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ABC transporters in Arthropods: genomic comparison and role in insecticide transport
and resistance
Dermauw, W.; Van Leeuwen, T.B.S.
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Insect Biochemistry and Molecular Biology
DOI:
10.1016/j.ibmb.2013.11.001
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Citation for published version (APA):
Dermauw, W., & Van Leeuwen, T. (2014). ABC transporters in Arthropods: genomic comparison and role in
insecticide transport and resistance. Insect Biochemistry and Molecular Biology, 45, 89-110. DOI:
10.1016/j.ibmb.2013.11.001
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Download date: 16 Jun 2017
Insect Biochemistry and Molecular Biology 45 (2014) 89e110
Contents lists available at ScienceDirect
Insect Biochemistry and Molecular Biology
journal homepage: www.elsevier.com/locate/ibmb
Review
The ABC gene family in arthropods: Comparative genomics and role
in insecticide transport and resistance
Wannes Dermauw a, **, Thomas Van Leeuwen a, b, *
a
b
Laboratory of Agrozoology, Department of Crop Protection, Faculty of Bioscience Engineering, Ghent University, Coupure Links 653, B-9000 Ghent, Belgium
Institute for Biodiversity and Ecosystem Dynamics, University of Amsterdam, Amsterdam, The Netherlands
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 4 November 2013
Received in revised form
6 November 2013
Accepted 6 November 2013
About a 100 years ago, the Drosophila white mutant marked the birth of Drosophila genetics. The white
gene turned out to encode the first well studied ABC transporter in arthropods. The ABC gene family is
now recognized as one of the largest transporter families in all kingdoms of life. The majority of ABC
proteins function as primary-active transporters that bind and hydrolyze ATP while transporting a large
diversity of substrates across lipid membranes. Although extremely well studied in vertebrates for their
role in drug resistance, less is known about the role of this family in the transport of endogenous and
exogenous substances in arthropods. The ABC families of five insect species, a crustacean and a chelicerate have been annotated in some detail. We conducted a thorough phylogenetic analysis of the seven
arthropod and human ABC protein subfamilies, to infer orthologous relationships that might suggest
conserved function. Most orthologous relationships were found in the ABCB half transporter, ABCD, ABCE
and ABCF subfamilies, but specific expansions within species and lineages are frequently observed and
discussed. We next surveyed the role of ABC transporters in the transport of xenobiotics/plant allelochemicals and their involvement in insecticide resistance. The involvement of ABC transporters in
xenobiotic resistance in arthropods is historically not well documented, but an increasing number of
studies using unbiased differential gene expression analysis now points to their importance. We give an
overview of methods that can be used to link ABC transporters to resistance. ABC proteins have also
recently been implicated in the mode of action and resistance to Bt toxins in Lepidoptera. Given the
enormous interest in Bt toxicology in transgenic crops, such findings will provide an impetus to further
reveal the role of ABC transporters in arthropods.
Ó 2014 The Authors. Published by Elsevier Ltd. Open access under CC BY-NC-ND license.
Keywords:
Traffic ATPases
Primary carriers
Allelochemicals
Multidrug resistance
Detoxification
Phase III
Arthropoda
P-gp
MDR
MRP
BCRP
SUR
Bt toxins
1. Introduction
All cells and organelles are separated from the external milieu by
lipid membranes and need transporters to traffic a wide diversity of
compounds across these membranes (Higgins, 1992). The importance of membrane transport is illustrated by the fact that in
Escherichia coli and humans about 10% and 4% of the total number of
genes encode transport-related proteins (Blattner et al., 1997;
* Corresponding author. Laboratory of Agrozoology, Department of Crop Protection, Faculty of Bioscience Engineering, Ghent University, Coupure Links 653, B9000 Ghent, Belgium. Tel.: þ32 92646154.
** Corresponding author. Tel.: þ32 92646155.
E-mail addresses: [email protected] (W. Dermauw), thomas.
[email protected] (T. Van Leeuwen).
Hediger et al., 2013). Among the latter, the ABC (ATP-binding
cassette)1 protein family is one of the largest transporter families
and is present in all kingdoms of life. The majority of these ABC
proteins function as primary-active transporters, requiring the
binding and hydrolysis of ATP to transport substrates across lipid
membranes. A functional ABC transporter, previously also referred to
as traffic ATPase, consists of two cytosolic nucleotide-binding domains (NBDs) that bind and hydrolyze ATP, and two integral transmembrane domains (TMDs) (Fig. 1A). The NBD domain contains
several conserved sequences like a Walker A and Walker B motif, Qloop, H-motif and the ABC-signature motif (LSGGQ-motif). The
transmembrane domains consist of 5e6 transmembrane helices and
1
ABC, ATP-binding cassette; NBD, nucleotide-binding domain; TMD, transmembrane domain; FT, full transporter; HT, half transporter; BBB, bloodebrain
barrier; MRPs, multidrug-resistance associated proteins; MDR, multidrug-resistance protein, P-gp, P-glycoprotein; SUR, sulfonylurea receptor; TMR, tetramethylrosamine; Trp, Tryptophan; ML, macrocyclic lactone; OP, organophosphates;
BPU, benzoylphenylureas.
0965-1748 Ó 2014 The Authors. Published by Elsevier Ltd. Open access under CC BY-NC-ND license.
http://dx.doi.org/10.1016/j.ibmb.2013.11.001
90
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
A
2
TMD1
TMD2
2
OUT
2
IN
3
4
5
8
6
9 10 11 12
NBD
NBD
NH2
COOH
B
step I
step II
step III
step IV
N BD
N BD
NBD
NBD
ATP
ATP
NBD
ATP
ATP
NBD
NBD
IN
NBD
OUT
Fig. 1. ABC full transporter structure and the ATP-switch model for the transport cycle of an ABC transporter (exporter-type) A) Typical structure of an ABC full transporter
containing two TMDs, TMD1 (blue) and TMD2 (yellow) each containing 6 transmembrane (TM) segments, and two NBDs, NBD1 (red) and NBD2 (green). In the case of “long” MRPs an
additional TMD (TMD0) is present at the N-terminus (Deeley et al., 2006). ABC half transporters have only one TMD and one NBD and need to homo or heterodimerize to form a
functional unit. B) The ATP-switch mechanism (Higgins and Linton, 2004; Linton and Higgins, 2007). The transport cycle is started by binding of a substrate (purple cross) to a highaffinity pocket formed by the TMDs (blue and yellow pentagon). Subsequently, a conformational change is transmitted to the NBDs (green and red), facilitating ATP (orange oval)
binding and closed NBD-dimer formation. The closed NBD dimer induces on its turn a major conformational change in the TMDs, with TMDs rotating and opening toward the
outside, initiating substrate translocation (step II). ATP hydrolysis initiates dissolution of the closed NBD dimer, resulting in further conformational changes in the TMDs (step III).
Finally, phosphate and ADP release restores the transporter to the open NBD-dimer conformation (step IV).
provide substrate specificity. The four domains of a functional
transporter can be present in one protein (Full Transporter, FT) or
spread over multiple proteins, for example one NBD and TMD per
protein (half transporter, HT). In the case of the latter, ABC proteins
need to homo or heterodimerize to form a functional ABC transporter (Higgins, 1992; Higgins and Linton, 2004; Rees et al., 2009).
Based on the sequence similarity of the NBDs, the ABC protein family
has been divided into eight subfamilies, denoted by the letters AeH
(Fig. 2). Of particular note, the ABCH subfamily has at present only
been identified in arthropod genomes and in zebrafish (Dean et al.
2001; Popovic et al., 2010). Eukaryotic ABC transporters mediate
the efflux of compounds from the cytoplasm to the outside of the cell
or into organelles, while bacterial ABC transporters can also facilitate
import of substances. However, not all ABC proteins do play a role in
transport. In the human ABC protein family, for example, ABC proteins acting as ion channel, receptor or with a role in translation,
have been characterized (Dean et al., 2001; Rees et al., 2009).
Currently, the ATP-switch mechanism is the favored model for
the transport cycle of ABC transporters (George and Jones, 2012;
Higgins and Linton, 2004; Linton and Higgins, 2007). In this model,
the transport cycle is started by binding of substrates to the TMDs.
Subsequently a conformational change is transmitted to the NBDs,
facilitating ATP binding and closed NBD-dimer formation. The closed
NBD dimer in turn induces a major conformational change in the
TMDs, with TMDs rotating and opening toward the outside, initiating substrate translocation. In a final step, ATP is hydrolyzed to
initiate transition of the closed NBD dimer to the open NBD dimer
and to return the transporter to its basal state (Fig. 1B). ABCtransporters traffic an array of substrates covering amino-acids,
sugars, lipids, inorganic ions, polysaccharides, metals, peptides, toxic
metabolites and drugs. Because of their ability to transport drugs,
many human ABC members are also important players in multidrug
resistance of cancer cells against chemotherapeutics: the multidrugresistance proteins (MDRs) or P-glycoproteins (P-gps) belonging to
the ABCB subfamily, the multidrug-resistance associated proteins
(MRPs) from the ABCC subfamily and the breast cancer resistance
protein (BCRP, ABCG subfamily member). The P-gp ABCB1 was in fact
identified as the first ABC transporter to be overexpressed in
multidrug resistant tumor cell lines (Kartner et al., 1985; Riordan
et al., 1985). Compared to bacterial, nematode and human ABC
transporters, the knowledge about arthropod ABC transporters is
still limited. At present, the ABC families of seven arthropod species,
spanning more than 400 arthropod ABC proteins, have been studied
in detail (Broehan et al. 2013; Dean et al., 2001; Dermauw et al.,
2013a; Liu et al., 2011; Roth et al., 2003; Sturm et al., 2009; Xie
et al., 2012). Nevertheless, the precise function of only a few
arthropod transporters has been unveiled. Drosophila melanogaster
white and its insect orthologues are probably the most thoroughly
characterized ABC transporters and are involved in the uptake of
pigment precursors in the developing eye (Ewart and Howells, 1998).
We performed a thorough phylogenetic analysis with a dataset
of well-annotated arthropod and human ABC transporters to infer
orthologous relationships, which in turn can suggest the role and
function of arthropod proteins. We further focus on biochemical
and molecular methods that can be deployed to study arthropod
ABC transporters, and give an overview of their documented role in
xenobiotic transport and resistance.
2. Comparison of ABC subfamilies in arthropods
Despite the fact that more than 60 arthropod genomes have
been sequenced, the ABC protein family was annotated and studied
in detail in only seven species: the dipterans D. melanogaster and
Anopheles gambiae, the honeybee Apis mellifera, the silk moth
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
91
Fig. 2. A schematic tree depicting the phylogenetic relationships between ABC subfamilies. NBDs of ABC proteins belonging to different subfamilies as defined by Dermauw et al.
(2013a) were used in a ML analysis. Similar to previous studies (e.g. Liu et al., 2011) C-terminal NBDs of the ABCC subfamily clustered together with NBDs of the ABCB subfamily.
Bombyx mori, the flour beetle Tribolium castaneum, the spider mite
Tetranychus urticae and the waterflea Daphnia pulex (Broehan et al.,
2013; Dean et al., 2001; Dermauw et al., 2013a; Liu et al., 2011; Roth
et al., 2003; Sturm et al., 2009; Xie et al., 2012) (Table 1). Among
these species, T. urticae has the largest number of ABC genes (103)
followed by T. castaneum (73) and D. pulex (65), while A. mellifera
has only 41 ABC genes (Table 1). The large number of ABC genes in
T. urticae is mainly due to lineage-specific expansions of ABCC,
ABCG and ABCH genes, while in T. castaneum and D. pulex lineagespecific expansions of ABCC and ABCH genes have been documented (Broehan et al., 2013; Dermauw et al., 2013a; Sturm et al.,
2009) (Table 1). In order to infer orthologous relationships between arthropod ABC proteins, we performed a maximum likelihood analysis for each ABC subclass using complete ABC protein
sequences from Homo sapiens and the before mentioned arthropod
species (for details see Supplementary Text).
2.1. The ABCA family
Phylogenetic analysis of the ABCA subfamily revealed a clear
orthologous relationship between D. melanogaster CG31731,
A. gambiae AGAP001523, T. castaneum TcABCA-7 and TcABCA-UA,
A. mellifera AmABCA2 and B. mori BGIBMGA004187/
BGIBMGA004188 (Supplementary Fig. 1, Table 2). This group of
insect ABCA FTs cluster together with high bootstrap support with 7
ABCA FTs of T. urticae. Except for CG31731, reported to be downregulated in the salivary glands of an E93 mutant of D. melanogaster
(Dutta, 2008), no information is available on the function of these
arthropod ABCA transporters. Other orthologous relationships
were found between A. gambiae AGAP010416, A. mellifera
AmABCA3, T. castaneum TcABCA-6A, B. mori BGIBMGA012789 and
D. pulex Dappudraft_346971 and human ABCA5, 6, 8e10 and between A. gambiae AGAP010582, D. melanogaster CG34120,
Table 1
Gene numbers in ABC subfamilies of H. sapiens and seven arthropod species.a
ABC subfamily
H. sapiens
D. melanogaster
A. gambiae
T. castaneum
A. melliferab
B. moric
D. pulex
T. urticae
A
B FT
B HT
C
D
E
F
G
H
TOTAL
12
4
7
12
4
1
3
5
0
48
10
4
4
14
2
1
3
15
3
56
9
1
4
13
2
1
3
16
3
52
10
2
4
35
2
1
3
13
3
73
3
1
4
9
2
1
3
15
3
41
9
5
4
15
2
1
3
13
3
55
4
2
5
7
3
1
4
24
15
65
9
2
2
39
2
1
3
23
22
103
a
Numbers were derived from Sturm et al. (2009), Liu et al. (2011), Xie et al. (2012), Broehan et al. (2013) and Dermauw et al. (2013a).
AmABCB5 and AmABCB6 from Liu et al. (2011) were not taken into account as they were incomplete (less than 270 AA), their respective genes were located on unplaced
scaffolds and had best BLASTp hits (E-value e100) with ABC transporters of bacteria.
c
The number of B. mori ABC genes for each ABC subfamily represents the highest number of B. mori genes found for each ABC subfamily in either Liu et al. (2011) or Xie et al.
(2012), some B. mori ABC genes might be incomplete and might possibly be combined into one gene (e.g. BGIBMGA007792/BGIBMGA007793 (Atsumi et al., 2012),
BGIBMGA007784/BGIBMGA007785 and BGIBMGA4187/BGIBMGA04188).
b
92
Table 2
Orthologous relationships between ABC proteins of Arthropoda and H. sapiens.e
Dma
Ag
Tcb
Am
Bm
Dp
Tu
Hs
Function?
Reference
ABCA
e
CG34120
AGAP010416
AGAP010582
TcABCA-6A
e
AmABCA3
AmABCA1
BGIBMGA012789
BGIBMGA009503
Dappudraft_346971
e
e
tetur27g01890
hABCA5/6/8-10
hABCA12/13
Wenzel et al., 2007,
Liu and Lehmann 2008; Akiyama,
2013
CG31731
AGAP001523
TcABCA-7A
TcABCA-UA
AmABCA2
BGIBMGA004187
BGIBMGA004188
e
(7)c
e
Lipid transportd?
ABCA12 is a keratinocyte lipid transporterd;
GAL4 changes CG34120 expression,
independent of UAS
CG31731 is downregulated in E93 mutants
ABCB HT
CG7955
CG4225
CG1824
CG3156
AGAP006364
AGAP002278
AGAP006273
AGAP002717
TcABCB-5A
TcABCB-6A
TcABCB-7A
TcABCB-4A
AmABCB1
AmABCB2
AmABCB7
AmABCB3
BGIBMGA012743
BGIBMGA005473
BGIBMGA004142
BGIBMGA008523
Dappudraft_347270
Dappudraft_347268
Dappudraft_347266
Dappudraft_347275
Dappudraft_347276
tetur32g01330
e
tetur17g02000
e
hABCB7
hABCB6
hABCB8
hABCB10
Mitochondrial ABC transporters with roles in
biogenesis of cytosolic ironesulfur clusters,
heme biosynthesis, iron homeostasis,
protection against oxidative stressd; CG4255 is
involved in cadmium tolerance; RNAi silencing
of TcABCB-5A results in severe developmental
defects during pupation and abortive pupaladult molting; role in pesticide resistance?
Zutz et al., 2009; Sooksa-Nguan
et al., 2009; Bariami et al., 2012;
Broehan et al., 2013
ABCC
sur
AGAP009799
e
AmABCC2
BGIBMGA006882
Dappudraft_442500
tetur11g05990
Subunit of the ATP-sensitive potassium channel
Nasonkin et al., 1999
CG6214
(dMRP)
CG7806
AGAP009835
AGAP008437
AGAP007917
TcABCC-9A
AmABCC9
(3)c
Dappudraft_347281
(23)c
AmABCC5
BGIBMGA010636
Dappudraft_347323
tetur03g07840
Drosophila CG6214 is capable of transporting
several substrates of its human orthologs
Transport of taxanesd; CG7806 is upregulated
1.56-fold in D. melanogaster upon exposure to
ecdysone; conserved function in metazoa?
Tarnay et al., 2004; Szeri et al., 2009
TcABCC-4A
hABBC8/9
(SUR1/2)
hABCC1/2/3/6
(MRP1, 2, 3 and 6)
hABCC10 (MRP7)
AGAP002071
AGAP000440
TcABCD-6A
TcABCD-9A
AmABCD1
AmABCD2
BGIBMGA004616
BGIBMGA012688
Dappudraft_347330
Dappudraft_347326
tetur05g06640
tetur35g01360
hABCD1/2
hABCD3
Import of long branched chain acyl-coA into
peroxisomed
Morita and Imanaka, 2012
AGAP002182
TcABCE-3A
AmABCE1
BGIBMGA010129
Dappudraft_189585
tetur30g0140
hABCE1
Essential role in ribosome biogenesis and
translation regulation
Andersen and Leevers, 2007
AGAP012249
TcABCF-2A
AmABCF1
BGIBMGA007869
tetur29g00620
hABCF1 (ABC50)
Essential role in translation regulationd
Paytubi et al., 2009
CG9281
CG9330
ABCG
CG3327
(E23)
AGAP002693
AGAP012005
TcABCF-5A
TcABCF-9A
AmABCF3
AmABCF2
BGIBMGA002004
BGIBMGA006964
Dappudraft_347357
Dappudraft_347363
Dappudraft_304799
Dappudraft_347354
tetur20g02610
tetur32g00490
hABCF2
hABCF3
e
TcABCG-8A
AmABCG1
e
Dappudraft_347416
tetur17g02510
e
Hock et al., 2000; Broehan et al.,
2013,
White
AGAP000553
TcABCG-9B
AmABCG2
BGIBMGA002922
(9)c
(9)c
e
Brown
AGAP007655
e
e
e
TcABCG-9A
Dappudraft_347393
e
e
CG11069
CG31121
Atet
AGAP000506
AGAP001333
AGAP002051
AGAP002050
AGAP001858
BGIBMGA002581
BGIBMGA002712
BGIBMGA002924
e
Scarlet
AmABCG11
AMABCG12
AmABCG10
TcABCG-9D
TcABCG-9C
TcABCG-4D
AmABCG15
AmABCG13
AmABCG3
BGIBMGA000472
BGIBMGA000220
BGIBMGA005094
Dappudraft_300887
Dappudraft_258299
Dappudraft_347444
tetur01g16290
tetur01g16280
e
hABCG5
hABCG8
hABCG1/4
CG3164
AGAP009850
TcABCG-4C
AmABCG5
BGIBMGA005202
e
e
e
20E induced ABC transporter, injection of
TcABCG-8A dsRNA results in molting defects
and developmental arrest
Best characterized ABC transporter: transport of
eye pigment precursors, role in courtship
behavior, uptake of uric acid, transport of cyclic
GMP and transport of biogenic amines.
Transport of eye pigment precursor; transport
of biogenic amines
Transport of eye pigment precursor;transport of
biogenic amines
Role in biliary excretiond
Role in biliary excretiond
ABC transporter expressed in trachea of
D. melanogaster
CG3164 knockdown line results in lethal
phenotype in pupa; essential in D. melanogaster
lipid metabolism; transport of lipids from
epidermal cells to the cuticle in T. castaneum;
injection of TcABCG-4C dsRNA in young females
Beckstead et al., 2007; HopperBorge et al., 2009
Zhang and Odenwald, 1995; Ewart
and Howells 1998; Borycz et al.,
2008; Evans et al., 2008; Komoto et
al., 2009; Broehan et al., 2013
Ewart and Howells 1998, Borycz et
al., 2008
Ewart and Howells 1998, Borycz et
al., 2008, Tatematsu et al., 2011
Graf et al., 2003
Graf et al., 2003
Kuwana et al., 1996
Ohmann 2012, Broehan et al., 2013,
Liu et al., 2011
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
ABCD
CG2316
CG12703
ABCE
CG5651
(pixie)
ABCF
CG1703
Dutta, 2008
?
e
e
e
BGIBMGA010825
AmABCH2
TcABCH-9A
AGAP002060
CG11147
Species abbreviations: Dm, D. melanogaster, Ag, A. gambiae, Tc, T. castaneum, Am, A. mellifera, Bm, B. mori, Dp, D. pulex, Tu, T. urticae, Hs, H. sapiens.
b
Broehan et al. (2013)-naming.
c
Number of orthologues that has been found in this arthropod species, gene names of orthologues can be found in the phylogenetic analysis of the corresponding ABC family.
d
Function of human ortholog.
e
Orthologous genes should 1) at least be present in four out of five insect species or five out of seven arthropod species and 2) be clustered with a bootstrap value of at least 80%; one-to-one orthologues in arthropods and H.
sapiens are underlined.
e
e
e
BGIBMGA010760
AmABCH1
TcABCH-9C
AGAP003680
CG9990
TcABCH-9B
AGAP002638
ABCH
CG33970
a
Mummery-Widmer et al., 2009,
Zhang et al., 2009, Broehan et al.,
2013
e
Qin et al., 2005
Upregulated in adult fruit flies after cold
hardening
Similar to CG3164/TcABCG-4C
e
e
e
BGIBMGA010726
e
e
e
e
e
e
BGIBMGA005203
BGIBMGA010557
BGIBMGA010555
AmABCG9
AmABCG8
AmABCG14
TcABCG-4B
e
TcABCG-4A
e
(5)c
AGAP009464
CG4822
CG9664
CG17646
AmABCH3
Liu et al., 2011
Liu et al., 2011
Buchmann et al., 2007, Liu et al.,
2011
Pedra et al., 2004, Ellis and Carney
2010
e
e
e
e
TcABCG-4F
AGAP008889
CG31689
AmABCG6
e
AGAP009463
CG5853
TcABCG-4G
AmABCG4
BGIBMGA005226
e
e
reduces number of eggs and eggs fail to hatch;
20E induced ABC transporter in B. mori
Downregulation of CG5853 following neuronal
AP1 overexpression; 20E induced ABC
transporter in B. mori
CG31689 is overtranscribed in a D. melanogaster
DDT-resistant line; downregulated in
D. melanogaster male heads after mating
20E induced ABC transporter in B. mori
20E induced ABC transporter in B. mori
Insertion of a engineered transposable Pelement (EP) upstream of CG17646 altered the
triglyceride content of D. melanogaster; 20E
induced ABC transporter in B. mori
Etter et al., 2005, Liu et al., 2011
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
93
A. mellifera AmABCA1, B. mori BGIBMGA009503, T. urticae
tetur27g01890 and human ABCA12 and 13 (see Supplementary
Fig. 1). Little is known about the function of human ABCA5, 6, 8e
10 but several studies have suggested roles of these transporters in
lipid transport (Wenzel et al., 2007). Human ABCA12 on the other
hand has been thoroughly characterized and is known as a keratinocyte lipid transporter (Akiyama, 2013). The role of the
arthropod orthologues of these human ABCAs is unknown, but they
might play a role in lipid transport. Finally, we also identified a
well-supported group of insect-specific ABCA proteins, containing
lineage-specific expansions in B. mori, T. castaneum, D. melanogaster
and A. gambiae (see Supplementary Fig. 1). To date, only one study
has investigated the function of ABC proteins in this insect-specific
ABCA group. Injection of dsRNA targeting TcABCA-9A or TcABCA-9B
of T. castaneum resulted in 30% mortality during the pupaeadult
molt and pupae and pharate adults showed severe wing defects
and elytra shortening (Broehan et al., 2013). Future studies might
confirm that this group of insect-specific ABCAs is involved in
(wing) development.
2.2. The ABCB family
The ABCB subfamily consists of half and full transporters (HTs
and FTs, respectively, see above). The phylogenetic analysis of
ABCB FTs, also referred to as P-gps, assigned H. sapiens, D. pulex,
T. urticae and insects into separate clades, confirming an earlier
hypothesis by Sturm et al. (2009) that this subfamily diversified
through lineage-specific duplications (Fig. 3A). Within the insect
ABCB FT clade, 2 main groups can be observed. In the first
Drosophila mdr50 clusters with TcABCB-3A, TcABCB-3B and
BGIBMGA007494, while in the second Drosophila mdr65, CG10226
and mdr49 cluster with A. gambiae AGAP005639, A. mellifera
AmABCB4 and B. mori BGIBMGA009452, BGIBMGA000724,
BGIBMGA000725 and BGIBMGA011228 (Fig. 3A). The ABCB FTs in
D. melanogaster have been extensively studied as mdr65 is
required for chemical protection of the fruit fly brain by creating a
bloodebrain barrier (BBB) (Mayer et al., 2009) and mdr49 plays an
essential role in germ cell migration (Ricardo and Lehmann, 2009).
Furthermore, it was demonstrated that exposure of
D. melanogaster flies to polycyclic aromatic hydrocarbons led to an
induced ABCB FT/P-gp expression (Vaché et al., 2006, 2007).
Chahine and O’Donnell (2009) revealed that D. melanogaster adult
flies raised on a diet containing 0.1 mM of the antifolate drug
methotrexate, showed a significant increased expression of
mdr49, mdr50 and to a lesser extent mdr65, in both the Malpighian tubules and the gut. A recent study reported the interaction
between P-gps and polyglutamine mediated pathogenesis in
D. melanogaster. Using a D. melanogaster transgenic strain
expressing repeats of glutamine, it was demonstrated that
increased ABCB FT/P-gp levels reduces eye degeneration, while
reduction of ABCB FT/P-gp levels resulted in an increase of the
disease (Yadav and Tapadia, 2013). Interestingly, D. melanogaster
and other arthropod ABCB FTs have also frequently been linked to
insecticide transport and/or resistance ((Buss and Callaghan,
2008), this study, see Section 4). Contrary to ABCB FTs, clear
orthologous relationships were identified among arthropod ABCB
HTs (Fig. 3B). Human ABCB6e8 each have one ortholog in insects
and D. pulex (Fig. 3B, Table 2) while one ortholog and two coorthologues (Dappudraft_347275 and Dappudraft_347276) of human ABCB10 were found in insects and D. pulex, respectively. Of
particular note, two previously annotated ABCB HTs of A. mellifera
(AmABCB5 and AmABCB6 (Liu et al., 2011)) were not included in
our analysis as they were incomplete (less than 270 AA), their
respective genes were located on unplaced scaffolds and had best
BLASTp hits (E-value e100) with ABC transporters of bacteria,
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A
B
7
Fig. 3. Phylogenetic analysis of ABCB full and half transporters of H. sapiens and seven arthropod species. A) ABCB full transporters, B) ABCB half transporters. For procedure and
details of phylogenetic analysis see Supplementary Text. Color and shape codes are as follows: blue diamond, D. melanogaster, red diamond, A. gambiae, brown diamond,
T. castaneum, green diamond, B. mori, yellow diamond, A. mellifera, purple square, D. pulex, gray triangle, T. urticae and black circle, H. sapiens. The scale bar represents in A) 0.2 and
in B) 0.5 amino-acid substitutions per site.
potentially indicating contamination (data not shown). Contrary to
insects and D. pulex, only two orthologues of human ABCB HTs
(ABCB7 and ABCB8) were identified in T. urticae. Finally, no
arthropod orthologues of human ABCB2 and ABCB3, both involved
in antigen processing (Trowsdale et al., 1990), and ABCB9 were
identified in our evolutionary analysis (Fig. 3, Table 2). The group of
evolutionary conserved ABCB HTs (orthologues of human ABCB6e
8 and 10) is known as mitochondrial ABC transporters with roles in
the biogenesis of cytosolic ironesulfur clusters, heme biosynthesis,
iron homeostasis and protection against oxidative stress (Zutz
et al., 2009). It was suggested recently that human ABCB6 is
located in other cellular compartments than the mitochondria, but
the precise identity, substrate specificity and functionality has not
yet been well determined (Chavan et al., 2013). Hardly any functional information is available on the function of arthropod
orthologues of human ABCB HTs. Sooksa-Nguan et al. (2009)
showed through heterologous expression in Schizosaccharomyces pombe that the D. melanogaster ortholog of human ABCB6,
CG4225 also known as D. melanogaster heavy metal tolerance
factor 1 (DmHMT-1), could confer resistance against cadmium.
CG7955 was downregulated in D. melanogaster lines selected for
increased resistance to chill coma stress (Telonis-Scott et al., 2009).
On the other hand, Broehan et al. (2013) found that injection of
TcABCB-5A dsRNA into pre-pupae of T. castaneum resulted in 1)
severe developmental defects during pupaeadult molt causing
lethality and 2) sterile females with ovaries failing to produce eggs.
Nevertheless, based on the clear orthologous relationships between human and arthropod ABCB HTs and the fact that many of
the arthropod ABCB HTs are predicted to be targeted to the
mitochondria (predicted by TargetP 1.0 (Emanuelsson et al., 2007),
data not shown), it is likely that arthropod ABCB HTs have similar
roles as their counterparts in humans.
2.3. The ABCC family
Human ABCC proteins are FTs with functionally diverse functions such as ion transport (e.g. human ABCC7, the cystic fibrosis
transmembrane conductance regulator), cell-surface receptor activity (e.g. human ABCC8 and 9, sulfonylurea receptors) and
translocation of a broad range of substrates like drugs, cyclic nucleotides, endogenous/exogenous compounds and their glutathione conjugates (Dean et al., 2001; Keppler, 2011; Leslie et al.,
2005; Schinkel and Jonker, 2003). Because of their ability to
extrude drugs with broad specificity, many ABCCs were historically
named multidrug-resistance associated proteins (MRPs), with
“long” MRPs (human ABCC1-3, 6 and 10/MRP1e3, 6 and 7) having
an extra N-terminal transmembrane spanning domain (TMD0)
compared to “short” MRPs (human ABCC4, 5, 11 and 12/MRP4, 5, 8
and 9) (Deeley et al., 2006). Human MRP1 was the first MRP to be
cloned and was identified in a multidrug resistant lung cancer cell
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
95
Fig. 4. Phylogenetic analysis of ABCC proteins of H. sapiens and seven arthropod species. For procedure and details of phylogenetic analysis see Supplementary Text. For color and
shape codes see Fig. 3. The scale bar represents 0.5 amino-acid substitutions per site. One-to-one ABCC orthologues found in H. sapiens and seven arthropod species are shaded in
grey, while examples of species-specific ABCC expansions are indicated with an arch.
line (Cole et al., 1992; Cole, 2013). The various MRPs differ from Pgps (ABCB FTs) by the ability to actively transport a broad range of
glucuronate, sulfate and glutathione (GSH)-conjugated organic
anions. It has also been shown that several MRPs act in synergy
with several phase II conjugating enzymes like the GSH-Stransferases (GSTs) and UDP-glycosyltransferases (UGTs) to confer
resistance to electrophilic drugs and carcinogens (Leslie, 2012; Liu
et al., 2012; Sau et al., 2010). Intriguingly, the two arthropod species with the highest number of ABCCs, T. urticae and T. castaneum,
also have a clear expansion of the GST repertoire (Grbi
c et al., 2011;
Shi et al., 2012). The T. urticae genome harbors an expansion of muclass GSTs that are not present in insects and were until recently
believed to be vertebrate specific (Grbi
c et al., 2011). Future studies
should reveal if there is a coordinated action between the high
number of GSTs and the many ABCCs in T. urticae and T. castaneum.
Next to the transport of conjugated compounds, human MRPs can
also transport unconjugated compounds, including the chemotherapeutics vinblastine, etoposide, daunorubicin, and doxorubicin,
in co-transport with GSH (Ballatori et al., 2009; Leslie et al., 2005;
Loe et al., 1998). In our phylogenetic analysis of the ABCC subfamily, 23 T. urticae ABCC transporters of which the majority has a
TMD0, (Dermauw et al., 2013a) clustered with D. melanogaster
CG6214, A. gambiae AGAP009835 and AGAP008437, A. mellifera
AmABCC9, T. castaneum TcABCC-9A, B. mori BGIBMGA010330,
BGIBMGA010331 and BGIBMGA010332 and a group of human
“long” MRPs (MRP1e3 and 6) (Fig. 4). Using recombinant expression in Sf9 cells and vesicular assays, it was reported that
D. melanogaster CG6214 (also named dMRP) shares the biochemical
properties of its human orthologues, such as the transport of bestradiol 17-b-D-glucuronide, leukotriene C4 and calcein (Szeri
et al., 2009; Tarnay et al., 2004). It has also been shown that
alternative splicing results in multiple isoforms for D. melanogaster
CG6214 and its orthologues in A. gambiae and Trichoplusia ni, which
might generate functional diversity for this ABCC transporter
(Grailles et al., 2003; Labbé et al., 2011; Roth et al., 2003).
Furthermore, Chahine and O’Donnell (2009, 2011) found that dietary exposure to the chemotherapeutic agent methotrexate or the
P450-monooxygenase inhibitor piperonylbutoxide resulted in an
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upregulation of CG6214 in the Malpighian tubules of
D. melanogaster. This group also discovered that reduction in the
expression of CG6214 was correlated with reduced secretion of the
chemotherapeutic daunorubicin (Chahine et al., 2012). Less is
known about the other arthropod orthologues of human ABCC1e3
and 6. Recently, Yoon et al. (2011) reported that the ortholog of
CG6214 in Pediculus humanus humanus (PhABCC4) was more highly
expressed after exposure to the pesticide ivermectin and that injection of PhABCC4 dsRNA altered the sensitivity of P. humanus to
ivermectin (see also Section 4.2.2).
The evolutionary analysis also identified two clear orthologous
relationships between arthropod ABCCs and human ABCCs. One
ortholog of a human “long” MRP, human ABCC10/MRP7, was present in all arthropods included in our analysis (Fig. 4, Table 2). In
contrast to human MRP1, not much is known about the function of
human ABCC10. This MRP is distinct from other human ABCCs in
that it does not transport typical human MRP substrates like
glutathione and sulfate conjugates. Unlike other MRPs, human
MRP7 is able to confer resistance to taxanes (Hopper-Borge et al.,
2009). Nevertheless, the presence of one-to-one orthologues
with arthropods, suggests a more conserved function of this ABCC
protein in humans and arthropods. The human sulfonylurea receptors (human ABCC8/SUR1 and human ABCC9/SUR2) clustered
with D. melanogaster Sur, A. gambiae AGAP009799, B. mori
BGIBMGA006882, A. mellifera AmABCC2, D. pulex Dappudraft_442500 and T. urticae tetur11g05990 (Fig. 4, Table 2). Sulfonylurea receptors assemble with inwardly rectifying potassium
(Kir) subunits, to form ATP-sensitive potassium channels (KATP).
These channels are involved in multiple physiological processes,
with roles in glucose homeostasis, ischemic protection and innate
immunity (Akrouh et al., 2009; Eleftherianos et al., 2011). Surprisingly, a clear ortholog of human SUR could not be identified in
the genome of T. castaneum (see also the OrthoDB group of
arthropod SURs (EOG7HFCZ2) (Waterhouse et al., 2011)). In addition, we could not identify such orthologues in transcriptome data
of other coleopterans (as determined by a tBLASTn search against
coleopteran ESTs in NCBI and using human SUR1 and 2 as query),
suggesting that SUR was lost in the coleopteran lineage. However,
as orthologues of Kir are present in T. castaneum (as determined by
a BLASTp search against the T. castaneum proteome using
D. melanogaster Irk1-PB as query) other ABC transporters might
assemble with Kir to form a functional ATP-sensitive potassium
channel. Although some reports suggest that SURs are the direct
target of insecticidal benzoylphenylureas, a group of commercially
used chitin synthesis inhibitors, this theory is no longer favored
(see Section 4.4.1).
A number of ABCC proteins of D. pulex (Dappudraft_347292,
Dappudraft_347295 and Dappudraft_347548) and A. mellifera
(AmABCC4) clustered with a group of “short” human MRPs (human
ABCC5, 11 and 12) that are characterized as transporters of (cyclic)
nucleotides/nucleosides and their analogs (Guo et al., 2003; Reid
et al., 2003) (Fig. 4). It has also been found that a SNP in the human ABCC11 gene is the determinant for the human earwax type
(Yoshiura et al., 2006) and that human ABCC11 is essential for the
secretion of odorants and their precursors from apocrine sweat
glands (Martin et al., 2009). Finally, human ABCC4 and ABCC7/CFTR
clustered with a large clade of arthropod ABCC proteins. Within this
clade, 14 T. urticae ABCC transporters formed a sister-group of
Dappudraft_347288 and a large insect-specific clade containing
lineage-specific expansions of D. melanogaster and T. castaneum
ABCC transporters. The function of members from this large
arthropod ABCC transporter clade is mainly unknown. T. urticae
ABCC proteins tetur14g02290, tetur14g02310, tetur14g02320,
tetur14g02330 and tetur23g02452 were reported to be downregulated in diapausing T. urticae females. Diapausing spider mites
do not feed and the reduction in plant allelochemicals uptake may
be linked to the downregulation of these ABCC genes (Bryon et al.,
2013). Next, it has been demonstrated that D. melanogaster
CG10505 contributes to metal homeostasis while D. melanogaster
CG14709 regulates responsiveness to O2 deprivation and development under hypoxia (Huang and Haddad, 2007; Yepiskoposyan
et al., 2006). Recently, Shah et al. (2012) also showed that
D. melanogaster CG4562 was upregulated in Cyp6g1 knockdown
flies, suggestive of molecular interaction within a detoxification
network. The most enigmatic member from this large arthropod
ABCC-group clade is probably B. mori BGIBMGA007792e
BGIBMGA007793 (these proteins were considered as separate
transporters by Liu et al., 2011, but should be combined to form one
full transporter (Atsumi et al., 2012)). An insertion in this B. mori
transporter has been clearly linked to resistance against the Bacillus
thuringiensis (Bt) Cry1A toxin but its precise role in Bt toxin mode of
action is however unknown (see Section 4.2.2). Remarkably, no
clear orthologues of this lepidopteran transporter were detected in
the other arthropod species included in our analysis (Fig. 4) and this
might help explain why Cry1A toxins are exclusively toxic against
Lepidoptera (de Maagd et al., 2001).
2.4. The ABCD subfamily
The ABCD subfamily consists of HTs that need to dimerize to
form a functional transporter. In humans they are involved in the
import of long and branched chain acyl-CoA molecules into the
peroxisome (Morita and Imanaka, 2012). Both T. urticae and all
insect species have two ABCD genes, while D. pulex and humans
have three and four ABCD genes respectively (Table 2). The clear
orthologous relationships between arthropod ABCDs and human
ABCD1/2 and ABCD3 suggest that their function is conserved
(Fig. 5A, Table 2).
2.5. The ABCE and ABCF family
ABC proteins from the ABCE and ABCF family have 2 NBDs but
lack TMDs and are involved in biological processes other than
transport. The ABCE1 protein is essential in all eukaryotes examined to date and plays a fundamental role in ribosome biogenesis
and translation regulation (Andersen and Leevers, 2007; Barthelme
et al., 2011). Contrary to human ABCE1 (also called Rli1p or OABP),
human ABCF1 (also called ABC50) is probably only involved in
translation regulation (Paytubi et al., 2009), while the precise role
of ABCF1 and ABCF2 is unclear. In our phylogenetic analysis, one
ortholog of human ABCE1, ABCF1e3 was found for each arthropod
species (Fig. 5B, Table 2). Based on these clear orthologous relationships, arthropod ABCE and ABCF proteins probably have an
analogous function as their human orthologues. An essential role of
ABCE and ABCF proteins was recently demonstrated for
T. castaneum, where TcABCE-3A or TcABCF-2A dsRNA injection in
penultimate larvae resulted in 100% mortality (Broehan et al.,
2013).
2.6. The ABCG family
The ABCG family is present in most metazoan species, plants
and fungi but while in metazoans only ABCG HTs have been reported to date, several ABCG FTs (also termed pleiotropic drug
resistance proteins (PDRs)) have been identified in plant and fungi
(Kovalchuk and Driessen, 2010; Verrier et al., 2008). ABCG HTs have
a typical reverse domain organization, with the NBD being localized to the N-terminal side of the TMD, and need to dimerize to
form a functional transporter. The principal substrates of four out of
five human ABCG proteins are endogenous and mostly dietary
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
A
97
B
Fig. 5. Phylogenetic analysis of ABCD, E and F proteins of H. sapiens and seven arthropod species. A) ABCD proteins, B) ABCE and F proteins. For procedure and details of phylogenetic
analysis see Supplementary Text. For color and shape codes see Fig. 3. The scale bar represents 0.5 amino-acid substitutions per site.
lipids, while human ABCG2 (breast cancer resistance protein, BCRP)
has a much broader substrate specificity and is known as a multidrug efflux pump (Kerr et al., 2011; Tarr et al., 2009). In our
phylogenetic analysis we identified for each arthropod species an
ortholog of human ABCG5 and 8. Similar to human ABCG5/ABCG8,
the arthropod orthologues (Fig. 6, Table 2), are all found juxtaposed
in a head-to-head orientation (Dermauw et al., 2013a). In humans,
ABCG5 and 8 are glycoproteins that form obligate heterodimers.
They limit the absorption of plant sterols and cholesterol from the
diet and promote secretion of plant sterols and cholesterol from
liver cells into the bile (Graf et al., 2003; Kerr et al., 2011; Tarr et al.,
2009). Based on their head-to-head orientation and clear orthologous relationships with human ABCG5 and ABCG8, these arthropod
ABCGs probably have a similar role as their human orthologues.
Phylogenetic analysis reveals that D. melanogaster white
clusters with T. castaneum TcABCG-9B, B. mori BGIBMGA002922,
A. mellifera AmABCG2, A. gambiae AGAP000553, and nine ABCG
proteins of both T. urticae and D. pulex. D. melanogaster scarlet has
two orthologues in A. gambiae and one in T. castaneum, B. mori,
A. mellifera and D. pulex while D. melanogaster brown clusters with
A. gambiae AGAP007655, A. mellifera AmABCG11 and AmABCG12,
and B. mori BGIBMGA002581 and BGIBMGA002712 (Fig. 6, Table 2).
The D. melanogaster ABCG genes white, scarlet and brown are at
present the best studied arthropod ABCG genes. The
D. melanogaster white gene product heterodimerises with gene
products of either D. melanogaster scarlet or brown to give functional transporters that are involved in the uptake of pigment
precursors in cells of Malpighian tubules and developing compound eyes. Mutation in genes encoding these D. melanogaster
ABCG proteins results in flies with eye-color phenotypes (white,
scarlet and brown) distinct from the wild-type brownered-eyecolor phenotype (Ewart et al., 1994; Ewart and Howells, 1998;
Mackenzie et al., 1999). D. melanogaster white mutants with a
white-eye color, were the first genetic Drosophila mutants to be
isolated and marked the beginning of the Drosophila genetics era
(Green, 1996; Morgan, 1910), but the ABC-transporter function has
been attributed to the D. melanogaster white protein only 25 years
ago (Mount, 1987). The role of the T. castaneum and B. mori ortholog
of D. melanogaster white in eye pigmentation has also been
experimentally validated (Broehan et al., 2013; Komoto et al.,
2009). In contrast to insects, both D. pulex and T. urticae have
nine co-orthologues of D. melanogaster white. Dimerization between these co-orthologues in T. urticae might result in translocation of pigment precursors to the eyes, while in D. pulex
dimerization between these co-orthologues or between these coorthologues and the D. pulex ortholog of scarlet, might result in
transport of pigment precursors to the eyes respectively. However,
as many other functions have been attributed to D. melanogaster
white and its insect orthologues (Borycz et al., 2008; Zhang and
Odenwald, 1995; Evans et al., 2008; Komoto et al., 2009), these
98
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T. urticae expanded
ABCG clade
1:1
D. pulex expanded
ABCG clade
Fig. 6. Phylogenetic analysis of ABCG proteins of H. sapiens and seven arthropod species. For procedure and details of phylogenetic analysis see Supplementary Text. For color and
shape codes see Fig. 3. The scale bar represents 0.5 amino-acid substitutions per site. One-to-one ABCG orthologues found in H. sapiens and seven arthropod species are shaded in
grey, while examples of species-specific ABCG expansions are indicated with an arch.
proteins might also have alternative functions. Noteworthy, the
T. castaneum ortholog of D. melanogaster brown, involved in
transport of red pigment (pteridine) precursors, could not be
identified in our phylogenetic analysis. The absence of a clear
D. melanogaster brown ortholog in T. castaneum has also been
indirectly demonstrated by Broehan et al. (2013), since injection of
TcACBG-9A dsRNA (T. castaneum ortholog of D. melanogaster scarlet
which is involved in transport of brown pigment (ommochrome)
precursors) produces a white-eye instead of a red-eye phenotype.
The latter phenotype is in line with Lorenzen et al. (2002) which
suggested that the Tribolium larval eyespot and adult eye are only
pigmented by ommochromes, whose precursors in D. melanogaster
are normally transported by heterodimers of white and scarlet, and
not pteridines, whose precursors in D. melanogaster are transported
by heterodimers of white and brown.
A clear ortholog of D. melanogaster CG3327, also named E23
(Early gene at 23), was identified in A. mellifera (AmABCG1),
T. castaneum (TcABCG-8A), D. pulex (Dappudraft1_347416) and
T. urticae (tetur17g02510) (Fig. 6, Table 2). In D. melanogaster E23 is
a 20-OH ecdysone (20E) induced ABC transporter, probably capable
of regulating the 20E response (Hock et al., 2000). Recently, a
similar function has been attributed to the T. castaneum ortholog of
D. melanogaster E23 (TcABCG-8A) by Broehan et al. (2013). Bryon et
al. (2013) also reported that tetur17g02510 was upregulated in
diapausing T. urticae females compared to non-diapausing females.
As T. urticae most likely uses ponasterone A instead of 20E as a
molting hormone (Grbic et al., 2011), future research should point
out if the T. urticae ortholog of E23 has a similar function as its
arthropod counterparts.
Liu et al. (2011) suggested that BGIBMGA010557 is a B. mori
ortholog of D. melanogaster E23. This orthologous relationship,
however, could not be deduced from our phylogenetic analysis but
instead B. mori BGIBMGA010557 clusters with a group of poorly
characterized insect ABCG proteins. On the other hand, Liu et al.
(2011) showed that B. mori BGIBMGA010557 (together with
BGIBMGA005226,
BGIBMGA005203,
BGIBMGA005202
and
BGIBMGA010555) is expressed in the midgut and is upregulated by
20E during molting and pupation. For these five B. mori ABCG
proteins we could identify a clear ortholog for almost all insect
species included in our analysis (Fig. 6, Table 2). This might suggest
that in insects more than one ABCG protein can be induced by 20E
and consequently that several ABCG proteins might mediate or act
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
downstream of 20E responses. The role of these insect ABCG proteins in insect development has also been shown by Broehan et al.
(2013). Injection of TcABCG-4C (T. castaneum ortholog of B. mori
BGIBMGA005202) dsRNA in T. castaneum larvae resulted in arrested
development and rapid death. In addition to this phenotype,
TcABCG-4C dsRNA injected females laid fewer eggs that failed to
hatch. Cryosections of TcABCG-4C dsRNA injected larvae also
showed that lipids were depleted in both the larval fat body as the
larval epicuticle. Based on these findings, Broehan et al. (2013)
suggested that TcABCG-4C is involved in transporting lipids to the
cuticle and is essential for the formation of a waterproof barrier in
the epicuticle.
Except for T. urticae, we also identified a clear ortholog of human
ABCG1 and ABCG4 in all arthropods included in our analysis (Fig. 6,
Table 2). The function of human ABCG4 is not well-known while
human ABCG1 has a critical role in controlling intracellular sterol
homeostasis in a variety of cells (Kerr et al., 2011; Tarr et al., 2009).
Arthropod orthologues of human ABCG1 have been poorly characterized, with D. melanogaster Atet reported to be expressed in the
trachea (Kuwana et al., 1996). Finally, we also identified a lineagespecific expansion of ABCG proteins in both D. pulex and T. urticae,
but found no arthropod orthologues of human ABCG2. No information is available on the function of these expanded groups in
D. pulex and T. urticae, although in T. urticae some of these
responded to xenobiotic exposure (Dermauw et al., 2013a). Human
ABCG2 is known to transport a whole gamut of substrates,
comprising anticancer drugs (Kerr et al., 2011; Tarr et al., 2009).
Because of this feature of the human ABCG2 proteins, arthropod
ABCGs have also been put forward as potential candidates in
xenobiotic resistance (Buss and Callaghan, 2008; Labbé et al., 2011)
(see Section 4.2).
2.7. The ABCH subfamily
The ABCH subfamily was first identified in D. melanogaster and is
not present in mammals, plants and fungi (Dean et al., 2001;
Kovalchuk and Driessen, 2010; Verrier et al., 2008). Next to arthropods, the zebrafish Danio rerio has also been reported to
contain an ABCH transporter (Popovic et al., 2010) while other
teleost fish lack this type of ABC transporter (Liu et al., 2013). In our
analysis clear orthologous relationships were detected between all
insect ABCH transporters, with each insect species having three
ABCH transporters (Table 2, see Supplementary Fig. 2). The
diamond-back moth Plutella xylostella has been reported to contain
twoefour copies (depending on the presence or absence of allelic
variation in the dataset) of the D. melanogaster ABCH protein
CG9990 (You et al., 2013). In contrast to insects, D. pulex and
T. urticae have a relatively high number of ABCH proteins (15 and
22, respectively). Both D. pulex and T. urticae ABCHs clustered into
separate clades (Table 2, see Supplementary Fig. 2), suggesting that
the diversity of ABCH proteins in these species has been due to
lineage-specific expansions ((Dermauw et al., 2013a), this study).
Similar to ABCG proteins, ABCH proteins are HTs and have a reverse
domain organization, with the NBD at the N-terminal side of the
TMD. Their function, however, is poorly understood.
D. melanogaster CG33970 is 2-fold upregulated after cold hardening
of adult flies (Qin et al., 2005) while six ABCH genes were differentially expressed in diapausing T. urticae females compared to
non-diapausing females (Bryon et al., 2013). D. melanogaster
CG9990 null mutants and an RNAi line that silences D. melanogaster
CG9990 are both lethal (Mummery-Widmer et al., 2009; Zhang
et al., 2009) while TcABCH-9C dsRNA injection in T. castaneum
larvae resulted in similar phenotypes as injection with TcABCG-4C
(see Section 2.6). Of peculiar interest, a P. xylostella ortholog of
D. melanogaster CG9990 ((You et al., 2013), this study) was the most
99
upregulated ABC transporter in two pesticide resistant strains (see
Section 4.2).
3. Inferring the involvement of ABC transporters in
xenobiotic transport: methods and assays
Human ABC transporters such as ABCB1 (P-gp, MDR1), ABCC2
(MRP2) and ABCG2 (BCRP) are extremely well studied for their
importance in the absorption, distribution and excretion of drugs and
therapeutic agents, and consequently a number of tools has been
devised and critically reviewed to study ABC transporters in vivo and
in vitro (Glavinas et al., 2008; Lin and Yamazaki, 2003; Zhang et al.,
2003). Many of the described assays and techniques are directly
applicable in the study of arthropod ABC transporters, in particular
the in vitro assays using cells and cellular membrane preparations.
3.1. Whole cell based assays
Whole cell based assays essentially measure the efflux or
accumulation of compounds in intact cells. Transporters can either
be naturally present, for example when investigating drug resistance cell lines after selection, or can be functionally expressed to
high levels in a variety of cells lines by transfection and transformation methods. The active transport of compounds (substrates)
mediated by ABC transporters out of cells results in detectable
lower rate of substrates accumulation, a lower concentration at the
steady state equilibrium or a faster rate of substrate elimination in
pre-loaded cells. Many variations in assay type and a multitude of
methods to detect and quantify substrate depletion or accumulation have been described. Substrates are often labeled (by a fluorochrome or radiolabel), but direct and high throughput LC-MS
methods have also been devised successfully. This type of assays is
often used in an indirect set-up. For this purpose, well-known and
validated fluorescent dye substrates (such as rhodamine 123, Calcein AM, Hoechst 33342, tetramethylrosamine (TMR)) are used in
competition with candidate substrates (Lebedeva et al., 2011). Cells
expressing ABC proteins are incubated or pre-loaded with these
tractable substrates and eliminated these dyes faster compared to
cells lacking ABC-transporter expression. Candidate substrates that
are transported are identified by their potential to interfere with
dye transport kinetics. In a similar set-up, inhibitors of ABC transporters can be identified.
Cellular assays have also been devised using cell mono-layers, as
this is of particular importance when in vivo pharmacological barriers such as intestinal epithelial cells are taken into account in drug
transport. However, in such experiments cell lines forming a
confluent monolayer with very tight junctions are needed, which
complicates these experiments (Proctor et al., 2010).
3.2. Vesicular assays
Vesicular transport assays are membrane based assays that are
often used to detect and quantify the translocation of molecules by
ABC transporters. The power of the assay is based on the revolutionizing work of Steck and Kant (1972), who describe the preparation of ‘inside out vesicles’, which can be separated from other
membrane fragment such as open membrane lamellae and ‘right
side out’ vesicles by density gradient centrifugation. In these vesicles, the ATP-binding site and substrate binding site of the ABC
transporters are facing outwards (to the assay buffer solution).
Consequently, the transport of substrates into vesicles can be
studied, which is essentially more straightforward compared to
quantifying substrate depletion or deviations from steady state
equilibrium in whole cell based assays. After substrate incubation,
vesicle preparations are isolated from the assay solution by rapid
100
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
filtration using glass fiber filters or nitrocellulose membranes and
the amount of substrates inside vesicles can be quantified by
similar methods as described above (Krumpochova et al., 2012).
This assay has been successfully performed with a number of
membrane preparations from different sources, including insect
cells. Similar to whole cell based assays, vesicular membrane assays
can also be used in an indirect set-up where test compounds
interfere with the uptake of known and easy to track model substrate molecules (Glavinas et al., 2008; Krumpochova et al., 2012;
Lin and Yamazaki, 2003; Zhang et al., 2003).
3.3. ATPase assays
The ATPase assay is another type of in vitro assay that is used
frequently. It is based on the link between ATP hydrolysis and substrate transport. In these assays, the hydrolysis of ATP during substrate transport is followed by quantification of the liberated
inorganic pyrophosphate. Compounds that are transported (or
interfere with transport, such as inhibitors) modulate this ATPase
activity (Litman et al., 1997). The rather low sensitivity of the assay
requires high levels of ABC proteins and thus inside out vesicle
preparations, prepared from transfected or transformed cell lines,
are most frequently used. In this set-up, also open lamellae in vesicular preparations contribute to the measured activity. ATPase
activity is vanadate sensitive, and a fast transport of substrates results in a reproducible and strong increase of ATPase activity. Substrates that are transported slowly, do not generally cause a
detectable increase in ATPase activity, but can be identified when
used as competitive substrates in an assay with fast transported
model substrates (Glavinas et al., 2008; Lin and Yamazaki, 2003;
Zhang et al., 2003). These cell and membrane assays are conceptually simple and interpretation of results is usually straightforward.
3.4. Nucleotide trapping assay
The nucleotide trapping assay is based on the occurrence of a
transition state complex during transport that contains an
occluded nucleotide in the nucleotide-binding site (Urbatsch
et al., 1995). This state can be stabilized by the addition of
vanadate. The amount of nucleotides trapped is proportional to
the rate of transport, and can be quantified by incubations with
32
P labeled azido ATP. This method is especially useful for ABC
transporters where ATPase activity is not stimulated by substrate
addition and where vesicular assays cannot be used (for example
for the so called ABC flippases, like human ABCB4) (Smith et al.,
2000).
3.5. Fluorescence spectroscopic tools
A number of fluorescence spectroscopic tools has also been
described to study ABC transporters and to detect potential substrates. Transporters can be labeled with sulfhydryl-specific fluorescent probes on the two conserved Cys residues in the Walker A
motif of the nucleotide-binding domain. The fluorescent groups are
covalently linked and upon nucleotide binding (ATP, ADP and analogs) a saturable quenching of fluorescence can be detected,
indicative of transport (Sharom et al., 2001; Sreeramulu et al.,
2007). Another technique is based on the conservation of tryptophan (Trp) residues in proteins across the ABC family. Within purified ABC transporters, those Trp residues contribute to
fluorescence which is quenched upon binding of trinitrophenyl
nucleotides. Trp fluorescence is also very sensitive to binding of
substrates, inhibitors and other modulators (Liu et al., 2000;
Sharom et al., 2001).
3.6. In vivo assays
Next to the in vitro assays described above, a number of in vivo
tools has also been used to link ABC-transporter activity to a
certain drug resistant phenotype, such as the generation of
(double) knock-out and mutant mice and rat strains (reviewed in
Lin and Yamazaki, 2003; Zhang et al., 2003). However, given the
lack of orthologues relationships between most ABC proteins
across species (and especially between vertebrates and invertebrates) (Table 2), such models are of limited use as a technique to assign function to arthropod ABC proteins. Unfortunately,
this is also true for most ABC transporters between different insect
species (Table 2) and limits the use of D. melanogaster as an
arthropod genetic model, except for the rare cases where clear
orthologues can be identified. Proof of principle that gene knockout by P-element insertion or by constructing RNAi lines can be
used to study ABC-transporter function in Drosophila has been
recently delivered (Chahine et al., 2012). More generally, reverse
genetics by RNAi (either by injection or oral administration) can
be a powerful tool in assessing ABC function in non-classical
models, as was recently very elegantly demonstrated for
T. castaneum (Broehan et al., 2013).
Last, chemosensitizers that inhibit ABC transporters have been
discovered a long time ago and have found clinical applications in
overcoming multidrug resistance in humans (reviewed in Choi,
2005), but they are also potentially useful experimental tools in
linking a resistant phenotype to the action of an ABC transporters. Chemosensitizers or synergists are administered
together with a toxicant or drug, and restore the toxicity or efficacy both in vivo and in vitro. One of the first compounds
discovered was verapamil, a phenylalkylamine Ca2þ and Kþ
channel blocker with wide clinical use. Verapamil was first
shown to reverse multidrug resistance in neoplastic cells and
chloroquine resistant Plasmodium falciparum (Martin et al., 1987).
It has since then been widely used in the study of mainly human
P-gp where it acts as a competitive substrate inhibitor, but it is
less potent in inhibiting human MRP1, although it stimulates
MRP1 mediated glutathione transport (Loe et al., 2000). Although
verapamil has been used as a synergist in insecticide resistance,
the efficiency and specificity of verapamil as a competing substrate or inhibitor in distantly related arthropod ABC transporters
is still largely unknown.
3.7. Genetic mapping
The association between ABC transporters and xenobiotic
resistance is often based on synergism of toxicity, combined with
the observation of increased expression (see Table 3). However,
single point mutations in a transmembrane domain can modulate
transport and selectivity of ABC transporters. A SerePhe substitution for example drastically altered the overall degree of drug
resistance and specificity conveyed by mdr1 and mdr3 in mice
(Gros et al., 1991). If synergism points out the role of ABC transporters in resistance, such point mutations cannot be detected by
expression analysis and unbiased approaches are needed. The
availability of genetic mapping tools in Drosophila and some lepidopteran species including B. mori, have proven powerful in finding
resistance loci associated with ABC transporters (Begun and
Whitley, 2000; Heckel, 2012) (see also Section 4.4.2). Until
recently, such approaches were limited to arthropods where classical genetic resources are available, such as genetic linkage maps.
However, it was recently shown how classical bulk segregant
analysis can be modified to allow ultra high resolution mapping in
any arthropods species for which classical genetic resources are
sparse, but a genome sequence is available ((Van Leeuwen et al.,
Table 3
An overview of reported associations between ABC transporters and insecticide/acaricide transport and resistance.
E/syna
Pesticide
Species
ABC subfamily
Carbamates
Pirimicarb
M. persicae
ABCG/ABCHb
Thiodicarb
H. virescens
ABCB FT (P-gp)
v
v
Abamectin
Abamectin
R. microplus
D. melanogaster
?
ABCB FT (P-gp)
v
v
v
v
Avermectin
D. melanogaster
ABCB FT (P-gp)
v
v
Emamectin
benzoate
Ivermectin
L. salmonis
ABCB FT (P-gp)/?
v
v
P. humanus
humanus
ABCB_unknown/
ABCC/ABCG
v
v
C. pipiens
R. microplus
?
ABCB HT
v
v
v
S. scabiei
C. riparius
ABCC
?
v
Moxidectin
Acetamiprid
R. microplus
A. mellifera
?
?
Imidacloprid
A. mellifera
Thiacloprid
Macrocyclic
lactones
Neonicotinoids
Organochlorines
Evidence
Reference
Silva et al., 2012
v
v
Increased expression (microarray) in adult aphids upon
exposure to pirimicarb
Quinidine synergism (synergism ratio of 12.5 and 1.8 in
resistant and susceptible strain, respectively); increased P-gp
expression in thiodicarb resistant larvae (Western Blot)
Cyclosporin A and MK571 synergism
Increased P-gp expression (Western Blot) and increased P-gp
related vanadate-sensitive ATPase activity in abamectin
resistant strain; verapamil synergism; higher P-gp expression
(Western Blot, C219 antibodies) in the humoral/CNS interface in
abamectin resistant strain
Induction of P-gP expression (Western Blot) and increased P-gp
related ATPase activity upon exposure of S2 cells to avermectin;
avermectin enhances cellular efflux of rhodamine 123 from S2
cells
Verapamil synergism of emamectin benzoate (EMB); induction
of P-gp expression (qPCR) upon exposure to EMB
Increased expression (qPCR) of six ABC transporters in female
lice 4 h post-ivermectin exposure by both immersion and
contact applications; injection of PhABCC4 dsRNA (P. humanus
ortholog of CG6214) resulted in increased sensitivity to
ivermectin; verapamil synergism
Verapamil synergism in 4th instar larvae
Cyclosporin A and MK571 synergism, increased expression
(qPCR) of RmABCB10 in ivermectin (IVM) resistant females and
RmABCB10 transcription was upregulated in IVM-resistant
females in response to IVM feeding
Increased expression (qPCR) upon exposure to ivermectin
Verapamil and cyclosporin synergism; ivermectin stimulates
ATPase activity in homogenate fractions of C. riparius larvae
Cyclosporin A and MK571 synergism
Verapamil synergism
?
v
Verapamil synergism
A. mellifera
?
v
Verapamil synergism
Thiametoxam
B. tabaci
ABCG
DDT
A. arabiensis
D. melanogaster
ABCB FT (P-gp)/ABCG
ABCB FT (P-gp)/ABCC
D. melanogaster
ABCG
C. pipiens
H armigera
?
ABCB FT (P-gp)
v
R. microplus
P. xylostella
?
ABCA/ABCC/ABCG/
ABCH/ABCF
?
v
Endosulfan
Organophosphates
Chlorpyrifos
Coumaphos
A. mellifera
E/exp
E/oth
v
v
v
v
v
v
v
v
v
v
v
v
v
Increased expression (microarray) in thiamethoxam resistant
strain
Increased expression (qPCR) in a DDT-resistant strain
Increased expression (qPCR) in a DDT-resistant strain (91-R),
RNAi results in 91-R flies being more susceptible to DDT,
verapamil synergism (synergism ratio of 10 and 1.1 in resistant
and susceptible strain, respectively)
Increased expression (microarray) in DDT-resistant strains (91R and Wisconsin)
Verapamil synergism in 4th instar larvae
Stimulation of ATPase activity; TMR transport declined with
increasing concentrations of insecticide; Trp quenching
Cyclosporin A and MK571 synergism
Increased expression in chlorpyrifos resistant strain (RNA-seq)
Verapamil and oxytetracycline synergism
Lanning et al., 1996a,b
Pohl et al., 2012
Luo et al., 2013a
Luo et al., 2013b
Igboeli et al., 2012; Tribble
et al., 2007
Yoon et al., 2011
Buss et al., 2002
Pohl et al., 2011, 2012
Mounsey et al., 2007
Podsiadlowski et al., 1998
Pohl et al., 2012
Hawthorne and Dively
2011
Hawthorne and Dively
2011
Hawthorne and Dively
2011
Yang et al., 2013a,b
Jones et al., 2012
Strycharz 2010, Strycharz
et al., 2013
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
Pesticide class
Pedra et al., 2004
Buss et al., 2002
Srinivas et al., 2005; Aurade
et al., 2006, 2010b
Pohl et al., 2012
You et al., 2013
Hawthorne and Dively
2011
(continued on next page)
101
Pesticide class
Pesticide
Species
ABC subfamily
Ethylparaoxon
H. armigera
Methylparathion
E/oth
Evidence
Reference
ABCB FT (P-gp)
v
Aurade et al., 2010b
H. armigera
ABCB FT (P-gp)
v
Monocrotophos
H. armigera
ABCB FT (P-gp)
v
Temephos
A. caspius
A. aegypti
?
ABCB FT (P-gp)
Triazophos
N. lugens
ABCC
Cyfluthrin
C. lectularius
?
Cypermethrin
C. pipiens
H. armigera
?
ABCB FT (P-gp)
C. lectularius
?
v
A. gambiae
T. ni
ABCC
ABCB FT (P-gp)
v
v
C. lectularius
?
v
Fenvalerate
H. armigera
?
Stimulation of ATPase activity; TMR transport declined with
increasing concentrations of insecticide; Trp quenching
Stimulation of ATPase activity; TMR transport declined with
increasing concentrations of insecticide; Trp quenching
Stimulation of ATPase activity; TMR transport declined with
increasing concentrations of insecticide; Trp quenching
Verapamil synergism (by factor 3.5)
Increased expression (qPCR) in larvae upon exposure to
temephos; RNAi of P-gp resulted in a significant increase in
temephos toxicity to larvae, verapamil synergism
Increased expression (SSH and qPCR) in nymphs exposed to
sublethal concentrations of triazophos
RNAi resulted in reduced resistance to beta-cyfluthrin in a
deltamethrin resistant strain
Verapamil synergism in 4th instar larva
Stimulation of ATPase activity; TMR transport declined with
increasing concentrations of insecticide; Trp quenching
Increased expression of eight ABC transporters (RNA-seq and
qPCR) in a deltamethrin resistant strain; an ABCG transporter
was the highest upregulated ABC-transporter
Increased expression (RNA-seq) in deltamethrin resistant strain
Limited increased expression (qPCR) in insects fed on a
deltamethrin containing artificial diet
Increased expression of four ABC transporters in a deltamethrin
resistant strain (RNA-seq and qPCR)
Stimulation of ATPase activity
Fluvalinate
A. mellifera
?
Pyrethroids
A. aegypti
ABCB HT/ABCC/ABCG
Diflubenzuron
Cry1A toxin
A. caspius
Lepidoptera
?
ABCA/ABCB/ABCC/ABCD/ABCGb
Fipronil
Multiresistant
Multiresistant
P. xylostella
T. urticae
H. armigera
ABCA/ABCC/ABCG/ABCH
ABCC/ABCH
ABCB FT (P-gp)
Deltamethrin
Diverse
a
b
E/syna
v
v
E/exp
v
v
v
v
v
v
v
v
Verapamil and oxytetracycline synergism
v
v
v
v
v
v
v
Increased expression (microarray and qPCR) in pyrethroid
resistant strains; ABCB4 gene has a higher copy number in
pyrethroid resistant strains
Verapamil synergism (by factor 16.4)
A deletion or insertion in the ABCC2 gene is correlated with both
higher Bt resistance levels and loss Cry1A binding to midgut
membranes; eight ABC genes were differentially expressed
between Cry1Ac-resistant and -susceptible larvae of P. xylostella
Increased expression (RNA-seq) in fipronil resistant strain
Increased expression (microarray) in two multiresistant strains
Increased P-gp expression (Western Blot) in multiresistant
larvae
Type of evidence was divided into three categories: E/syn, ABC inhibitor synergism; E/exp, increased expression of ABC genes; E/oth, other type of evidence.
ABC subfamily was determined in this study and based on the top BLASTp hits in the NCBI non redundant protein database.
Srinivas et al., 2005; Aurade
et al., 2006, 2010b
Srinivas et al., 2005; Aurade
et al., 2006, 2010b
Porretta et al., 2008
Figueira-Mansur et al., 2013
Bao et al., 2010
Zhu et al., 2013
Buss et al., 2002
Srinivas et al., 2005; Aurade
et al., 2006, 2010b
Mamidala et al., 2012
Bonizzoni et al., 2012
Simmons et al., 2013
Zhu et al., 2013
Srinivas et al., 2005, Aurade
et al., 2006
Hawthorne and Dively
2011
Bariami et al., 2012
Porretta et al., 2008
Heckel 2012, Lei et al., 2013
You et al., 2013
Dermauw et al., 2013a,b
Srinivas et al., 2004
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
Pyrethroids
102
Table 3 (continued )
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
2012), see also Section 4.4.1). This technique relies on next generation re-sequencing of selected populations and seems especially
timely with thousands of arthropods genomes being sequenced
currently (i5K-Consortium, 2013).
4. Arthropod ABC transporters in xenobiotic transport and
resistance
4.1. ABC transporters as players in detoxification pathways
Arthropods have developed a variety of mechanisms to cope
with toxic compounds. These mechanisms are generally classified
in mechanisms of decreased response to toxic substances, often
referred to as pharmacodynamic mechanisms (interaction of toxic
substances with their target site) and mechanisms of decreased
exposure also called pharmacokinetic mechanisms, which include
altered penetration, distribution, metabolism and excretion of the
toxic substances. In most cases, high levels of toxicant resistance
in arthropods is due to alterations in the sensitivity of the target
site caused by point mutations (pharmacodynamic mechanism),
or the efficient metabolism or sequestration of the toxic compound (pharmacokinetic mechanism). The overall detoxification
process can be divided into three phases, phase IeIII. In Phase I
reactions, enzymes such as P450 monooxygenases and carboxylesterases activate the toxic molecules and make them more
reactive and water soluble by introducing hydroxyl, carboxyl and
amino groups, which often results in an increased conjugation
with endogenous molecules by phase II enzymes such as GSTs or
UPD-glycosyltransferases. Finally, in phase III the polar compounds or conjugates can be transported out of the cell by cellular
transporters and it is in this phase that ABC transporters play an
important role. In some cases cellular transporters such as ABCs
directly and efficiently transport toxicants out of the cell without
enzymatic modifications, thereby preventing the exertion of
toxicity in the cell or organism (sometimes also referred to as
“Phase 0”) (Despres et al., 2007; Kennedy and Tierney, 2013). ABC
transporters have been implicated in resistant phenotypes in
many animals, including both vertebrates and invertebrates, and
are in the latter group probably best studied in nematodes
(Ardelli, 2013; Lespine et al., 2012). Fewer cases have been reported in arthropod resistance, although the interest in this family
is steadily growing, especially with the availability of more
sequence data and the availability of genetic tools in organisms
other than Drosophila. A survey of cases where the involvement of
ABC transporters in arthropod resistance is suggested is presented
in Table 3 and discussed below in the light of the strength of the
evidence.
4.2. ABC transporters associated with insecticide transport and
resistance
ABC transporters have been associated with transport of and/or
resistance to 27 different insecticides/acaricides, belonging to nine
distinct chemical classes and several mode of action groups: carbamates, macrocyclic lactones, neonicotinoids, organophosphates,
pyrethroids, cyclodienes, benzoylureas, phenylpyrazoles, and DDT
(reviewed in (Buss and Callaghan, 2008) and updated in this study,
Table 3). ABCB FTs/P-gps followed by ABCCs and ABCGs were the
most reported ABC subfamilies to be involved in pesticide transport
and/or resistance. The evidence pointing toward a role in transport
or resistance is most often based on the quantification of transcript
or protein levels, and by synergism studies using ABC inhibitors
that are poorly characterized in arthropods. In some cases, ATPase
assays revealed the simulation of ATP hydrolysis upon contact with
insecticides, which is considered indicative of transport. Vesicular
103
transport assays that directly quantify insecticide transport have
not been reported until present, although many of the methods
using Sf9 cells to recombinantly express ABC transporters and
prepare vesicles are directly relevant for arthropod studies (see
Section 3).
4.2.1. Carbamates and organophosphates
Lanning et al. (1996a) showed for the first time a link between
an arthropod ABC transporter and resistance against an insecticide, the carbamate thiodicarb (Table 3). Using human monoclonal
P-gp antibodies, it was demonstrated that ABCB FTs/P-gps were
highly expressed in the cuticle of larvae of the tobacco budworm
Heliothis virescens and that P-gps were overexpressed in thiodicarb resistant larvae of this species. Furthermore it was found
that inhibition of ABCB FTs/P-gps with quinidine decreased the
LD50 of thiodicarb for resistant H. virescens larvae by a factor of
12.5, while that of susceptible larvae only decreased by a factor of
1.8. In a follow-up paper, it was further shown that thiodicarb
treatment in combination with quinidine increased C14 labeled
thiodicarb accumulation 2-e3-fold as compared to thiodicarb
treatment alone (Lanning et al., 1996b). Other reports have associated ABC transporters with carbamate resistance, but without
further characterization. Genes encoding an ABCG and ABCH
transporter were among the 183 upregulated genes in adults of
the green peach aphid Myzus persicae exposed to pirimicarb. The
upregulation of these ABC transporters was however not validated
by qRT-PCR and no synergism tests were performed (Silva et al.,
2012). Several studies have also reported on the ABCB FT/P-gpmediated efflux of organophosphates (OPs) (Aurade et al., 2006,
2010b; Srinivas et al., 2005). Aurade et al. (2006, 2010b) found
that ethylparaoxon, monocrotophos and methylparathion at a
concentration of 100, 100 and 50 mM stimulated purified P-gpmediated ATPase activity by 20%, 25% and 30% in the tobacco
budworm Helicoverpa armigera, respectively, suggestive of P-gp
meditated transfer of these compounds (see Section 3). In this
study, it was also shown by Trp fluorescence quenching that
ethylparaoxon and methylparathion interact with H. armigera Pgp with relatively high affinity (Kd values of 25 and 50 mM). Finally,
the authors also succeeded in reconstituting H. armigera P-gp into
proteoliposomes and showed that ethylparaoxon, monocrotophos
and methylparathion inhibit P-gp-mediated transport of the
fluorescent reporter dye tetramethylrosamine (TMR) in a concentration dependent manner. Whether OPs compete with TMR
as substrates or act as non-competitive inhibitors was not
demonstrated. Interestingly, in a follow-up study it was shown
that cholesterol could increase the ethylparaoxon or methylparathion stimulated H. armigera P-gp ATPase activity in reconstituted proteoliposomes by 10e20% (Aurade et al., 2012). Another
example of interaction between ABCB FTs/P-gps and OPs was
recently documented for the mosquito Aedes aegypti (FigueiraMansur et al., 2013). An 8-fold increase of an A. aegypti ABCB
FT/P-gp gene was observed in A. aegypti larvae 48 h after treatment with 0.0007 mg/L temephos. Furthermore, temephos
treatment of A. aegypti larvae in combination with a sublethal
dose of verapamil resulted in a decrease of the LC50 of temephos
by a factor of 1.24 and temephos toxicity increased by 57% in P-gp
silenced larvae. Similarly, Porretta et al. (2008) showed that P-gp
inhibition by verapamil in the closely related Aedes caspius
resulted in LD50 reduction by a factor of 3.5. These results suggest
a role for P-gp in efflux of temephos in Aedes sp., but whether
these P-gps are also involved in temephos resistance in these
species remains to be determined. Finally, a link between ABC
transporters in OP efflux has also been suggested for two other
arthropod species. In a chlorpyrifos resistant strain of P. xylostella
ABCA/C/G/H and F transporter(s) were more highly expressed
104
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
compared to a susceptible strain as determined by RNA-seq (You
et al., 2013) while in the brown planthopper Nilaparvata lugens
SSH and qRT-PCR revealed a significant increase of an ABCC
transporter in nymphs exposed to sublethal concentrations of
triazophos (Bao et al., 2010). However, whether increased
expression is related to transport and resistance has not been
demonstrated.
4.2.2. Macrocyclic lactones
Resistance to macrocyclic lactones (MLs), such as abamectins,
ivermectin and moxidectins, in arthropods and nematodes is in
most cases linked to several genetic factors. Both mutations in
target-site genes (e.g. in glutamate-gate chloride channels), as well
as quantitative changes in the expression of detoxification enzymes
and ABC transporters have been reported to be involved in ML
resistance (Dermauw et al., 2012; Pu et al., 2010; Wolstenholme and
Kaplan, 2012) (Table 3). In mammals, it has been clearly demonstrated that ivermectin is a substrate of P-gps as human MDR-cells
accumulate less 3H ivermectin (Pouliot et al., 1997; Schinkel et al.,
1994). The role of ABC transporters in ML transport and/or resistance in nematodes has been thoroughly characterized, with the
majority of MLs stimulating ABCB FT/P-gp-mediated efflux of the
reported substrate Rhodamine 123 (Kerboeuf and Guégnard, 2011).
In contrast, the involvement of ABCB FT/P-gps in ML transport and/
or resistance in arthropods is not well documented. Podsiadlowski
et al. (1998) reported for the first time the synergistic action of
ABCB FT/P-gp-transporter inhibitors on the toxicity of a ML in an
insect species. Sublethal concentrations of verapamil (30 mM) or
cyclosporin A (3 mM) reduced the LC50 of ivermectin for the midge
Chironomius riparius with a factor 2.7 and 2.8, respectively.
Furthermore, it was shown that P-gps were located in the anal
papillae of C. riparius larvae and that ivermectin at a concentration
of 0.1e1 mM stimulated ABCB FT/P-gp-mediated ATPase activity in
homogenate fractions of C. riparius larvae. Similar to C. riparius,
100 mM verapamil has also been shown to synergistically enhance
the toxicity of ivermectin in the mosquito Culex pipiens. Comparison
of synergism ratios between resistant and susceptible strains did not
reveal any differences, indicating that ABC transporters did not
contribute to resistance (Buss et al., 2002). In 2012, Igboeli et al.
found that concurrent exposure of the sea lice Lepeophtheirus salmonis (Crustacea: Copepoda) to 10 mM verapamil and 300 mg/L
emamectin benzoate (EMB) caused significantly higher mortality
compared to exposure of L. salmonis to EMB alone. In addition, ABCB
FT/P-gp expression was induced in adult sea lice exposed to 1 mg/L
EMB. Induced expression of ABC transporters upon exposure to
ivermectin was also observed in females of the body louse
P. humanus and the scabies mite Sarcoptes scabiei (Mounsey et al.,
2007, 2006; Yoon et al., 2011). For the body louse, it was also
shown that injection of dsRNA of P. humanus ABCC4 (the most highly
expressed ABC gene after exposure to ivermectin) into female lice
increased their sensitivity to ivermectin with 20e30% compared to
buffer injected lice (Yoon et al., 2011). However, the low number of
lice injected (n ¼ 15) and the discrepancy between assessing toxicity
(12 h post injection) and the reduction in transcript levels (48 h post
injection) require further validation.
The role of ABC transporters in ML resistance has at present only
been thoroughly investigated in two arthropod species, the cattle
tick Rhipicephalus microplus and the fruit fly D. melanogaster (Luo
et al., 2013a; Pohl et al., 2011, 2012). Pretreatment with 5 mM of
ABC-transporter inhibitors cyclosporin A and MK571 reduced the
LC50 of ivermectin, abamectin and moxidectin in field collected
resistant R. microplus larvae with a factor of 1.2e1.5, while toxicity
was not affected in a susceptible laboratory strain. Resistant
R. microplus females showed a higher constitutive expression of an
ABCB HT gene (RmABCB10) compared to the susceptible strain.
Furthermore, exposure of tick females to ivermectin resulted in an
upregulation of RmABCC10 in the resistant strain while transcript
levels did not significantly change in females of the susceptible
strain (Pohl et al., 2011, 2012). ABCB HTs are located in the mitochondria and are believed to play a role in iron homeostasis and
protection against oxidative stress. However, contrary to ABCB FTs
they have rarely been linked with xenobiotic resistance (Elliott and
Ai-Hajj, 2009; Zutz et al., 2009).
The most convincing evidence for the involvement of ABC
transporters in ML resistance has been published recently by Luo
et al. (2013a,b). It was shown that avermectin stimulated P-gp
ATPase activity and efflux of Rhodamine 123 in Drosophila derived
S2 cells and that ABCB FTs/P-gp expression was induced by avermectin in a dose- and time-dependent manner. Next, it was shown
that avermectin induced P-gp expression was caused by an elevation of intracellular calcium levels, both directly by activating calcium ion channels, and indirectly by activating chloride channels.
Of particular interest, a well-known chloride channel agonist
pentobarbital enhanced ivermectin induced P-gp expression in a
concentration dependent manner. Induced expression by the
closely related chemical structure phenobarbital of two ABCB FTs/
P-gps and an ABCC transporter (CG6214) in D. melanogaster flies
was recently also demonstrated by Misra et al. (2011). In a followup study of Luo et al. (2013a,b), the role of ABCB FTs/P-gps in abamectin resistance in D. melanogaster was investigated using a 7-fold
abamectin resistant strain that was selected from a susceptible
laboratory strain (Canton-S). ABCB FT/P-gp expression and P-gp
ATPase activity in the resistant strain was increased 2-e3-fold
compared to the susceptible strain. Treatment with verapamil
reduced P-gp ATPase activity in the adults of the resistant strain by
89% compared to 31% in the susceptible strain, while abamectin
LD100 values for resistant and susceptible larvae were reduced by
84% and 33% in the same treatment. Through confocal microscopy
and immunofluorescent staining with human anti-P-gp C219 antibodies, it was also observed that P-gp expression in the bloode
brain barrier (BBB) in heads from adults of the resistant strain was
higher than that of the susceptible strain. Finally, avermectin
treatment resulted in an increased P-gp expression in the BBB of
the resistant strain, compared to a decline in the susceptible strain.
Taken all this data into account, it is likely that P-gps are involved in
(low-levels) of abamectin resistance in D. melanogaster.
4.2.3. Organochlorines
Associations between DDT resistance and ABC transporters are
scarce. An ABCG transporter gene was significantly overtranscribed
in resistant strains of D. melanogaster and Anopheles arabiensis
(Jones et al., 2012; Pedra et al., 2004). Recently, Strycharz et al.
(2013) determined that D. melanogaster flies from a highly resistant DDT strain (91-R) excreted w4-fold more DDT and its metabolites than flies from the susceptible strain. A topical pretreatment
with verapamil reduced the LC50 value of the resistant flies by 10fold while no effect on the LC50 of susceptible flies was observed.
In a preliminary report, Strycharz et al. also showed that two ABCB
(mdr50 and mdr65) and one ABCC (dMRP/CG6214) transporter
gene(s) were overexpressed (w1.3-fold) in the resistant
D. melanogaster strain compared to the susceptible strain. RNAi
knockdown of these genes in resistant flies also increased their
susceptibility to DDT (Strycharz, 2010; Strycharz et al., 2010). Taken
together, ABC transporters are most likely involved in the increased
excretion of DDT in D. melanogaster (91-R) resistant flies, hereby
lowering the concentration of DDT reaching the target. The role of
ABC transporters in transport/resistance of other organochlorines
has only been marginally studied. Verapamil increased toxicity to
endosulfan in 4th instar larvae of the mosquito C. pipiens (Buss
et al., 2002) while endosulfan stimulated ATPase activity and
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
105
Table 4
An overview of reported associations between ABC transporters and allelochemical transport.
Allelochemical
Species
ABC subfamily
Evidence
Reference
?
T. urticae
ABCA/ABCC/ABCE/ABCG/ABCH
?
ABCGa
Grbi
c et al., 2011,
Dermauw et al., 2013a
Robert et al., 2013
Cardenolide
Dendroctonus
ponderosae
M. sexta
Colchicine
D. melanogaster
ABCB FT(P-gp)
Curcumin
H. armigera
ABCB FT(P-gp)
Flavonoids
H. armigera
ABCB FT(P-gp)
Nicotine
M. sexta
?
Vinblastine
M. sexta
?
Increased expression (RNA-seq) upon transfer from preferred host
(bean) to a more challenging host (tomato)
Increased expression (RNA-seq) in adults that were fed on lodgepole
pine host tree tissues compared to adults that were starved
Verapamil inhibits P-gp-mediated efflux of digoxin at the perineurium
of nerve cords
Expression of mdr49 in brain and gut of larvae is enhanced upon
colchicine feeding (as determined by Northern blotting);
Drosophila mdr49 deletion mutants showed increased sensitivity to
colchicine
Curcumin mixture inhibited the activity of H. armigera P-gp ATPase by
80e90% at 100 mM concentration, addition of the curcuminoid mixture
collapsed the tetramethylrosamine concentration gradient generated
by H. armigera P-gp ATPase
Flavonoids (morin, quercetin and phloroglucinol) inhibited H. armigera
P-gp ATPase activity and bind to H. armigera P-gp with high affinity
Verapamil inhibits accumulation of nicotine in the lumen of Malpighian
tubules; ABC transporters were downregulated (microarray) in M. sexta
larvae fed on tobacco plants transformed to silence nicotine
biosynthesis compared to larvae that fed on wild-type tobacco plants
Verapamil inhibits accumulation of vinblastine in the lumen of
Malpighian tubules
a
?
Petschenka et al., 2013
Tapadia and Lakhotia
2005; Wu et al., 1991
Aurade et al., 2010a
Aurade et al., 2011
Murray et al., 1994,
Gaertner et al., 1998,
Govind et al., 2010
Gaertner et al., 1998
ABC subfamily was determined in this study and based on the top BLASTp hit in the NCBI non redundant protein database.
inhibited transport of tetramethylrosamine in H. armigera (Aurade
et al., 2006, 2010b; Srinivas et al., 2005). Sreeramulu et al. (2007)
observed similar effects of endosulfan on ABCB FTs/P-gps of human multidrug resistant cells.
4.2.4. Pyrethroids
For three different insect species, pyrethroid resistance has also
been associated with the upregulation of ABC-transporter genes
(Bariami et al., 2012; Bonizzoni et al., 2012; Mamidala et al., 2012;
Zhu et al., 2013). Bariami et al. (2012) showed by microarray experiments that a P450 monooxygenase (Cyp9J26) and an ABCB
transporter gene (ABCB4), were upregulated in a pyrethroid resistant strain of the dengue vector A. aegypti. Furthermore, quantitative PCR revealed that increased transcript levels of these A. aegypti
genes were associated with a w7-fold increase in gene copy
number. A. aegypti ABCB4 belongs to the ABCB half transporters,
which have only very rarely been associated with xenobiotic
resistance but are thought to be mitochondrial ABC transporters
with roles in the biogenesis of cytosolic ironesulfur clusters, heme
biosynthesis, iron homeostasis and protection against oxidative
stress (Elliott and Ai-Hajj, 2009; Zutz et al., 2009). In another
transcriptome study, RNA-seq analysis and qPCR revealed that four
ABC transporters (ABC8e11) were upregulated in a deltamethrin
resistant strain of Cimex lectularius when compared to a susceptible
strain. In addition, RNAi mediated knockdown of two of these ABC
genes (ABC8 and 9) resulted in an increased susceptibility to betacyfluthrin in deltamethrin resistant female bed bugs (Zhu et al.,
2013). However, in some species there is little to no relationship
between pyrethroid exposure and increased expression of ABCB
FTs/P-gp as recently shown by Simmons et al. (2013) for larvae of
T. ni. Besides alterations in ABCB FT/P-gp expression levels, pyrethroid stimulation of ATPase activity and verapamil synergism of
pyrethroid toxicity has been reported for several insect species
(Aurade et al., 2006, 2010b; Buss et al., 2002; Hawthorne and
Dively, 2011; Srinivas et al., 2005).
4.2.5. Neonicotinoids, diflubenzuron and fipronil
Only three studies have associated ABC transporters with
neonicotinoid transport and/or resistance. Microarray gene
expression studies revealed that an ABCG transporter gene was
upregulated in adult Bemisia tabaci flies of a thiamethoxam recent
strain (Yang et al., 2013a,b). Hawthorne and Dively (2011), on the
other hand, showed that verapamil synergized honeybee mortality
to three neonicotinoids: acetamiprid, imidacloprid and thiacloprid
but necessary experiments to link this finding to neonicotinoid
resistance were not performed. Verapamil has also been shown to
enhance the toxicity of diflubenzuron, a chitin synthesis inhibitor,
by a factor of 16.4 in 4th instar larvae of the mosquito A. caspius
(Porretta et al., 2008). Finally, several ABC transporters (belonging
to five different ABC subfamilies) were upregulated in a P. xylostella
strain resistant against fipronil, an insecticide belonging to the
chemical class of phenylpyrazoles (You et al., 2013).
4.3. ABC transporters involved in transport of plant secondary
compounds (allelochemicals)
For several plant pathogenic fungi, it has been shown that ABC
transporters mediate cellular tolerance to natural toxic plant
compounds and/or virulence (Coleman and Mylonakis, 2009;
Gardiner et al., 2013). Compared to fungi however, transport of
plant allelochemicals by arthropod ABC transporters has only been
marginally studied (Sorensen and Dearing, 2006) (Table 4). About
15 years ago it was shown that verapamil inhibited transport of
nicotine and vinblastine to the lumen of Malpighian tubules of the
tobacco hornworm Manduca sexta, indicating the involvement of an
ABCB FT/P-gp transporter in the excretion of alkaloids in this lepidopteran species. A similar ABC transporter for nicotine excretion
was also suggested to regulate transport over the BBB of the same
insect species (Gaertner et al., 1998; Murray et al., 1994). In line
with the latter findings, Govind et al. (2010) reported that ABC
transporters were downregulated in M. sexta larvae that fed on
tobacco plants that were transformed to silence nicotine biosynthesis compared to larvae that fed on wild-type tobacco plants.
Arthropod ABCB FT/P-gps have also been reported to be involved in
colchicine transport, a secondary metabolite from plants of the
genus Colchicum (autumn crocus) and a well-known human P-gp
inhibitor (Seelig and Landwojtowicz, 2000). D. melanogaster mdr49
deletion mutants showed increased sensitivity to colchicine (Wu
et al., 1991) while Tapadia and Lakhotia (2005) showed that
colchicine induced mdr49 expression in D. melanogaster larvae.
106
W. Dermauw, T. Van Leeuwen / Insect Biochemistry and Molecular Biology 45 (2014) 89e110
More recently, the group of Aurade et al. (2010a, 2011) found that
several secondary plant compounds are potent inhibitors of P-gp
activity. A curcumin mixture, extracted from Turmeric (Curcuma
longa) powder, inhibited H. armigera P-gp ATPase activity by 90%
and inhibited TMR transport in proteoliposomes containing
H. armigera P-gp. In addition, it was found that H. armigera P-gp also
showed large tryptophan fluorescence quenching in the presence
of 10 mM of this curcuminoid mixture, suggesting that curcuminoids may bind with ATP-binding sites and steroid acting regions.
Similar effects were also reported for the flavonoids morin, quercetin, and phloroglucinol (Aurade et al., 2011). Interestingly,
H. armigera resistant larvae fed with diet supplemented with ethylparaoxon or cypermethrin (100 mM g1 diet), insecticides known
to stimulate H. armigera P-gp ATPase activity (see Sections 4.2.1 and
4.2.4), and 500 mM morin or quercetin, showed a significant
reduction in weight gain and survival rate compared to H. armigera
larvae that were fed with diet supplemented with either ethylparaoxon, cypermethrin, morin or quercetin alone (Aurade et al.,
2011). Furthermore, Petschenka et al. (2013) showed by immunostaining that an ABCB FT/P-gp transporter is expressed in the
perineurium of the oleander hawkmoth, Daphnis nerii, and that
verapamil suppressed the efflux of the cardenolide digoxin from
the perineurium in both D. nerii and M. sexta, suggesting that a
ABCB FT/P-gp transporter mediates the efflux of digoxin. Finally, it
was found for the mountain pine beetle Dendroctonus ponderosae
that an ABCG transporter was upregulated in adults fed on lodgepole pine host tree tissue compared to adults that were starved
(Robert et al., 2013).
4.4. ABC transporters as targets for insecticides
4.4.1. SUR and benzoylphenylureas
Next to the transport of insecticides and allelochemicals, ABC
transporters have also been studied as target sites for insecticides.
Probably the most well-known ABC transporter in arthropod toxicology is the sulfonylurea receptor (SUR, ABCC family, see Section
2.3 for detailed description), a conserved protein with orthologues
in many invertebrate and vertebrate species including humans. In
arthropods, SUR has been put forward as the target site of benzoylphenylureas (BPUs), a class of selective insecticides that inhibit
chitin synthesis in insects and mites. The interference of BPUs with
chitin synthesis was uncovered soon after their development and
commercialization, as the lead compound diflubenzuron was
shown to inhibit in vivo the incorporation of UDP-N-acetylglucosamine (the building blocks of the chitin polymer) into chitin (Post
and Vincent, 1973). However, all further attempts to document a
direct biochemical effect on any of the enzymes in the chitin synthesis pathway failed in the mid 1980s (Matsumura, 2010). The
group of Matsumura has pioneered BPU mode of action studies and
put forward the hypothesis of an interaction between BPUs and
SUR (reviewed in Matsumura, 2010). However, biochemical evidence of such interaction is still missing and counter arguments for
SUR as the target site of BPUs have recently been accumulating.
First, although Drosophila is very sensitive to diflubenzuron, the
SUR mRNA is not detected in the epidermis where chitin synthesis
takes place (Gangishetti et al., 2009). More convincingly, by taking
advantage of genetic resources in Drosophila, Meyer et al. (2012)
show that SUR is dispensable for chitin synthesis in Drosophila, as
larvae with completely eliminated SUR function display a normal
cuticular architecture. In addition, although the beetle T. castaneum
is a model insect for studying the effects of chitin synthesis inhibitors such as diflubenzuron (Merzendorfer et al., 2012), a SUR
homolog has not been detected in this species or any other coleopteran ((Broehan et al., 2013), see also Section 2.3). Furthermore,
RNAi knockdown of ABC transporters in T. castaneum did also not
result in any phenotype with cuticle defects (Broehan et al., 2013).
Together, these studies would argue against SUR as the target site of
BPU. In contrast, the processive glycosyltransferase chitin synthase
1 (CHS1) has been shown to be the target site of etoxazole in the
mite T. urticae. Although etoxazole is a diphenyl oxazoline acaricide
that belongs to a different chemical family, the symptoms of
poisoning strongly resemble those of BPUs in insects. Van Leeuwen
et al. (2012) developed a population-level bulk segregant mapping
method to map a monogenic, recessive etoxazole resistance with
high resolution to a single locus in the T. urticae genome. Additional
genetic evidence further supported a mutation in a conserved Cterminal domain of CHS1, which is highly conserved and might
serve a non-catalytic but essential function. The authors suggest
that CHS1 may be a target site for BPUs in insects as well, although
similar mutations as documented in T. urticae have not been reported yet for any insect species.
4.4.2. ABCC2 and Bt mode of action and resistance
Very recently, an ABC transporter (ABCC2) has been shown to be
involved in resistance to insecticidal protein toxins from the bacterium B. thuringiensis (Bt toxins) (Heckel, 2012). The mode of action of Bt toxins (and the resistance mechanisms to Bt in insects)
are of huge commercial value, given that millions of hectares have
been planted with transgenic Bt crops such as maize and cotton
(Tabashnik et al., 2013). Independent studies in several insect
species including H. virescens, T. ni, P. xylostella, have associated
mutations in ABCC2 with Bt resistance (Atsumi et al., 2012; Baxter
et al., 2011; Gahan et al., 2010), recently reviewed by Heckel (2012).
The most convincing study by Atsumi et al. (2012) used germ line
transformation of the silkworm B. mori to provide functional evidence that a single tyrosine insertion in the extracellular loop of a
TMD was sufficient to confer Bt resistance. Recently it was also
shown, through ectopical expression of B. mori ABCC2 in Sf9 cells,
that B. mori ABCC2 functions as a receptor for Cry1A toxins (Tanaka
et al., 2013). The mode of action of Bt toxins is a complex multi-step
process, which involves activation of the pro-toxin by proteases
after interaction with 12-cadherin domain proteins, formation of an
oligomeric pre-pore structure and several binding steps that
involve several midgut membrane proteins (see Bravo et al., 2011;
Gahan et al., 2010). It has been proposed that ABCC2 plays a role
in the insertion of the pre-pore structure into the midgut membrane. The transporter in the open state is exposed to the outside of
the cell (see Section 1) and could interact with extended helices of
the pre-pore Bt structure, followed by irreversible insertion into the
membrane upon closure (Heckel, 2012). Given the interest in Bt
resistance mechanisms, it can be expected that the precise mechanism of this interaction will be the subject of extended future
research.
Acknowledgments
T.V.L. and W.D. are postdoctoral fellows of the Fund for Scientific
Research Flanders (FWO). We thank Gunnar Broehan and Sheng Li
for providing T. castaneum and B. mori ABC sequences, respectively.
Appendix A. Supplementary data
Supplementary data related to this article can be found at http://
dx.doi.org/10.1016/j.ibmb.2013.11.001.
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