Báo cáo khoa học: Identification of an atypical insect olfactory receptor subtype highly conserved within noctuids potx

Identification of an atypical insect olfactory receptor
subtype highly conserved within noctuids
´
Isabelle Brigaud1, Nicolas Montagne2, Christelle Monsempes1, Marie-Christine Francois1 and
¸
Emmanuelle Jacquin-Joly1
1 INRA, UMR PISC, UMR-A 1272, Versailles, France
´
2 UPMC Universite Paris 6, UMR PISC, Paris, France

Keywords
Lepidoptera; Noctuidae; olfaction; olfactory
receptor; phylogeny
Correspondence
E. Jacquin-Joly, INRA UMR 1272
INRA-UPMC PISC Physiologie de l’Insecte:
Signalisation et Communication, route de
Saint-Cyr, F-78000 Versailles, France
Fax: (33) 1 30 83 31 19
Tel: (33) 1 30 83 32 12
E-mail:
(Received 24 June 2009, revised 31 July
2009, accepted 3 September 2009)
doi:10.1111/j.1742-4658.2009.07351.x

Olfaction is primarily mediated by the large family of olfactory receptors.
Although all insect olfactory receptors share the same structure with seven
transmembrane domains, they present poor sequence homologies within
and between species. As the only exception, Drosophila melanogaster
OR83b and its orthologues define a receptor subtype singularly conserved
between insect species. In this article, we report the identification of a new

subtype of putative olfactory receptors exceptionally conserved within noctuids, a taxonomic group that includes crop pest insects. Through homology-based molecular cloning, homologues of the previously identified
OR18 from Heliothis virescens were identified in the antennae of six noctuid species from various genera, presenting an average of 88% sequence
identity. No orthologues were found in genomes available from diverse
insect orders and selection pressure analysis revealed that the noctuid
OR18s are under purifying selection. The OR18 gene was studied in details
in the cotton leafworm, Spodoptera littoralis, where it presented all the
characteristic features of an olfactory receptor encoding gene: its expression
was restricted to the antennae, with expression in both sexes; its developmental expression pattern was reminiscent of that from other olfactory
genes; and in situ hybridization experiments within the antennae revealed
that the receptor-expressing cells were closely associated with the olfactory
structures, including pheromone- and non-pheromone-sensitive structures.
Taken together, our data suggest that we have identified a new original
subtype of olfactory receptors that are extremely conserved within noctuids
and that might fulfil a critical function in male and female noctuid chemosensory neurones.

Introduction
The insect noctuid family includes devastating agricultural pests. As nocturnal animals, they depend strongly
on olfactory cues to detect food and mates. Therefore,
their olfactory system is an attractive target for their
control. Odour reception is primarily mediated by the
large family of olfactory receptors (ORs) that ensure

the specificity of the olfactory receptor neurone (ORN)
responses. ORs are expressed on the surface of ORN
dendrites that are housed in morphofunctional units,
distributed along the antennae – the olfactory sensilla.
Intense efforts to identify insect ORs are currently
being undertaken, as their G-protein-coupled receptor

Abbreviations

GPCR, G-protein-coupled receptor; OR, olfactory receptor; ORN, olfactory receptor neurones; PR, pheromone receptor; qPCR, quantitative
real-time PCR.

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(GPCR)-like structure may open up the way for the
design of agonist and ⁄ or antagonist molecules based
on the pharmacological know-how accumulated on
GPCRs.
ORs were first discovered in vertebrates in 1991 [1],
but, because of extreme sequence divergence, insect
ORs were not discovered until the Drosophila melanogaster genome was sequenced [2–5]. Since then, insect
OR-encoding genes have been mainly identified
through bioinformatics analysis of complete or partial
available genomic databases. Complete or partial sets
of OR genes are now available from various insects,
including 12 species of the Drosophila genus [6], the
mosquitoes Anopheles gambiae [7] and Aedes aegypti
[8], the lepidopterans Bombyx mori [9,10] and Heliothis virescens [11,12], the honeybee Apis mellifera [13]
and the red flour beetle Tribolium castaneum [14,15].
All insect ORs identified so far share common features:
like GPCRs, they belong to the superfamily of seven
transmembrane domain receptors; they are exclusively

expressed in chemosensory organs; and the expressing
cells are located beneath the chemosensory sensilla.
However, they present a pronounced intra- as well as
interspecific sequence diversity (20–40% sequence identity) [16]. This poor sequence conservation has halted
industrial interest, precluding the elaboration of broadspectrum products for crop protection.
As an exception, one particular set of insect ORs
defines a unique subtype of receptors [17]. This subtype
groups receptors singularly conserved between insect
species, the so-called D. melanogaster OR83b (DmelOR83b) orthologues. Their high conservation level
(60–90% sequence identity) has allowed the isolation
of their counterparts in various insect orders through
homology cloning [17–22], whereas this strategy has
failed for most other OR types. The DmelOR83b
protein does not appear to encode an OR per se, but
heterodimerizes with conventional ORs, enabling their
correct trafficking and functionality [23,24]. Interestingly, DmelOR83b orthologues can retain their function when expressed in D. melanogaster, although there
is as yet no direct proof of their function in vivo in any
other species apart from D. melanogaster [25].
Although, at first glance, this receptor subtype could
appear to be the ideal universal target to disturb insect
olfaction, members are also found in beneficial insects
such as the honeybee [17], thus precluding the use of
molecules interfering with OR83b receptors.
Another subfamily of insect ORs, restricted to
moths, can also be distinguished but with lower conservation. This family consists of the pheromone receptors (PRs) that share an average of 40% identity.
Members have been identified in several moth species,
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thanks to genomic data analyses [11,26,27], differential
screening [9,28] and homology cloning strategies [29].

These receptors are predominantly male specific, and
some have been shown to respond to pheromones
[9,29,30]. As most of the moth species are severe crop
pests, disruption of the moth pheromone communication system, through the use of synthetic pheromones,
is currently an efficient strategy. The design of compounds affecting PRs may allow the development of
novel strategies, but their relatively high divergence in
sequence will require a species-specific approach.
In this article, we used homology cloning strategies to
identify a new moth subtype of highly conserved candidate ORs, the OR18 subtype. Six full-length cDNAs
encoding ORs highly related to the H. virescens OR18
[11] were identified in representative noctuid species: the
Amphipyrinae Spodoptera littoralis and Sesamia nonagrioides, the Heliothinae Helicoverpa zea and Helicoverpa armigera, the Noctuinae Agrotis segetum, and
the Hadeninae Mamestra Brassicae (all crop pests).
Interestingly, gene subtype members could be identified
only in noctuids, and we present evidence that these
receptors are under purifying selection.
A detailed study of S. littoralis OR18 revealed typical features of insect ORs. Its expression is restricted
to the antennae and it is expressed late in development
and in association with olfactory sensilla. Taken
together, our data suggest that this new original subtype of ORs might play a specific role related to noctuid ecology and its conservation may offer a single
target for noctuid control.

Results and Discussion
The cloning of six H. virescens OR18 homologues
in noctuid species revealed a new highly
conserved subtype of candidate ORs
Through homology cloning strategies, six full-length
cDNAs related to the H. virescens OR HvirOR18,
were identified from six species representative of the
noctuid family. The encoded proteins are 398–400

amino acids long and were named SlitOR18 (S. littoralis), MbraOR18 (M. brassicae), HzeaOR18 (H. zea),
HarmOR18 (H. armigera), SnonOR18 (S. nonagrioides) and AsegOR18 (A. segetum). The Phobius tool
revealed high probability for the occurrence of five to
seven transmembrane domains depending on the proteins, but close examination of the hydropathy profiles
and sequence alignment suggested the occurrence of
seven transmembrane domains for all, occurring at
similar positions (Fig. 1A,B). OR18 sequences are
characterized by an extraordinarily high sequence

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I. Brigaud et al.

An atypical olfactory receptor subtype in noctuids

A

I

II

III

IV

V

VI

V

VII

B

SlitOR18
N-ter

DmelOR83b

EC2
C-ter

Cell
membrane

C-ter

IC2

N-ter
Fig. 1. Noctuid OR18 sequences and predicted membrane topology. (A) Alignment of amino acid sequences deduced from the HvirOR18,
HzeaOR18, HarmOR18, SnonOR18, MbraOR18, AsegOR18 and SlitOR18 cDNAs. Amino acids identical in the maximum sequences are
marked with grey shading. Arrows indicate the positions of the primers used in RT-PCR for gene fragment amplifications. Transmembrane
domains I–VII identified from Phobius [41] () are indicated. (B) Representation of the transmembrane topology prediction for SlitOR18 compared with DmelOR83b. Black, transmembrane domains; EC2, second extracellular loop; IC2, second intracellular loop.

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identity, ranging from 82% (e.g. SlitOR18 ⁄ SnonOR18
and SlitOR18 ⁄ AsegOR18 comparisons) to 98–99%
(e.g. HvirOR18 ⁄ HzeaOR18, HvirOR18 ⁄ HarmOR18
and HarmOR18 ⁄ HzeaOR18 comparisons). Within the
OR18 sequences, the overall identity reached 77% and
even 92% homology. Apart from the DmelOR83b
subtype, such a remarkable sequence conservation of
candidate ORs across species from different genera has
not been observed previously.
A phylogenetic analysis was run using a non-exhaustive repertoire of OR sequences identified in various
insect orders (Fig. 2A). Three subtypes ⁄ subfamilies
of insect ORs were clearly defined: the already wellcharacterized OR83b subtype, the known moth PR

subfamily and a new subtype of insect ORs formed by
HvirOR18 and its homologues identified in this study.
Like the members belonging to the OR83b and PR
subfamilies, the OR18 candidates clustered into a
monophyletic group, clearly distinct from the other
insect ORs and supported by the bootstrap values
(Fig. 2A,B). These observations suggest that members
of the OR18 subtype are orthologues.
OR18 orthologues were found only in noctuids
and are under purifying selection
Interestingly, this well-supported group contained only

lepidopteran sequences (in red, Fig. 2A). This raised the

A

B

6540

Fig. 2. (A) Unrooted neighbour-joining tree
of ORs from Lepidoptera (in red) and from
species representative of Hymenoptera
(Apis mellifera, yellow), Diptera (Drosophila melanogaster, green) and Coleoptera
(Tribolium castaneum, blue). Bootstrap
support values are based on 1000 replicates. Nodes with high bootstrap support
(over 90%) are marked by open circles and
those with support between 70% and 90%
are marked by filled circles. The branch
length is proportional to the genetic
distance. Three clades, grouping sequences
from different species, are clearly visible:
the already known moth pheromone receptor and OR83b clades, and a new clade
grouping the OR18 sequences identified in
this study. (B) Detail of the clade containing
the OR18 subtypes. Note that all of these
sequences only belong to the order
Lepidoptera.

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I. Brigaud et al.

An atypical olfactory receptor subtype in noctuids

question of whether related receptor types may also exist
in insects other than noctuids. To approach this question, we used both RT-PCR experiments and blast
searches against available insect genomes. Despite
several attempts, RT-PCR performed with total RNA
from the antennae of the Bombycidae B. mori and the
Crambidae Ostrinia nubilalis – both representative
species of two other Lepidoptera families – gave no
amplification. In blast searches against the National
Center for Biotechnology Information (NCBI) DNA
and protein databases, or directly towards partial or
complete sequenced genomes, the most significant
sequence matches did not exceed 36%, in accordance
with the absence of an amplicon in the B. mori RT-PCR
experiments. As complete OR repertoires are now established in species representative of the major holometabolous insect orders (Diptera, Hymenoptera, Coleoptera,
Lepidoptera) and none of these ORs appears to share
significant identity with OR18 sequences, we concluded
that the OR18 subtype may be restricted to some
lepidopteran species, including the noctuids.
Selection pressure on the OR18 gene subtype has
been studied by comparative analysis of synonymous
(dS) and nonsynonymous (dN) nucleotide divergence.
This approach allows for the testing of evolutionary
selection scenarios, acting on protein coding sequences.
Table 1 compares the dN ⁄ dS values of the noctuid
OR18 and immediately related OR genes. dN ⁄ dS values are low for the OR18 gene subtype (0.009–0.131).
Assuming that all nucleotides have an equal probability of changing over evolutionary time, the observation

of a disproportionate number of synonymous changes
(dN ⁄ dS < 1) suggests purifying selection for noctuid
OR18s, confirmed by statistical analysis (P < 0.05).
This is consistent with the strong conservation of
OR18 genes across noctuids and also with a potential
essential functional role. Indeed, such a low ratio
has also been observed for the OR83b gene, whose

essential function in insect olfaction is well established
[23,24]. Although dN ⁄ dS ratios are relatively high for
chemosensory genes across insects, such purifying
selection has been proposed to be the main force governing the evolution of chemosensory genes within the
D. melanogaster group (reviewed in [31]). Thus, we
extend such an evolutionary scenario within more distantly related insect species. To date, only a few OR
genes have been identified in Lepidoptera. Further OR
gene identification within Lepidoptera, and particularly
within noctuids, may reveal that purifying selection is
a more common force in the evolution of OR genes.
Like the OR18 subtype, other highly conserved OR
subtypes may emerge.
Detailed analyses of SlitOR18 revealed common
features with insect ORs
The polyphagous S. littoralis is an example of a crop
pest. The molecular characterization of its OR18 gene,
SlitOR18, revealed expected features of genes belonging to the OR superfamily, thus confirming the newly
identified genes as candidate OR encoding genes.
First, real-time PCR was used as a quantitative
method to compare SlitOR18 expression levels in different tissues (male and female antennae, brain, proboscis, abdomen and legs). As illustrated in Fig. 3,
SlitOR18 expression was restricted to the antennae,
with negligible expression in the other tissues tested.

Within antennae, SlitOR18 was found to be almost
equally expressed in males (55%) and females (43%).
Second, the developmental expression pattern of
SlitOR18 was established using RT-PCR on head ⁄
antennal RNA from different stages of development.
As shown in Fig. 4A, SlitOR18 expression was
detected only in late pupal stages (starting 2 days
before emergence) and adulthood. No expression was
observed in embryos, fifth instar larvae heads and

Table 1. dN ⁄ dS in the OR18 subtype. Values in bold italic indicate purifying selection (P < 0.05) that occurred in all the OR18 subtypes.
SlitOR18
MbraOR18
SnonOR18
AsegOR18
HzeaOR18
HarmOR18
HvirOR18
HvirOR20
BmorOR30
BmorOR33
BmorOR34

MbraOR18

SnonOR18

AsegOR18

HzeaOR18

HarmOR18

HvirOR18

HvirOR20

BmorOR30

BmorOR33

0.12
0.097
0.131
0.112
0.112
0.092
0.746
1.289
1.623
1.68

0.054
0.053
0.086
0.098
0.082
0.726
0.95
1.359

1.424

0.038
0.094
0.105
0.1
0.858
1.147
1.264
1.355

0.09
0.082
0.078
0.65
1.253
1.296
1.366

0.009
0.031
0.72
1.195
1.312
1.418

0.039
0.749
1.17
1.255

1.317

0.723
1.212
1.322
1.434

0.866
1.378
1.41

0.584
0.566

0.378

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SlitOR18 transcript level
relative to level in male antennae

An atypical olfactory receptor subtype in noctuids

I. Brigaud et al.

1.5

1

0.5

0

Fig. 3. Expression of SlitOR18 in different adult tissues using
real-time PCR. Expression levels were calculated relative to the
expression of the rpL8 control gene, expressed as the ratio
ESlitOR18(DCT)SlitOR18 ⁄ ErpL8(DCT)rpL8 [45] and are reported relative to
the level in male antennae.

Pupae

Adult
48
E+

E+

1
E-

2
E-

5
E-

L5

Eg

gs

12

h

h

Larvae

A

750
650
600
550

SlitOR18

gD

cD

NA

B

NA

rpL8

2000
1500
1000
500
Fig. 4. Temporal expression pattern of SlitOR18 using RT-PCR. (A)
RT-PCRs were performed using the SlitOR18 primer pair and RNAs
isolated from the heads of fifth instar larvae (larvae L5), antennae
from pupae collected x days before emergence (pupae E-x) and
antennae from adults collected y hours after emergence (adult
E + yh). rpL8, control. (B) PCRs were performed on adult antennal
cDNA (cDNA) and genomic DNA (gDNA), revealing different-sized
amplification products, as a control to exclude any gDNA contamination in cDNA samples. PCR products were analysed on agarose
gels and visualized by UV illumination after ethidium bromide staining. The positions of marker bands (bp) are indicated.

pupae antennae, collected 5 days before emergence.
This pattern of expression is similar to that of previously characterized olfactory genes in Lepidoptera
antennae [19,32,33] and coincides with the maturation
of the olfactory system.
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Third, in situ hybridization was performed to investigate more deeply the expression pattern of SlitOR18 in
the adult male antennae. In S. littoralis, antennae are
filiform and segmented. The dorsal side is covered with
scales, whereas the ventral side carries different morphological ⁄ functional types of sensilla, including the sensilla
chaetica, the sensilla styloconica and the sensilla trichodea (Fig. 5A). Two types of sensilla trichodea have been
described: long and short, the latter being enriched in

the middle of the ventral side. These sensilla are devoted
to olfaction, the long ones being mainly tuned to pheromone components and the short ones responding to
both pheromone components and other chemicals [34].
A SlitOR18 sense strand probe gave no signal (Fig. 5B).
Antisense probe hybridizations were clearly restricted to
the sensilla side of the antenna, with no signal on the
scale side (Fig. 5C). Labelled spots were clearly
restricted to the bases of the olfactory sensilla of the
trichodea type (Fig. 5C,D). No staining could be
observed at the base of either the sensilla chaetica
(Fig. 5E), known to be involved in mechano ⁄ contact
chemoreception [35], or the sensilla styloconica
(Fig. 5F), known to be involved in taste, suggesting that
the expression of SlitOR18 should be confined to the
olfactory sensilla. As long and short sensilla trichodea
are often intermingled in this species, their distinction
was difficult in optical sections. In some sections, entire
long sensilla were visible, allowing us to clearly assign
the expression of SlitOR18 to long sensilla trichodea
(Fig. 5C,D). In sections through the middle of the
ventral surface (as in Fig. 5F), the abundance and distribution of the labelled spots suggest SlitOR18 expression
in the short sensilla trichodea as well. Thus, SlitOR18
seems to be expressed in different functional types of
olfactory sensilla, including pheromone-sensitive and
non-pheromone-sensitive. Taken together, our data
argue for a role of SlitOR18 in olfactory processes.
Possible function for OR18
The relationship between OR sequences and functions
is the focus of intense research, and functional orthologues of ORs have been established only for the
OR83b and PR subtypes. Our data suggest that OR18

could play a critical role in olfaction within noctuids.
To generate appropriate adaptive behaviours, insects
need to sample salient features of the broad chemical
environment. Thus, the simplest hypothesis would be
that OR18, expressed almost equally in the antennae
of both sexes, would respond to odorants particularly
relevant to noctuid ecology. Although OR18 sequences
did not cluster within the PR clade (Fig. 2A), the
SlitOR18 expression pattern in male antennae suggests

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I. Brigaud et al.

v

d
or

An atypical olfactory receptor subtype in noctuids

sst

lst

A

B

lst

C

lst

sch
ssty

trachea

Cuticule

sc

sc
D

sc
E

lst

F

st
ssty

sch

Fig. 5. Expression pattern of SlitOR18 in adult male antennae by in situ hybridization. (A) Scheme of a longitudinal section of two antennal
segments showing the distribution of scales (sc) on the dorsal side (d), and long and short sensilla trichodea (lst, sst), sensilla chaetica (sch)
and sensilla styloconica (ssty) on the ventral side (v). or, ornamentations. (B–F) Longitudinal sections of hybridized adult male antennae. (B)
Sense probe control. (C) Consecutive antennal segments with antisense probe staining restricted to the ventral side carrying olfactory sensilla, with no labelling on the dorsal scaled side. (D) Detail of long sensilla trichodea showing intense labelling at the base (black arrows).
Sensilla chaetica (E) and styloconica (F) are unstained (white arrows). Scale bars, 200 lm (A–C); 10 lm (D–F).

expression in pheromone-sensitive sensilla. Thus, their
function as pheromone receptors could not be
excluded. Alternatively, OR18 may be expressed in an
ORN co-compartmentalized with a PR-expressing
ORN within the same sensillum, but unresponsive to
any pheromone-related odorants. Indeed, such a situation has been described recently in H. virescens male
antennae [36], giving new insights into the complex sex
pheromone detection system of moths [37]. Interestingly, OR18s share common features with the nonconventional OR83b orthologues: they are highly
conserved; they are expressed at the bases of olfactory
sensilla with different functional properties; and their
protein structure exhibits a particularly long loop
between transmembrane segments IV and V (Fig. 1B),
a feature not observed in conventional insect ORs.

However, OR18 may not fulfil a general function in
insect olfaction like OR83b, as the OR18 subtype is
only found in noctuids. In DmelOR83b, the long loop
between transmembrane domains IV and V is intracytoplasmic (IC2) as a result of inverse topology of the
receptor compared with classical GPCRs (Fig. 1B),
and it has been proposed to link OR83b to the intracellular transport machinery [24]. The OR18 fourth
loop presents no homology with DmelOR83b IC2, and
a PROSITE pattern search resulted in no hit with any
known protein motif. In addition, although the membrane topology of OR18 proteins has not been investigated experimentally to date, Phobius predicted an
extracellular localization for this loop in all the OR18

sequences, defining it as the second extracellular loop
(EC2, Fig. 1B), which is incompatible with a function

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in intracellular trafficking. In mammals, the EC2 loop
of certain types of GPCR may be critical for ligand
binding and affinity [38]. Thus, if the classical GPCR
topology of OR18 were confirmed, OR18 EC2 might
also play an important role in ligand binding. Further
functional analyses will help to answer these questions
and to define the exact role of the OR18 subtype.
The discovery of an original subtype of extremely
conserved ORs opens up new routes for the understanding of insect OR evolution. In particular, further
insect genome sequencing, some already under way,
may confirm that the OR18 subtype is restricted to
noctuids and may reveal other new conserved subtypes. From an applied point of view, the OR18 subtype offers a single target for the design of molecules
to interfere with the olfactory process in widespread
species – at least among the noctuids – but preserving
beneficial insects that lack such receptors.

Materials and methods
Insect rearing and tissue collection

S. littoralis, M. brassicae, H. armigera and S. nonagrioides
were reared on a semi-artificial diet in the laboratory at
20 °C and 60–70% relative humidity, under a 16 h : 8 h
light : dark cycle until emergence. A. segetum, B. mori and
O. nubilalis pupae were generously provided by C. Lofsted
ă
(Lund University, Lund, Sweden), C. Royer (UNS, Lyon,
France) and F. Marion-Poll [Physiologie de l’Insecte: Signalisation et Communication (PISC), Paris, France], respectively. Antennae from H. zea were generously provided by T.
Baker (Pennsylvania State University, University Park, PA,
USA). Tissues from S. littoralis whole embryos, L5 larvae
(heads), different pupal stages (male antennae) and adults
(male and female antennae and male brains, proboscis, abdomen, legs) were dissected and used for cDNA synthesis. For
in situ hybridization, S. littoralis male antennae were fixed
overnight in 4% paraformaldehyde at 4 °C, dehydrated in
methanol and stored at )20 °C until use.

RNA extraction and cDNA synthesis
For RT-PCR, total RNAs were extracted from the different
tissues with TRIzolÒ reagent (Invitrogen, Carlsbad, CA,
USA). After a DNase1 step treatment (Promega, Madison,
WI, USA), single-stranded cDNAs were synthesized from
1 lg of total RNAs with 200 U of M-MLV reverse transcriptase (Clontech, Mountain View, CA, USA) using buffer
and protocol supplied in the Advantag RT-for-PCR kit
(Clontech).
For 5¢ and 3¢ rapid amplification of cDNA ends (RACE)
PCR, cDNAs were synthesized from 1 lg of male antennal
RNA at 42 °C for 1.5 h, with SuperScriptÔ II reverse

6544

transcriptase (200 U, Gibco BRL, Invitrogen), using the
3¢-CDS primer (for 3¢ RACE) or the 5¢-CDS primer and
the SMARTƠ II oligonucleotide (for 5¢ RACE), supplied
in the SMARTƠ RACE cDNA amplification kit (Clontech), following the manufacturer’s instructions.
For quantitative real-time PCR (qPCR), RNAs were
extracted from S. littoralis male and female antennae (10 of
each sex), brains (5), proboscis (5), abdomens (2) and legs
(6) with the RNeasyÒ MicroKit (Qiagen, Hilden, Germany),
which included a DNase treatment. Single-stranded cDNAs
were synthesized from 1 lg of total RNAs as above.

Molecular cloning of H. virescens OR18
homologues
Two degenerate primers were designed from the H. virescens
OR18 amino acid sequence (accession number: AJ748333
[11]): OR18F (5¢-GTGCTYTRTTTCCTATTTATGCTGG-3¢)
and OR18R (5¢-GTAATCAAAGTGAAGAARGARTAAG
AAG-3¢). They were used for PCR amplifications of A. segetum, H. zea, H. armigera, M. brassicae, S. nonagrioides,
S. littoralis, B. mori and O. nubilalis antennal cDNA
templates with PCR Mastermix (Promega) through 40 cycles
at 94 °C for 30 s, 50 °C for 30 s and 72 °C for 1 min. A
single  740 bp fragment was amplified for all species, except
for B. mori and O. nubilalis (no amplification). After gel
purification (GenEluteÔ; Sigma-Aldrich, St Louis, MO,
USA), fragments were cloned into the pCRÒII-TOPOÒ
plasmid (Invitrogen). Recombinant plasmids were isolated
by mini preparation (QIAprep Spin Miniprep Kit, Qiagen),
and both strands were sequenced (Biofidal, Vaulx-en-Velin,
France). The 3¢ and 5¢ regions of the cDNAs were amplified
by 3¢ and 5¢ RACE, using the AdvantageÔ 2 polymerase mix

(Clontech) and the Universal Primer Mix versus the following gene-specific primers: 5¢RACE primer (used for all species),
5¢-GTGCTYTRTTTCCTATTTATGCTGG-3¢; 3¢RACE primers:
Aseg3¢Race, 5¢-CTGGCATGGGGCTAGTCGTCTTCGAC
ATGG-3¢; Mbra3¢Race, 5¢-CCGGGATGGGGCTCATCG
TCTTCAATATGG-3¢; Snon3¢Race, 5¢-CCGGGATGGAT
CTTGTCGTCTTTGACATGG-3¢; Harm3¢Race, 5¢-CCGG
TATGGGGCTTGTGGTCTTCAACATGG-3¢; Hzea3¢Race,
5¢-CCGGCATGGGGCTTGTGGTCTTCAACATGG-3¢;
Slit3¢Race, 5¢-CTGGGATGGGCATAGTGGTGTTTAAT
ATGG-3¢. Touchdown PCRs were performed as follows:
1 min at 94 °C, five cycles of 30 s at 94 °C and 3 min at
72 °C, then five cycles of 30 s at 94 °C, 30 s at 70 °C and
3 min at 72 °C, then 30 cycles of 30 s at 94 °C, 30 s at 68 °C
and 3 min at 72 °C, followed by a final elongation step of
10 min at 72 °C. The PCR products were cloned, sequenced
on both strands and analysed as described above. By merging
overlapping 3¢, 5¢ and internal fragment sequences, six
full-length cDNAs encoding putative open reading frames
were generated and named SlitOR18 (S. littoralis), MbraOR18 (M. brassicae), HzeaOR18 (H. zea), HarmOR18
(H. armigera), SnonOR18 (S. nonagrioides) and AsegOR18

FEBS Journal 276 (2009) 6537–6547 ª 2009 The Authors Journal compilation ª 2009 FEBS


I. Brigaud et al.

(A. segetum). Nucleotide sequence data are available in the
GenBank database under the accession numbers: SlitOR18,
EU979124; MbraOR18, EU979123; HzeaOR18, EU979121;
HarmOR18, EU979122; SnonOR18, EU979119; AsegOR18,

EU979120.

Sequence analyses and phylogenetic inference
Gene sequence analyses and database comparisons were
performed using the blast program [39]. OR18 homologue
sequences were searched in GenBank and in insect available
genomes via the blastp program on protein databases
(Anobase: NCBI Map Viewer at .
gov/mapview/map_search.cgi?taxid=7165, RefSeq protein database; Beebase: />blast/blast.html, PreRelease2_protein database; Beetlebase:
protein sequences downloaded from .
edu/pub/BeetleBase/3.0/; Butterflybase: terfly
base.ice.mpg.de/, All species, Protein database; Flybase:
http://flybase.bio.indiana.edu/, Annotated Protein database;
Silkbase: Silkworm
Annotated Protein database). Alignment was performed
using clustalW2 [40]. Transmembrane topology was predicted with the phobius tool [41], and protein motifs were
searched against all patterns stored in the PROSITE pattern
database. For the phylogenetic analysis, the seven noctuid
OR18 amino acid sequences were included in a dataset containing full-length lepidopteran OR sequences, together with
OR sequences identified from genomes of D. melanogaster,
A. mellifera and T. castaneum. Owing to the large number of
putative OR sequences, identified from this last species, only
the sequences of ORs expressed in adult tissues were included
[15]. After alignment and removal of nonconserved residues,
the dataset contained 418 taxa and 378 characters. An unrooted tree was inferred from this dataset by the neighbourjoining method, with distance correction based on a Dayhoff
PAM matrix, as implemented in mega4 software [42]. Node
support was assessed by a bootstrap procedure based on
1000 replicates.
For synonymous (dS) and nonsynonymous (dN ⁄ dS)
substitution calculations, nucleotide sequences were aligned

with their corresponding amino acid sequences using Tranalign (EMBOSS). dN ⁄ dS values were calculated using the
Nei–Gojobori model [43] with Jukes–Cantor correction, as
implemented in mega4 [42]. Evolutionary selection was
assessed using the Z-test (mega4) with the alternative
hypothesis of purifying selection (dN < dS).

Developmental studies in S. littoralis
The temporal expression pattern of SlitOR18 was analysed
by RT-PCR carried out on cDNAs from whole embryos,
larvae heads, pupae antennae and adult antennae, using the
degenerated primer pair and the PCR conditions mentioned
above, generating a 750 bp band. The ribosomal protein L8

An atypical olfactory receptor subtype in noctuids

gene (rpL8) was used as an RNA extraction control, as
described previously [44], generating a 580 bp fragment. In
parallel, PCR was conducted with the same primer pair on
genomic DNA, as a control to exclude contamination of
the RNA preparations with genomic DNA. This PCR led
to the amplification of a 1500 bp band (Fig. 4B). The
750 bp bands obtained in the different cDNA samples then
specifically reflected transcript amplification. Amplification
products were loaded on 1.5% agarose gels and visualized
by ethidium bromide staining.

qPCR analyses in S. littoralis tissues
Gene-specific primers for SlitOR18 (SlitOR18F: 5¢-GCTG
GGACCTTGATGAGTATTG-3¢; SlitOR18R: 5¢-CACGC
ATTGGACGCAGTTATAG-3¢) and the endogenous control rpL8 (rpL8F: 5¢-ATGCCTGTGGGTGCTATGC-3¢;

rpL8R:
5¢-TGCCTCTGTTGCTTGATGGTA-3¢)
were
designed using Beacon Designer 4.0 software (Bio-Rad,
Hercules, CA, USA), yielding PCR products of 150 and
210 bp, respectively. qPCR mix was prepared in a total
volume of 20 lL with 10 lL of Absolute QPCR SYBR
Green Mix (ABgene, Epsom, UK), 5 lL of diluted cDNA
(or water for the negative control, or RNA for controlling
for the absence of genomic DNA) and 200 nm of each
primer. qPCRs were performed on S. littoralis cDNAs prepared from male antennae, female antennae, male brains,
proboscis, abdomen and legs using an MJ Opticon Monitor
Detection System (Bio-Rad). The PCR programme began
with a cycle at 95 °C for 15 min, followed by 40 cycles of
20 s at 95 °C, 15 s at 55 °C and 20 s at 72 °C. Melting
curves were built every 0.2 °C from 50 to 95 °C with a 2 s
hold, and allowed an assessment of the purity of the PCRs.
Standard curves were generated by a five-fold dilution
series of a cDNA pool evaluating primer efficiency
E [E = 10()1 ⁄ slope)]. All experiments included a no-template
control, two replicates of biological samples and dilution
points. SlitOR18 expression levels were calculated relative
to the expression of the rpL8 control gene and expressed as
the ratio ESlitOR18(DCT)SlitOR18 ⁄ ErpL8(DCT)rpL8 [45].

SlitOR18 expression pattern in antennae
Digoxygenin-labelled RNA sense and antisense probes
(427 bp long) were reverse transcribed in vitro from PCR
fragments amplified from the recombinant plasmid SlitOR18-pCRÒII-TOPOÒ with M13 Forward and M13
Reverse primers, using T7 and SP6 RNA polymerases (Promega) and following the recommended protocol. SlitOR18

RNA probes were then purified on RNA G50 Sephadex
columns (Quick Spin columns; Roche Applied Science,
Indianapolis, IN, USA). The hybridization protocol was
performed on whole-mount pieces of antennae, as described
previously [46]. After hybridization and embedding, longitudinal sections (6 lm) were prepared and counter-stained

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6545


An atypical olfactory receptor subtype in noctuids

I. Brigaud et al.

with acridine orange and photographed. Pictures were digitized and processed using AdobeÒ PhotoshopÒ 7.0 (Adobe
Systems Inc., San Jose, CA, USA).

Acknowledgements
We thank Fabien Tissier (UMR PISC, INRA
Versailles, France) for help with insect rearing, Hadi
Quesneville (URGI, INRA Versailles, France) for help
with dN ⁄ dS calculations, David Tepfer (Pessac, INRA
Versailles, France) for English improvement, and
Christer Lofsted (Lund University, Lund, Sweden),
ă
Corinne Royer (UNS, Lyon, France), Frederic MarionPoll (PISC, Paris, France) and Tom Baker (Pennsylvania State University, University Park, PA, USA) for
providing insects or antennae from the different species
used in this work. This work was supported by INRA,
´

Universite Paris VI, ACI JC5249 funding, as well as an
ACI JC doctoral fellowship to Isabelle Brigaud.

References
1 Buck L & Axel R (1991) A novel multigene family may
encode odorant receptors: a molecular basis for odor
recognition. Cell 65, 175–187.
2 Clyne PJ, Warr CG, Freeman MR, Lessing D, Kim J &
Carlson JR (1999) A novel family of divergent seventransmembrane proteins: candidate odorant receptors
in Drosophila. Neuron 22, 327–338.
3 Gao Q & Chess A (1999) Identification of candidate
Drosophila olfactory receptors from genomic DNA
sequence. Genomics 60, 31–39.
4 Vosshall LB, Amrein H, Morozov PS, Rzhetsky A &
Axel R (1999) A spatial map of olfactory receptor
expression in the Drosophila antenna. Cell 96, 725–736.
5 Robertson HM, Warr CG & Carlson JR (2003)
Molecular evolution of the insect chemoreceptor gene
superfamily in Drosophila melanogaster. Proc Natl Acad
Sci USA 100(Suppl 2), 14537–14542.
6 Guo S & Kim J (2007) Molecular evolution of Drosophila odorant receptor genes. Mol Biol Evol 24, 1198–1207.
7 Hill CA, Fox AN, Pitts RJ, Kent LB, Tan PL, Chrystal
MA, Cravchik A, Collins FH, Robertson HM &
Zwiebel LJ (2002) G protein-coupled receptors in
Anopheles gambiae. Science 298, 176–178.
8 Bohbot J, Pitts RJ, Kwon HW, Rutzler M, Robertson
HM & Zwiebel LJ (2007) Molecular characterization of
the Aedes aegypti odorant receptor gene family. Insect
Mol Biol 16, 525–537.
9 Sakurai T, Nakagawa T, Mitsuno H, Mori H, Endo Y,

Tanoue S, Yasukochi Y, Touhara K & Nishioka T
(2004) Identification and functional characterization of
a sex pheromone receptor in the silkmoth Bombyx mori.
Proc Natl Acad Sci USA 101, 16653–16658.

6546

10 Wanner KW, Anderson AR, Trowell SC, Theilmann
DA, Robertson HM & Newcomb RD (2007) Femalebiased expression of odourant receptor genes in the
adult antennae of the silkworm, Bombyx mori. Insect
Mol Biol 16, 107–119.
11 Krieger J, Grosse-Wilde E, Gohl T, Dewer YM,
Raming K & Breer H (2004) Genes encoding candidate
pheromone receptors in a moth (Heliothis virescens).
Proc Natl Acad Sci USA 101, 11845–11850.
12 Krieger J, Raming K, Dewer YM, Bette S, Conzelmann
S & Breer H (2002) A divergent gene family encoding
candidate olfactory receptors of the moth Heliothis virescens. Eur J Neurosci 16, 619–628.
13 Robertson HM & Wanner KW (2006) The chemoreceptor superfamily in the honey bee, Apis mellifera: expansion of the odorant, but not gustatory, receptor family.
Genome Res 16, 1395–1403.
14 Abdel-Latief M (2007) A family of chemoreceptors in
Tribolium castaneum (Tenebrionidae: Coleoptera). PLoS
ONE 2, e1319.
15 Engsontia P, Sanderson AP, Cobb M, Walden KK,
Robertson HM & Brown S (2008) The red flour beetle’s
large nose: an expanded odorant receptor gene family
in Tribolium castaneum. Insect Biochem Mol Biol 38,
387–397.
16 Rutzler M & Zwiebel L (2005) Molecular biology of
insect olfaction: recent progress and conceptual models.

J Comp Physiol A Neuroethol Sens Neural Behav Physiol
191, 777–790.
17 Krieger J, Klink O, Mohl C, Raming K & Breer H
(2003) A candidate olfactory receptor subtype highly
conserved across different insect orders. J Comp Physiol
A 189, 519–526.
18 Xia Y & Zwiebel LJ (2006) Identification and characterization of an odorant receptor from the West Nile virus
mosquito, Culex quinquefasciatus. Insect Biochem Mol
Biol 36, 169–176.
19 Malpel S, Merlin C, Francois MC & Jacquin-Joly E
(2008) Molecular identification and characterization of
two new Lepidoptera chemoreceptors belonging to the
Drosophila melanogaster OR83b family. Insect Mol Biol
17, 587–596.
20 Lu B, Wang N, Xiao J, Xu Y, Murphy RW & Huang
D (2009) Expression and evolutionary divergence of the
non-conventional olfactory receptor in four species of
fig wasp associated with one species of fig. BMC Evol
Biol 9, 43.
21 Melo AC, Rutzler M, Pitts RJ & Zwiebel LJ (2004)
Identification of a chemosensory receptor from the
yellow fever mosquito, Aedes aegypti, that is highly
conserved and expressed in olfactory and gustatory
organs. Chem Senses 29, 403–410.
22 Pitts RJ, Fox AN & Zwiebel LJ (2004) A highly
conserved candidate chemoreceptor expressed in both
olfactory and gustatory tissues in the malaria vector

FEBS Journal 276 (2009) 6537–6547 ª 2009 The Authors Journal compilation ª 2009 FEBS

I. Brigaud et al.

23

24

25

26

27

28

29

30

31

32

33

34

Anopheles gambiae. Proc Natl Acad Sci USA 101, 5058–
5063.
Larsson MC, Domingos AI, Jones WD, Chiappe ME,

Amrein H & Vosshall LB (2004) Or83b encodes a
broadly expressed odorant receptor essential for
Drosophila olfaction. Neuron 43, 703–714.
Benton R, Sachse S, Michnick SW & Vosshall LB
(2006) Atypical membrane topology and heteromeric
function of Drosophila odorant receptors in vivo. PLoS
Biol 4, e20.
Jones WD, Nguyen TA, Kloss B, Lee KJ & Vosshall
LB (2005) Functional conservation of an insect odorant
receptor gene across 250 million years of evolution.
Curr Biol 15, R119–R121.
Nakagawa T, Sakurai T, Nishioka T & Touhara K
(2005) Insect sex-pheromone signals mediated by
specific combinations of olfactory receptors. Science
307, 1638–1642.
Krieger J, Grosse-Wilde E, Gohl T & Breer H (2005)
Candidate pheromone receptors of the silkmoth
Bombyx mori. Eur J Neurosci 21, 2167–2176.
Patch HM, Velarde RA, Walden KK & Robertson HM
(2009) A candidate pheromone receptor and two odorant receptors of the hawkmoth Manduca sexta. Chem
Senses 34, 305–316.
Mitsuno H, Sakurai T, Murai M, Yasuda T, Kugimiya
S, Ozawa R, Toyohara H, Takabayashi J, Miyoshi H &
Nishioka T (2008) Identification of receptors of main
sex-pheromone components of three Lepidopteran
species. Eur J Neurosci 28, 893–902.
Grosse-Wilde E, Gohl T, Bouche E, Breer H & Krieger
J (2007) Candidate pheromone receptors provide the
basis for the response of distinct antennal neurons to
pheromonal compounds. Eur J Neurosci 25, 2364–2373.

Sanchez-Gracia A, Vieira FG & Rozas J (2009) Molecular evolution of the major chemosensory gene families
in insects. Heredity 103, 208–216.
Gyorgyi TK, Roby-Shemkovitz AJ & Lerner MR (1988)
Characterization and cDNA cloning of the pheromonebinding protein from the tobacco hornworm, Manduca sexta: a tissue-specific developmentally regulated
protein. Proc Natl Acad Sci USA 85, 9851–9855.
Rogers ME, Sun M, Lerner MR & Vogt RG (1997)
Snmp-1, a novel membrane protein of olfactory neurons
of the silk moth Antheraea polyphemus with homology
to the CD36 family of membrane proteins. J Biol Chem
272, 14792–14799.
Ljungberg H, Anderson P & Hansson BS (1993)
Physiology and morphology of pheromone-specific
sensilla on the antennae of male and female Spodoptera

An atypical olfactory receptor subtype in noctuids

35

36

37

38

39

40

41

42

43

44

45

46

littoralis (Lepidoptera: Noctuidae). J Insect Physiol 39,
253–260.
Keil TA (1999) Morphology and development of the
peripheral olfactory organs. In Insect Olfaction
(Hansson BS ed.), pp. 5–47. Springer, Berlin.
Krieger J, Gondesen I, Forstner M, Gohl T, Dewer Y
& Breer H (2009) HR11 and HR13 receptor-expressing
neurons are housed together in pheromone-responsive
sensilla trichodea of male Heliothis virescens. Chem
Senses 34, 469–477.
Baker T (2009) Nearest neural neighbors: moth sex
pheromone receptors HR11 and HR13. Chem Senses
34, 465–468.
Chao TH, Ember JA, Wang M, Bayon Y, Hugli TE &
Ye RD (1999) Role of the second extracellular loop of
human C3a receptor in agonist binding and receptor
function. J Biol Chem 274, 9721–9728.
Altschul SF, Gish W, Miller W, Myers EW & Lipman
DJ (1990) Basic local alignment search tool. J Mol Biol
215, 403–410.

Larkin MA, Blackshields G, Brown NP, Chenna R,
McGettigan PA, McWilliam H, Valentin F, Wallace IM,
Wilm A, Lopez R et al. (2007) Clustal W and Clustal X
version 2.0. Bioinformatics 23, 2947–2948.
Kall L, Krogh A & Sonnhammer EL (2004) A
combined transmembrane topology and signal
peptide prediction method. J Mol Biol 338, 1027–
1036.
Tamura K, Dudley J, Nei M & Kumar S (2007)
MEGA4: Molecular Evolutionary Genetics Analysis
(MEGA) software version 4.0. Mol Biol Evol 24, 1596–
1599.
Nei M & Gojobori T (1986) Simple methods for
estimating the numbers of synonymous and nonsynonymous nucleotide substitutions. Mol Biol Evol 3, 418
426.

Maă beche-Coisne M, Merlin C, Francois M-C,
Queguiner I, Porcheron P & Jacquin-Joly E (2004)
Putative odorant-degrading esterase cDNA from the
moth Mamestra brassicae: cloning and expression
patterns in male and female antennae. Chem Senses 29,
381–390.
Pfaffl MW (2001) A new mathematical model for
relative quantification in real-time RT-PCR. Nucleic
Acids Res 29, e45.
Jacquin-Joly E, Bohbot J, Francois MC, Cain AH &
Nagnan-Le Meillour P (2000) Characterization of the
general odorant-binding protein 2 in the molecular
coding of odorants in Mamestra brassicae. Eur J
Biochem 267, 6708–6714.

FEBS Journal 276 (2009) 6537–6547 ª 2009 The Authors Journal compilation ª 2009 FEBS

6547