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Genetic architecture of the Tetragonula carbonaria species complex of Australian stingless bees (Hymenoptera: Apidae... Article in Biological Journal of the Linnean Society · June 2014 DOI: 10.1111/bij.12292
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Biological Journal of the Linnean Society, 2014, ••, ••–••. With 4 figures
Genetic architecture of the Tetragonula carbonaria species complex of Australian stingless bees (Hymenoptera: Apidae: Meliponini) RUTE M. BRITO1,2*, FLÁVIO O. FRANCISCO2,3, SIMON Y. W. HO2 and BENJAMIN P. OLDROYD2 1
Instituto de Genética e Bioquímica, Universidade Federal de Uberlândia, Av. Pará 1720, 2E sala 34, Uberlândia, MG 38400-902, Brazil 2 School of Biological Sciences, University of Sydney, Sydney, NSW 2006, Australia 3 Departamento de Genética e Biologia Evolutiva, Instituto de Biociências, Universidade de São Paulo, Rua do Matão 277 – sala 320, São Paulo, SP 05508-090, Brazil Received 17 December 2013; revised 25 February 2014; accepted for publication 28 February 2014
A species complex is a group of closely related species whose ecological or morphological boundaries are sufficiently vague that delimiting one species from another is difficult. In Australia, a group of four stingless bee species – Tetragonula carbonaria Smith, Tetragonula hockingsi Cockerell, Tetragonula mellipes Friese, and Tetragonula davenporti Franck – form a species complex in which gross morphology is clinal and overlapping. The species are most readily distinguished by the morphology of their brood combs. Here we genetically characterize bees sampled in areas where the species do and do not have contact. Our data corroborate previous evidence that T. hockingsi and T. carbonaria are genetically distinct and that there are two genetically distinct groups of T. hockingsi – one in the north and the other in the south of Queensland. Curiously, northern populations of T. hockingsi, which are allopatric to T. carbonaria, are genetically closer to T. carbonaria than are southern populations of T. hockingsi, which are in sympatry with T. carbonaria. We detected three hybrid colonies that appear to have arisen because of anthropogenic movement of T. hockingsi colonies from north to south of Queensland where males mated with local T. carbonaria queens. We discuss the status of T. davenporti, a recently described species cryptically similar to T. hockingsi from south-east Queensland. © 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••.
ADDITIONAL KEYWORDS: hybridization – microsatellites – mitochondrial DNA – species complex – Tetragonula.
INTRODUCTION The concept of a ‘species’ is central to our classifications of biodiversity, but a ‘species’ is often difficult to define (reviewed in Wheeler & Meier, 2000; de Queiroz, 2007; Hausdorf, 2011). This is particularly the case for a group of closely related taxa in which the species-defining characters lack clear morphological delimitations but are nonetheless sufficiently divergent in developmental biology (Frohlich et al.,
*Corresponding author. E-mail:
[email protected]
1999) or genotype (e.g. Fernández et al., 2012) to be considered as separate species. In this case taxonomists may refer to the group of species as a ‘species complex’ (Bickford et al., 2007). Species complexes often show morphological clines along their geographical distributions (e.g. Franck et al., 2004) and may have overlapping distributions (Alexandrino et al., 2005). Regions in which the members of a species complex come into contact can provide important insights into the processes that underlie speciation and reinforcement (Barton & Hewitt, 1989; Beekman et al., 2008). Estimates of genetic admixture between the members
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
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of a species complex can be used to infer the existence (or absence) of pre- or postzygotic barriers to hybridization between the species and to uncover the mechanisms that are driving the speciation process (Coyne & Orr, 1989, 2004). When no hybrids can be identified in a zone of contact, it can be inferred that two fully allopatric species are now in contact following recent range expansions (Moritz et al., 2000). On the other hand, if hybrids can be identified, the contact zone can be regarded as a hybrid zone and is probably an area of secondary contact between two (sub)species that were geographically isolated in the past but are again in contact (Hewitt, 1988). In such cases, the accumulation of divergent traits through time has been insufficient to prevent hybridization upon secondary contact. If there are no barriers to hybridization, the gene pool of the species complex will rehomogenize and the incipient species will be lost (Seehausen et al., 2008). However, if there are barriers to mating or if hybrids have reduced fitness relative to the parental types, the hybrid zone is regarded as a tension zone in which gene flow between the incipient species is impeded and speciation is reinforced (Beekman et al., 2008). In Australia, a group of stingless bees comprising Tetragonula mellipes, Tetragonula carbonaria, Tetragonula hockingsi, and Tetragonula davenporti provides an interesting example of a species complex (Rasmussen & Cameron, 2010). Species of this group were formerly placed in the genus Trigona (Jurine) but Moure (1961) erected the genus Tetragonula to delineate the Old World Trigona-like taxa from the New World Trigona. Moure’s nomenclature has been adopted by the most recent and comprehensive catalogue of Australasian stingless bees (Rasmussen, 2008) and by Michener (2013), and we have used it here. The first three species of the T. carbonaria species complex show strikingly different nest architectures (Dollin, Dollin & Sakagami, 1997; see illustrations in Franck et al., 2004). Tetragonula mellipes builds its brood cells as an irregular brood comb and constructs a prominent, 5-cm entrance tube of wax. Tetragonula carbonaria builds a spiral brood comb and deposits wax and resin ornaments around the entrance. Tetragonula hockingsi builds its brood cells in an open semi-comb structure and rarely deposits wax or resin on the nest entrance. Tetragonula davenporti is a cryptic species that builds nests identical to those of T. hockingsi (Franck et al., 2004). Morphologically, only T. mellipes is significantly divergent from the other species. Uniquely for the group, the posterior corbicular fringe is pale (Dollin et al., 1997). The remaining species of the T. carbonaria complex are identified by head width: T. hockingsi is slightly larger than T. carbonaria and
T. davenporti. However, head width is not diagnostic because there is a north–south cline in size along the east coast of Queensland (Dollin et al., 1997). Therefore, nest architecture is the only speciesdefining character for T. hockingsi and T. carbonaria. Tetragonula davenporti is distinguishable from T. hockingsi and T. carbonaria based on four diagnostic characters: (1) the colour of the hairs on the head; (2) the presence or absence of hairs in the malar space (the area between the eye and the mouth); (3) the presence or absence of hairs on the sixth tergite; and (4) the darker coloration of T. hockingsi (Franck et al., 2004). Curiously, Franck et al. (2004) identified a few hybrids between T. carbonaria and T. davenporti based on analyses of microsatellite loci and morphological characters. Tetragonula davenporti is a rare species that is restricted to a small region south of Brisbane (Fig. 1). We wished to further characterize this cryptic species, delineate its range more precisely, and understand its relationship to T. carbonaria and T. hockingsi. To address these points, we extended sampling of both species at the contact area in southern Queensland (QLD) and compared their genetic profiles at nine microsatellite loci and two mitochondrial gene sequences with samples from areas where only one species is extant.
MATERIAL AND METHODS SAMPLING, SPECIES IDENTIFICATION AND DNA EXTRACTION Tetragonula mellipes does not occur in the Australian states of New South Wales (NSW) or QLD, but is confined to the Northern Territory and is not included in this study. In QLD and NSW we collected samples of T. carbonaria (N = 110), T. hockingsi (N = 123), and T. davenporti from the original nests sampled by Franck et al. (2004) (N = 2; TD01 and TD02). We collected at seven major sites, encompassing the entire range of the three species (Dollin et al., 1997), from beekeepers (N = 187) and off flowers by sweep netting or from wild colonies (N = 48), as follows: Cape York QLD (a known contact zone between T. carbonaria and T hockingsi, Franck et al., 2004), Townsville QLD (allopatric T. hockingsi), Mackay QLD (T. hockingsi), Rockhampton QLD (T. hockingsi), Mount Perry QLD (contact zone), Brisbane QLD (the contact zone and the original range of T. davenporti), and Sydney NSW (allopatric T. carbonaria) (Fig. 1 and Table 1). One worker per nest was collected when sampling from colonies, or one foraging worker per sampling site was collected within a radius of 1 km when sampling off flowers. We also included two Austroplebeia australis specimens (from Brisbane and
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
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Figure 1. Map of Australia showing collection sites grouped in seven major regions (see Table 1) and the geographical distribution of Tetragonula carbonaria (diamonds) and Tetragonula hockingsi (circles): Cape York (York), QLD (T. hockingsi); Townsville (Twns), QLD (T. hockingsi); Mackay (Mack), QLD (T. hockingsi); Rockhampton (Rock), QLD (T. hockingsi); Mount Perry (Mt Perry), QLD (both species); Brisbane, QLD (T. hockingsi, T. carbonaria, and T. davenporti); and Sydney, NSW (T. carbonaria). NSW, New South Wales; NT, Northern Territory; QLD, Queensland; SA, South Australia; TAS, Tasmania; VIC, Victoria; WA, Western Australia. (Colour figure available online.)
Rockhampton) as an outgroup for the network analysis. Samples were preserved in 70% ethanol during fieldwork, washed in the laboratory (in 50 mM Tris, pH 8.0, 10 mM NaCl, and 2 mM MgCl2), and stored at −80 °C until required for DNA extraction. When possible, specimens (N = 43) were identified by the morphology of their brood comb (Franck et al., 2004). When the brood comb was not available, for example when the sampled colony was in a house cavity or for bees collected off flowers, we identified the species of the sampled insect from its head width and other morphological characters, such as the mesoscutal shape and pilosity of the mesosomal side (Dollin et al., 1997; Franck et al., 2004). The two T. davenporti colonies from the same hives as those sampled by Franck et al. (2004) have been continuously occupied by bees since Franck’s sampling in 1999. For these colonies we performed a more detailed morphological examination based on the criteria specified by Franck et al. (2004). We extracted DNA from the thorax of one individual per nest (or per sampling site for foragers) using a high-salt method (Aljanabi & Martinez, 1997).
GENOTYPING
AND GENETIC ANALYSES OF
MICROSATELLITE LOCI
We genotyped each worker at nine microsatellite loci using primers designed from T. carbonaria (Tc7.13, Tc1.20, Tc4.63, Tc3.155, Tc4.214, Tc4.287, and Tc3.302) (Green, Franck & Oldroyd, 2001) and from Tetragonisca angustula (Tang60 and Tang70) (Brito et al., 2009). Reverse ‘Tc’ primers were labelled with one of the following fluorescent dyes: 6-FAM (SigmaAldrich), NED, PET, and VIC (Life Technologies). Forward ‘Tang’ primers had been designed with a non-specific tag sequence (5′-CCTGGCGACTCCT GGAG-3′) according to Schuelke (2000). Polymerase chain reaction (PCR) amplifications were performed in 5-μL volumes containing 1 × reaction buffer, 1.5 mM MgCl2, 0.1 mM deoxyribonucleotide triphosphates (dNTPs), 0.4 μM forward primer and 0.4 μM labelled reverse primer (or 0.0125 μM tagged F primer, 0.125 μM regular reverse primer, and 0.125 μM tag sequence labelled with 6-FAM, NED, PET, or VIC), 1% glycerol, 0.5 units of TAQ-Ti DNA polymerase (Fisher Biotec), and 1 μL (10–20 ng) of DNA template.
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Table 1. Geographical regions, sampling sites, and geographical coordinates of sampling sites of Tetragonula specimens: samples were pooled in major geographical regions according to their distribution along the east coast of Australia Geographical region
Sampling site
Coordinates
Cape York QLD (Cape York)
Walkamin Tolga Cardwell Townsville Ayr Mackay Rockhampton Mount Perry
17° 17° 18° 19° 19° 21° 23° 25°
07′S 13′S 15′S 16′S 34′S 02′S 20′S 08′S
145° 145° 146° 146° 147° 149° 150° 151°
Miles Tara Cecil Plains Highvale Hatton Valle Bundamba Brassall Brisbane Elanora Sydney Blue Mountains
26° 27° 27° 27° 27° 27° 27° 27° 28° 33° 33° 33° 33° 33° 33° 33° 33° 33° 33° 33° 33°
39′S 16′S 31′S 22′S 31′S 36′S 35′S 31′S 07′S 29′S 35′S 36′S 39′S 40′S 42′S 44′S 45′S 46′S 47′S 47′S 39′S
150° 11′E 150° 27′E 151° 11′E 152° 49′E 152° 29′E 152° 48′E 152° 44′E 153° 01′E 153° 28′E 151° 19′E 151° 01′E 150° 44′E 150° 00′E 150° 51′E 151° 05′E 151° 03′E 151° 00′E 151° 04′E 151° 01′E 151° 03′E 150 °58′E
Northern QLD (Townsville) Mackay QLD (Mackay) Middle QLD (Rockhampton) Mt Perry QLD (Mount Perry) Southern QLD (Brisbane)
NSW (Sydney)
25′E 28′E 01′E 47′E 24′E 09′E 32′E 37′E
Species
N
T. hockingsi T. carbonaria
41 1
T. hockingsi
11
T. hockingsi T. hockingsi T. hockingsi T. carbonaria T. hockingsi T. carbonaria T. davenporti
8 31 6 12 30 65 2
T. hockingsi T. carbonaria
1 40
NSW, New South Wales; QLD, Queensland. N, number of samples collected in each major region.
The cycling conditions used were those of Gloag et al. (2008) for Tc primers. For Tang primers, the cycling conditions consisted of an initial denaturation step of 8 min at 96 °C followed by 35 cycles of denaturation at 94 °C for 30 s and 48 °C for 60 s for annealing of Tang60, or 94 °C for 30 s and 53 °C for 60 s for annealing of Tang70, and elongation at 72 °C for 60 s. There was a final elongation step at 72 °C for 10 min. The PCR products labelled with different dyes were pooled and diluted 1 : 10, and 1 μL of each diluted PCR pool was added to 10 μL of LIZ 500 size standard (Life Technologies) and formamide (1 : 100). Samples were run on a 3130xl Genetic Analyzer (Life Technologies). Bees were genotyped using GENEMAPPER v3.7 (Applied Biosystems, USA). We used GENALEX v6.4 (Peakall & Smouse, 2006) to generate genetic diversity indices for the number of alleles per locus (NA), number of private alleles, and observed (HO) and expected (HE) heterozygosities.
We calculated Nei’s unbiased genetic distance (Nei, 1978), which is more appropriate for small numbers of samples, between species and between sampling sites, and used Principal Coordinate Analysis (PCoA) (Peakall & Smouse, 2006) to perform visual assessment of the genetic similarity of bees within each species. GENEPOP v4.1 (Rousset, 2008) was used to perform exact tests of departure from Hardy– Weinberg equilibrium, heterozygote deficiency, and linkage disequilibrium for each sampling site and species combination. Sequential Bonferroni corrections (Rice, 1989) were used to adjust significance levels for these tests. Measures of population structure and admixture were obtained using STRUCTURE v2.3.3 (Pritchard, Stephens & Donnelly, 2000) with parameters set to ‘USEPOPINFO’ (off), ‘POPFLAG’ (on), and ‘LOCPRIOR’ (off). We first analysed all specimens as a single population, setting the number of estimated
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
HYBRIDIZATION IN AUSTRALIAN STINGLESS BEES populations (k) as 1–7 (as there were seven collection sites) under an admixture model, and correlated allele frequencies with a burn-in of 105 Markov chain steps, sampling from 106 steps, and 10 iterations for each k (Cabria et al., 2011). Initially we expected three major groups, corresponding to T. carbonaria, T. davenporti, and T. hockingsi. However, the most likely value of k (highlighted by STRUCTURE HARVESTER v0.6.92 (Earl & VonHoldt, 2012) was k = 2, and k = 3 was not supported. Therefore, we re-ran the analysis with all specimens as a single population based on k = 2. From this second analysis, individuals with Q values (the estimated membership coefficient for each individual, in one of the clusters) of less than 95% were considered as potential hybrids (Schwartz & Beheregaray, 2008). Hybrid Q values always showed higher estimated membership of one of the clusters, T. hockingsi > T. carbonaria (Th > Tc) or vice versa (Tc > Th). A third analysis was then performed, either with all samples included or with only T. hockingsi samples included; in this analysis we designated major collection sites as populations (Table 1). Again we used k = 1–7 under admixture models and correlated allele frequencies with a burn-in of 105 Markov chain steps, sampling from 106 steps, and 10 iterations for each k. This third analysis was used to confirm which, if any, subpopulations contained hybrids. The program CLUMPP v1.1.2 (Jakobsson & Rosenberg, 2007) was used to align the 10 repetitions of the best k. The program DISTRUCT v1.1 (Rosenberg, 2004) was then used to display the results, produced by CLUMPP, graphically. We generated simulated hybrid and backcross genotypes between T. carbonaria and T. hockingsi using HYBRIDLAB (Nielsen, Bach & Kotlicki, 2006) to determine if any of our putative hybrid genotypes was more consistent with being F1 or backcross hybrids of an interspecific cross. For the parental generation we used 23 ‘pure’ genotypes of each species from the Brisbane (T. hockingsi) and Sydney (T. carbonaria) populations. We then generated 50 virtual F1 hybrids and used these genotypes as a parental stock to generate in-silico backcrosses to either T. carbonaria or T. hockingsi.
SEQUENCING We used mtD6 (C1-J-1718) (5′-GGAGGATTTGGAA ATTGATTAGTTCC-3′) and mtD9 (C1-N-2119) (5′CCCGGTAAAATTAAAATATAAACTTC-3′) universal mitochondrial primers for insects (Simon et al., 1994) to amplify 500 bp of the cytochrome c oxidase subunit 1 (COI) gene and 16Sar with 16Sbr designed from the Apis mellifera 16S ribosomal RNA (16S) gene of the mitochondria to amplify 600-bp fragments (Simon et al., 1994). Reactions were carried out in 50 μL
5
containing 1 × reaction buffer, 3 mM MgCl2, 0.4 mM dNTPs, 0.2 μM each primer, 0.5 units of TAQ-Ti DNA polymerase (Fisher Biotec), and 2.5 μL (25–50 ng) of DNA template. Cycling conditions consisted of a denaturation step of 5 min at 94 °C followed by 35 cycles of denaturation at 94 °C for 60 s; 80 s at 42 °C for primer annealing, 64 °C for 120 s for elongation. An additional elongation step at 64 °C for 10 min was added at the end. The PCR products were purified using Exo (0.1 U): SAP (2 U) enzymes, in 1.8 μL of Tris (50 mM), per 20 μL of PCR product and then sent to the Australian Genome Research Facility, Australia, or Macrogen, South Korea, for sequencing. Electropherograms were checked by eye and sequences were aligned and concatenated using GENEIOUS v5.1 (Drummond et al., 2010). We analysed haplotypes using DNASP v5 (Librado & Rozas, 2009) to estimate haplotype diversity (Hd) and nucleotide diversity (π). NETWORK v4.5.0.0 was used to build a haplotype network using the median-joining method (Bandelt, Forster & Röhl, 1999).
RESULTS MICROSATELLITES All microsatellite loci analysed were highly polymorphic, with no significant linkage disequilibrium observed between any pair of loci (Bonferroniadjusted P-value = 0.0014). We observed high molecular diversity, with allele numbers ranging from four to nine per locus for T. hockingsi and from four to 19 for T. carbonaria. The mean HO and HE values were 0.396 and 0.403, respectively, for T. hockingsi, and were 0.532 and 0.482 for T. carbonaria (Table 2). Significant heterozygote deficiency was detected in the Cape York, Townsville, and Brisbane sampling areas for T. hockingsi (P < 0.05) but not for T. carbonaria at any site. When all samples of all species were analysed as a single population with k = 1–7 subpopulations, STRUCTURE supported k = 2 as the most likely number of subgroups (Fig. 2A). Samples were then assigned to their collection sites and the analysis was re-run with k = 1–7 groups. Again, the most likely number of subgroups was 2, reinforcing the conclusion that our samples comprised just two species (and a few hybrids), irrespective of sampling site. Bees collected from Cape York to Rockhampton were classified in the same group (T. hockingsi). One exception was a T. carbonaria hive purchased from Brisbane and kept in a yard in Walkamin, QLD (Fig. 3A, arrow 1). Conversely, all samples collected in NSW were classified in the second group (T. carbonaria), with the singular exception of a wild colony found in a compost bin
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
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Table 2. Genetic variability at nine microsatellite loci of Tetragonula hockingsi and Tetragonula carbonaria from all seven major areas studied T. hockingsi
T. carbonaria
Locus
NA
PA
HO
HE
NA
PA
HO
HE
Tc7.13 Tc1.20 Tc4.214 Tc3.155 Tc4.63 Tc3.302 Tc4.287 Tang60 Tang70 Mean
9 9 4 9 8 6 6 8 5 7.1
5 5 1 2 5 1 2 1 – 2.4
0.517 0.408 0.360 0.346 0.414 0.192 0.556 0.372 0.402 0.396
0.554 0.499 0.351 0.332 0.414 0.210 0.499 0.369 0.402 0.403
5 5 6 10 19 6 5 10 4 7.8
1 3 3 5 15 1 1 4 – 3.7
0.684 0.272 0.192 0.496 0.810 0.366 0.416 0.785 0.766 0.532
0.574 0.247 0.191 0.535 0.691 0.372 0.450 0.688 0.592 0.482
HO, observed heterozygosity; HE, expected heterozygosity; NA, number of alleles; PA, number of private alleles.
Figure 2. Genetic structure of the Tetragonula carbonaria species complex of stingless bees along the eastern Australian coast, inferred using Bayesian analysis in the program STRUCTURE. Δk values (k = 1–7) suggested k = 2 as the most likely number of subpopulations. Each individual is represented by a vertical line. y-axis, Q values estimated for each individual. (A) All samples of all species considered as a single population. Green, individuals classified as pure Tetragonula hockingsi based on STRUCTURE; red, pure Tetragonula carbonaria individuals. Individuals with Q values less than 95% were considered hybrids. (B) Principal Coordinate Analysis (PCoA) of the T. carbonaria species complex was based on the genetic distance of microsatellite genotypes. PCoA1 and PCoA2 represent the first two factorial components that together explain 81% of the observed variation. TD, Tetragonula davenporti. (Colour figure available online.)
in Sydney (Fig. 3A, arrow 2) that grouped with T. hockingsi. Head measurements agreed with this result. This is the first report of T. hockingsi in NSW. Mount Perry and Brisbane are contact zones between T. carbonaria and T. hockingsi (Fig. 1). Indi-
viduals with hybrid nuclear genotypes were observed in Brisbane (Tc > Th) (N = 3). Samples of ‘T. davenporti’ (N = 2) were genotypically most similar to T. carbonaria; TD01 showed a Q value of 95% and spiral brood, and TD02 showed a Q value of 80% and classic T. hockingsi brood. When we ran STRUCTURE
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
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Figure 3. Structure of the Tetragonula carbonaria species complex of stingless bees inferred using Bayesian analysis in STRUCTURE. Each individual is represented by a vertical line. y-axis, Q values estimated for each individual. (A) All samples of both species, with k = 3 as the number of subpopulations. Tetragonula hockingsi from northern Queensland were assigned to the dark-green group, T. hockingsi from southern Queensland were assigned to the light-green group, and T. carbonaria samples were added to the red group. (B) T. hockingsi populations for k = 2 as the number of subpopulations. Samples of T. hockingsi from northern Queensland were assigned to the dark-green group, whereas those from the south were assigned to the light-green group. Arrow 1, T. carbonaria hive in the Walkamin area purchased from Brisbane; arrow 2, T. hockingsi hive in Sydney area, found in a compost bin. Hybrids are indicated with asterisks above arrows 3–7: 18, 188, 210, TD01, and TD02, respectively. Mack, Mackay; Mt Perry, Mount Perry; Rock, Rockhampton; Twns, Townsville; York, Cape York. (Colour figure available online.)
using k > 2, we observed a split between T. hockingsi from northern (York, Townsville, Mackay) and southern (Rockhampton, Mt Perry and Brisbane) QLD (Fig. 3A), but not between T. davenporti and T. carbonaria. We included the three hybrid samples in the group named ‘hybrids’ in subsequent analyses. Nei’s unbiased genetic distance between T. hockingsi and T. carbonaria is high (2.312). No genetic distance was observed between T. carbonaria and ‘hybrids’ (0), in contrast to the comparison of T. hockingsi with ‘hybrids’ (1.139). The pairwise distances between T. davenporti and T. carbonaria, T. davenporti and T. hockingsi, and T. davenporti and ‘hybrids’, were 0.649, 1.599, and 0.877, respectively. The PCoA of populations of T. hockingsi and T. carbonaria based on microsatellite genotypes showed that most variation (71%) is explained by species-specific genotypes. Hybrids from the Brisbane area grouped with T. carbonaria. The T. davenporti sample, TD01, grouped with T. carbonaria, whereas TD02 was intermediate between T. hockingsi and T. carbonaria species groups, but more similar to T. carbonaria than to T. hockingsi (Fig. 2B). When T. hockingsi samples, as identified from the first STRUCTURE analysis, were analysed separately using k = 1–7, the most likely number of groups, based on Δk-values from STRUCTURE HARVESTER, was 2. This provides further evidence that the T. hockingsi population we sampled in north QLD (York and Townsville) is genetically distinct from
the T. hockingsi population in southern QLD (Rockhampton, Mt Perry, and Brisbane), and that Mackay is a hybrid zone comprising individuals bearing genotypes of both major T. hockingsi groups, or even hybrids (Fig. 3B). Unexpectedly, T. carbonaria samples are less differentiated from northern T. hockingsi than from T. hockingsi samples from Brisbane (Table 3). Both T. hockingsi and T. carbonaria are closer to hybrids than to each other (Table 3). The results from the STRUCTURE analysis with k = 3 suggest that our ‘T. davenporti’ samples were more parsimoniously assigned to T. carbonaria than to a third group. We generated simulated F1 hybrid genotypes using HYBRIDLAB with genotypes of pure stocks of T. hockingsi from Brisbane and T. carbonaria from Sydney. These simulated genotypes had Q values of 50% after STRUCTURE analysis with k = 2. On the other hand, backcross hybrids showed Q values varying from 70 to 98% when an F1 hybrid was backcrossed to T. hockingsi or T. carbonaria, similar to those observed in the group we classified as being ‘hybrids’.
SEQUENCING We successfully sequenced 333 bp of COI and 452 bp of 16S mitochondrial genes from 183 of the 235 samples used for microsatellite analyses, including all hybrids (18, 188, and 210) and the ‘T. davenporti’
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
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Table 3. Pairwise population matrix of unbiased Nei’s genetic distances between Tetragonula hockingsi collected in different regions, and Tetragonula carbonaria, ‘hybrids’ and Tetragonula davenporti NQ
Twns Mack Rock Mt Perry Brisbane Carb Hyb Tdav
SQ
York
Twns
Mack
Rock
Mt Perry
Brisbane
Carb
Tdav
0.132 0.205 0.353 0.349 0.310 1.956 1.100 1.240
0 0.102 0.170 0.166 0.139 2.123 1.080 1.266
0 0.094 0.089 0.061 2.979 1.235 2.076
0 0 0.002 2.645 1.295 2.033
0 0.012 2.727 1.367 2.003
0 2.541 1.225 1.905
0* 0.650†
0.877** 0
Values in bold indicate distances between T. carbonaria and northern T. hockingsi populations, and values underlined indicate distances between T. carbonaria and southern T. hockingsi populations. Values in italic highlight distances between T. hockingsi populations and hybrids (Hyb). *Distance between T. carbonaria (Carb) and hybrids. **Distance between hybrids and T. davenporti (Tdav). Light grey and dark grey shading highlight distances between T. davenporti and northern and southern T. hockingsi populations, respectively. †Distance between T. davenporti and T. carbonaria. NQ, northern Queensland; SQ, southern Queensland. Figure 4. Network relating 43 haplotypes based on 183 concatenated cytochrome c oxidase subunit 1 (COI) (333 bp) and 16S ribosomal RNA (16S) (452 bp) mitochondrial DNA sequences from Tetragonula carbonaria, Tetragonula hockingsi, and hybrids, including Tetragonula davenporti, and two samples from the outgroup Austroplebeia australis. Circle sizes represent haplotype frequencies. Unbroken lines represent a single mutational step. Numbers in parentheses represent the number of mutational steps between adjacent haplotypes. Colours represent species assignments based on microsatellite loci. Red, T. carbonaria; blue, T. davenporti; dark green, T. hockingsi from northern Queensland; light green, T. hockingsi from southern Queensland; yellow, A. australis. Asterisks indicate haplotypes observed in hybrids 18 and 210 (H12) and 188 (H32). (Colour figure available online.) ▶
samples (TD01 and TD02). Concatenated sequences contained 129 polymorphic sites, with high values of haplotype (gene) diversity (0.897) and nucleotide diversity per site (0.05714). The sequences represented 41 haplotypes, with a G + C content of 26.3%. No haplotypes were shared between species. All bees classified as hybrids based on the nuclear genome had mitochondrial haplotypes that clustered within T. carbonaria (Fig. 4). Individuals 18 and 210, which were classified as hybrids based on their nuclear DNA, carried the most frequent T. carbonaria haplotype (H12). The mitochondrial haplotype of hybrid 188 also clustered within those of T. carbonaria (H32). A network relating all haplotypes (Fig. 4) provides further evidence of population subdivision between T. hockingsi from northern and southern QLD. Tetragonula carbonaria haplotypes are closer to T. hockingsi from northern QLD than to T. hockingsi from southern QLD, reinforcing this unexpected conclusion from our microsatellite data. Interestingly, haplotype H29 is present in both north T. hockingsi samples from Mackay and south T. hockingsi from Rockhampton, Brisbane, and Sydney. The anomalous
distribution of this haplotype may have arisen from anthropogenic movements of T. hockingsi from north to southern QLD. Samples of ‘T. davenporti’ showed a unique haplotype (H30) with 20 more private nucleotide substitutions than any other individual in our study. However, this degree of difference is similar to that seen between some pairs of haplotypes within T. hockingsi (e.g. H29 and H6; Fig. 4). Austroplebeia australis haplotypes were placed at the end of a long branch extending from the northern T. hockingsi group (Fig. 4), indicating that T. carbonaria is nested within the diversity of T. hockingsi.
MORPHOLOGY
OF
T. DAVENPORTI
HYBRID SAMPLES
Our T. davenporti samples were from the same hives designated as T. davenporti by Franck et al. (2004). We examined six workers from each of the two colonies for the four diagnostic features of T. davenporti (Franck et al., 2004) in the two putative T. davenporti colonies. In all specimens the hairs on the vertex were white, characteristic of T. hockingsi. There was an absence of hairs in the malar space, characteristic of
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
HYBRIDIZATION IN AUSTRALIAN STINGLESS BEES
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
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T. davenporti. The sixth tergite was covered in hairs, characteristic of T. hockingsi, and the overall coloration was light brown, characteristic of T davenporti. These results suggest that the colonies are now hybrids between T. davenporti and another species.
DISCUSSION Our data, and those of others (Dollin et al., 1997; Franck et al., 2004; Rasmussen & Cameron, 2007), show that T. hockingsi and T. carbonaria are valid species. Franck et al. (2004) reported significant genetic structuring between T. hockingsi from northern and southern QLD, leading them to erect a fourth species in the T. carbonaria species complex, namely T. davenporti, restricted to south QLD (Elanora). The widely divergent haplotype of the T. davenporti hybrid colonies corroborate that T. davenporti is rare but is also a valid species. Our microsatellite data show that there is significant genetic structuring of the T. hockingsi population between north and south QLD. However, our STRUCTURE analysis supports just two, not three, groups within the bees we sampled, meaning that we failed to find two species among bees with open broodcomb morphology. Thus, the southern forms of T. hockingsi that we sampled should not be regarded as T. davenporti. It seems that T. davenporti has a restricted range in southern QLD that we failed to sample and ‘pure’ examples. Potentially the natural range of T. davenporti lies outside the Elanora area, and the colonies sampled by Franck (and later by ourselves) were transported to the area. We observed individuals with nuclear genotypes that suggest hybridization between T. hockingsi and T. carbonaria, and between T. carbonaria and T. davenporti. Global phylogenies show that T. hockingsi and T. carbonaria are closely related (Rasmussen & Cameron, 2007, 2010) and our data reinforce the hypothesis that biological speciation is as yet incomplete within the T. carbonaria species complex because hybridizations occur (Franck et al., 2004). The level of admixture estimated for the hybrid T. davenporti samples was 60–95%, which is consistent with an F1 hybrid backcrossed to T. carbonaria. This may suggest that in the time period 2001–2009, between the first collection of T. davenporti (Franck et al., 2004) and our current collection from the same colonies, queens were replaced and F1 hybrid queens mated with local T. carbonaria males. We have little information on the longevity of Meliponini queens (Roubik, 1992; Carvalho-Zilse & Kerr, 2004), but honeybee queens can live for up to 5 years (Roubik, 1992). The detailed morphology of the current ‘T. davenporti’ colonies, and the spiral brood shape of TD02, also
suggests that they are now hybrids because they differ substantially from the first description (Franck et al., 2004) and yet are unlike either ‘pure’ T. carbonaria or T. hockingsi. As a whole, T. carbonaria is more genetically distinct from T. hockingsi from southern QLD than it is from T. hockingsi from northern QLD. In the mitochondrial data, we found T. carbonaria nested within the diversity of northern T. hockingsi. The reasons behind this pattern are unclear. One possibility is incomplete sorting of mitochondrial lineages, whereby the T. carbonaria lineage is more closely related to northern T. hockingsi than to southern T. hockingsi simply by chance. Incomplete lineage sorting appears to be widespread and leads to inconsistencies between gene trees and species trees (Tajima, 1983). Another possibility is that the mitochondrial DNA of T. carbonaria has been replaced with that of northern T. hockingsi, perhaps by a selective sweep. Although we did not find indigenous hives of T. carbonaria in Walkamin, Cardwell, Townsville, Ayr, and Mackay, Dollin et al. (1997) reported that T. carbonaria is sympatric to T. hockingsi in northern QLD as well as in southern QLD. This sympatry could allow some degree of gene flow between northern T. hockingsi and T. carbonaria, and the mitochondrial DNA in T. carbonaria has been replaced with that of T. hockingsi before the southward dispersal of T. carbonaria along the east coast of Australia. This is a less probable hypothesis, but the microsatellite results do not rule it out. Potentially, the southern T. hockingsi population is and was more reproductively isolated from T. carbonaria by ecological constraints than the northern population, reducing opportunities for gene flow between the incipient species. Hybridization has been previously reported for morphologically similar Melipona (Nascimento, Matusita & Kerr, 2000), Tetragonula (Franck et al., 2004), and Tetragonisca (Francisco et al., 2014) stingless bee species. Males of different species have been reported in the same drone congregations in Brazil (Velthuis, 2006). In Tetragonula, drone congregations are often seen as swarms of hundreds of males circling in front of a nest. Drone congregations comprise males from many tens of unrelated colonies (Sommeijer & de Bruijn, 1995; Cameron, Franck & Oldroyd, 2004; Kraus, Weinhold & Moritz, 2007). As T. carbonaria and T. hockingsi are closely related species, aggregations of males of one species could potentially attract males from the other species. We are unaware of any description of male genitalia for these species (see Dollin et al., 1997), but we suggest that morphological incompatibility is an unlikely prezygotic barrier. The nest architectures of T. carbonaria and T. hockingsi are unique in this genus, and are not
© 2014 The Linnean Society of London, Biological Journal of the Linnean Society, 2014, ••, ••–••
HYBRIDIZATION IN AUSTRALIAN STINGLESS BEES seen in related Tetragonula species from Asia (Dollin et al., 1997; Rasmussen & Cameron, 2007, 2010). This peculiarity suggests that the Australian species complex diverged within Australia after migration of an ancestral species from South-East Asia (Franck et al., 2004). Brito et al. (2012) showed that small changes in the rules followed by comb-building Tetragonula bees can result in radically different comb structures. While nest architecture remains the most obvious species-diagnostic trait, it is uncertain whether more frequent hybridizations, mediated by anthropogenic colony movements, may blur the associations between nuclear genotype and nest architecture. Species boundaries in the T. carbonaria complex are not fully established, as we observe hybrids between species. However, the few hybrids we observed showed allelic characteristics of groups that are not in sympatry: T. carbonaria collected in Brisbane and Sydney; and T. hockingsi from northern QLD. We suggest that trading of hives between beekeepers in northern and southern QLD has brought together genotypes that have not yet developed prezygotic barriers, thereby facilitating artificial hybridization. Thus, we speculate that the sympatric variants of southern T. hockingsi and T. carbonaria have evolved barriers to hybridization, but when geographically distant variants are brought together anthropogenically, hybridization is possible. We conclude that the hybrids observed here are not natural, and that T. carbonaria and T. hockingsi are separate species. Keepers of stingless bees need to be aware that moving colonies across great distances can interfere in the natural course of speciation, creating unnatural hybridization events.
ACKNOWLEDGEMENTS We are grateful to Adrian Lewis, Alan Ashhurt, Alan Mathews, Carlos Alonso, Cecil Heather, Charlie Roberts, Collin Web, Glen Brook, Helen Wallace, John Klumpp, Keith Paul, Les Falhaber, Malcom Johnston, Margaret Tracey, Martyn Robson, Megan Halcroft, Paul Anderson, Peter Davenport, Robert Luttrell, Roderick Yates, Russel Zabel, Steve Bonney, Tom Carter, Wayne Allen, and Willian White for their help in collecting samples. We would like particularly to thank Dr Anne Dollin for her help discussing the project, in obtaining and identifying samples; Dr Tim Heard and Adrian Lewis for providing samples and driving us around showing us bee hives and introducing us to beekeepers. We also would like to thank Julianne Lim for technical assistance. RMB was supported by a Postdoctoral Fellowship (Conselho Nacional de Desenvolvimento Científico e Tecnológico – CNPq, PDE: 201470/2008-0) and an Australian
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Endeavour Awards Research Fellowship. FOF was supported by a doctoral scholarship (2008/08546-4 FAPESP). Additional funding was provided by an Australian Research Council Grant to BPO. The authors have no conflicts of interest to declare.
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SHARED DATA Sequence data have been submitted to GenBank under the accession numbers JX304919 to JX305294. The sequence alignment has been deposited on the Sydney eScholarship Repository (http://hdl.handle.net/2123/ 10445).
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