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General and Comparative Endocrinology 169 (2010) 65–74 Contents lists available at ScienceDirect General and Comparative Endocrinology journal homepage: www.elsevier.com/locate/ygcen Identification and expression of mRNAs encoding bursicon in the plesiomorphic central nervous system of Homarus gammarus Jasmine H. Sharp a, David C. Wilcockson b, Simon G. Webster a,* a b School of Biological Sciences, Bangor University, Bangor, Gwynedd LL57 2UW, UK Institute of Biological, Environmental and Rural Sciences, Aberystwyth, Ceredigion SY23 3DA, UK a r t i c l e i n f o Article history: Received 13 May 2010 Revised 7 July 2010 Accepted 21 July 2010 Available online 5 August 2010 Keywords: Arthropods Bursicon Crustaceans Developmental expression Ecdysis In-situ hybridisation Quantitative PCR Neurohormones Homarus gammarus a b s t r a c t Ecdysis in arthropods is a complex process, regulated by many neurohormones, which must be released in a precisely coordinated manner. In insects, the ultimate hormone involved in this process is the cuticle tanning hormone, bursicon. Recently, this hormone has been identified in crustaceans. To further define the distribution of bursicon in crustacean nervous systems, and to compare hormone structures within the sub-phylum, cDNAs encoding both bursicon subunits were cloned and sequenced from the nervous system of the European lobster, Homarus gammarus, and expression patterns including those for CCAP determined using in-situ hybridisation, quantitative RT-PCR and immunohistochemistry. Full-length cDNAs encoded bursicon subunits of 121 amino acids (Average Mr: 13365.48) for Burs a, 115 amino acids (Average Mr: 12928.54) for Burs b. Amino acid sequences were most closely related to those of crabs, and for Burs b the sequence was identical to that of the American lobster, Homarus americanus. Complete colocalisation with CCAP in the VNC was seen. Copy numbers burs a, burs b and CCAP mRNAs were between 0.5 and 1.5 105 for both bursicon subunits, 0.5–6 105 per cdn neurone for CCAP. The terminal abdominal ganglia (AG 6–8) contained about 52 cdn-type neurons, making it the largest bursicon producing region in the CNS. Double labelling IHC using recombinant Carcinus Burs a and CCAP antisera demonstrated complete co-localisation in the VNC. On the basis of the results obtained, it is proposed that CCAP and bursicon release occur simultaneously during ecdysis in crustaceans. Ó 2010 Elsevier Inc. All rights reserved. 1. Introduction The hormonal control of somatic changes and behaviour during ecdysis in arthropods is a precisely coordinated process. For insects, this process has been the subject of intense research (reviews, Truman, 2005; Kim et al., 2006; Ewer, 2007; Žitňan et al., 2007), and a complex temporal series of neurohormones including – and this list is probably incomplete – pre-ecdysis triggering hormone (PETH), ecdysis triggering hormone (ETH), kinins, diuretic hormones (DHs), myoinhibitory peptides (MIPs), eclosion hormone (EH), crustacean cardioactive peptide (CCAP)1 and burs* Corresponding author. Fax: +44 01248 371644. E-mail address: [email protected] (S.G. Webster). 1 Abbreviations used: AG, abdominal ganglion; AVLC, anterior ventral–lateral cell cluster; Burs a-IR, Bursicon a-immunoreactivity; CCAP, crustacean cardioactive peptide; CCAP-IR, CCAP immunoreactivity; cdn, CCAP-IR descending neurone; CG, cerebral ganglion; cnc, CCAP-IR neurosecretory cell; CNS, central nervous system; DIG11-UTP, digoxygenin-11-uridine-50 -triphosphate; EDC, 1-ethyl-3-(3-dimethyl aminopropyl)-carbodiimide; EST, expressed sequence tag; GSP, gene specific primer; IHC, immunohistochemistry; IPTG, isopropyl b-D-1-thiogalactopyranoside; ISH, in-situ hybridisation; ORF, open reading frame; PBS, phosphate-buffered saline; PLC, posterior-lateral cell cluster; PMLC, posterior medial–lateral cell cluster; qRT-PCR, quantitative RT-PCR; RACE, rapid amplification of cDNA ends; RT-PCR, reverse transcription PCR; SDS–PAGE, sodium dodecyl sulphate polyacrylamide gel electrophoresis; SOG, sub-oesophageal ganglion; TG, thoracic ganglion; UTR, untranslated region. 0016-6480/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.ygcen.2010.07.006 icon act in a tightly controlled cascade before, during and after ecdysis as recently exemplified by an RNAi study on the flour beetle, Tribolium castaneum (Arakane et al., 2008). Since many of the recent advances in our understanding of these processes in insects have been made using the genetic resources of model insects, it is unsurprising that our knowledge of analogous events and hormones involved in ecdysis of genetically intractable crustaceans is by comparison, extremely limited. Emergence from the exoskeleton at the start of ecdysis is initiated by a massive release of crustacean hyperglycaemic hormone (CHH) from paraneurones in the fore and hind-gut in Carcinus maenas, which leads to dipsogenesis and rapid swelling (Chung et al., 1999; Webster et al., 2000). This is immediately followed by a large, rapid release of CCAP from the pericardial organs (Phlippen et al., 2000), which probably initiates stereotyped motor patterns involved in active ecdysis (escape from the old cuticle) in the same way as has been suggested for insects (Gammie and Truman, 1997). The existence of peptides that initiate pre-ecdysial and other ecdysial behaviours, such as PETH, ETH and EH have not been established in crustaceans, excepting the presence of a transcript encoding an ETH-like molecule in the water flea, Daphnia pulex (Gard et al., 2009). Despite these differences, the insect cuticle tanning hormone bursicon, which has long been known to play a pivotal role in tanning and 66 J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 melanisation of the insect cuticle (Cottrell 1962a,b; Fraenkel and Hsiao, 1962, 1965), seems to have a widespread, possibly universal occurrence in arthropods. Following the identification and full characterisation of the bursicon as a heterodimeric cystine knot protein encoded by CG13419 (burs or burs a) and CG15284 (pburs or burs b) (Luo et al., 2005; Mendive et al., 2005), database searches revealed the presence of a burs a-like transcript in the Daphnia arenata EST database. Information from this sequence, together with those available from insects allowed us to identify cDNAs (using a strategy involving degenerate PCR 50 and 30 RACE) encoding both bursicon subunits in D. arenata and the shore crab C. maenas, thus firmly establishing bursicon as a hormone common to both subphyla (Wilcockson and Webster, 2008). For insects, many, but not all bursicon immunoreactive neurons in the CNS co-localise with CCAP (review, Honegger et al., 2008), occasionally some are immunoreactive to only one bursicon monomer (Luo et al., 2005; Dai et al., 2008), and it has been proposed from molecular dissection via enhancer trapping in Drosophila that the differential distribution of bursicon and CCAP expressing neurons forms a neural network which controls the sequential activation of bursicon release during the ecdysis programme (Luan et al., 2006a,b). Additionally, for holometabolous insects, adult emergence is associated with a peak in expression and release of bursicon, together with subsequent apoptosis of most of the abdominal neurons that express CCAP (and which co-express bursicon) following eclosion (Ewer et al., 1998; Draizen et al., 1999). For crustaceans expression patterns of mRNA and the neurons expressing bursicon and CCAP are only known for the shore crab C. maenas (Wilcockson and Webster, 2008). Whilst that study suggested that transcripts for both bursicon subunits and CCAP were co-expressed throughout the CNS, the apomorphic nature of the fused abdominal ganglia of the crab (which contain the majority of bursicon expressing neurons) did not easily allow detailed analysis. We therefore reasoned that the more plesiomorphic central nervous system of the lobster, where abdominal ganglia are clearly defined would be ideal to resolve issues concerning co-localisation of bursicon and CCAP. Furthermore, since previous neuronal mapping of CCAP in the ventral nervous system of crayfish (Audehm et al., 1993; Trube et al., 1994) recorded anatomies of each CCAP expressing neuron in every ganglion, and our previous cloning and sequencing of mRNA encoding lobster CCAP (Chung et al., 2006) it would now be possible to determine steady state mRNA copy number per neuron, to accurately determine ratios of bursicon transcript number in relation to CCAP by qRT-PCR in each ganglion of the ventral nerve cord. Here we report nucleotide and amino acid sequences encoding both bursicon subunits in the European lobster Homarus gammarus, and expression patterns of both bursicon and CCAP in individual ganglia of the CNS using in-situ hybridisation, immunohistochemistry (IHC) and quantitative RT-PCR. 2. Materials and methods 2.1. Animal collection, tissue preparation Adult H. gammarus (ca. 600 g) were purchased from local fishermen (Anglesey, UK), individually held in a recirculating seawater aquarium under ambient conditions of temperature and photoperiod and fed chopped fish and squid ad libitum. Additionally, juvenile lobsters (ca. 60 mm total length) were grown from post-larvae (National lobster Hatchery, Padstow, Cornwall, UK) for immunohistochemical studies of whole mounted VNC. After deep anaesthetisation (>60 min) on ice, nervous systems (eye stalks, cerebral ganglion, thoracic ganglia 1–5, abdominal ganglia 1–5, terminal ganglia) were dissected in chilled saline, snap-frozen in liquid N2 and stored at 80 °C. For in-situ hybridisation nervous systems were fixed immediately in 4% paraformaldehyde (PFA) in phosphatebuffered saline (PBS). Following overnight fixation, tissues were dehydrated through a graded methanol/PBS series and stored for 2–3 days at room temperature in methanol before use. Tissues dissected for whole mount immunohistochemistry were fixed in 4% paraformaldehyde in PBS containing 1% 1-ethyl-3-(3-dimethyl aminopropyl)-carbodiimide (EDC) overnight at 4 °C prior to extensive washing in PBS. 2.2. RNA extraction and cDNA synthesis Total RNA was extracted using TRIzol (Invitrogen, Carlsbad, CA, USA), followed by treatment with DNase1 (37 °C, 1 h, TURBO DNAfree, Ambion, TX, USA) and quantification (ND-1000, NanoDrop Technologies, Wilmington, DE, USA). For rapid amplification of cDNA ends (RACE), mRNA was isolated from extracted total RNA using poly-dT Dynabeads (Dynal, Oslo, Norway) according to the manufacturer’s instructions and stored in 10 mmol l1 Tris at 80 °C. Approximately 50 ng mRNA was reverse transcribed in a 20 ll reaction. For 30 RACE, mRNA samples were reverse transcribed (50 °C, 50 min) using SuperScript III RT (Invitrogen, Carlsbad, CA, USA) and primed with the Gene Racer 30 oligo(dT) adapter primer (Invitrogen) according to the manufacturer’s instructions. For 50 RACE, mRNA was dephosphorylated, decapped, ligated to a 50 RACE RNA oligo (Invitrogen) and reverse transcribed using SuperScript III with random hexamers according to the manufacturer’s instructions. Samples were then treated with 2 U RNase H (37 °C, 20 min). For degenerate PCR, mRNA was reverse transcribed (50 °C, 50 min) using SuperScript III RT and primed with random hexamers. For qRT-PCR total RNA was reverse transcribed using a Taqman High Capacity cDNA synthesis kit (Applied Biosystems, Foster City, CA, USA). Briefly, 5 ng of total RNA was reverse transcribed using random hexamer primers in 20 ll reaction volumes according to the manufacturer’s instructions. qRT-PCR cRNA standards were reverse transcribed simultaneously with the RNA samples. 2.3. Primers A complete list of primers and their identifying abbreviations is provided in Table 1. 2.4. Degenerate PCR of cDNA encoding burs a Degenerate primers previously used to identify cDNA encoding burs a in C. maenas (Wilcockson and Webster, 2008), were used to identify cDNA fragments encoding burs a of H. gammarus. PCRs were performed using the following conditions: 12.5 ll AmpliTaq Gold Master Mix (Applied Biosystems), 9 ll water, 1.25 ll (100 lmol l1) forward and reverse primers (4F GCVPKPIP, 11R MCRPCTSIE; 1 ll cDNA template. Amplification conditions were: 1 cycle 94 °C 9 min; 5 cycles of 94 °C 30 s, 63 °C 30 s; 5 cycles of 94 °C 30 s, 60 °C 30 s; 25 cycles of 94 °C 30 s, 57 °C 30 s, 72 °C 45 s and final extension at 72 °C for 10 min. A second PCR was performed using fully nested primers (8F ERSCMCCQE, 9R CMCRPCTSI) using the following conditions: 1 cycle 94 °C 4 min, 35 cycles of 94 °C 30 s, 58 °C 30 s, 72 °C 45 s, final extension at 72 °C for 7 min. One microlitre of first round PCR was used as template. PCR products were electrophoresed on 2% agarose gels and bands of the expected size excised and extracted using a gel purification kit (Perfectprep Gel Cleanup, Eppendorf AG, Hamburg, Germany). 2.5. Rapid amplification of cDNA ends (RACE) Using sequence information obtained from cloning and sequencing degenerate PCR products, and using available sequence J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 67 Table 1 Primers used for bursicon sequence identification, production of cRNA probes and quantitative PCR. Primer name Sequence Burs a Carcinus degenerate 4F GCVPKPIP 11R MCRPCTSIE 8F ERSCMCCQE 9R CMCRPCTSI CGGCTGCGTGCCCAARSCNATHCC TCGATGGAGGTCCAGGGNCKRCACATRC GACCGCTCCTGCATGTGYYGYCARGA TGGAGGTGCAGGGGCKRCACATRCANT Burs a Homarus 30 RACE HomA RACE F1 HomA RACE F1N GCCAAGAGTCGGGGGAACGGGAAGC GAGTCGGGGGAACGGGAAGCTTCTG Burs a Homarus 50 RACE HomA RACE R1 HomA RACE R1N TCAATGGGTGCCCGTGTCAAGATC GGTGCCCGTGTCAAGATCTTCCTC Burs a Homarus ISH Bursa Bursa Bursa Bursa TAATACGACTCACTATAGGGAGAACTTCGCCCTCCCACCCATTA ACTTCGCCCTCCCACCCATTA TAATACGACTCACTATAGGGAGATAGCGATCTCTTGAGCCAGGACAG TAGCGATCTCTTGAGCCAGGACAG Burs a Homarus qRT-PCR Hom_alphaF Hom_alphaR alphapro GACGCGGAGGAAGATCTTGA TGTGCAGGGACGACACATG ACGGGCACCATTGA Burs b Homarus 30 RACE HomB RACE F1 HomB RACE F1N GAATGTGAGACTCTTCCCTCAACC GAGACTCTTCCCTCAACCATACAC Burs b Homarus 50 RACE HomB RACE R1 HomB RACE R1N CGCCACACTTGAAGCACTGAC CAGGGAGTGAGTAAAGGGGTGAG Burs b Homarus ISH BursB BursB BursB BursB TAATACGACTCACTATAGGGAGAAGGCGATATGACTTGGAATGT AGGCGATATGACTTGGAATGT TAATACGACTCACTATAGGGAGATTTGAGTAAGGGAGGGATGTAT TTTGAGTAAGGGAGGGATGTAT Burs b Homarus qRT-PCR Hom_betaF Hom_betaR betapro GAGGACTTGGCCGTCAACAA TGACTGAAGGTTGGACTTTGGA FAM-TGTGAGGGAGCCTGTG CCAP Homarus qRT-PCR Hom_CCAPF Hom_CCAPR ccappro AAGCCAAACTGTCGGAGCAA CGCATAGCTGCGCATCAT FAM-TCCAGAGCAAGATGG RPL18 Homarus qRT-PCR RPL18 F RPL18 T7F RPL18 R RPL18 T7R Hom_RPLF Hom_RPLR RPLpro CGAAAAGTGATCAGGCGGGAGCCG TAATACGACTCACTATAGGGAGACGAAAAGTGATCAGGCGGGAGCCG CCTTCTGCGACCCTGTACCAGCAGAG TAATACGACTCACTATAGGGAGACCTTCTGCGACCCTGTACCAGCAGAG ACCGCCCACCTCTGTCTCT TGACGGCCTTGTTTCTTTGC FAM-TCCCGTCTCGTACGCC CCAP Homarus ISH CCAP CCAP CCAP CCAP TATCGGTGACTTGCTGGAGGGTAA TAATACGACTCACTATAGGGAGA TATCGGTGACTTGCTGGAGGGTAA TAATACGACTCACTATAGGGAGA GTTTGGGGAATGGGGGAGTGG GTTTGGGGAATGGGGGAGTGG T7 F F T7 R R T7 F F T7 R R F T7 F T7 R R information from an EST of Homarus americanus encoding burs b (Accession No. CN854188) gene specific primers (GSP) were designed for 30 and 50 RACE. For 30 RACE, PCR was performed as follows: 12.5 ll AmpliTaq Gold Master Mix (Applied Biosystems), 9 ll water, 1.25 ll (10 lmol l1) HomA RACE F1 or HomB RACE F1 primer for burs a and burs b, respectively, 1.25 ll 30 GeneRacer primer (Invitrogen), 1 ll 30 RACE cDNA template. PCR conditions were: 1 cycle 94 °C 9 min; 5 cycles of 94 °C 30 s, 63 °C 30 s; 5 cycles of 94 °C 30 s, 60 °C 30 s; 25 cycles of 94 °C 30 s, 57 °C 30 s, 72 °C 45 s and final extension at 72 °C for 10 min. A second nested PCR using primer HomA RACE F1N or HomB RACE F1N primer for burs a and burs b, respectively, was performed in the following reaction mixture: 22.5 ll Megamix Blue (Helena Biosciences, Sunderland, UK), 1.25 ll nested GSP (10 lmol l1), 1.25 ll 30 GeneRacer nested primer, 1 ll first round PCR product. PCR conditions were: 1 cycle of 94 °C 4 min; 35 cycles of 94 °C 30 s, 58 °C 30 s, 72 °C 45 s and final extension at 72 °C for 10 min. PCR products were electrophoresed on 2% agarose gels and bands of the expected size excised and extracted as described above. For 50 RACE, PCR was performed as follows: 12.5 ll AmpliTaq Gold Master Mix (Applied Biosystems), 9 ll water, 1.25 ll (10 lmol ll) HomA RACE R1 or HomB RACE R1 (burs a and burs b, respectively), 1.25 ll (10 lmol l1) 50 GeneRacer primer (Invitro- gen), 1 ll 50 RACE cDNA template. PCR conditions were identical to those for 30 RACE. Primer HomA RACE R1N or HomB RACE R1N was paired with the 50 nested GeneRacer primer for nested PCR under identical conditions to those used for the nested 30 RACE. 2.6. Cloning and sequencing of PCR products Purified PCR products were ligated into a pCR4-TOPO vector (Invitrogen) and transformed (TOP-10F’ (Invitrogen) according to the manufacturer’s instructions. Plasmid DNA was purified (Fastplasmid Mini, Eppendorf) and sequenced (MWG Biotech) from positive clones containing inserts of the correct size as determined by EcoR1 digestion (5 U for 1 h at 37 °C) and agarose gel electrophoresis. 2.7. In-situ hybridisation: Homarus burs a, burs b and CCAP PCR products from an amplification using Homarus ISH primers (see Table 1) were electrophoresed and bands extracted as described above. These amplicons served as template for subsequent PCRs with primers containing T7 phage promoter sequences (Bursa T7 F, Bursa T7 R; BursB T7 F, BursB T7 R for burs a and burs b, respectively, CCAP T7 F, CCAP T7 R). DNA templates for making digoxygenin (DIG)-labelled cRNA probes for in-situ hybridisation 68 J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 were prepared by PCR using either a forward or reverse Homarus ISH T7 primer paired with the complementary Homarus ISH primer for the generation of sense and antisense cRNA. PCR reagents were as follows: 45 ll Megamix Blue (Helena Biosciences), 2.5 ll Homarus ISH T7 primer, 2.5 ll Homarus ISH primer (both 10 lmol l1), 1 ll template (above). PCR conditions were: 94 °C 4 min; 35 cycles of 94 °C 30 s, 58 °C 30 s, 72 °C 45 s and final extension at 72 °C for 10 min. After confirmation of specific amplification of correctly sized fragments on agarose gels, PCR products from three reactions were concentrated by centrifugation on YM30 columns (Millipore, Billerica, MA, USA). DNA (100–200 ng) was subsequently used for in vitro transcription using a MegaShortScript kit (Ambion) according to the manufacturer’s instructions, but the transcription reaction was modified to include DIG-11-UTP. Transcription conditions were 1 ll each of CTP, GTP, ATP, 0.5 ll UTP (all 75 mmol l1), 2 ll 10 lmol l1 DIG-11-UTP (Roche Diagnostics GmbH, Mannheim, Germany). Transcript quality was tested as detailed previously (Wilcockson and Webster, 2008). Paraformaldehyde fixed, methanol dehydrated ganglia were rehydrated (100%, 66%, 33%, 0% methanol/PBST) and incubated in 200 lg ml1 proteinase K (Roche) in PBS (7 min). Tissues were then washed in PBST (3 10 min) and post-fixed in 4% paraformaldehyde (60 min), followed with washing in PBST (3 10 min). Tissues were pre-hybridised with hybridisation solution (50% formamide, 1.3 SSC, 5 mmol l1 EDTA, 50 lg ml1 tRNA, 0.2% Tween 20, 0.5% CHAPS, 100 lg ml1 heparin) (30 min, 50 °C) prior to hybridisation in fresh hybridisation solution containing 1 ng ll1 DIG-labelled cRNA probe (sense or antisense, 18 h at 50 °C). Post-hybridisation washes were performed as follows: 2 SSC, 50% formamide (10 min, 50 °C, 2), 0.2 SSC (10 min, 50 °C, 2), PBST/0.2 SSC (33%, 66%, 100%) (5 min, RT). Tissues were washed (3 10 min, RT) in 1 TE containing 0.5 mol l1 NaCl (TNE) before treating with 20 lg ml1 RNase A in TNE (10 min, RT). Following RNase treatment, ganglia were blocked in 1% BSA in PBST for 2 h prior to overnight incubation in 1:5000 anti-DIG alkaline phosphatase (Roche) in PBST containing 0.1% BSA. Tissues were then washed extensively in PBST (5 min, 10 min, 2 15 min, 2 30 min, 60 min), followed by TMNT (10 min) and developed in NBT/BCIP. Reactions were terminated with distilled water when satisfactory colour development was observed. Ganglia were then carefully dissected to remove the perineural sheath and tissue surrounding the labelled somata (since the labelled neurons were otherwise not visible) and mounted in 80% glycerol/PBS. Digitally acquired microscopic images were prepared using ImageJ 1.42, Adobe Photoshop 7.0 and CorelDraw 8.0 software. 2.8. Quantitative RT-PCR Homarus burs a, burs b, CCAP and the reference gene rpl18 cRNA templates for the generation of qPCR standard curves were prepared by PCR using one Homarus T7 primer paired with one nonT7 Homarus primer (see Table 1), followed by in vitro transcription with T7 RNA polymerase (MEGAshortscript, Ambion) according to the manufacturer’s instructions. Run-off transcripts were purified on 6 mol l1 urea–polyacrylamide (10%) gels. Bands of the expected size were excised and eluted overnight at room temperature in Probe Elution Solution (Ambion). Eluates were ethanol precipitated, resuspended in TE, quantified by spectrophotometry (NanoDrop ND-1000), and converted to copy number by multiplying moles per sample by Avogadro’s number. Standards were diluted to 1 1011 copies per ll in TE and stored at 80 °C until use. Quantitative PCR was performed on an Applied Biosystems 7900 thermocycler using Taqman Universal PCR mix (Applied Biosystems). Reaction volumes were 20 ll, with 200 nmol l1 of one of the following primer pairs: Hom_alphaF/Hom_betaR, Hom_betaF/ Hom_betaR, Hom_CCAPF/Hom_CCAPR, Hom_RPLF/Hom_RPLR. Each reaction contained 1 ll cDNA. Specific amplification was detected by 250 nmol l1 FAM-labelled Taqman MGB hydrolysis probes (alphapro, betapro, ccappro, RPLpro). Standards were run on each plate in 10-fold serial dilutions in the range 1 108 to 1 103 copies per reaction. All reactions were run in duplicate. Cycling conditions were 95 °C 10 min, 40 cycles of 95 °C 15 s, 60 °C 60 s. PCR efficiencies were determined using the formula E = 1 + 10(1/slope) where ‘slope’ refers to the gradient of the line plotted from the Ct value/log copies RNA. In all cases, PCR efficiencies were >90%. 2.9. Immunohistochemistry and recombinant hormone production Following fixation, ganglia were washed extensively in PBS containing 0.1% Triton X-100, 0.01% sodium azide (PTX). CCAP immunohistochemistry was performed using anti-CCAP (Code 2TB, Dircksen and Keller, 1988). For bursicon, the antiserum raised in rabbit against recombinant Carcinus Burs a, was used (see below). Incubations were performed for 3 days at 4 °C in PTX at dilutions of 1:500. Following extensive washing in PTX, ganglia were incubated (overnight, 4 °C in Alexa Fluor 568 goat anti-rabbit IgG, 1:1000 (Invitrogen), then washed extensively in PTX. For double staining, these preparations were then incubated (3 days, 4 °C) in 1:100 Carcinus anti-burs a conjugated to Dylight Fluor 488 (Thermo Scientific, Wilmington, USA). After extensive washing in PTX, ganglia were mounted in VectaShield (Vector Laboratories, Burlingame, USA), and examined by confocal microscopy using a Zeiss LSM 510 instrument. Proprietary software was used for stacked projection analysis. Between 10 and 15 consecutive (1.5 lm thick optical slices) images were collected for each projection. Recombinant Burs a was produced from PCR amplified product to which NcoI (50 ) and XhoI (30 ) restriction sites had been added (forward: GGCCATGGGGACGAGTGTTCTCTCCGCCTG, reverse: TATACTCGAGCTATTTGAGGAAGGGAACGCTGTCC) followed by directional cloning into pENTR11, recombination into pDEST17 via Gateway cloning (Invitrogen). Competent cells (BL21-CodonPlus (DE3)-RIPL (Stratagene, La Jolla, USA) were transfected according to the manufacturer’s instructions, and expression induced with 1 mM IPTG (6 h, 37 °C). Cells were harvested, and inclusion body protein extracted and solubilised with B-PER and inclusion body solubilisation reagent (Thermo Scientific, Rockford, USA). Recombinant protein was affinity purified (HisPur Cobalt 6 His 1 ml column, Thermo Scientific), and purity of eluate confirmed via SDS–PAGE, Western blotting and detection with anti-polyHistidine (anti-mouse, 1:1000, 1 h, RT, Sigma), sheep anti-mouse peroxidase, 1:15,000, 1 h, RT, GE Healthcare Ltd., UK) and chemiluminescent detection (Amersham Hyperfilm ECL). Following extensive dialysis and concentration, recombinant Burs a was solubilised in 3 M urea, quantified by spectrophotometry, and antibodies raised commercially in rabbits (Eurogentec, Seraing, Belgium). 3. Results 3.1. Characterisation of cDNAs Using a strategy involving identification of partial cDNA sequences for burs a by fully degenerate PCR, 30 and 50 RACE and using GSPs derived from an EST for H. americanus burs b, full-length cDNA sequences encoding both bursicon subunits were obtained for H. gammarus. Conceptually translated peptide sequences, including mature peptide and signal peptides identified using Signal P 3.0 (http://www.cbs.dtu.dk/services/SignalP/) in comparison with other crustacean bursicon sequences are shown in Fig. 1, and compared with other arthropods, in Fig. 2. H. gammarus mRNA encoding the Burs a precursor consists of a 420 nt ORF encoding a J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 69 Fig. 1. Alignments and comparisons of conceptual translations of crustacean bursicon a and b subunit precursor peptides derived from cDNA sequences. Water flea, Daphnia arenata; Green shore crab, Carcinus maenas; Blue crab, Callinectes sapidus; European lobster, Homarus gammarus; American lobster, Homarus americanus. Putative signal peptides are underlined. Amino acids that are identical in all are boxed in grey. Accession Nos. Bursicon a/b: D. arenata, EU139431/EU139430; C. maenas EU139428/ EU139429; C. sapidus EU677191/EU677190; H. gammarus HM113369/113370. H. americanus –/CN854188. Note. For C. sapidus Burs b a Cys residue has been annotated at position 46. Since this is at odds with the Ser at this position for all other bursicons, this ambiguity most likely represents a sequencing miscall |(tgt-tct). 19 amino acid signal and 121 amino acid Burs a subunit (Average Mr: 13365.48, with reduced Cys residues). For Burs b a 408 nt ORF encodes a 21 amino acid signal and 115 amino acid Burs b subunit (Average Mr: 12928.54, with reduced Cys residues). Nucleotide and protein sequences have been submitted to EMBLGenBank databases: Accession Nos. for H. gammarus burs a, HM113369; burs b, HM113370. 3.2. Expression of CCAP, burs a and burs b mRNA in the CNS of H. gammarus by in-situ hybridisation Whole mount in-situ hybridisation of cerebral, sub-oesophageal, thoracic, abdominal and terminal ganglia using DIG-labelled antisense cRNA probes for CCAP, burs a and burs b showed intense hybridisation signals. For sense controls, specific hybridisation was never observed (results not shown). A summary of the hybridisation profiles is shown in Fig. 3. For all three transcripts, apparent co-localisation was striking, excepting the singular expression of CCAP transcripts in the brain (Fig. 3m) and optic ganglia, where bursicon subunit expression was never observed. On the dorsal surface of the sub-oesophageal ganglion, three pairs of strongly hybridising large (80 lm) and small (20 lm) perikarya, and anteriorly, two pairs of small (20 lm) perikarya corresponding to each neuromer were observed, for burs a, burs b and CCAP (Fig. 3a, e and i, respectively). In each thoracic ganglion, essentially similar patterns of expression of pairs of large and small perikarya were seen (Fig. 3b, f and j). For abdominal ganglia 1–5, two pairs of large and one pair of small perikarya were observed (Fig. 3c, g and k). In the terminal ganglion three pairs of large, particularly intensely hybridising perikarya were always observed in the anteriodorsal area. A further group of three ventrolateral cells were seen in the central region of the terminal ganglion (Fig. 3d, h and l). However, the most striking feature was the large number of intensely hybridising neurons on both sides of the medial posterior region of this ganglion (Fig. 3 d, h and l). Whilst it was difficult to determine the exact cell number, due in part to the thickness and position of the neurons in these large preparations which hindered both probe penetration and visualisation, in most preparations, approximately 20 large perikarya could be seen on each side of this ganglion. Thus, the terminal ganglion contained around 52 large perikarya expressing all three transcripts. It was notable that the small hybridising neurones seen in the other ganglia were never observed in the terminal ganglion (see Fig. 4). 3.3. CCAP, Burs a and Burs b expression in the CNS of H. gammarus by immunohistochemistry Whole mount immunohistochemistry on CNS from juvenile H. gammarus using heterologous antisera raised against recombinant C. maenas Burs a revealed immunopositive somata in the CNS (Fig. 3n–s). Burs a-immunoreactivity (IR) was always much poorer (with very high background) than for CCAP probably as a result of low IgG titre, low cross reactivity, or poor labelling efficiency of antiserum conjugate. Bursicon IR was specific: in that preabsorbtion controls showed that immunoreactivity of bursicon expressing neurones in the of thoracic ganglia of C. maenas could be completely abolished by preabsorbtion (4 °C, overnight) of 1 ll of antiserum with ca. 1 nmol of recombinant Burs a, and that the antiserum recognised only a single bursicon-containing peak in HPLC separated pericardial organ extracts and single ca. 14 kDa band from Western blotting of thoracic ganglia and pericardial organ extracts (own unpublished results). Patterns of CCAP-IR and Burs a-IR in the VNC were strikingly similar to those for CCAP and burs a expression detected by in-situ hybridisation. Each thoracic ganglion contained a dorsal pair of large (50 lm) and small (20 lm) CCAP-IR and Burs a-IR (Fig. 3p and q), whilst two large and one small perikarya were observed in AG 1–5 (Fig. 3n and o). In the sub-oesophageal ganglion five pairs of one large and one small perikarya exhibited both Burs a and CCAP immunoreactivity (Fig 3r and s). Immunoreactive neurons could not be consistently observed in the terminal ganglion, 70 J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 Fig. 2. Sequence alignments of bursicon a and b subunit proteins in arthropods. Identical residues are boxed in grey. Gaps have been added to maximise identity. Cysteine residues are indicated by arrows. Only species in which both subunits have been fully identified are included. Accession numbers for insect and tick sequences, bursicon a/b: Drosophila melanogaster, AY672905/AY823257; Anopheles gambiae, AY735443/AY823259; Apis mellifera, AM420631/AM420632; Tribolium castaneum, DQ138189/DQ138190; Acyrthosiphon pisum (http://www.aphidbase.org), APD05277/APD13715; Manduca sexta, DQ09449/DQ291147; Bombyx mori, BN000691/BN000690; Pediculus humanis corporis, XP_002430782/XP_002430781; Ixodes scapularis, XP_002407512/XM_002407469. Note. For Pediculus, reanalysis of the gene structure of bursicon b showed that predicted intron 3 of the gene (nts 481,291–481,401) contained the start of exon 3. The correct intronic sequence, which contains the GT-AG splice donor and acceptor sites is from nts 481,291 to 481,379 thus, this revised exon 3 sequence codes for the gap in the predicted protein sequence, which is at odds with all other Burs b sequences and accounts for residues ECFCCREK which are missing in XP_002430781. presumably because of the thickness of the preparation, position of the immunoreactive cells and weak fluorescent signals relative to strong background. Nevertheless, double immunolabelling using anti-CCAP and anti-Burs a antisera sequentially did suggest complete co-localisation of CCAP and Burs a in the sub-oesophageal, thoracic and abdominal ganglia. Burs a-IR perikarya were never observed in the brain or eyestalks. The lack of axonal labelling in all of the preparations was noteworthy. Some labelling was present immediately adjacent to the soma but it was impossible to determine any neuroarchitecture. 3.4. Expression analysis of CCAP, burs a and burs b mRNA by quantitative RT-PCR To accurately quantify the expression of each mRNA in the individual ganglia we performed qRT-PCR with Taqman chemistry. Data were initially normalised to those of the reference gene RPL18, but since this was obviously related to tissue mass, a more useful normaliser might be one involving cell number per ganglion. Thus, transcript abundance was normalised against the number of large perikarya (i.e., per cell), as shown by in-situ hybridisation. J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 71 Fig. 3. Expression of bursicon a, b and crustacean cardioactive peptide (CCAP) transcripts in the central nervous system of H. gammarus: In-situ hybridisation. ((a–d) Labelling with antisense DIG-burs a cRNA probe). (a) Whole mount of sub-oesophageal ganglion. (b) First thoracic ganglion. Arrows point to small (20 lm) perikarya adjacent to large (80 lm) strongly hybridising perikarya. (c) First abdominal ganglion. Arrows show small, weakly hybridising perikarya adjacent to pairs of strongly hybridising large perikarya. (d) Terminal abdominal ganglion. Arrows show group of three large cells (80 lm) at anterior. ((e–h) labelling with antisense DIG-burs b cRNA probe). (e) Suboesophageal ganglion. (f) First thoracic ganglion. (g) First abdominal ganglion. (h) Terminal abdominal ganglion. Arrows point to lateral groups of three hybridising perikarya. ((i–l) labelling with DIG-CCAP cRNA probe). (i) Sub-oesophageal ganglion, showing three pairs of large and small perikarya (arrows), anteriorly, two pairs of small perikarya (arrowheads). (j) First thoracic ganglion. (k) First abdominal ganglion. (l) Terminal abdominal ganglion. Arrow points to lateral group of three large perikarya. (m) Ventral view of whole mount of cerebral ganglion showing five pairs of large, strongly hybridizing (50 lm) perikarya in an anterior position. Posterior to these are four pairs of small, indistinct (ca. 15 lm) cells (arrows). In all cases, control (sense) DIG- labelled cRNA probes of showed no specific hybridisation signals. (n–s) Immunohistochemistry, red: anti-CCAP, green, anti-rburs a (C. maenas). (n and o) Double labelling of perikarya in first abdominal ganglion. (p and q) thoracic ganglion. (r and s) Sub-oesophageal ganglion. Scale bars: 1 mm (a, e, i and m), 500 lm (b, c, d, f, g, h, k and l) 250 lm (n, o, p, q, r and s), 50 lm. Fig. 4. Expression of CCAP (black), burs a (pale grey) and burs b (dark grey) mRNA in the central nervous system ganglia of H. gammarus. SOG, sub-oesophageal ganglion; T1– T5, thoracic ganglia 1–5; AG1–AG5, abdominal ganglia 1–5; TG, terminal ganglion. Measurements of mRNA copy numbers were expressed per large perikaryon (cnc-type1 neuron: nomenclature, Dircksen and Keller, 1988), as shown by whole mount in-situ hybridisation (Fig. 3). Error bars represent + 1SEM. N = 4 Differences in copy number between ganglia were not statistically significant for any transcript. All tissues were taken from intermoult specimens. All standards and samples were assayed in duplicate. Assuming similar rates of transcription by the small compared to large perikarya, the contribution of small perikarya, which have a volume approximately 2% of the large perikarya, is negligible. Cell numbers per ganglion were sub-oesophageal ganglion (SOG), six; thoracic ganglia (T), two; abdominal ganglia (AG), four; terminal ganglion (TG), ca. 52. burs a and burs b transcripts levels were quite 72 J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 similar, varied little between ganglia, and mean transcript numbers per cell were between 0.5 and 1.5 105 copies per cell. There were no significant differences in expression of all three transcripts between ganglia (two-way ANOVA p = 0.48). However, CCAP transcript number, although somewhat variable (0.5 105–6 105 copies per cell), was significantly higher than burs a or burs b (ANOVA, p < 0.05). These differences could not be attributed to differences in PCR efficiency (burs a, slope 3.35, 98% efficiency; burs b, slope 3.55, 91% efficiency, CCAP, slope 3.37, 98% efficiency). 4. Discussion In this study we first identified cDNAs (mRNAs) encoding both bursicon subunits in the European lobster, H. gammarus, using an approach (for burs a) based on degenerate PCR, using sequence information from other crustacean bursicons to design appropriate primers, and also, for burs b using primers designed from sequence data from the closely related lobster species, H. americanus. Peptide sequences for both H. gammarus bursicon subunits show high sequence similarity to the previously described crustacean bursicons of D. arenata, C. maenas (Wilcockson and Webster, 2008) and Callinectes sapidus (J.S. Chung, unpublished); with 67% and 58% sequence identity for Burs a and Burs b, respectively (Fig. 1). All Cys residues are located in identical positions for all crustacean sequences. Interestingly, the amino acid sequences for H. americanus and H. gammarus bursicon b subunits are identical. Whilst this might be unsurprising, considering their relatedness, in the two closely related portunid crabs – C. maenas, and C. sapidus, whilst both bursicon a subunits are identical, there are three amino acid differences in the bursicon b subunit sequences. However, for both lobster species, the similarity of not only protein, but nucleotide sequences is extraordinary, for the DNA sequence encoding the ORF there are only 13 nucleotide substitutions and three deletions (a valine deletion in the signal of H. americanus). A summary of arthropod bursicon sequences, where full, unambiguous annotation of both subunits has been documented, is given in Fig. 2. Whilst sequence identity is clearly evident on a phylogenetic basis (the lowest sequence identity is seen in the arachnid, Ixodes scapularis), it is noteworthy that in all the crustacean Burs b sequences there has been a deletion of D/E, (compared to insects) 19 residues from the C-terminus. Quite a number of bursicon-like molecules have now been identified in a variety of invertebrates, not only in arthropods, but also from the echinoderm Strongylocentrotus purpuratus (Van Loy et al., 2007), mollusc, Lottia gigantea, (Veenstra, 2010) and a hemichordate, Saccoglossus kowalevskii (Accession No. XP-002732831). Thus, since these molecules seem to be found amongst both protostomian and deuterostomian lineages, their origin is presumably quite ancient. Indeed, structurally related cystine knot proteins (CKPs) are widespread in vertebrates and invertebrates (reviews, Hearne and Gomme, 2000; Vitt et al., 2001). However, it should be stressed, if this is necessary, that it is extremely unlikely that these bursicon-like molecules will have equivalent functions to the arthropod hormones, in these animals. However, for crustaceans, analogous roles for bursicon, as for insects seem likely. Homogenates of abdominal ganglia of Homarus have been shown to be active in the Sarcophaga cuticle tanning assay (Kostron et al., 1995). Likewise, HPLC purified fractions of pericardial organs from C. maenas, showing immunoreactivity to Burs a are biologically active in this assay (own unpublished results). Bursicon expression, measured both qualitatively by ISH, and quantitatively by qRT-PCR has only recently been determined in the crab C. maenas (Wilcockson and Webster, 2008). In this study all neurons expressing CCAP mRNA, excepting those in the brain, also co-expressed mRNA that encodes both bursicon subunits. Fur- thermore, expression of all three transcripts was invariant throughout the moult cycle. This situation contrast vividly with that seen in insects where expression patterns of CCAP and bursicon are not always coincident, and expression patterns vary during development (review, Honegger et al., 2008). Additionally since the abdominal ganglion of crabs is relatively small, and fused to the thoracic ganglion, in our previous study, we could not precisely map the positions of the bursicon and CCAP expressing neurones in the AG, and thus it was possible that CCAP and bursicon might not be completely co-localised, since we were unable to perform ISH double hybridisations. To obtain more precision regarding anatomical localisation of CCAP and bursicon co-expressing neurons, we reasoned that the plesiomorphic nervous system of the lobster, with its large well-defined abdominal ganglia would be a useful model for ISH studies, particularly since the anatomy of CCAP neurons in the related astacurans (Astacus astacus, Orconectes limosus) have been mapped in great detail (Audehm et al., 1993; Trube et al., 1994). Whole mount in-situ hybridisation patterns for all three transcripts ( CCAP, burs a, burs b) were identical for neurons in the sub-oesophageal, thoracic, abdominal and terminal ganglia. Additionally, using CNS tissue from juvenile lobsters, we observed complete co-localisation of CCAP and Burs a peptides in the thoracic and abdominal ganglia (1–5) by immunohistochemistry, so also proving that these neurons produce both transcript and translated protein. Neurons that only expressed CCAP mRNA were exclusive to the brain. This is in accord with our results for C. maenas (Wilcockson and Webster, 2008) but in H. gammarus five pairs of large CCAP expressing midline neurons were observed at the anterior ventral margins of the protocerebrum compared to a single pair in C. maenas. In contrast, for insects, early studies showed bursicon-IR neurons in the brain of Periplaneta americana, Manduca sexta and Drosophila using an antibody raised against a bursicon fragment (Honegger et al., 2002). Whilst this result could not be confirmed for M. sexta in more recent studies using much more specific antisera raised against a Drosophila Burs a peptide or recombinant Burs a and b (Dai et al., 2008), the latter two antisera do label brain neurons in P. americana (which are not CCAP-IR), and these project to the corpora cardiaca (Honegger, unpublished, cited in Honegger et al. (2008)). Comparison of neurons exhibiting hybridisation for all three transcripts in lobsters, compared to homologous CCAP containing neurons in crayfish (Audehm et al., 1993; Trube et al., 1994) revealed similarities, such as the presence of five pairs of neurons in each hemineuromer of the SOG ganglion and TG: for the neuromers corresponding to maxilliped 1–3 in the SOG and for thoracic ganglia 1–5, they could clearly be identified (in terms of size) as cnc (large type 1, ca. 80 lm) and cdn (small type 2, ca. 20 lm) neurons (terminology according to Audehm et al. (1993)). In the anterior two maxillary neuromers of the SOG, these neurons could not be differentiated in this way, and the single pair of neurons associated with the mandibular neuromer seen in crayfish was never observed. In the first five abdominal ganglia expression of all three transcripts was essentially identical to that seen in crayfish (two pairs of cdn, one pair of cnc neurons), but for the terminal ganglion there were some differences. The arrangement of the three groups of neurons hybridising with all three transcripts (anterior-lateral ventral cell cluster, AVLC; posteriorlateral cell cluster, PLC; and posterior median lateral cell cluster, PMLC (terminology according to Trube et al. (1994)) were broadly similar, and indicate that this ganglion is fused from AG 6 to 8 as previously comprehensively discussed by the above authors. However, we could not identify small cdn-type neurones in the AVLC or PLC – all appeared to be of a diameter indicative of cnc-type neurons. Additionally, in the crayfish each PMLC contains six CCAP expressing neurones, whereas in the lobster, approximately 20 neurons were seen. The presence of large cnc-type neurons in J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 the terminal ganglion is reminiscent of the situation in C. maenas, where small type neurons are absent. However, a caveat might be that the somewhat harsh processes involved in ISH could introduce artefacts regarding apparent neuronal size; despite the observation of small cdn-type neurons in the SOG, TG and AG1–5. Thus, full identification of the cell types which depends on comprehensive analysis of their projection patterns awaits the availability of suitable antisera. Apropos this, it was surprising that we could not trace projections of CCAP (or Burs a) neurons by ICC in our preparations. Whilst the juvenile lobsters used were of equivalent sizes to crayfish to minimise issues related to poor penetration of antibody, axon profiles were never observed, despite strong labelling of perikarya for CCAP (but not using the heterologous C. maenas Burs a antibody). Nevertheless, since the release sites of the cnc neurons in crayfish is the large and diffuse dorsal area of the perineural sheath, extending the entire length of the VNC and extending to the posterior and dorsal telson flexor muscles (Audehm et al., 1993; Trube et al., 1994), the same morphology in lobsters would ensure that co-release of CCAP and bursicon into the circulation would be rapid, as it would from the equivalent release site (the pericardial organs, in crabs). If the distributions of CCAP and bursicon expressing neurons in crustaceans are compared to those of insects, then it is obvious that there are fundamental differences. In holometabolous insects, changes in bursicon expression are obviously related to development: in larval Manduca, bursicon is only expressed in Cells 27 (cnc type) of the first abdominal ganglia, but in pharate pupae Cells 27 of all abdominal ganglia express bursicon (Dai et al., 2008). Similarly the homologues of Cells 27 in the first four abdominal ganglia (i.e., 8 cells) of Drosophila express bursicon in larvae, but in pharate adults the number increases to 14 (Luan et al., 2006a,b), and in both M. sexta and Drosophila programmed cell death of most of the abdominal CCAP-IR neurones occurs after eclosion (Ewer et al., 1998; Draizen et al., 1999). However for hemimetabolous insects such as P. americana, the pattern of bursicon (and CCAP expressing neurons) remains similar throughout development. It has been suggested that this would be expected in these insects, since the nature of the cuticle remains essentially unchanged during development, in contrast to holometabolous insects (Honegger et al., 2008). Although the patterns of development of bursicon immunoreactive neurons during embryogenesis and larval development in crustaceans are unknown, burs a, b expression is seen in C. maenas embryos at about 50–70% development (Wilcockson and Webster, 2008) which parallels the appearance of CCAP transcripts and peptide (Chung et al., 2006). Furthermore the serial iteration of CCAP immunoreactive neurones (cdn and cnc types) in the sub-oesophageal ganglia and thoracic ganglia 1–5 together with segmental nerve projections to the PO is seen in embryonic lobsters at a similar stage (E79) of development (Pulver and Marder, 2002). Given this background, it would be interesting to determine the expression patterns of bursicon and CCAP during embryonic and larval development, particularly with reference to adult neuronal morphology. As with hemimetabolous insects, there is no overtly dramatic change in body form or cuticle plasticity in decapod metamorphosis (compared to holometabolous insects) thus it seems possible that patterns of larval bursicon and CCAP containing neurons will be very similar to those of the adult. In insects, it is known that some neurons only display immunoreactivity to one of the bursicon subunits (Honegger et al., 2002; Luo et al., 2005; Dai et al., 2008). Whilst our qualitative (ISH) studies on H. gammarus and C. maenas indicated that all neurones in the VNC expressing CCAP also expressed both bursicon transcripts and translated products, suggesting that this might not occur in crustaceans, the plesiomorphic nature of the lobster VNC allowed us to estimate approximate levels of transcript per neuron, to see whether there were any segment specific changes in gene expres- 73 sion, or large differences in ratios of bursicon subunit expression that might indicate the existence of bursicon homodimers. Since the total volume of the small cdn-type neurons is only about 2% of that of the cnc type, assuming similar rates of transcription in both small and large cells, the contribution to overall gene expression is essentially from the large cnc-type neurons. Thus, approximate transcript number per (cnc) cell could be estimated by qRT-PCR of cDNA derived from identified ganglia. This approach showed that transcript number for both burs a and burs b mRNAs were similar in all ganglia. This contrasts with the situation in C. maenas, where burs a transcripts seem to be about three times more abundant than those of burs b in the adult CNS and during the latter stages of embryogenesis (Wilcockson and Webster, 2008) CCAP expression was quite variable, but significantly greater than that of bursicon. However, the generalisation that expression of all three transcripts is essentially similar is probably justified. An important difference between the patterns of synthesis of bursicon in insects and crustaceans is obviously related to life history and developmental processes. In Drosophila levels of both bursicon subunit transcripts rise during puparium formation, peaking at the pharate adult stage and thereafter decline rapidly (Luo et al., 2005). This pattern is of course entirely in keeping with the established roles of this hormone in wing expansion and maturation, and cuticle sclerotization and melanisation (review, Honegger et al., 2008). Since expression of bursicon is constitutive in C. maenas and H. gammarus, and bearing in mind that in these crustaceans (indeed for most decapods) moulting continues throughout adult life, release patterns of this hormone must be fundamentally different to those of insects, Furthermore, since all neurones in the CNS express both CCAP and bursicon, these hormones will be co-released in the same terminal boutons in the pericardial organs and (for lobsters) dorsal perineural sheath. Since bursicon and CCAP are co-packaged in the same dense cored vesicles in neurosecretory neurons of the abdominal neurons of P. americana (Woodruff et al., 2008), it is likely that this situation will prevail in crustaceans. Since CCAP is released from the PO of crustaceans in a massive surge at the start of active ecdysis (Phlippen et al., 2000), bursicon should also be released at the same time. Whilst co-release would certainly aid efficient circulation of bursicon via CCAP’s first established cardioacceleratory action (Stangier et al., 1987), a co-release would also seem counterintuitive, since during ecdysis CCAP should act before bursicon. However, whilst the downstream targets for bursicon are currently unknown in crustaceans, it is entirely possible that its effects might well be long lived. In this context the recent studies on effects of recombinant bursicon on gene transcription in Drosophila are fascinating. Gene expression following hormone injection, measured by microarray and quantitative PCR showed that a diverse collection of 87 genes, of extraordinarily diverse function, are regulated by bursicon, 13 of which were identified as being involved in cuticle sclerotization. (An et al., 2008). Thus, a hypothetical scenario involving temporal separation of signal, (i.e., long lasting post-ecdysis effects of bursicon on gene expression, versus short-term behavioural changes exerted by CCAP during ecdysis) despite co-release of both hormones is an attractive one. A central role for CCAP initiating ecdysis motor behaviour associated with cuticle shedding has been suggested from studies on M. sexta (Gammie and Truman, 1997), and in Drosophila genetic ablation of CCAP neurons (which also express bursicon) cause failure of pupal ecdysis and wing expansion defects (Park et al., 2003). However, new layers of complexity are constantly being discovered: recent studies show that subsets of CCAP neurons are active at different times during ecdysis. Neurons that co-express bursicon (NCCAP-c929) are recruited late in the ecdysis sequence (Kim et al., 2006), and are under the neural control of those that only produce CCAP (NCCAP) (Luan et al., 2006a,b). Furthermore, the 74 J.H. Sharp et al. / General and Comparative Endocrinology 169 (2010) 65–74 excitability of the abdominal neurons that co-express CCAP and bursicon is modulated by a pair of co-expressing neurons in the sub-oesophageal ganglion (BSEG) which send projections to the abdominal ganglia (and elsewhere). It has been suggested that activity of these cells during ecdysis lead to bursicon release from the AG, and other behaviours such as air swallowing (Peabody et al., 2008). Additionally, these authors have shown that rickets flies lacking bursicon receptor functionality have defects in bursicon release patterns and post-eclosion wing epidermal cell apoptosis. These studies highlight the complexity and subtlety of neural networks involved in ecdysis in a genetically tractable animal, and with relevance to intractable crustacean models, shows how release patterns of bursicon might be controlled. 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