Cannabinoid receptors in invertebrates

Abstract

Two cannabinoid receptors, CB1 and CB2, are expressed in mammals, birds, reptiles, and fish. The presence of cannabinoid receptors in invertebrates has been controversial, due to conflicting evidence. We conducted a systematic review of the literature, using expanded search parameters. Evidence presented in the literature varied in validity, ranging from crude in vivo behavioural assays to robust in silico ortholog discovery. No research existed for several clades of invertebrates; we therefore tested for cannabinoid receptors in seven representative species, using tritiated ligand binding assays with [3H]CP55,940 displaced by the CB1-selective antagonist SR141716A. Specific binding of [3H]CP55,940 was found in neural membranes of Ciona intestinalis (Deuterstoma, a positive control), Lumbricusterrestris (Lophotrochozoa), and three ecdysozoans: Peripatoides novae-zealandiae (Onychophora), Jasus edwardi (Crustacea) and Panagrellus redivivus (Nematoda); the potency of displacement by SR141716A was comparable to measurements on rat cerebellum. No specific binding was observed in Actinothoe albocincta (Cnidaria) or Tethya aurantium (Porifera). The phylogenetic distribution of cannabinoid receptors may address taxonomic questions; previous studies suggested that the loss of CB1 was a synapomorphy shared by ecdysozoans. Our discovery of cannabinoid receptors in some nematodes, onychophorans, and crustaceans does not contradict the Ecdysozoa hypothesis, but gives it no support. We hypothesize that cannabinoid receptors evolved in the last common ancestor of bilaterians, with secondary loss occurring in insects and other clades. Conflicting data regarding Cnidarians precludes hypotheses regarding the last common ancestor of eumetazoans. No cannabinoid receptors are expressed in sponges, which probably diverged before the origin of the eumetazoan ancestor.

Full text

Cannabinoid receptors in invertebrates
J. M. McPARTLAND,* J. AGRAVAL,D. GLEESON,àK. HEASMAN§ & M. GLASS
*GW Pharmaceuticals, Ltd., Salisbury, Wiltshire, UK
Department of Pharmacology, University of Auckland, Auckland, New Zealand
àLandcare Research NZ Ltd, Auckland, New Zealand
§Cawthron Institute, East Nelson, New Zealand
Introduction
Two cannabinoid receptors have been described to date,
CB1 and CB2. They are G-protein-coupled receptors
(GPCRs) named after their exogenous ligand, D
9
-tetra-
hydrocannabinol (THC) (Mechoulam et al., 1998). CB1 is
primarily expressed in the central nervous system,
whereas CB2 occurs in leukocytes and immune tissues
(Felder & Glass, 1998). Orthologs of cannabinoid
receptors are expressed in mammals, birds, reptiles,
amphibians, and fish (McPartland, 2004). Within the
vertebrate lineage, cannabinoid receptors track a ‘classic’
evolutionary pattern, with gene duplication in teleost
fish followed by paralog divergence (Yamaguchi et al.,
1996). The evolution of cannabinoid receptors in inver-
tebrates has been disputed, due in part to conflicting
in vivo,in vitro, and in silico evidence. The validity of this
evidence is hierarchical, as indicated by Elphick &
Egertova (2001), and delineated below:
Level 0: in vivo studies, behavioural changes evoked by
cannabis extracts;
Level I: in vivo studies, behavioural changes evoked by
specific cannabinoid ligands;
Correspondence: J. M. McPartland, 53 Washington Street Ext., Middlebury,
VT 05753, USA.
Tel.: +1 802 388 8303; fax: +1 802 399 8304; e-mail: mcpruitt@verizon.net,
jmcpartland@unitec.ac.nz
366 J. EVOL. BIOL. 19 (2006) 366–373ª2005 EUROPEAN SOCIETY FOR EVOLUTIONARY BIOLOGY
Keywords:
cannabinoid;
ecdysozoa;
endocannabinoid;
evolution;
G-protein coupled receptor;
invertebrates.
Abstract
Two cannabinoid receptors, CB1 and CB2, are expressed in mammals, birds,
reptiles, and fish. The presence of cannabinoid receptors in invertebrates has
been controversial, due to conflicting evidence. We conducted a systematic
review of the literature, using expanded search parameters. Evidence
presented in the literature varied in validity, ranging from crude in vivo
behavioural assays to robust in silico ortholog discovery. No research existed for
several clades of invertebrates; we therefore tested for cannabinoid receptors
in seven representative species, using tritiated ligand binding assays with
[
3
H]CP55,940 displaced by the CB1-selective antagonist SR141716A. Specific
binding of [
3
H]CP55,940 was found in neural membranes of Ciona intestinalis
(Deuterstoma, a positive control), Lumbricus terrestris (Lophotrochozoa), and
three ecdysozoans: Peripatoides novae-zealandiae (Onychophora), Jasus edwardi
(Crustacea) and Panagrellus redivivus (Nematoda); the potency of displacement
by SR141716A was comparable to measurements on rat cerebellum. No
specific binding was observed in Actinothoe albocincta (Cnidaria) or Tethya
aurantium (Porifera). The phylogenetic distribution of cannabinoid receptors
may address taxonomic questions; previous studies suggested that the loss of
CB1 was a synapomorphy shared by ecdysozoans. Our discovery of cannabi-
noid receptors in some nematodes, onychophorans, and crustaceans does not
contradict the Ecdysozoa hypothesis, but gives it no support. We hypothesize
that cannabinoid receptors evolved in the last common ancestor of bilaterians,
with secondary loss occurring in insects and other clades. Conflicting data
regarding Cnidarians precludes hypotheses regarding the last common
ancestor of eumetazoans. No cannabinoid receptors are expressed in sponges,
which probably diverged before the origin of the eumetazoan ancestor.
doi: 10.1111/j.1420-9101.2005.01028.x

Level II: in vitro studies, effects of cannabinoid ligands
upon signal transduction effectors, or immunohisto-
chemical studies using tagged antibodies raised against
CB1;
Level III: in vitro studies, tritiated ligand binding assays, or
PCR cloning techniques;
Level IV: in silico studies, ortholog identification in
whole-genome sequences.
In vivo studies provide the lowest levels of evidence.
For example, Parkinson (1640), described a Cannabis
extract exerting in vivo behavioural effects, ‘…poured
into the holes of earthwormes, will draw them forth, and
fishermen have used this feate to get wormes to baite
their hookes’. This evidence is nearly anecdotal, given
the polypharmaceutical content of crude Cannabis. The
behavioural effects elicited by Cannabis could have been
caused by an entourage of compounds acting at a variety
of targets. The next level of evidence (Level I) appraises
the behavioural effects of specific cannabinoid receptor
agonists and antagonists. For example, Buttarelli et al.
(2002) exposed the planaria worm Dugesia gonocephala to
WIN55212-2, which elicited dose-dependent changes in
motor behaviour, and this effect was attenuated by the
CB1 antagonist SR141716A.
In vitro assays offer more precise levels of evidence, such
as the Level-II study of THC’s effects at the neuromuscular
junction of the lobster Homarus americanus (Turkanis &
Karler, 1988). This study provided indirect proof of
cannabinoid receptors, because THC could have affected
the neuromuscular junction via other lobster protein
targets or via membrane disrupting effects. Immunohis-
tochemical assays that use tagged antibodies raised against
mammalian CB1 also represent Level-II evidence, suscep-
tible to Type-I error (false positives due to cross reactions
with other proteins). On the other hand, detection of
specific binding sites with [
3
H]CP55,940 and other tritiated
cannabinoids represents Level-III evidence, especially
when the potency of binding displacement is compared
to mammalian receptors (Elphick & Egertova, 2001).
Cloning of CB1 genes by reverse-transcription polymerase
chain reaction (RT-PCR) technology is Level III, possibly
prone to Type-I error: a cannabinoid receptor gene cloned
from the leech Hirudo medicinalis (Stefano et al., 1997b)
was contested as an artifact of PCR contamination
(Elphick, 1998). Similarly, a cannabinoid gene cloned
from the fruit fly Drosophila melanogaster (Abbott, 1990)
has been refuted by a tritiated ligand binding study
(McPartland et al., 2001) and in silico studies (Elphick &
Egertova, 2001, McPartland et al., 2001).
Whole genome in silico studies provide a rigorous level
of evidence, Level IV. Thanks to high-throughput
sequencing and advances in computational biology, the
entire genomes of many organisms have been sequenced
and deposited in internet-accessible databases. Using this
approach, no orthologs of CB1 or CB2 were found in the
genome of the fruit fly D. melanogaster (Elphick &
Egertova, 2001; McPartland et al., 2001) and the
nematode Caenorhabditis elegans (Elphick & Egertova,
2001; McPartland et al., 2001). Unfortunately the in silico
approach poorly assesses invertebrates, because only four
invertebrate genomes have been sequenced, namely sea
squirt, Ciona intestinalis (a deuterostome), and three
protostomes: fruit fly, D. melanogaster; mosquito, Anoph-
eles gambiae; and nematode, C. elegans. To wit, all three
protostomes are members of the Ecdysozoa clade, a
recently described clade of invertebrates (Aguinaldo
et al., 1997). No genomes have been sequenced from
the other major protostome clade (the Lophotrochozoa),
or from basal animals, such as cnidarians and sponges.
Evidence suggests cannabinoid receptors evolved in basal
animals, based on a Level-III study of the cnidarian Hydra
vulgaris (De Petrocellis et al., 1999). Yet the receptors
evidently lack expression in D. melanogaster and C. elegans,
as aforementioned. This led McPartland et al. (2001) to
hypothesize that cannabinoid receptors evolved in early
metazoans, at least 525 million years ago (MYA), but
were secondarily lost in the Ecdysozoa.
The purpose of this study was three-fold: (1) conduct a
systematic review of the literature concerning cannabi-
noid receptors in invertebrates; (2) utilize this literature
database to identify subgroups of invertebrates that have
not been studied; (3) search for cannabinoid receptors in
organisms representative of subgroups lacking data.
Rather than using RT-PCR technology, which has been
problematic when cloning cannabinoid receptors, we
utilized tritiated ligand binding assays, coupled with
comparisons to mammalian receptors. Tritiated ligand
binding results were then scrutinized within the context
of evidence assembled in the systematic review, weighed
with respect to levels of evidence.
Materials and methods
Systematic review
MEDLINE (1966-June 2005) and AGRICOLA (1990-June
2005) were searched using MeSH keywords alone or in
various Boolean combinations: invertebrate, cannabi-
noids, cannabinoid receptors, THC, cannabis, marijuana.
All reports were scanned for supporting citations; antece-
dent sources were retrieved, except for doctoral disserta-
tions. Unindexed conference proceedings and textbooks
were scanned by hand. Data validity was assessed by
source (peer-reviewed journal article vs. chapter in edited
book vs. conference proceeding abstract), experimental
methodology (segregated by levels of evidence, Table 1)
and the frequency of independent observations.
Tissue sampling
Based on the literature review, seven groups were
selected for experimentation: a deuterostome (positive
control), lophotrochozoan, three sub-clades within the
Ecdysozoa. a cnidarian, and a sponge. The UNITEC
Cannabinoid receptors in invertebrates 367
J. EVOL. BIOL. 19 (2006) 366–373 ª2005 EUROPEAN SOCIETY FOR EVOLUTIONARY BIOLOGY

institutional review board approved the study protocol
prior to the experiment. Seven species were assayed:
(1) Sea squirt, C. intestinalis (Chordata-Urochordata),
30 individuals collected from infralittoral mussel beds in
the Marlborough Sounds, and couriered to Auckland in
sea water on ice. Harvested tissues included neural
ganglia and neural gland, testis, intestine (which may
include peyer’s patch-like lymphoid tissues), and hemo-
lymph (cells that exhibit immune functions) mixed with
heart tissue. (2) Earthworm, Lumbricus terrestris (Lophot-
rochozoa-Annelida-Oligochaeta), about 100 individuals
were purchased from Biosuppliers, Ltd, commercial
insectary, Auckland. The cephalic end was opened at
the prostomium and cerebral ganglia removed from
around the anterior pharynx. (3) Velvet worm, Peripato-
ides novae-zealandiae (Ecdysozoa-Onychophora-Peripati-
dae), 25 individuals were collected from a protected
location near Auckland (under permit by Landcare
Research, NZ Ltd). The cephalic end was decapitated
and adhesive secretions from head glands removed. (4)
Rock lobster, Jasus edwardi (Ecdysozoa-Arthropoda-
Crustacea), one individual, sourced from Sea World,
Auckland. Brain tissue (subesophageal ganglion) was
dissected from the dorsal aspect of the head and thorax.
Identity of the ganglion was confirmed by tracing the
optic nerves caudally. (5) Beer mat nematode, Panagrellus
redivivus (Ecdysozoa-Nematoda-Rhabditida), several hun-
dred individuals were purchased from Biosuppliers, Ltd,
commercial insectary, Auckland. Whole organisms were
sonicated and then homogenized. (6) Common variable
Table 1 Summary of in vivo,in vitro,and
in silico evidence for the presence (+) or
absence ()) of cannabinoid receptors in
invertebrates (n.d. indicates no data).
Taxa Level IV Level III Level II Level I Level 0
DEUTEROSTOMA
Chordata-Vertebrates
Primates, rodents, etc. (+)
1
(+)
2
(+)
2
(+)
3
(+)
3
Chordata-Tunicates
Sea squirt Ciona intestinalis (+)
4
(+)
5,6
(+)
5
n.d. n.d.
Echinodermata
Sea urchin Strongylocentrus spp. n.d. (+)
7
(+)
8)14
n.d. n.d.
PROTOSTOMA
Lophotrochozoa
Earthworm Lumbricus terrestris n.d. (+)
6
n.d. n.d. (+)
15
Leech Hirudo medicinalis n.d. (+)
16,17
())
1,18
(+)
16,19)22
n.d. n.d.
Planaria Girardia tigina n.d. n.d. (+)
23,24
n.d. n.d.
Mussel Mytilus edulis n.d. (+)
16
(+)
19.21
Sea slug Aplysia californica n.d. (–)
25
(+)
26
n.d. n.d.
Ecdysozoa
Fruit fly Drosophila melanogaster (–)
1,27,28
(–)
27
(+)
29
n.d. n.d. n.d.
Mosquito Anopheles gambiae (–)
28
n.d. n.d. n.d. n.d.
Honeybee Apis melifera n.d. (–)
28
n.d. n.d. (–)
30
Ant Formica pratensis n.d. (–)
31
n.d. n.d. (–)
30
Locust Shistocerca gregaria n.d. (+)
32
n.d. n.d. n.d.
Nematode Caenorhabditis elegans (–)
1,33
n.d n.d. n.d. n.d.
Mat nematode Panagrellus redivivus n.d. (+)
6
n.d. n.d. n.d.
Lobster Homarus americanus n.d. (+)
34
n.d. n.d. n.d.
Rock lobster Jasus edwardi n.d. (+)
6
n.d. n.d. n.d.
Velvet worm P. novaen.d.zelandiae n.d. (+)
6
n.d. n.d. n.d.
CNIDARIA
Hydra Hydra vulgaris n.d. (+)
35
n.d. (+)
35
n.d.
Sea anomone Actinothoe albocincta n.d. (–)
6
n.d. n.d. n.d.
PORIFORA
Sponge Tethya aurantium n.d. (–)
6
n.d. n.d. n.d.
1
Elphick & Egertova (2001);
2
Felder & Glass (1998);
3
Chaperon & Theibot (1999);
4
Elphick
et al. (2003);
5
Matias et al. (2005);
6
current study;
7
Chang et al. (1993);
8
Schuel et al. (1987);
9
Chang et al. (1991);
10
Chang & Schuel (1991);
11
Schuel et al. (1991a);
12
Schuel et al. (1991b);
13
Schuel et al. (1994);
14
Berdyshev (1999);
15
Parkinson (1640);
16
Stefano et al. (1996);
17
Stefano et al. (1997b);
18
Elphick (1998);
19
Stefano et al. (1997a);
20
Salzet et al. (1997);
21
Stefano et al. (1998);
22
Matias et al. (2001);
23
Lenicque et al. (1972);
24
Buttarelli et al.
(2002);
25
Howlett et al. (1990);
26
Acosta-Urquidi & Chase (1975);
27
McPartland et al. (2001);
28
McPartland (2004);
29
Howlett et al. (2000);
30
McPartland et al. (2000);
31
Waser (1999);
32
Egertova et al. (1998);
33
McPartland & Glass (2001);
34
Turkanis & Karler (1988);
35
De
Petrocellis et al. (1999).
368 J. M. McPARTLAND ET AL.
J. EVOL. BIOL. 19 (2006) 366–373 ª2005 EUROPEAN SOCIETY FOR EVOLUTIONARY BIOLOGY

sea anomone, Actinothoe albocincta (Cnidaria, Anthozoa),
one individual was collected from the intertidal zone of a
rocky pool at Piha Beach. Tissues from around the oral
disc were dissected, a combination of peri-oral sphincter
muscles, tentacles, and actinopharynx tissues. (7) Golf
ball sponge, Tethya aurantium (Porifera-Epipolasida-
Tethyidae), six individual colonies were collected from
the infralittoral zone at Piha Beach (Auckland, New
Zealand). Individuals were dissected in half and squeezed
to express cellular material from spiculae.
Binding assays
Organisms were cold-anaesthetized by chilling for
30 min at 4 C, then moved to a dissection tray atop a
bed of dry ice (frozen CO
2
) and instantly frozen or
asphyxiated with sublimated CO
2
gas. Dissected tissue
were immediately homogenized in ice-cold lysis buffer
(10 m
MM
MOPS, 1 m
MM
EDTA, 10 l
MM
AEBSF pH 7.5), and
sequentially centrifuged to obtain a pellet of cell mem-
branes, as described previously (McPartland et al., 2001).
Membrane protein concentrations were determined via
standardized ‘Biorad D
c
’ assay following the manufac-
turers’ protocol. Ligand binding assays used established
methods (Kearn et al., 1999; McPartland et al., 2001),
utilizing [
3
H]CP55,940 (NEN Life Sciences), a synthetic
CB1 and CB2 receptor agonist. Nonspecific binding was
determined by displacement with SR141716A, a CB1
selective antagonist. Briefly, membrane samples (15–
40 lg) were incubated with 6.3 n
MM
[H
3
]CP55,940
(158.00 Ci mmol
)1
), in the presence or absence of
100 n
MM
SR141716A in TME assay buffer (50 m
MM
Tris,
2m
MM
MgCl2, 1 m
MM
EDTA, 5 mg mL
)1
BSA for 90 min at
37 C. The incubation was terminated by addition of
200 lL ice cold TME buffer, and samples were filtered
though a printed filtermat A (GF-C) filter (Perkin Elmer)
presoaked in 0.1% polyetheleneimine, and washed twice
with 300 lL TME buffer on an Inotech cell harvester. The
filter was then dried prior to the addition of Multilex
(melt-on scintillant; Perkin Elmer) and was counted for
5 min in a Wallac Microbeta Trilux (Perkin Elmer).
Experiments were performed twice in triplicate. Positive
binding results were then extended by displacement
analysis for three representative organisms, L. terristus,
P. redivivus, and P. novae-zealandiae. Membranes were
incubated with 6.3 n
MM
[
3
H]CP55,940 in the presence of
increasing concentrations of SR141716A (10
)6
to
10
)11
MM
). Samples were filtered and counted as above.
Displacement curves were generated by nonlinear regres-
sion utilizing Graph Pad Prism, Version 4.0.
Results
Systematic review
Publications regarding cannabinoid receptors and inver-
tebrates were located in peer-reviewed journals (n¼25),
chapters in edited books (n¼3), conference proceedings
abstracts (n¼2), and older, nonpeer reviewed sources
(n¼2). The total (n¼32) did not include conference
abstracts that were subsequently republished as peer-
reviewed articles, nor did the tally include dissertations.
In terms of levels of evidence, the literature distributed in
a skewed histogram: Level 0, n¼3; Level I, n¼6; Level
II, n¼14; Level III, n¼13; Level IV, n¼6. Note that
several publications reported several experiments, at
several levels of evidence. A summary of these publica-
tions is presented in Table 1, arrayed phylogenetically.
Binding assays
Specific, displaceable binding of [
3
H]CP55,940 was
observed in all deuterostomes, lophotrochozoans, and
ecdysozoans; no specific binding was observed in cnidar-
ian and poriferan species (Table 2). Nondisplaceable
binding levels were comparable between all tissues.
Percentage specific binding was highest in C. intestinalis
neural tissue and haemolymph, followed by tissues from
P. redivivus, L. terrestris, P. novae-zealandiae, and J. edwardi
(Table 2). Low receptor binding was observed in C. intes-
tinalis testis and intestine tissues.
In order to determine the binding characteristics of
these putative receptors, full competition binding assays
were performed for L. terristus,P. novae-zealandiae, and
P. redivivus (Fig. 1), Log IC50 values obtained from
displacement curves were )11.20 ± 0.12
MM
,)10.93 ±
0.28
MM
, and )11.21 ± 0.2
MM
, respectively. For the pur-
poses of comparison, binding was performed under
identical conditions on rat cerebellum, which has estab-
lished high expression of CB1 receptors, and produced a
Log IC50 value of )10.17 ± 0.12
MM
(Fig. 1).
Table 2 Concentrations of [
3
H]CP55,940 specific binding sites
observed in membrane preparations from seven invertebrates and
rat cerebellum.
Species, tissue fmol/mg Specific binding (%)
Tethya aurantium ND )2±8
Actinothoe albocincta ND )1±3
Peripatoides novae-zealandiae 90 ± 11 52 ± 3
Panagrellus redivivus 55±5 58±1
Jasus edwardi 104±3 41±2
Lumbricus terristus 78±5 56±2
Ciona intestinalis–neural tissue 134 ± 3 76 ± 10
Ciona intestinalis–intestine ND 11 ± 11
Ciona intestinalis–haemolymph 35 ± 2 60 ± 2
Ciona intestinalis–testis 30 ± 6 45 ± 14
Rat cerebellum 648 ± 16 48 ± 1
Results are presented as the mean ± SEM, for two experiments
performed in triplicate. The % specific binding represents the
proportion of the total binding observed that could be displaced by
100 n
MM
SR141716A. Specific binding not significantly different to
zero was classed as no receptors being detected (ND).
Cannabinoid receptors in invertebrates 369
J. EVOL. BIOL. 19 (2006) 366–373 ª2005 EUROPEAN SOCIETY FOR EVOLUTIONARY BIOLOGY

Discussion
Tritiated ligand binding assays detected specific cannabi-
noid binding in all organisms tested except sea anemone
(A. albocincta) and sponge (T. aurantium). The receptors
were identified by binding of the nonspecific cannabi-
noid agonist [
3
H]CP55,940 displaced by the highly CB1
selective antagonist SR141716A, and therefore are con-
sistent with cannabinoid CB1, but not CB2 receptors. In
three of the organisms tested, earthworm (L. terristus),
velvet worm (P. novae-zealandiae), and mat nematode
(P. redivivus), sufficient tissue was available to carry out
competition binding assays, in comparison to a well
characterized CB1 ortholog in rat cerebellar tissue. In all
three organisms, a high-affinity binding interaction was
observed for SR141716A at various concentrations,
consistent with rat cerebellar tissue and characteristic of
CB1 receptors (Fig. 1).
The phylogenetic distribution of cannabinoid receptors
may address larger taxonomic questions. Invertebrate
taxonomy has recently been updated by the Ecdysozoa
hypothesis, which classifies protostome invertebrates by
their ability or inability to molt (Aguinaldo et al., 1997).
Under the Ecdysozoa arrangement, animals are grouped
into five clades: the Deuterostomes (vertebrates as well as
some invertebrates such as sea squirts and sea urchins),
Lophotrochozoans (protostomes that do not molt: anne-
lids, mollusks, platyhelminths), Ecdysozoans (protos-
tomes that molt: insects, crustaceans, nematodes,
onychophorans), Cnidarians (hydras, sea anemones,
and other nonbilaterian animals), and Poriferans (spon-
ges, primitive multicellular animals whose cells do not
form tissues or organs). Prior in vitro and in silico studies
indicated a lack of cannabinoid receptors in insects and
nematodes, which led McPartland et al. (2001) to propose
that cannabinoid receptors were secondarily lost in the
Ecdysozoa. However, our new binding studies suggest
that cannabinoid receptors are present in three species
representing three classes of animals within the Ecdyso-
zoa, P. novae-zealandiae (Ecdysozoa-Onychophora),
J. edwardi (Ecdysozoa-Crustacea), and P. redivivus
(Ecdysozoa-Nematoda). The presence of cannabinoid
receptors in onychophorans, crustaceans, and nematodes
does not contradict the Ecdysozoa hypothesis, but gives it
no support.
We also conducted ligand-binding studies upon a
lophotrochozoan, cnidarian, and sponge. These clades
had not been represented by good evidence in our
systematic review. Unique methods of reviewing the
literature (e.g. sourcing agricultural databases and hand-
scanning unindexed publications) proved successful. The
systematic review located 32 publications regarding
cannabinoid receptors in invertebrates, whereas 15 pub-
lications were cited by Elphick & Egertova (2001)
(including dissertations), and 14 publications were cited
by Salzet & Stefano (2002). The following discussion will
be presented under the Ecdysozoa arrangement, grouped
into five clades (see Table 1).
Deuterostomes
The high-affinity [
3
H]CP55,940 binding site we found in
C. intestinalis is presumably the cannabinoid receptor
ortholog that Elphick et al. (2003) characterized as the
descendant of a receptor that predated the CB1–CB2
duplication event. Our results with [
3
H]CP55,940 dis-
placed by the highly CB1 selective antagonist
SR141716A suggest the ancestral sequence may have
functioned more like present-day CB1 than CB2.
Another recent [
3
H]CP55,940 binding study of C. intes-
tinalis (Matias et al., 2005) reported a lower percentage
specific binding in cerebral ganglion than we report
herein. Evidence for cannabinoid receptors in sea urchins
included a slew of Level-II studies on Strongylocentrotus
purpuratus sperm cells (Berdyshev, 1999; Chang et al.,
1991; Chang & Schuel, 1991; Schuel et al.,
1991a,b,1994,1987), plus a Level-III study-specific bind-
ing of [
3
H]CP55,940, with dose–dependent displacement
by THC (Chang et al., 1993).
Lophotrochozoans
We found specific binding of [
3
H]CP55,940 in neural
membranes of L. terristus, extended by measuring the
potency of binding displacement with SR141716A (Log
IC50 ¼)11.20 ± 0.12
MM
), comparable to measurements
performed under identical conditions on rat cerebellum
(Log IC50 ¼)10.17 ± 0.12). These results lends credi-
bility to Level-I evidence regarding earthworms
(Parkinson, 1640), and Level-II evidence regarding
planaria worms (Buttarelli et al., 2002; Lenicque et al.,
1972). Our evidence also reflects upon the controversy
involving a related annelid, the leech H. medicinalis.
Evidence suggests the leech expresses cannabinoid
receptors, based on Level-II studies (Stefano et al.,
1996,1997a,1998; Salzet et al., 1997; Matias et al.,
2001), and a tritiated ligand binding study that used
–14 –13 –12 –11 –10 –9 –8
0
50
100
Lumbricus terristus
Peripatoides
novae-zealandiae
Panagrellus redivivus
Rat c ereb ellum
log
(SR141716A)M
CP55
(
940 bound)
(% of maximum)
Fig. 1 Displacement of 6.3 n
MM
[
3
H]CP55,940 by increasing con-
centrations of SR141716A in 40 lgofLumbricus terristus,Peripatoides
novae-zealandiae, Panagrellus redivivus and rat cerebellum. Values are
expressed as mean ± SEM for values normalized to binding in the
absence of SR141716A in all cases.
370 J. M. McPARTLAND ET AL.
J. EVOL. BIOL. 19 (2006) 366–373 ª2005 EUROPEAN SOCIETY FOR EVOLUTIONARY BIOLOGY

an unusual ligand, [
3
H]anandamide (Stefano et al.,
1996). PCR cloning of a cannabinoid receptor gene
(Stefano et al., 1997b) has been contested (Elphick,
1998; Elphick & Egertova, 2001).
Studies on mollusks (mussel, Mytilus edulis and sea
slug, Aplysia californica) produced four positive studies
and one negative study: M. edulis cannabinoid suppres-
sion of dopamine release, antagonized by SR141716A
(Level II, Stefano et al., 1997a); M. edulis cannabinoid-
stimulated release of nitric oxide, blocked by SR141716A
(Level II, Stefano et al., 1998); M. edulis tritiated ligand
binding study (Level III, Stefano et al., 1996); A. califor-
nica cannabinoid suppression of nerve cell excitability
(Level II, Acosta-Urquidi & Chase, 1975); whereas
Howlett et al. (1990) reported no specific binding of
[
3
H]CP55,940 in A. californica (Level III). The binding
assay used by Howlett and colleagues may have lacked
power (displacement with weaker agonists, not
SR141716A), they also reported no binding in a verteb-
rate, which seems unlikely (a lamprey called Ichthyomy-
zon intercostus but probably I. unicuspis).
Ecdysozoans
Level-IV studies indicate that at least two insects,
D. melanogaster and A. gambiae, lack cannabinoid
receptors (Elphick & Egertova, 2001; McPartland,
2004; McPartland et al., 2001). This agreed with one
tritiated ligand binding study that reported no specific
binding of [
3
H]CP55,940 or [
3
H]SR141716A in a panel
of species spanning the Insecta: D. melanogaster, Apis
mellifera, Gerris marginatus, Spodoptera frugiperda, and
Zophobas atratus (McPartland et al., 2001), but disagreed
with two positive tritiated ligand binding studies.
Egertova et al. (1998) reported 5% specific binding of
[
3
H]CP55,940 in the locust Schistocerca gregaria, but
questioned their findings. Howlett et al. (2000) detected
specific binding of [
3
H]CP55,940 in D. melanogaster,
although the binding was not displaced by CB1-specific
SR141716A or CB2-specific SR144528. In a Level-II
study, the moth Pieris brassicae demonstrated beha-
vioural changes when exposed to THC (Rothschild &
Fairbairn, 1980). But nearly identical behaviour was
elicited by cannabidiol (CBD), a ligand with little
affinity for CB receptors, suggesting behavioural
changes were not mediated by cannabinoid receptors.
Waser (1999) fed THC to the ant Formica pratensis and
no change in behaviour was noted, even though the
drug was absorbed and reached concentrations of
800 pg THC per ant brain. McPartland et al. (2000)
reported no change in behaviour in honeybee
(A. melifera) and ant (Formica sp.) that fed upon high-
THC plants vs. high-CBD plants.
Regarding crustaceans, THC suppressed neuromuscu-
lar junction activity in the lobster H. americanus (Level-II
evidence, Turkanis & Karler, 1988). This positive study
was supported by our tritiated ligand binding study
with the rock lobster J. edwardi. The only onychophoran
examined to date has been the velvet worm P. novae-
zealandiae, whose neural tissues exhibited high-affinity
binding of [
3
H]CP55,940 displaced by SR141716A, at
potencies consistent with rat cerebellar tissue. Our
tritiated binding assay also produced positive results with
the nematode P. redivivus (Fig. 1). This conflicts
with negative Level-IV studies of the related nematode
C. elegans (Elphick & Egertova, 2001; McPartland et al.,
2001). The discrepancy may be due to C. elegans’s
stripped-down genome, resulting in a high rate of
character loss (Copley et al., 2004); perhaps another
‘idiosyncrasy’ of C. elegans is the loss of cannabinoid
receptor genes.
Cnidarians
De Petrocellis et al. (1999) presented strong evidence for
cannabinoid receptors in H. vulgaris (Cnidaria, Class
Hydroza), the most primitive animal with a nervous
system. Cannabinoids induced a Hydra feeding response,
blocked by SR141716A (Level II), and specific, displace-
able cannabinoid binding sites were detected with
[
3
H]SR141716A (Level III). This conflicts with our results
on a related species, A. albocincta (Cnidaria, Class Antho-
zoa). We cannot explain the discrepancy, although
A. albocincta was the most difficult specimen to identify
neural tissue for dissection, perhaps resulting in a Type-II
error.
Poriferans
Systematic review of the literature uncovered no publi-
cations regarding evidence of cannabinoid receptors in
sponges. We found no specific binding of [
3
H]CP55,940
in T. aurantium (Porifera-Epipolasida-Tethyidae).
Conclusions
New information provided by our tritiated ligand binding
study, together with previous findings located in a
systematic review of the literature, indicates the presence
of cannabinoid receptors in all major subdivisions of
bilaterians (deuterostomes, lophotrochozoans, and
ecdysozoans). This information suggests cannabinoid
receptors evolved in the last common ancestor of
bilaterians. Conflicting data regarding Cnidarians (pre-
sent in Hydra, absent in A. albocincta) precludes us from
commenting upon the last common ancestor of eumeta-
zoans. No cannabinoid receptors are expressed in spon-
ges, which probably diverged before the origin of the
eumetazoan ancestor.
The most parsimonious explanation for the lack of
cannabinoid receptors in insects and some nematodes is
secondary loss, Secondary simplification in the Ecdyso-
zoa is also seen in Hox genes (de Rosa et al., 1999),
steroid receptors (Escriva et al., 2000), b-thymosin
Cannabinoid receptors in invertebrates 371
J. EVOL. BIOL. 19 (2006) 366–373 ª2005 EUROPEAN SOCIETY FOR EVOLUTIONARY BIOLOGY

orthologs (Manuel et al., 2000), hedgehog genes (Kang
et al., 2003), and the Wnt gene family (Kusserow et al.,
2005). Loss of cannabinoid receptors is not a synapo-
morphic character shared by all ecdysozoans, however,
as shown by our results with P. novae-zealandiae,
J. edwardi, and P. redivivus. Selection pressures leading
to the loss of cannabinoid receptors in insects and some
nematodes is open to conjecture, as is the functional
relevance of this loss. Transgenic ‘knockout mice’ that
lack CB1 receptors have been generated, and they
survive and reproduce, but they suffer increased mor-
bidity and premature mortality (Zimmer et al., 1999).
CB1 knockout mice show greater aggression, anxiogenic-
like behaviour, depressive-like behaviour, anhedonia,
and they develop fear of newness (Martin et al., 2002).
Lack of cannabinoid receptors in knockout mice and in
insects demonstrates that the receptors may profoundly
alter consciousness, but the receptors did not evolve as a
prerequisite for consciousness. After all, cannabinoid
receptors are absent in honeybees, so the receptors are
not required for spatial memory, goal-directed desires,
elements of deception, and symbolic communication
through dance.
It is worth noting that insects continue to biosynthe-
size sn-2 arachidonyl glycerol (2-AG), an endogenous
ligand of CB1 and CB2, even in the absence of the
receptors (McPartland et al., 2001). This supports Hoyle
(1999), who hypothesized that there is greater evolu-
tionary pressure to conserve ligands than to conserve
receptors. However, insects do not biosynthesize anand-
amide (AEA), the other endogenous lignad of CB1 and
CB2. AEA is a metabolite of arachidonic acid that arises at
the sn-1 position of membrane phospholipids; 2-AG is
metabolized from arachidonic acid at the sn-2 position of
phospholipids (Mechoulam et al., 1998). Insects produce
very little arachidonic acid, especially at the sn-1 position
(Stanley-Samuelson & Pedibhotla, 1996). This may
explain the dearth of AEA in insects, but does it explain
the loss of receptors?
In summary, we hypothesize that the evolution of
cannabinoid receptors was linked to the evolution of
multi-cellular animals. This agrees with the current
employment of cannabinoid receptors in cell-to-cell
communication. The loss of cannabinoid receptors in
some organisms remains an enigma. Future studies are
clearly needed, particularly focusing upon the Nematoda
and Cnidaria, where enhanced phylogenetic resolution is
required.
Acknowledgments
This work was funded by an unrestricted grant from GW
Pharmaceuticals, with additional financial support from a
UNITEC (NZ) AssocProf research grant. We thank Patricia
Pruitt, DVM, for Ciona dissections, Michelle Kelly, PhD,
for Tethya recommendations, and Katherine Blake-
Palmer for technical support.
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Cannabinoid receptors in invertebrates 373
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