Download Identification and expression of mRNAs encoding bursicon in the

General and Comparative Endocrinology 169 (2010) 65–74
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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
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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
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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,
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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
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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
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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. Accordingly,
one of next stages of research in crustaceans aimed at understanding the hormonal control of ecdysis should be to simultaneously
measure both CCAP and bursicon contents of individual ganglia
and circulating hormone levels during precisely staged periods of
ecdysis and post-ecdysis.
Acknowledgments
This work was funded by the Biotechnology and Biological Sciences Research Council (BBSRC), Grant Reference No. BBE0231261.
The authors thank Dr. M. Ehrhardt (University of Manchester) for
his invaluable help producing recombinant bursicon.
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