, 2005). DmDopEcR is unusual in that it can be activated by both catecholamines and ecdysteroids (Fig. 18.3). As discussed above, this could be due to the two sets of agonists binding competitively to overlapping topographical sites on the outer surface of the receptor. However, it could also be due to the agonists binding noncompetitively to two distinct non-overlapping sites that could interact allosterically.

Rapid Non-Genomic Effects

E

DA CG18314

G1 G2

Cyclic AMP AKT (PI3Kinase )

MAPKinase

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Fig. 18.3 The orphan Drosophila GPCR (CG18314) responds to both ecdysteroids and to the catecholamine dopamine and has been renamed DmDopEcR (Srivastava et al., 2005). DmDopEcR shows ‘agonist-specific coupling’ whereby the catecholamine dopamine can couple the receptor to an increase in cyclic AMP levels together with an activation of the PI3Kinase pathway, as judged by the increased phosphorylation of Akt. Ecdysteroids can produce rapid non-genomic effects through this receptor by coupling it to the activation of the MAPKinase pathway and by inhibiting the actions of dopamine. The receptor has a much higher affinity for ecdysteroids compared to dopamine (See Color Plates)

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It is very difficult to distinguish between these two possibilities experimentally for GPCRs (Christopoulos and Kenakin, 2002). It appears unlikely that the present case represents a classical allosteric interaction since both the ecdysteroids and the catecholamines appear to be able to activate the receptor. However, rare cases of allosteric agonism have been described for GPCRs, such as the chemokine receptor CXCR4 (Sachpatzidis et al., 2003). Further experimentation will be required to definitively distinguish between these two possibilities. The relationship between the results obtained from simple receptor binding studies and those from the efficacy of functional coupling of GPCRs to different second messenger systems is difficult to understand (Kenakin, 2002, 2004). Current ideas such as ‘agonist-specific coupling’ (agonist trafficking) (Evans et al., 1995; Kenakin, 1995) and the concept of an ‘ensemble’ of different agonist-induced receptor conformations, each with their own specific signaling capabilities (Kenakin, 2002, 2004) are challenging the classical ideas of GPCR signaling. Thus, the ability of PoA to functionally displace dopamine from the receptor to inhibit the stimulation of cyclic AMP levels at much lower concentrations than could produce a significant displacement of [3H] PoA binding is likely to be a reflection of the higher affinity of the receptor for steroids than catecholamines and a reflection of the insensitivity of the binding assay. Thus, it could be argued that even at these very low concentrations PoA is still able to produce a conformational change in the receptor that is able to block dopamine actions through the receptor. This is supported by recent models of receptor activation which infer that there are an infinite number of receptor conformations, which further suggests that the binding affinity of a ligand does not necessarily predict its efficacy (Kenakin, 2002, 2004; Kenakin and Onaran, 2002). As such, these models imply that if a ligand binds to the receptor, it will produce a bias in the available conformations of the receptor (Kenakin, 2002, 2004; Kenakin and Onaran, 2002). Future studies using radioactively labeled steroid ligands with much higher binding affinities may help to resolve these issues. The DmDopEcR demonstrates ‘agonist-specific coupling’ (Fig. 18.3). Dopamine was able to activate the PI3 kinase pathway via the receptor as assessed by Akt phosphorylation, in addition to increasing cyclic AMP levels. Ecdysone did not increase Akt phosphorylation but was able to activate the MAP kinase pathway, as assessed by the phosphorylation of ERK-2. Thus, it is likely that the binding of different agonists to the receptor produces different combinations of responses due to the induction of different receptor conformations by the different agonists (Evans et al., 1995; Kenakin, 1995). A number of other Drosophila GPCRs also display ‘agonist-specific coupling’ including the D1-like dopamine receptor DopR99B (Reale et al., 1997), the Oct/Tyr receptor (Robb et al., 1994), the short neuropeptide F receptor (Reale et al., 2004) together with a range of vertebrate adrenergic and neuropeptide activated GPCRs (Spengler et al., 1993; Airriess et al., 1997). In Drosophila the MAP kinases have been implicated in a number of processes, including proliferation of cells in the imaginal disc in Drosophila eyes (Kurada and White, 1999) and differentiation of photoreceptor cells (Sawamoto and Okano, 1996; Karim and Rubin, 1998; Kurada and White, 1999). They are believed to have roles in both cell survival and cell death (Karim and Rubin, 1998; Kurada

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and White, 1999). Several studies have also demonstrated the activation patterns of the MAP kinases and have shown that they are active during the development of Drosophila embryos and larval tissues (Knust, 1996; Gabay et al., 1997; Bier, 1998). In addition, the prothoracicotrophic hormone (PTTH), which regulates the titer of E and 20E, has also been shown to activate ERK in the prothoracic gland (Rybczynski et al., 2001; Rybczynski and Gilbert, 2003). This activation of ERK, also suggested to be via a GPCR, is implicated in the rapid regulation of ecdysteroid synthesis in Manduca sexta (Rybczynski et al., 2001; Rybczynski and Gilbert, 2003). In addition, both the PI3 kinase pathway (Scheid and Woodgate, 2001) and catecholamines (Pendelton et al., 1997) are known to have important roles in the control of Drosophila development. In the future it will be of interest to see if both dopamine and the ecdysteroids can signal or modulate each other’s actions through DmDopEcR in the intact animal. The expression pattern of DmDopEcR suggests that the receptor may play a variety of important roles during development, as well as in the functioning of the adult nervous system (Srivastava et al., 2005). Using in situ hybridization with digoxigenin-labelled antisense riboprobes we initially saw expression in stage 14–15 embryos in the developing ducts of the salivary glands and in the primary constriction of the midgut. In these regions the receptor is likely to be involved with the control of cell proliferation and differentiation but its exact role in these regions is yet to be elucidated. However, this expression was transient and had largely disappeared by late stage 17 embryos when strong expression was observed in clumps of cells in the developing nervous system. Staining was particularly intense in two parallel bands of cells in the lower portion of the ventral ganglia corresponding with the distribution of the neuroblasts that eventually give rise to most of the motorneurons and large interneurons of the adult nervous system. Staining was also observed in the anterior and dorsal portions of the cerebral hemispheres. This intense staining of the nervous system was maintained through the first and second larval instars. However, it had largely disappeared by the third larval instar, except for a narrow band of cells extending from the anterior dorsal to the posterior ventral regions of the brain lobes in the region of the outer optic anlagen. The latter structure is composed of neuroblasts which give rise to the cortices of the lamina and the distal medulla (Meinertzhagen and Hanson, 1993). Expression was also seen in a line of cells in the eye-antennal imaginal disk in the region of formation of the morphogenetic furrow. This suggests that DmDopEcR could well be involved in the control of the progression of the morphogenetic furrow since this process requires E, but not the nuclear EcR (Brennan et al., 2001). Equally, it is interesting to speculate that the many surges of ecdysteroids released during insect development (Riddiford, 1993; Truman, 1996) might serve to turn off dopamine-mediated signaling via DmDopEcR and initiate specific patterns of neurogenesis and differentiation appropriate to particular stages of development. This could perhaps occur via a rapid ecdysteroid mediated inhibition of NO signaling, such as has been described during optic lobe neurogenesis in the optic lobes of Manduca (Champlin and Truman, 2000). A PCR analysis of DmDopEcR expression (Srivastava et al., 2005) reveals strong expression in adult heads with

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a possible sexual dimorphism with increased expression in females compared to males. This suggests a possible modulatory role for ecdysteroids via DmDopEcR during neuronal processing, which might parallel the well known non-genomic effects of estrogen on neuronal signaling, dendritic spine formation and memory in vertebrates (Srivastava et al., 2008; Woolley, 2007).

18.5

Future Directions

It seems very likely that the rapid, non-genomic actions of ecdysteroids, in parallel with the similar actions of vertebrate steroids, will be mediated by a wide range of receptors including nuclear receptors expressed close to the plasma membrane or at other intracellular locations where they can be activated by lipid soluble ecdysteroids, together with true plasma membrane expressed GPCRs, ion channels and other membrane associated proteins. The challenge now is to identify which receptor mechanisms underlie the various non-genomic actions of ecdysteroids that have been described in this chapter, as well as those likely to be described in the future as a greater awareness of non-genomic actions of ecdysteroids becomes more widespread. In terms of future work on increasing our understanding of the functional role of DmDopEcR, it will be important to increase our detailed knowledge of the relationship between the binding sites for dopamine and ecdysteroids on this receptor and to determine the amino acid residues involved in these interactions. Such information will be of much use in the development of potential novel insect control agents but it will also be important in terms of increasing our understanding of the interactions of steroids with GPCRs in general. It seems likely that genetic overexpression and knockout studies, particularly time dependent studies in specific tissues, will lead to an increased understanding of the functional roles of this receptor at different stages during development and in the functioning of the adult nervous system. In vertebrate systems, although only two GPCRs have been cloned to date that are reported to directly interact with steroids (estrogen and progesterone, see above), it seems very likely that some of the rapid non-genomic actions reported for other vertebrate steroids may also be associated with direct interactions with GPCRs. Equally, it would seem very likely that DmDopEcR will not be the only insect GPCR to interact with ecdysteroids. It will be of much interest to see if the homologues of DmDopEcR identified in other species, including, Anopheles gambiae, Apis mellifera, Bombyx mori and Caenorhabditis elgans (see Srivastava et al., 2005), also bind and are activated by ecdysteroids or if DmDopEcR is an evolutionary unique GPCR in this respect. Equally, it will be of interest to determine if any other families of insect GPCRs have also evolved to respond to either ecdysone or to other insect ecdysteroids. Acknowledgments The work reported in this chapter was supported by the BBSRC via The Babraham Institute.

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Chapter 19

Ecdysone Receptors of Pest Insects – Molecular Cloning, Characterisation, and a Ligand Binding Domain-Based Fluorescence Polarization Screen Lloyd D. Graham, Wynona M. Johnson, Donya Tohidi-Esfahani, Anna Pawlak-Skrzecz, Marianne Bliese, George O. Lovrecz, Louis Lu, Linda Howell, Garry N. Hannan, and Ronald J. Hill Abstract EcR- and USP-encoding cDNAs of four pest insects (Lucilia cuprina, Myzus persicae, Bemisia tabaci, Helicoverpa armigera) were cloned from highquality lambda cDNA libraries and sequenced. Cognate EcR-USP cDNA pairs were shown to express functional ecdysone receptors in transfected cells. The amino acid sequences of the EcR ligand binding domains (LBDs) were employed in conjunction with those known for other arthropods to construct a phylogenetic tree. Affinity tagged EcR-USP LBD heterodimers were co-expressed efficiently in insect cells using a baculovirus vector. The recombinant EcR and USP DE/F segments from each species associated spontaneously to form heterodimers that bound ecdysteroids with high affinity. An E/F segment pair (constructed only for H. armigera) also associated spontaneously to form a functional heterodimer, but one with ligand binding affinities several times lower than its DE/F counterpart. A fluorescein-inokosterone conjugate was synthesized and used to develop a novel ligand binding assay based on fluorescence polarization. This assay can be used in place of the classical [3H]-ponasterone A binding assay, and is ideally suited to high-throughput screening. The ligand binding data obtained in vitro using recombinant LBD heterodimers reflect the ability of agonists to induce ecdysone receptor controlled transgene expression in recombinant mammalian cells; in vitro binding data can also reflect the potency of ligands to act as insecticides.

L.D. Graham, D. Tohidi-Esfahani, A. Pawlak-Skrzecz, G.N. Hannan, and R.J. Hill () CSIRO Molecular & Health Technologies, Sydney Laboratory, PO Box 184, North Ryde, NSW 1670, Australia W.M. Johnson, M. Bliese, and L. Howell CSIRO Molecular & Health Technologies, Ian Wark Laboratory, Bag 10, Clayton South, Victoria 3169, Australia G.O. Lovrecz and L. Lu CSIRO Molecular & Health Technologies, Parkville Laboratory, 343 Royal Parade, Parkville, Victoria 3052, Australia e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Keywords Ecdysone receptor • pest • phylogeny • ligand binding domain • recombinant • fluorescence polarization • high-throughput screening • insecticide

19.1

Introduction

Evidence for a hormonal factor controlling insect moulting and metamorphosis came from the elegant pioneering experiments of Kope (1922), Wigglesworth (1934) and Fraenkel (1934). The existence of a specific receptor for the hormone ecdysone was formally postulated by Ashburner et al. (1973), and a few years later the physical reality of the receptor in Drosophila melanogaster was demonstrated by the rigorous characterisation of its ability to bind tritium-labelled ponasterone A by Yund et al. (1978) and Maroy et al. (1978). Somewhat surprisingly, ecdysone receptors have also been shown to be the site of action for the dibenzoylhydrazine class of environmentally-friendly insecticides (Wing, 1988; Dhadialla et al., 1998). Molecular cloning of the EcR gene (Koelle et al., 1991), and the realisation that the ecdysone receptor is actually a heterodimer of the EcR protein with the USP protein product of the usp gene (Thomas et al., 1993; Yao et al., 1993), paved the way for production of sufficient recombinant receptor protein for detailed characterisation. This chapter describes a procedure we have employed for the molecular cloning of the EcR and USP subunits of ecdysone receptors from six insect pests drawn from four taxonomic orders. A phylogenetic analysis which incorporates the EcRs from these species is presented. Full-length proteins have been characterised in vivo by their ability to confer ecdysteroid responsiveness on reporter genes in mammalian cells. We have employed a baculovirus system in insect cells to express recombinant heterodimeric ligand binding domain (LBD) proteins in sufficient quantity to allow their purification and characterisation. We also describe the development of a novel ligand binding assay based on fluorescence polarization, which is suitable for high-throughput screening of chemical libraries.

19.2

Molecular Cloning and Characterisation of Ecdysone Receptors

Of the >30 EcRs that have been derived to date from arthropod species, only the first (that from Drosophila melanogaster; DmEcR) was isolated by protein purification from cells of the relevant insect (Luo et al., 1991) prior to the cloning of the cognate EcR cDNA (Koelle et al., 1991). For other arthropod species, where often little was known about the underlying genetics, cloning of the EcR- and USP-encoding cDNAs, together with their subsequent expression of recombinant subunits, has provided the primary route to studies of the functioning of the receptor.

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While several different approaches are now possible, we began cloning cDNAs encoding EcR and USP proteins from pest insect species that were relatively poorly understood by exploiting sequence information from known ecdysone receptors. Since the DNA binding domain (C-domain) is the most highly conserved region in these nuclear receptors, we designed degenerate primers specific for the DNA binding domain and used these to generate homologous probes by PCR, using either genomic DNA or a cDNA library as template. An advantage of the latter is that the amplification not only provides a probe but also confirms the presence of the relevant cDNA in the library, or at least sequences extending from the 3’- end to the region encoding the DNA binding domain. After cloning, the amplified C-domain probes were used to screen cDNA libraries for putative cDNA clones encoding EcR and USP proteins. High-quality cDNA libraries generally yielded clones that contained full-length open reading frames. Following the isolation of cDNAs anticipated to encode EcR and USP protein subunits from novel insect species, it is useful to undertake functional testing to demonstrate the major functions of a nuclear hormone receptor: evidence of DNA binding, hormone binding, nuclear localisation and transcriptional activation all provide reassurance of a successful outcome. While domain-based functions can be tested independently in vitro under defined conditions, the most comprehensive test involves simultaneous verification of all these properties via transgenic activation of ecdysteroid-inducible reporter genes in insect or mammalian cells. A particular advantage of using the latter for such testing is the fact that, as a concomitant of some 500 million years of evolutionary separation from insects, mammalian cells do not naturally contain ecdysone-responsive systems, and thus present a low level of background (Yang et al., 1995; Yao et al., 1993). This section (Section 19.2) relates the common approach that we have employed to clone and characterise EcR and usp cDNAs from six different insect species – Lucilia cuprina, Myzus persicae, Helicoverpa armigera, Bemisia tabaci, Nezara viridula and Bovicola ovis. Furthermore, we give examples of tests for overall function of ecdysone receptors in cultured mammalian cells.

19.2.1

Isolation of mRNA

For the isolation of mRNA encoding EcR and USP subunits, it is important to obtain insects (either fresh, or stored in liquid nitrogen) at a stage in the life-cycle when the concentration of EcR and usp mRNAs is expected to be elevated. Obviously, it is highly desirable to isolate RNA that has suffered minimal degradation and is predominantly full-length. Whilst other extraction methods can be used, we have found that very high-quality RNA can routinely be obtained from a range of insect tissues using stringent guanidinium isothiocyanate extractions (Okayama et al., 1987). Messenger RNA can be purified from total RNA preparations using a polyA+ mRNA isolation kit such as the one available from Promega.

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cDNA Library Construction

A cDNA library was constructed in Lambda ZAP II (Stratagene) using 5 µg of poly(A)+ RNA for each insect species, as follows. First-strand DNA was synthesised using oligo(dT) primers and, after second-strand synthesis, the cDNA was cloned into Lambda ZAP II. After packaging, a primary library of 1–2 × 106 pfu was obtained and subsequently amplified to about 1012 pfu. Once a high-quality cDNA library has been constructed, it provides a valuable resource that can be screened repeatedly for open reading frames encoding nuclear receptors and other proteins of interest from the insect. Amplification of cDNA in phage lambda incurs a mutations at 7.74 × 10−8 per base pair per DNA duplication (J. W. Drake, personal communication, 2007) which compares favourably with the rate of even the highestfidelity DNA polymerases used in PCR amplifications (for example, ca. 40 × 10−8, www.stratagene.com/amplification).

19.2.3

PCR Cloning of Homologous Probes and Library Screening

We screened phage libraries for EcR- and usp-containing clones using radiolabelled DNA probes that were homologous to the regions encoding the relevant DNA binding domains. For example, genomic DNA from Lucilia cuprina larvae was used as a template in a PCR amplification to prepare a DNA probe specific for an EcR from the sheep blowfly. Two degenerate oligonucleotides, 5’ CGG AAT TCC GCC TCI GGI TA(C/T) CA(C/T) TA(C/T) AA(C/T) GC 3’ and 5’ CGC GGA TCC (G/A)CA CTC CTG ACA CTT TCG (C/T)CT CA 3’, were designed using highly conserved regions in the C-domains of DmEcR and of CtEcR. Sequence data from two other members of the steroid receptor superfamily of D. melanogaster, namely Drosophila hormone receptor 3, DHR3 (Koelle et al., 1992) and Drosophila early ecdysone responsive gene, E75 (Segraves and Hogness, 1990), were taken into account in designing the primers so as to minimise the probability of cloning the L. cuprina homologs of these proteins. The resulting 130 bp PCR product was subcloned into pBluescriptSK+ (Stratagene), and its identity as an amplicon encoding the LcEcR DNA binding domain was confirmed by DNA sequence analysis. Screening of the amplified phage library was conducted by hybridization with 32P-labelled probe (106 cpm/ml) under low-stringency conditions followed by autoradiography. Usually, screening 1–2 million plaques from the amplified library was sufficient to identify EcR- or USP-encoding cDNAs. Positive plaques were purified to homogeneity and converted into pBluescriptSK− plasmids by the in vivo excision method using Exassist (Stratagene) helper phage, after which both strands of the inserts were sequenced. In our experience, the vast majority of clones isolated from high-quality cDNA libraries contained full-length EcR and

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USP open reading frames. For each subunit, complete open reading frames were sequenced from at least three independent library clones.

19.2.4

Functional Testing via Transactivation of a Reporter Gene in Mammalian Cell Culture

The coding region for each putative EcR- or USP-encoding cDNA was subcloned into the mammalian expression vector pSG5 (Stratagene) for use in transfection experiments to assay for ecdysone receptor activity. Mammalian CHO cells, which contain substantial levels of RXR (a mammalian homologue of insect USP capable of providing USP function) were employed to assay EcR activity in the absence of USP. Conversely, CV1 monkey kidney cells, which contain relatively low levels of RXR, were chosen for assessment of the activity of EcR-USP heterodimers. Transient transfections were conducted using DOTAP (Boehringer-Mannheim) at 15 µg/ml. For example, replicate 35 mm dishes of subconfluent CV1 cells were co-transfected with EcR and USP expression plasmids (based on pSG5) plus an inducible reporter plasmid expressing CAT under the control of an ecdysoneresponsive promoter, p(EcRE)5CAT. Also included, at 1 µg/ml, was an internal control plasmid expressing β-galactosidase, pPGKLacZ. For induction experiments, 1 µM of the ecdysone analogue ponasterone A (a gift from Dr Denis Horn) was added to cells 6 h after transfection. For control experiments, cells were treated only with ethanol. The CAT and β-galactosidase levels in extracts of cells were measured 48 h after transfection. Variations between experiments were controlled by normalising the level of CAT to β-galactosidase in the same extract (see Hannan and Hill, 1997, for further details). By way of example, the integrity of our LcEcR- and LcUSP-encoding cDNAs, along with their ability to function together to produce a heterodimeric receptor, were demonstrated in mammalian CV1 tissue culture cells as shown in Fig. 19.1. In attempting to test the functionality of our cloned full-length M. persicae EcR, we found that it was unable to effect reporter gene expression in mammalian cell systems even in the presence of ecdysteroids (not shown); this was probably because its A/B region (Pawlak-Skrzecz et al., in preparation) is more similar to the A isoform of D. melanogaster EcR, a version that has low or undetectable transcription-activating capability in mammalian cells (Mouillet et al., 2001). To enable ligand-inducible reporter gene expression mediated by the M. persicae EcR LBD, we constructed the chimeric receptor DmA/B-MpCDE EcR by substituting the A/B region from the B1 isoform of DmEcR (which is known to potentiate ecdysteroid-dependent transactivation in mammalian cells) for the corresponding segment of M. persicae EcR. For completeness, we also constructed the complementary chimera, MpA/B-DmCDEF EcR, by substituting the A/B region of M. persiace EcR for the corresponding segment of D. melanogaster EcR. These constructs were transfected into mammalian

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Fig. 19.1 Induction of LcEcR-controlled reporter gene expression in transfected mammalian cells. LcEcR, when partnered with LcUSP, can confer ecdysteroid inducibility on a reporter plasmid expressing CAT under the control of an ecdysone-responsive promoter p(EcRE)5CAT in CV1 mammalian cells. CV1 cells were cotransfected with combinations of the LcEcR expression plasmid, pVPLcEcR and the LcUSP expression plasmid, pSGLcUSP, in the presence or absence of 1 µM ponasterone A (Reproduced from Hannan and Hill, 2001. With permission of Elsevier)

CHO cells, which contain sufficient RXR, the mammalian homologue of USP, to potentially form functional heterodimers with EcR. Receptor incorporating the DmA/B domain (either native DmEcR or the chimeric DmA/B-MpCDEF) conferred ecdysteroid inducibility on the CAT reporter gene, as indicated by the data in Fig. 19.2 (Graham et al., 2007a). Like pUC18, MpA/B-DmCDEF EcR shows no induction of CAT expression, confirming that the inability of wildtype M. persicae EcR to transactivate in mammalian cells is due to the sequence of its A/B region. Interestingly, these experiments with chimeric receptors in mammalian cells indicated that the MpEcR LBD led to a stronger response with ponasterone A, whereas the DmEcR LBD mediated a greater response to muristerone A. These responses suggest stronger binding by ponasterone A to the hemipteran LBD and by muristerone A to the dipteran LBD, an issue discussed further in Section 19.6.

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Fig. 19.2 Induction of DmEcR- and MpEcR-controlled reporter gene expression in transfected mammalian cells. CHO cells were transfected transiently with EcR-expressing plasmids or a pUC18 negative control plasmid (horizontal axis categories). At the same time, the cells were co-transfected with a reporter plasmid providing EcR-controlled CAT expression and a third plasmid that reported the transfection efficiency. The expression of CAT was measured in response to various treatments: 10 µM ponasterone A (shown as “10 uM ponA” in the graph), 10 µM muristerone A (shown as “10 uM murA”), and a negative control containing solvent but no ecdysteroid (ethanol only, shown as “EtOH”). Expression data were plotted after normalization for transfection efficiency (vertical bars, mean values in arbitrary units; error bars, ± standard errors of the means) (Reproduced with permission of Elsevier from Graham et al., 2007a, where further details have been given)

19.3

Evolutionary Variation and Relationships

Evolutionary variation between nuclear receptors of different species may be analysed by three phylogenetic approaches, namely parsimony, maximum likelihood and distance. In this chapter, we have chosen primarily to apply the distance method to amino acid sequences. Our focus in this section, and indeed in the remainder of the chapter, is on the EcR LBD or E/F-region. In this section, we further narrow our analyses to consider only the E-region, i.e. the highly structured domain in which the ligand binding pocket resides. Neighbour-Joining is a distance method which measures, by comprehensive pair-wise comparisons, the relative differences between all sequences being analysed (Escriva et al., 1999). The output of this method can be presented in the form of an unrooted phylogenetic tree, where branch lengths are proportional to the number of amino acid differences (Fig. 19.3). Therefore, sequences which are much diverged from each other are separated by longer branches or ‘distances’,

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Fig. 19.3 Phylogenetic tree of EcR E-domains from various arthropods. The tree was constructed using the Neighbour-Joining method on ClustalW(accurate) alignments of 212 amino acid sites of 39 EcR E-domains. Sites with gaps were excluded from the analysis. The JTT (Jones et al., 1992) amino acid model with different rates among sites (gamma) was used. Numbers at nodes and near order names are bootstrap values out of 1,000 replicates, expressed as percentages. Diamonds indicate EcRs discussed in some detail in this chapter. Species names were abbreviated as follows: Diptera: Aa, Aedes aegypti, Aalb, Aedes albopictus, Ag, Anopheles gambiae, Cc, Ceratitis capitata, Ct, Chironomus tentans, Cv, Calliphora vicina, Dm, Drosophila melanogaster, Lc, Lucilia cuprina. Mecoptera: Panorpa germanica. Siphonaptera: Cfe, Ctenocephalides felis. Trichoptera: Hi, Hydropsyche incognita. Lepidoptera: Ba, Bicyclus anynana, Bm, Bombyx mori, Cf, Choristoneura fumiferana, Cs, Chilo suppressalis, Ha, Helicoverpa armigera, Hv, Heliothis virescens, Jc, Junonia coenia, Ms, Manduca sexta, Pi, Plodia interpunctella. Hymenoptera: Ap, Apis mellifera, Pd, Polistes dominulus. Coleoptera: Tc, Tribolium castaneum, Tm, Tenebrio molitor, Ld, Leptinotarsa decemlineata. Phthiraptera: Bo, Bovicola ovis. Orthopteroidea: Bg, Blatella germanica, Lm, Locusta migratoria. Hemiptera: Bt, Bemisia tabaci, Mp, Myzus persicae, Nv, Nezara viridula. Strepsiptera: Xv, Xenos vesparum. Chelicerata: Aam, Amblyomma americanum, Om, Ornithodoros moubata. Crustacea: Dma, Daphnia magna, Cm, Carcinus maenas, Gl, Gecarcinus lateralis, Mj, Marsupenaeus japonicus, Up, Uca pugilator. Branch lengths are proportional to sequence divergence; the scale at the bottom of the figure represents 0.1 differences per site

whereas sequences which are similar lie close to each other in the phylogenetic tree. The robustness of branch nodes or groupings within the tree can be measured by a resampling method called bootstrapping. Including a more conserved region

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(such as the C-domain, in the case of nuclear receptors) in the computation of a tree may also confirm phylogenetic relationships (Escriva et al., 1999; Laudet and Bonneton, 2005). The evolutionary robustness of a tree can be tested further by comparing Neighbour-Joining outputs with those from parsimony and maximum likelihood methods (refer to Escriva et al., 1999, for descriptions of each of these methods). We have performed phylogenetic analyses by Neighbour-Joining, parsimony, and maximum likelihood methods on EcR E-domains, including ones cloned in this laboratory, ones recently sequenced from hemimetabolous insects, and ones found in the Chelicerata and Crustacea. In agreement with previous findings by Bonneton et al. (2003, 2006), we have observed a distinct separation of the more diverged Mecopterida superorder (which includes Diptera and Lepidoptera) from those of other insects and arthropods (Tohidi-Esfahani et al., in preparation). Our Neighbour-Joining analysis found the EcR E-domains of non-mecopteridan insects, such as those of the Hymenoptera, Coleoptera, Orthopteroidea, and Hemiptera, to cluster within a major clade with significant bootstrap confidence (60%, Fig. 19.3). The E-domain of the recently cloned phthirapteran EcR from Bovicola ovis (Pollard et al., in preparation) clustered with those from fellow hemimetabolous insects with 51% confidence. In agreement with the findings of Bonneton and colleagues (2006), the EcR E-domain of the twisted-wing wasp parasite, Xenos vesparum, did not group with those of the Mecopterida, but rather with other insects. In our analysis, X. vesparum was found to have a weak association (35% bootstrap confidence) with that of M. persicae (Fig. 19.3). Apart from the Hemiptera, all taxonomic orders with more than one representative in our EcR E-domain analysis clustered separately with high bootstrap confidence (Fig. 19.3). This finding was confirmed by phylogenetic analysis of segments spanning the C- and part of the D-domains (Tohidi-Esfahani, in preparation). Hemipteran EcRs do not group with each other, or indeed with any other order, with significant bootstrap values. This may be a reflection of the divergence of EcR sequences within this order, or simply the current paucity of data from representative species. An interesting correlation was observed when relative phylogenetic distances between EcR E-domains of insects from different orders were compared to the corresponding sensitivities to the dibenzoylhydrazine insecticide, tebufenozide. This analysis will be presented and discussed in Section 19.6.

19.4

Baculovirus Expression of Ligand Binding Domain Heterodimers

Recombinant nuclear hormone receptors and their ligand binding domains (LBDs) have usually been expressed using E. coli as host, but poor solubility of the LBDs has often required them to be modified (Mossakowska, 1999). The yields published for LBDs purified (with dimerization partners, where appropriate) from

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recombinant bacteria range 0.5–17 mg/l culture (Apriletti et al., 1995; Gampe et al., 2000), with a median value of 8 mg/l culture (Li et al., 1997; Halling et al., 1999; Bourguet et al., 1995; Egea and Moras, 2001), although sometimes only ∼10% of the purified recombinant molecules were functional (Li et al., 1997). The only ecdysone receptor LBD heterodimer included in those examples had been modified: the F-region had been deleted from its EcR component, and a fusion partner had been provided for its USP (Halling et al., 1999). More recently, an ecdysone receptor LBD heterodimer was expressed adequately in E. coli only if a fusion partner was supplied for the EcR LBD and/or some surface residues were mutated (Billas et al., 2003). In contrast, baculovirus-infected insect cells had been shown to deliver a high proportion of functional molecules without any need for modification (Clagett-Dame and Repa, 1997; Juntunen et al., 1999), and yields of 0.2–0.5 mg/l culture were reported for a vitamin D receptor LBD purified from recombinant insect cells (Juntunen et al., 1999). Baculovirus expression might be expected to be particularly appropriate for recombinant LBDs from insect receptors. Using a baculovirus system (pFastBac Dual, Invitrogen) to co-express LBDs from insect ecdysone receptors, we obtained good yields of functional EcR-USP LBD heterodimers without any need for fusion partners or other modifications. Using recombinant baculovirus constructs, we co-expressed the DE/F segments of the EcR and USP subunits from each of four insect species (L. cuprina, M. persicae, B. tabaci, H. armigera) as affinity-tagged proteins (‘recombinant LBDs’) (Graham et al., 2007b). We also prepared a construct co-expressing tagged E/F segments from the H. armigera receptor subunits (Graham et al., 2007b). The sequences of the recombinant EcR segments are given (without affinity tags) in Fig. 19.4. Most of the baculovirus constructs produced approximately equal amounts of recombinant EcR and USP LBDs. Large-scale (5–6 l) fermentations of recombinant insect cells yielded 30–100 g wet cells, which typically contained 0.3–1.6 mg of recombinant protein per gram of cells. All of the recombinant segment pairs (including the H. armigera E/F pair) associated spontaneously with high affinity to form heterodimers that were active in the ligand binding assay, thereby indicating that neither ligand nor D-regions were essential for the formation of tightly-associated and functional LBD heterodimers. The first conclusion contrasts with the finding that recombinant LBDs from D. melanogaster EcR and USP associate only weakly in the absence of ligand (Lezzi et al., 2002; Yao et al., 1993), while the second conflicted with our expectation that some of the hinge region would be required for LBD heterodimerization and/or ligand binding (Perera et al., 1999; Miyamoto et al., 2001; Grebe et al., 2003; Palli et al., 2003). Yields of affinity-purified protein ranged 2.9–16.3 mg/l culture, with a median value of 7.6 mg/l culture (Graham et al., 2007b). Recombinant cell extracts containing the L. cuprina LBD heterodimer typically had a specific [3H]-ponasterone A binding activity approximately 10,000 times higher than that of L. cuprina embryo extracts. The highest expression level that we observed was 16.6 mg active LBD heterodimer per liter of culture (Graham et al., 2007b).

DmEcR_part LcEcR_part MpEcR_part BtEcR_part HaEcR_part

1 11 21 31 41 51 MRPECVVPENQCAMKRREKKAQKEKDKMTTSPSSQHGGNGSLASGGGQDFVKKEILDLMT MRPECVVPENQCAMKRREKKAQKEKDKIQTSVCAT-----E---------IKKEILDLMT --PECVVPEVQCAVKRKEKKAQREKDKPNSTTDIS-----------------PEII---K --PECVVPEFQCAVKRKEKKAQKDKDKPNSTTSCS-----------------PDGI---K -RPECVVPENQCAMKRKEKKAQREKDKLPVSTTTV----------------DDHMPPIMQ

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61 71 81 91 101 111 CEPP-----------QHATIP-LLPDEILAKCQARNIPSLTYNQLAVIYKLIWYQDGYEQ CEPP-----------SHPTCP-LLPEDILAKCQARNIPPLSYNQLAVIYKLIWYQDGYEQ IEPT-----------EMKIECGEPMIMGTPMPTVPYVKPLSSEQKELIHRLVYFQDQYEA QEID-----------PQRLDTDSQLLS------VNGVKPITPEQEELIHRLVYFQNEYEH CDPPPPEAARILECLQHEVVPRFLNEKLMEQNRLKNVPPLTANQKSLIARLVWYQEGYEQ * 121 131 141 151 161 171 PSEEDLRRIMS-----QPDENESQTDVSFRHITEITILTVQLIVEFAKGLPAFTKIPQED PSEEDLKRIMS-----SPDENESQHDASFRHITEITILTVQLIVEFAKGLPAFTKIPQED PSEKDMKRLTINNQNMDEYDEEKQSDTTYRIITEMTILTVQLIVEFAKRLPGFDKLVRED PSPEDIKRI-VN----AAPEEENVAEERFRHITEITILTVQLIVEFSKRLPGFDKLIRED PSEEDLKRVTQT---WQSDEDDEDSDMPFRQITEMTILTVQLIVEFAKGLPGFAKISQSD

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181 191 201 211 221 231 QITLLKACSSEVMMLRMARRYDHSSDSIFFANNRSYTRDSYKMAGMADNIEDLLHFCRQM QITLLKACSSEVMMLRMARRYDHNSDSIFFANNRSYTRDSYKMAGMADNIEDLLHFCRQM QITLLKACSSEAMMFRVARKYDITTDSIVFANNQPFSADSYNKAGLGDAIENQLSFSRFM QIALLKACSSEVMMFRMARRYDAETDSILFATNQPYTRESYTVAGMGDTVEDLLRFCRHM QITLLKACSSEVMMLRVARRYDAATDSVLFANNQAYTRDNYRKAGMAYVIEDLLHFCRCM

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241 251 261 271 281 291 FSMKVDNVEYALLTAIVIFSDRPGLEKAQLVEAIQSYYIDTLRIYILNRHCGDSMSLVFY YSMKVDNVEYALLTAIVIFSDRPGLEEAELVEAIQSYYIDTLRIYILNRHCGDPMSLVFF YNMKVDNAEYALLTAIVIFSSRPNLLDGWKVEKIQEIYLESLKAYVDNRD--RDTATVRY CAMKVDNAEYALLTAIVIFSERPSLSEGWKVEKIQEIYIEALKAYVENRR--KPYATTIF YSMMMDNVHYALLTAIVIFSDRPGLEQPLLVEEIQRYYLNTLRVYILNQNSASPRCAVIF

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301 311 321 331 341 351 AKLLSILTELRTLGNQNAEMCFSLKLKNRKLPKFLEEIWDVHAIPPSVQSHLQITQEENE AKLLSILTELRTLGNQNAEMCFSLKLKNRKLPKFLEEIWDVHAIPPSVQSHIQATQAEKARLLSVLTELRTLGNENSELCMTLKLKNRVVPPFLAEIWDVMP----------------AKLLSVLTELRTLGNMNSETCFSLKLKNRKVPSFLEEIWDVVS----------------GKILGILTEIRTLGMQNFNMCISLKLKNRKLPPFLEEIWDVADVSTTATPVVADSPAL--

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361 371 381 391 401 411 RLERAERMRASVGGAITAGIDCDSASTSAAAAAAQHQPQPQPQPQPSSLTQNDSQHQTQP ---AAQEAQATTSAISAAATSSSSINTSMATSSSSS-----LSPSAASTPNGGAVDYVGT

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Fig. 19.4 Multiple sequence alignment of EcR DE/F segments. DmEcR_part, partial sequence for D. melanogaster EcR; LcEcR_part, partial sequence for L. cuprina EcR; MpEcR_part, partial sequence for M. persicae EcR; BtEcR_part, partial sequence for B. tabaci EcR; HaEcR_part, partial sequence for H. armigera EcR. For convenience, recombinant DE/F and E/F segments are both referred to as recombinant ligand binding domains (LBDs). The first LBD residue in the H. armigera E/F construct is marked by an asterisk. Further details of this alignment can be found in the Supplementary Material that accompanies Graham et al. (2007b)

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Purification of Recombinant Ligand Binding Domain Heterodimers

IMAC capture via the His6-tag afforded the easiest and most effective purification of our recombinant LBD heterodimers. Further purification could be achieved by subjecting IMAC-purified recombinant proteins to FPLC ion exchange chromatography (Mono-Q) or gel filtration (Superdex-200). Both FPLC steps were efficient (> 65% yields) and inexpensive. It was possible to use both in sequence, but the combination of IMAC followed by a single FPLC step (typically Superdex-200) was usually sufficient (Graham et al., 2007b). To obtain apo-LBD heterodimer preparations for ligand binding experiments, the recombinant heterodimer was purified from cell lysates by IMAC in the absence of any ligand. However, ligand-free LBD heterodimers were fragile and easily became inactivated during preparative manipulations, with specific activity values suggesting that only 7–37% of the heterodimers in such preparations were functional (Graham et al., 2007b). Other researchers have estimated 50–60% functionality for their purified preparations of ecdysone receptor apo-LBD heterodimers (Halling et al., 1999). It has been remarked previously that wild-type ecdysone apo-receptors are inherently labile, but that they could be stabilized by adding a carrier protein (Lehmann and Koolman, 1988) or by loading them with hormone (Sage et al., 1986). Likewise, we found that our purified recombinant apoLBD heterodimers were stabilized by including BSA (0.5 mg/ml) as carrier protein and/or by saturating them with ponasterone A. X-ray crystal structures reveal that bound ponasterone A provides many interactions that stabilize the EcR LBD (Billas et al., 2003; Carmichael et al., 2005), and the buried ligand constitutes a necessary part of the E-domain’s hydrophobiccore (Wagner et al., 1995). Curiously, the ligand binding stoichiometry of purified apo-LBD heterodimer preparations could be increased several-fold by including the non-denaturing detergent CHAPS, which may increase the proportion of functional LBDs by rescuing ligand-incompetent conformations (Graham et al., 2007b). CHAPS seems to affect other recombinant nuclear hormone receptors in a similar way (Clagett-Dame and Repa, 1997). To prepare ligand-LBD heterodimer complexes for crystallization trials, recombinant cells were sonicated in the presence of excess ponasterone A and the recombinant complex was purified using IMAC followed by an FPLC chromatography step (usually Superdex-200) in the presence of saturating concentrations of ponasterone A. Proteins purified as ponasterone A-LBD heterodimer complexes contained a higher proportion of active molecules than those purified without ligand, with best estimates suggesting ∼70% of such populations were functional (Graham et al., 2007b). A substantial proportion of bacterially-expressed nuclear hormone receptor LBD populations also remained unliganded when they were purified in the presence of ligand (Egea and Moras, 2001). The molecular masses determined for each polypeptide by denaturing gel electrophoresis (SDS-PAGE) and mass spectrometry agreed well with the expected values. Figure 19.5 shows representative SDS-PAGE separations (Graham et al.,

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Fig. 19.5 SDS-PAGE of purified recombinant LBD heterodimers. Samples contained reducing agent and bands were visualized using Coomassie stain. M, marker proteins with molecular masses in kilodalton. Preparations were purified by IMAC followed by gel filtration (lanes 1–3) or by IMAC alone (lanes 4–5). L. cuprina heterodimer (lane 1); M. persicae heterodimer (lane 2); B. tabaci heterodimer (lane 3); H. armigera DE/F heterodimer (lane 4); H. armigera E/F heterodimer (lane 5). Band assignments, based on apparent molecular mass and immunostaining (Graham et al., 2007b), were as follows: intact recombinant EcR (E) and USP (U) subunits, nicked versions of recombinant EcR (A) and USP (B), the 75 kDa (C) and 57 kDa (F) contaminants mentioned in the text, and aggregated protein that failed to enter the resolving gel (G) (Reproduced from Graham et al., 2007b. With permission of Elsevier)

2007b). For all species, a small amount of Hsc70 (75 kDa; Fig. 19.5, band C) co-purified with the recombinant LBD heterodimers through the IMAC, Mono-Q and Superdex-200 steps. Moreover, IMAC-purified preparations of H. armigera E/F recombinant protein contained relatively large amounts of an α-tubulin contaminant (57 kDa; Fig. 19.5, band F) that was barely present in preparations of its DE/F counterpart; in addition, they often contained some high molecular mass material (Fig. 19.5, band G). A small proportion of the recombinant subunits of the L. cuprina and M. persicae heterodimers usually underwent proteolytic nicking near the C-terminus (Fig. 19.5, bands A and B). L. cuprina EcR was particularly susceptible, possibly because it contains a long and potentially nonessential C-terminal extension (F-domain) (Halling et al., 1999; Hu et al., 2003) not present in the other EcR LBDs. C-terminal nicking did not appear to compromise heterodimer assembly (Graham et al., 2007b). Freshly-purified B. tabaci LBDs appeared as a triplet in SDS-PAGE (Fig. 19.5, lane 3), and preparations of this protein underwent substantial proteolysis – including N-terminal proteolysis of the USP subunit – during prolonged crystallization trials (Carmichael et al., 2005; Graham et al., 2007b). In the absence of reducing agents at ≥4°C, disulfide bonds slowly formed both within and between certain LBDs, although this did not seem to impact greatly on ligand binding. Intramolecular disulfide bonds were observed for recombinant EcR subunits (L. cuprina and M. persicae) and USP subunits (L. cuprina), whereas intermolecular bonds were observed only for recombinant EcR subunits (all four

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insect species) (Graham et al., 2007b). The process seemed to be non-specific, and perhaps involved Cys residues located in relatively unstructured regions (D-, and possibly F-segments). Although thiol-specific alkylation (i.e., treatment with iodoacetic acid or iodoacetamide) could suppress this phenomenon, such treatments greatly increased mass microheterogeneity (Graham et al., 2007b). Our best crystallization results were obtained when unalkylated proteins were maintained in reducing environments (Carmichael et al., 2005). Native PAGE of purified L. cuprina [3H]-ponasterone A-LBD heterodimer complex gave a series of bands in which each band contained equimolar amounts of recombinant EcR and USP subunits (Fig. 19.6) (Graham et al., 2007b). The main band, which had the highest mobility, was presumed to be the ligand-EcR-USP complex. The slower-moving bands seem to represent dimers and trimers of this complex, whether present as non-covalent assemblies (faint bands in reduced samples) or as disulfide-mediated assemblies (stronger bands in air oxidized samples). [3H]-ponasterone A was associated specifically both with the main LBD heterodimer band and with the slower-moving ‘heterodimer oligomer’ bands (Fig. 19.6). In non-denaturing IEF gels, the DE/F heterodimers

Fig. 19.6 Non-denaturing PAGE of purified recombinant LBD heterodimers. The gel shows IMAC-purified L. cuprina ponasterone A-LBD heterodimer complexes prepared without reducing agent. Immunoblots using antibodies against FLAG alone (lane 1) or His6 tag alone (lane 2) produced an identical pattern, namely a major band corresponding to the ponasterone A-LBD heterodimer complex (A) and decreasing amounts of less mobile species (B, C) that are likely to be dimers and trimers, respectively, of the complex. The remaining lanes, which were visualized by Coomassie staining, include a reference lane of bovine serum albumin (lane 3) and a sample of [3H]-ponasterone A-LBD heterodimer complex (lane 4). The amount of [3H]-ponasterone A in horizontal slices of lane 4 was determined by scintillation counting (bar graph). For each of the bands A–C there is a peak of radioactivity (marked) whose relative magnitude reflects the relative band intensity seen in gels and blots. Clearly the ligand-EcR-USP complexes remain assembled in this native PAGE system (Reproduced from Graham et al., 2007b. With permission of Elsevier)

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focused in the range pH 6.2–7.1. For each species, the recombinant protein was divided between 4–7 closely related isoforms. Since the heterogeneity could not be related to post-translational modifications such as glycosylation or phosphorylation, its cause remains unclear. However, the presence of only three bands in the H. armigera E/F protein suggests that some isoforms may reflect discrete conformational options available to the D-domain (Graham et al., 2007b).

19.6

Ligand Binding Measured Using a Radioactive Ecdysteroid

In equilibrium binding studies, the Kd values for [3H]-ponasterone A binding by LBD heterodimers that included the hinge regions (i.e., DE/F heterodimers) were 1.0, 0.7, 1.2, and 2.5 nM for L. cuprina, M. persicae, B. tabaci and H. armigera, respectively (Graham et al., 2007a). Published Kd values for non-recombinant fulllength ecdysone receptors are in the same range (Dhadialla et al., 1998), suggesting that our recombinant DE/F heterodimers are fully functional. Our Kd values for the L. cuprina and H. armigera DE/F heterodimers are in good agreement with those reported for the binding of [3H]-ponasterone A to the ecdysone receptor from D. melanogaster (~3 nM) and to a recombinant form of the Spodoptera litura receptor (2.8 nM), respectively (Graham et al., 2007a). Indeed, we found that L. cuprina embryo extracts bound [3H]-ponasterone A with a Kd of 0.9 nM, almost identical to the value obtained with preparations of our recombinant L. cuprina LBD heterodimer (1.0 nM). In kinetic studies, [3H]-ponasterone A association and dissociation timecourses for the recombinant L. cuprina LBD heterodimer indicated kon ~ 4.1 × 106 M−1 min−1 and koff ~ 0.98–2.3 × 10−2 min−1. The ratio of these rate constants gives Kd = 2.4–5.6 nM, which is only slightly higher than the Kd value obtained from equilibrium binding studies (1.0 nM) (Graham et al., 2007a). Equilibrium binding experiments were performed to assess the ability of various ligands to compete with [3H]-ponasterone A for binding to the recombinant LBD heterodimers (Graham et al., 2007a). Competitive inhibition plots are shown in Section 13.2, along with corresponding mean Ki values. The binding affinity of the dibenzoylhydrazine ligand, tebufenozide, for recombinant heterodimers varied greatly between different taxonomic orders (Fig. 19.7), with Ki values ranging from 3.7 nM for the DE/F heterodimer from the lepidopteran H. armigera to > 448 µM for that from the hemipteran B. tabaci (Carmichael et al., 2005; Graham et al., 2007a). Our Ki values for DE/F heterodimers from the dipteran L. cuprina and lepidopteran H. armigera (170 and 3.7 nM, respectively) match closely those published for tebufenozide binding to extracts of cells from the dipteran D. melanogaster (192 nM) and lepidopteran Plodia interpunctella (3 nM) (Dhadialla et al., 1998). Our in vitro binding data clearly reflect the lepidopteran-selective toxicity of this insecticide, a feature well known from bioassays and field trials (Dhadialla et al., 1998; Smagghe and Degheele, 1994; Smagghe et al., 1996, 2002; Argentine et al., 2002). The not insignificant affinity of tebufenozide for the recombinant

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Fig. 19.7 Inhibition by tebufenozide of [3H]-ponasterone A binding to recombinant ecdysone receptor LBD heterodimers. Heterodimers were from the dipteran L. cuprina (filled squares), the hemipterans M. persicae (open triangles) and B. tabaci (filled circles), and the lepidopteran Helicoverpa armigera (open circles). The vertical axis shows [3H]-ponasterone A binding expressed as a percentage of that obtained without competing ligand, with s.e.m. as error bars (n = 2–4). The horizontal axis shows the concentration of the tebufenozide (nM) used with each DE/F heterodimer. For H. armigera, this axis also represents 0.3x the concentration of competing ligand used with the E/F heterodimer; the resulting horizontal displacement compensates for the poorer binding obtained with the E/F heterodimer and renders the E/F and DE/F curves coincident. Experiments used 1.3 nM [3H]-ponasterone A, with details as described by Graham et al., 2007a

L. cuprina LBD heterodimer (Ki ~170 nM) is consistent with its moderate toxicity (LC50 ~5 ppm) towards the larvae of this species (Graham et al., 2007a). The lack of interaction in vitro between tebufenozide and our two hemipteran LBD heterodimers (Ki > 132 µM) is consistent with the low binding affinity (Ki > 10 µM) reported for this compound with the ecdysone receptor of Bemisia argentifolii (Dhadialla et al., 2005), as well as its low toxicity towards heteropteran nymphs (Smagghe and Degheele, 1994) and those of B. tabaci (Elzen, 2004). Correlations between the binding affinity of tebufenozide for ecdysone receptors and the biological efficacy of this agent are discussed more broadly at the end of this section. The Ki values for ecdysteroid ligands varied less widely than those for tebufenozide, ranging from 0.1 to 240 nM for a set comprised of 20-hydroxyecdysone, ponasterone A, muristerone A, and inokosterone (Graham et al., 2007a). Of these, the Ki

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for muristerone A varied most (0.1–30 nM) and the Ki for 20-hydroxyecdysone varied least (66–240 nM). The scope of the latter resembles the 40–247 nM range reported elsewhere for binding of this ligand to extracts from dipteran, coleopteran and lepidopteran insect cells (Dhadialla et al., 1998). Although the IC50 values for non-radioactive ponasterone A (acting as a competitor) and Kd values for [3H]-ponasterone A (acting as a ligand) are obtained by methods that are wholly independent, both parameters rank the binding affinities of the recombinant heterodimers in the same order: M. persicae DE/F < L. cuprina DE/F < B. tabaci DE < H. armigera DE/F < H. armigera E/F. The Ki values for non-radioactive ponasterone A, whose calculation makes use of the relevant Kd value, also rank in this order. Our various datasets for ligand dissociation constants are therefore mutually consistent (Graham et al., 2007a). It is interesting that, for all of the ligands tested, the Kd, IC50 and Ki values for the H. armigera E/F heterodimer were 3.7-, 3.3- and 4.4-fold higher, respectively, than the corresponding values for its DE/F counterpart. These results suggest that D-domains do make some contribution to ligand binding by the H. armigera LBD heterodimer. Precedents indicate that the inclusion of inter-domain hinge regions can sometimes improve the stability and/or activity of recombinant domains (Graham et al., 2007a). For the dipteran L. cuprina LBD heterodimer, we found the order of ecdysteroid binding affinities in vitro to be muristerone A > ponasterone A >> 20-hydroxyecdysone (Graham et al., 2007a), which is similar to the ranking obtained in vitro with the dipteran D. melanogaster ecdysone receptor (Sage and O’Connor, 1985; Sage et al., 1986; Cherbas et al., 1988; Dhadialla et al., 1998; Dela Cruz et al., 2000). The ranking also reflects the relative potency of these ligands as inducers of transgene expression in mammalian cells when transcriptional control was provided by a recombinant dipteran EcR, whether taken from L. cuprina (Hannan and Hill, 1997; G.N. Hannan, unpublished data, 2000) or D. melanogaster (Christopherson et al., 1992; Yang et al., 1995; Albanese et al., 2000; Oberdörster et al., 2001). In contrast, for the hemipteran LBD heterodimers from M. persicae and B. tabaci, the order of ecdysteroid affinities in vitro was ponasterone A > muristerone A > 20-hydroxyecdysone (Graham et al., 2007a). Above, we noted a similar switch in ligand preference when examining data from transfected mammalian cells (Section 19.2). In mammalian cells harboring a reporter gene under the transcriptional control of an EcR, ponasterone A was a more potent inducer than muristerone A when the LBD came from the hemipteran M. persicae, whereas muristerone A was the more potent inducer with an LBD from the dipteran D. melanogaster (Fig. 19.2). Thus it would appear that ligand binding affinities determined in vitro can serve as a guide to inducer potency in mammalian cells. Although the binding affinity of a ligand for an ecdysone receptor in vitro is often reflected in its ability to activate the same receptor in transgenic mammalian cells, the same is not necessarily true for insect cells or tissues. For ecdysteroids, issues of ligand transport and metabolism (which underlie episodic changes in the concentrations of these molecules during insect life cycles) can disrupt the correlation between affinity and potency at or above the level of intact cells. Thus, for dipterans (e.g., D. melanogaster and L. cuprina), the in vitro binding affinities

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rank muristerone A ≥ ponasterone A >> 20-hydroxyecdysone (Graham et al., 2007a; Cherbas et al., 1988; Dhadialla et al., 1998), whereas their potency with intact insect cells is very different: ponasterone A > 20-hydroxyecdysone > muristerone A (Dinan et al., 1999; Baker et al., 2000; Oberdörster et al., 2001). A de-coupling of affinity from potency also occurs for lepidopterans (e.g., S. frugiperda and C. suppressalis), although in this case the disconnect appears only at the level of intact larvae (Fujita and Nakagawa, 2007). Non-ecdysteroid ligands may also be affected by pharmacokinetic phenomena. For example, dibenzoylhydrazines

pEC50 or pLD50 or 10*Phylog dist

10

8

6 pEC50 pLD50 10*Phylog dist

4

2

0

0

2

4 6 pKi or plC50

8

10

Fig. 19.8 Relationship between the binding affinity of tebufenozide for ecdysone receptors, its potency in bioassays, and phylogenetic distance of the EcR LBD from a lepidopteran reference sequence. The binding affinity of tebufenozide for ecdysone receptors (or recombinant forms thereof) from a range of insects is based on Ki or IC50 values determined in vitro (horizontal axis). The biological potency of tebufenozide in tissues or organisms of the corresponding insect relies upon EC50 values from imaginal disc evagination or molting assays (open circles), or uses LD50 values from larvicidal assays (filled squares). pKi, pIC50, pEC50 and pLD50 denote −log10[tebufenozide] for the relevant parameter, for which the tebufenozide concentration is expressed in moles per liter. The elongated gray boxes framing some symbols indicate instances where the plot uses a maximum value for a particular parameter as only an upper limit was available. Phylogenetic distance (gray filled triangles) reflects branch lengths in a Neighbor-Joining EcR E-domain tree rooted on C. fumiferana. The correlation coefficients are R = 0.67 (EC50), 0.90 (LD50), and 0.91 (phylogenetic distance). Data are for dipterans L. cuprina (Graham et al., 2007a), D. melanogaster (Dhadialla et al., 1998; Farkas and Slama, 1999), A. egypti (Beckage et al., 2004), C. tentans (Smagghe et al., 2002), C. vicina (Slama, 1995); lepidopterans S. exigua (Smagghe et al., 1999), C. suppressalis (Minakuchi et al., 2003), P. interpunctella (Dhadialla et al., 1998), C. fumiferana (Dhadialla et al., 1998), H. virescens (Allenza and Eldridge, 2007; Argentine et al., 2002), G. mellonella (Smagghe et al., 1996), S. littoralis (Smagghe et al., 2000); S. frugiperda (Nakagawa et al., 2000; Argentine et al., 2002); the coleopterans L. decemlineata (Smagghe et al., 1994, 1996; Ogura et al., 2005), A. grandis (Dhadialla et al., 1998); the orthopteran L. migratoria (Smagghe and DeGheele, 1994; Hayward et al., 2003), and the hemipteran B. tabaci (Graham et al., 2007a; Elzen, 2004). Additional data used in the calculations were cited by Dhadialla et al. (1998, 2005) or Graham et al., 2007a)

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are actively transported out of dipteran (D. melanogaster) but not lepidopteran (C. fumiferana and Malacosoma disstria) cell lines (Sundaram et al., 1998; Retnakaran et al., 2001). Transport and metabolism considerations probably explain why dibenzoylhydrazine binding affinity did not correlate with larvicidal activity for the coleopteran Leptinotarsa decemlineata (Ogura et al., 2005), and similar phenomena probably contribute to the development of dibenzoylhydrazine resistance in lepidopteran species (Retnakaran et al., 2001; Spindler-Barth and Spindler, 1998; Grebe et al., 2000; Smagghe et al., 2001). However, the action of some synthetic ligands on insect cells and organisms appears to be more straightforward than that of ecdysteroids. For example, C. suppressalis displays no de-coupling of dibenzoylhydrazine binding affinity from biological potency at any level (Minakuchi et al., 2003; Fujita and Nakagawa, 2007). Moreover, a survey of published data for tebufenozide confirms that its biological potency in a particular insect correlates positively with its in vitro binding affinity for the ecdysone receptor from that species (Fig. 19.8). As might be expected, the in vitro binding affinity of tebufenozide also shows a strong negative correlation with phylogenetic distance of the EcR LBDs from a lepidopteran reference sequence (Section 19.3 and Fig. 19.8).

19.7

Ligand Binding Measured by Fluorescence Polarization, and Development of an Automated Screen

Fluorescence-based assays offer benefits in safety, environmental impact, cost, speed and sensitivity over those that rely upon radioactivity (Sportsman and Leytes, 2000). For this reason, we synthesized and tested a wide variety of fluorescently-tagged ecdysteroids. An initial conjugate in which 7-diethylaminocoumarin was attached at the C3 position of inokosterone (MB4587) proved inactive: even at 80 µM, this compound failed to compete with 1.3 nM [3H]-ponasterone A for binding to recombinant LBD heterodimers from L. cuprina, M. persicae and B. tabaci. In contrast, conjugation of the same fluorophore to the C26 position of this ecdysteroid yielded a compound (MB4592, Fig. 19.9) that, under the same conditions, caused complete inhibition of [3H]-ponasterone A binding. Since the reactivity of the C26 primary alcohol made it most suitable for derivatisation, different fluorophore precursors were reacted with inokosterone to yield the conjugates shown in Fig. 19.9. Despite the size and variety of the fluorescent moieties, all of these C26 conjugates competed effectively with [3H]-ponasterone A for binding to recombinant LBD heterodimers (Graham et al., 2007a). With the recombinant LBD heterodimer from M. persicae, the Ki values for inokosterone MB4628, MB4592, MB4603 and MB4622 were all similar: 65, 40, 20, 40, and 104 nM, respectively. Since there would not be space to accommodate the bulky fluorophores within the ligand binding pocket (Chapter 13), we postulate that the fluorescent moieties project into solvent; this could easily perturb the placement of helix 12 in the EcR-ligand complex, leading the conjugates to act as antagonists rather than agonists of ecdysone receptor function.

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Fig. 19.9 Functional fluorescent inokosterone conjugates. The fluorophores (R) are attached at the C26 position of the ecdysteroid. Two isomers are possible for the fluorescein conjugate, MB4628, and these probably exist in equilibrium (Graham et al., 2007a). The fluorophores in MB4592, MB4603, and MB4622 are 7-diethylaminocoumarin, 7-methoxycoumarin, and dansyl, respectively. The graph shows that all of the compounds inhibited, to a comparable extent, the binding of [3H]-ponasterone A to the recombinant M. persicae LBD heterodimer. Titration curves are for inokosterone (filled circles), MB4628 (crosses), MB4592 (open squares), MB4603 (filled triangles), and MB4622 (open circles). Experiments used 2.2 nM [3H]-ponasterone A, with details as described by Graham et al., 2007a

Although all of the C26 conjugates were effective ligands, the wide availability of optical filters for detection of the fluorescein moiety made the fluorescein- inokosterone conjugate (MB4628, Fig. 19.9) particularly suitable for use in fluorescence-based assays. MB4628 was therefore used to develop a novel ligand binding assay based on fluorescence polarization (FP) (Graham et al., 2007a). A fluorimeter with polarizing filters can provide an overall polarization value (P) for samples at equilibrium which is low for free ligand (Pmin; rapid tumbling) and high for bound ligand (Pmax; slow tumbling). Advantages of FP assays include the fact that there is no need for bound ligand to be separated physically from free ligand, and the ability to conduct such assays in low volumes (Sportsman and Leytes, 2000; Checovich et al., 1995; Wedin, 1999).We therefore developed a novel assay in which the binding of MB4628

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to recombinant LBD heterodimers was monitored by FP. The polarization value for free MB4628 in standard FP assay buffer was 100 mP. Titrations of a fixed concentration of MB4628 (optimally, 36 nM) with an increasing concentration of recombinant LBD heterodimer gave a sigmoid curve where Pmax was 210–335 mP, depending on the protein preparation (Graham et al., 2007a). Since an assay window of 50 mP is considered adequate for high-throughput FP readers (Prystay et al., 2001), a dynamic range of 235 mP is excellent. The inclusion of 2 mM CHAPS could increase the dynamic range of assays where the original range was < 200 mP (Graham et al., 2007a), as seen for the B. tabaci LBD heterodimer preparation in Table 19.1. Competition assay conditions were designed so that the polarization value observed in the absence of a competing ligand was equal to Pmax for that LBD heterodimer preparation. Increasing the concentration of a non-fluorescent competitor progressively displaced greater amounts of MB4628, thereby decreasing the observed polarization value until it reached 100 mP (Fig. 19.10) (Graham et al., 2007a). The IC50 value for each titration was taken to be the concentration of competing ligand at the mid-point between Pmax and 100 mP. It is difficult to extract classical Ki values from FP data, so we report the IC50 values themselves (Table 19.1). The absolute values depend very much upon experimental conditions, but the relationship between IC50 datasets from different FP competition experiments can be established by the routine inclusion of a few reference compounds (Graham et al., 2007a). For each species of LBD heterodimer, the IC50 values from FP studies of competition with MB4628 (Table 19.1) ranked ligands in the same order of potency as the Ki values determined using competition with [3H]-ponasterone A (Section 19.6). Thus, for the recombinant L. cuprina LBD heterodimer, both platforms ranked ligand affinity as muristerone A > ponasterone A > 20-hydroxyecdysone > tebufenozide, whereas for the hemipteran LBD heterodimers, both platforms ranked ligand affinity as ponasterone A > muristerone A > 20-hydroxyecdysone. A log-log plot of IC50 (FP)

Table 19.1 Ligand IC50 values determined by FP (Reproduced from Graham et al., 2007a. With permission of Elsevier) Species LBD

L. cuprina DE/F

M. persicae DE/F

B. tabaci DE

[Protein] (µg/ml) CHAPS (mM) Pmax (mP)

57–82 0 340–350

2.1 0 325

7.8 0 225

4.7 2 325

Ecdysone 62,000 n.d.a n.d. n.d. 20-hydroxyecdysone 1,600 3,000 17,000 1,600 Ponasterone A 880 2 70 10 Muristerone A 640 300 500 50 Tebufenozide 21,000 n.d. n.d. n.d. The lower half of the table shows mean IC50 values, expressed in nanomoles per litre, under the conditions defined in the upper half of the table. [Protein] refers to the total concentration of IMAC-purified protein. a Not determined.

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350

Pmax 300

105 4

10

250

lC50 (nM)

Mp Bt Bt+CHAPS

1000

lC50

200 100

150

10 1

100 50 10−8

100 mP

K i (nM)

0.1 0.1

1

10−6

10

100

1000

0.0001

0.01

1

100

104

106

Competitor conc. (nM)

Fig. 19.10 Ligand competition measured by fluorescence polarization (FP) or radioactivity. Main figure: Competition with MB4628 monitored by FP. Different non-fluorescent ecdysteroids were assessed by FP for their ability to compete with MB4628 for binding to purified recombinant LBD heterodimer from B. tabaci. The ecdysteroids were 20-hydroxyecdysone (filled squares), ponasterone A (open circles), and muristerone A (filled circles). An IC50 value was estimated from each curve using the midpoint between the observed maximum polarization value (Pmax) and the theoretical minimum polarization value (100 mP). Inset: Ligand IC50 values (from FP assay) and Ki values (from [3H]-ponasterone A assay) for hemipteran LBD heterodimers. IC50 values obtained by competing non-radioactive, non-fluorescent ecdysteroids with MB4628 (measured by FP; vertical axis) are plotted against Ki values obtained by competing the same ligands with [3H]-ponasterone A (measured by radioactivity; horizontal axis). Data are for recombinant LBD heterodimers from M. persicae (filled squares) and B. tabaci (triangles). Some assays of the latter contained 2 mM CHAPS (filled symbols) (Reproduced with permission of Elsevier from Graham et al., 2007a, where further details have been given)

and Ki ([3H]-ponasterone A) values for the hemipteran LBD heterodimers gave a straight line (Fig. 19.10, inset). Although our FP competition assays required higher amounts of recombinant LBD heterodimer than the corresponding [3H]-ponasterone A assays, the higher quality of the FP signal meant that fewer replicate experiments were required (Graham et al., 2007a). Moreover, the FP assay volume can be reduced to 20 µl without compromising the quality of the data, which allows high-throughput screening (HTS) to be performed in 384-well plates. To date, however, we have mainly worked in 96-well plate format; liquid handling is performed by a Tecan Genesis Workstation 200 robot, and FP values are read using a PHERAStar platereader (BMG Labtechnologies, Germany).

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Overview and Conclusions

We expressed and purified recombinant DE/F heterodimers from the ecdysone receptors of four pests insects (L. cuprina, M. persicae, B. tabaci, and H. armigera) drawn from three taxonomic orders (Diptera, Hemiptera, and Lepidoptera). We found that ligand was not required for LBD heterodimer formation, although its presence did stabilize the proteins during purification. For ligand-free preparations, an in vitro binding assay based on [3H]-ponasterone A showed that the purified DE/F heterodimers bound a variety of ligands with dissociation constants similar to those reported for full-length ecdysone receptors. A H. armigera recombinant lacking both D-regions formed tightly-associated E/F heterodimers in the absence of ligand, but displayed ligand binding affinities approximately four times lower than its DE/F counterpart. In consequence, we routinely used recombinant DE/F heterodimers to compare the binding of potential ligands. To complement the [3H]-ponasterone A-binding assay, we developed an alternative assay in which the binding of a fluorescein-inokosterone conjugate is monitored by FP. The FP assay ranked the affinity of competitor ecdysteroids in the same order as the [3H]-ponasterone A binding assay. Although both assays can be used to guide the discovery of new ecdysone receptor ligands, the FP platform is particularly suited to high-throughput screening. The Ki values for a dibenzoylhydrazine ligand varied greatly (> 105-fold) between the recombinant LBD heterodimers. The Ki values for the ecdysteroids in the present study varied less widely (≤ 300-fold for any one ligand), with those for 20-hydroxyecdysone varying least (≤ 3.6-fold); presumably this reflects the need for this natural hormone to bind effectively to most, if not all, ecdysone receptors. Ponasterone A had a higher affinity than muristerone A for the two recombinant hemipteran LBD heterodimers examined in this study, whereas the reverse was true for the recombinant dipteran protein; the same preference was observed when these ligands were tested as inducers of ecdysone receptor-controlled gene expression in transfected mammalian cells. Our in vitro binding data indicated that tebufenozide bound most tightly to the H. armigera DE/F heterodimer, in keeping with expectations for this lepidopteran-selective insecticide. Although metabolic complexities often result in a mismatch between the in vitro binding affinity of ecdysteroids and their potency in insect cells or whole insects, the same is not necessarily true for non-steroidal agonists. For example, there is a broad correlation between the receptor binding affinity of tebufenozide and its biological effects on insects drawn from a wide range of taxonomic orders. In conclusion, our results indicate that the binding data obtained in vitro using recombinant LBD heterodimers can reflect the ability of agonists to induce ecdysone receptor-controlled transgenes in mammalian cells, and can also predict their efficacy as insecticides. Use of the FP assay in HTS should therefore facilitate the discovery of new and useful ecdysone agonists. In a complementary approach (Chapter 13), purified preparations of the B. tabaci ponasterone A-LBD heterodimer complex gave crystals that afforded the first atomic structure of an LBD heterodimer from a hemipteran ecdysone receptor. Structural comparisons of

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the ligand binding pocket in this EcR LBD with counterparts from other insects can be used to inform the rational design of new ligands for ecdysone receptors. HTS and structure-based design strategies can be highly synergistic, and their joint pursuit should expedite the discovery of specific EcR effectors based on novel chemistries. Furthermore, joint application of these receptor-based tools provides the means for integrating our growing knowledge of LBD sequence variation into the insecticide discovery process. In this way, the resource provided by natural variation in receptor structure offers a future pathway to finding the balance between, on the one hand, broadening the insecticide control spectrum commensurate with commercial realities, and, on the other, narrowing the spectrum in response to important safety and environmental concerns. The same tools also promise to underpin further improvements in the EcR-controlled expression systems currently being developed for use as ‘gene switches’ in gene therapy and transgenic crops.

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Chapter 20

SAR and QSAR Studies For In Vivo and In Vitro Activities of Ecdysone Agonists Yoshiaki Nakagawa, Robert E. Hormann, and Guy Smagghe

Abstract Insect molting is regulated by the steroid 20-hydoxyecdysone interacting with the ecdysone receptor (EcR) together with either the retinoid X receptor (RXR) or its homolog, ultraspiracle (USP). Similarly, the non-steroidal diacylhydrazines (DAHs) also bind to EcR, but regulate molting in a dysfunctional manner; they are therefore insecticidal. The four DAHs tebufenozide, methoxyfenozide, chromafenozide and halofenozide have been commercialized to control Lepidoptera and Coleoptera. DAH congeners with various substituents at both benzene rings were synthesized and their ecdysonergic activity in whole body, tissue, cell and protein was quantified. Insecticidal potency (whole body) was measured against three insect species: rice stem borer Chilo suppressalis, beet armyworm Spodoptera exigua, and Colorado potato beetle Leptinotarsa decemlineata. Substituent effects on the activity were analyzed using quantitative structure-activity relationship (QSAR) methods such as the classical (Hansch-Fujita) QSAR and comparative molecular field analysis (CoMFA). These QSAR methods were also applied to analyse in vitro ecdysonergic potency (tissue and cell level) and EcR binding (protein level). Molecular hydrophobicity was extracted as an important physicochemical property to activity at all biosystem levels. CoMFA results for activation of gene expression in the silkworm Bombyx mori are consistent with the milieu of the ligand binding pocket homology-modeled from the crystal structure of DAH-bound EcR of the tobacco budworm Heliothis virescens. Keywords QSAR • insecticides • larvicides • N,N ′-dibenzoyl-N-t-butylhydrazines • ecdysone analogues • ecdysone agonists • ecdysone receptor • receptor binding • CoMFA • diacylhydrazine • insect molting hormone • larvicidal activity • receptor binding activity Y. Nakagawa Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Kyoto 606-8502, Japan e-mail: [email protected] R.E. Hormann Intrexon Corporation, Norristown, PA 19403, USA e-mail: [email protected] S. Smagghe Laboratory of Agrozoology, Faculty of Bioscience Engineering, Ghent University, B-9000 Ghent, Belgium e-mail: [email protected] G. Smagghe (ed.), Ecdysone: Structures and Functions © Springer Science + Business Media B.V. 2009

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Introduction

Arthropods such as insects and crustaceans develop through repeated molting and metamorphosis, a process regulated by a molting hormone, 20-hydroxyecdysone (20E). In insects, prothoracicotropic glands secrete the 20E progenitors ecdysone or 3-dehydroecdysone (Kiriishi et al., 1990). These precursors are then oxidized to the active hormone 20E in the peripheral tissues (Gilbert and Warren, 2005). In addition to insects, the steroid hormone 20E is also found in non-arthropods (Romer, 1979; Mendis et al., 1983, 1984; Schallig et al., 1991) and even in plant species (Imai et al., 1967; Dinan et al., 2001a). In fact, a host of structurally-related steroids – phytoecdysteroids – are identified in plants (Dinan et al., 1999b, 2001a). Among them, ponasterone A isolated from Podocarpus nakaii, is the most potent ecdysone agonist (Nakanishi et al., 1966; Kobayashi et al., 1967). Phytoecdysones may deter phytophagous insects through disruption of molting. The role of 20E as an elicitor of gene expression was proposed by Ashburner et al. (1974). The identification of the ecdysone receptor (EcR) together with its heterodimerizing partner, ultraspiracle protein (USP), as the critical transcription factors was disclosed by Yao et al. (1993). In 2003 and subsequently, ligand-bound EcRs yielded to crystallization and x-ray diffraction, thereby enabling elucidation of both ligand-EcR interactions as well as the heterodimer interactions (Billas et al., 2003; Carmichael et al., 2005; Browning et al., 2007; Iwema et al., 2007). After the discovery and characterization of ecdysone (Huber and Hoppe, 1965; Karlson et al., 1965) academic and industrial investigators alike realized that steroidal molting hormone agonists might have insecticidal properties. Such substances were sought vigorously. However, among ecdysteroids, this endeavor was unsuccessful due to expense of synthesis and poor bioavailability related to ineffective cuticle penetration and rapid metabolism. In the mid-1980s, the diacylhydrazine (DAH) chemotype was discovered, paving the way for the development of novel potent insecticides with excellent specificity, ecotoxicology, and production economics (Wing, 1988; Wing et al., 1988; Hsu, 1991). Tebufenozide (Hsu et al., 1997) and methoxyfenozide (Carlson et al., 2001), representatives of the DAH subset, are the first and second commercialized insecticides of this class, both developed by Rohm and Haas in the U.S. (Dhadialla et al., 1998). Later, chromafenozide (ANS-118) containing the chroman ring system was developed in Japan (Tanaka et al., 2001; Sawada et al., 2003; Yanagi et al., 2006). These three compounds are selectively toxic to Lepidoptera. Halofenozide was also registered as the third Rohm and Haas insecticide in US to control not only Lepidoptera species but also Coleoptera such as Scarabeidae (Dhadialla et al., 1998). Structures of these four insecticides are shown in Fig. 20.1. Since DAHs are generally not very effective against non-Lepidoptera insects, new chemical scaffolds possessing different insect toxicity spectra have been sought. To date, several compound classes such as the amidoketones (Tice et al., 2003a, 2003b), oxadiazolines (Hormann et al., 2004), tetrahydroquinolines (Smith et al., 2003), and benzamides (Mikitani, 1996b) have been reported as new non-steroidal ecdysone agonists (Fig. 20.2). Amidoketones (Tice et al., 2003) and oxadiazolines (Hormann et al., 2004) are structurally related to DAHs and also have a similar spectrum of activity. The tetrahydroquinolines, on the other hand, are structurally quite different from DAHs and are toxic to the yellow fever mosquito Aedes aegypti but not to the

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Fig. 20.1 Commercialized DAHs

fruit fly Drosophila melanogaster, even though both species are grouped as Diptera (Palli et al., 2005). The activity spectrum of the benzamides has been explored in only a limited mode in Drosophila systems (Nakagawa, 2005). Structure-activity relationships of ecdysone agonists have been studied in the last two decades by several research groups. The toxicity of representative DAHs such as RH-5849, tebufenozide, methoxyfenozide, and halofenozide to pest insects, as well as in vitro activity such as EcR binding and tissue pharmacology was compared (Dhadialla et al., 1998; Smagghe et al., 2000; Carlson et al., 2001). In a further study, designed DAHs with assorted aromatic ring substitution were synthesized and their activity for ecdysone agonism was measured on several biosystem levels such as whole organisms (larvae) (Oikawa et al., 1994a, 1994b; Nakagawa et al., 1999, 2001, 2002b; Smagghe et al., 1999), tissues (Oikawa et al., 1993; Nakagawa et al., 1998), cells (Nakagawa et al., 2002b, 2002a) and receptor proteins (Minakuchi et al., 2003a, 2007; Ogura et al., 2005). Biological activities have been analyzed using quantitative-structure activity relationship (QSAR) methodologies such as multiple

CH3 C C N C H O O

O H C N N

Et H3CO

CH3 F

Et CH3

HN F N O

F CH3

Fig. 20.2 Non-natural, non-DAH ecdysone agonists

HO

O N

O H C N

CH3

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linear regression (Hansch-Fujita analysis) (Hansch and Fujita, 1964; Fujita, 1990) and three-dimensional QSAR (3D-QSAR) methods, including comparative molecular field analysis (CoMFA) (Cramer et al., 1988) and comparative molecular similarity indeces analysis (CoMSIA) (Klebe et al., 1994). A four-dimensional QSAR (Hopfinger et al., 1997) has also been developed for DAHs (Hormann et al., 2003) and steroids (Dinan et al., 1999). In general, the QSAR approach is useful not only to inspire the design of new compounds, but also to understand mode of action at the molecular level. Here we review the classical Hansch-Fujita analyses and multi-dimensional CoMFAs of ecdysone agonism by DAHs. The first section of this review compares QSARs derived for in vivo and in vitro activity against the rice stem borer Chilo suppressalis. The second part addresses inter-species QSARs using in vivo and in vitro activities to codify information about selective toxicity. The third section treats QSARs that describe ecdysone receptor (EcR) interactions across species. In the last section, ligand-receptor interactions in a modeled EcR are evaluated by a CoMFA model constructed for the EcR gene-switch inducing activity of a large set of DAHs. The chosen target variable determines the best use of the QSAR model: in vivo toxicity for design of insecticides and in vitro EcR binding for the understanding of ligand-receptor interactions. Comparison of in vivo with in vitro models for a common organism and ligand set assists understanding of the relationship between ligand structural features and the biosystem level, i.e., receptor vs. tissue vs. organism, of greatest on which these features have greatest impact. Finally, EcR ligand-receptor (or enzyme) interactions are analyzed by matching of QSAR fields with a modeled receptor.

20.2

20.2.1

Comparison of In Vivo and In Vitro QSARs for the Same Species of Lepidoptera, Chilo suppressalis In Vivo Larvicidal Activity

The insecticidal activity for a set of substituted DAH congeners (Fig. 20.3) against third instar larvae of the C. suppressalis was measured in terms of the dose required to give 50% mortality (LD50, mmol/insect) and expressed as pLD50, the reciprocal logarithm of LD50. (Table 20.1) (Oikawa et al., 1994a, 1994b). Correlation equations relating in vivo

Fig. 20.3 Substituted dibenzoylhydrazines

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Table 20.1 Larvicidal activity of DAH congeners against C. suppressalis, S. exigua, and L. decemlineata pLD50 (mmol/insect) No

X

Y

C. suppressalis

S. exigua

L. decemlineata

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43

H 2-F 2-Cl 2-Br 2-I 2-CF3 2-NO2 2-CH3 2-C2H5 2-C6H5 2-OCH3 2-OBusec 2-OCH2C6H5 2-SCH3 3-F 3-Cl 3-Br 3-I 3-CF3 3-NO2 3-CN 3-CH3 3-OCH3 4-F 4-Cl 4-Br 4-I 4-CF3 4-NO2 4-CN 4-CH3 4-But 4-C6H5 4-OCH3 4-O(CH2)3C6H5 2,3-Cl2 2-CH3–3-Cl 2,3-(CH3)2 2,4-Cl2 2,4-(CH3)2 2,5-Cl2 2-OCH3-5-Prn 2,5-(CH3)2

H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H H

6.27 6.24 6.83 6.88 6.99 6.90 6.80 5.82 6.03