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Figures 2–8 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figures 2–8: Alexandrium affine, LM. (2, 3) Chains of 8 and 3 cells, respectively. (4) An empty cell showing only the theca in ventral view. (5) Epitheca in ventral view showing the ventral pore (arrow) in the first apical plate (1′). (6) Epitheca with Po and 1′ showing the ventral pore (arrow). (7, 8) Po and posterior sulcal plate (Sp) (with a connecting pore), respectively.

Figures 67–69 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figures 67–69: Alexandrium tropicale, LM. (67) Pair of cells. (68, 69) Epitheca and hypotheca with plate tabulation.

Figure 1 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figure 1: Map with the sampling points where species of Alexandrium were found and the sites from which the established strains were isolated.

Figures 38–43 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figures 38–43: Alexandrium pseudogonyaulax, LM. (38) Cell in ventral view. (39) Empty cell in ventral view, showing 1′, 4′, 6″ and the large ventral pore (arrow). (40) Detail of Po with the foramen. (41–43) Epitheca in ventral view showing 1′, 4′, 6″, and ventral pore (arrow).

Figures 22–24 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figures 22–24: Alexandrium margalefii, LM. (22) General outline of a cell. (23) An empty cell in ventral view showing 1′ and 6″ and the ventral pore (arrow) in the first apical plate (1′). (24) Hypotheca with plate tabulation.

Figures 12–19 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figures 12–19: Alexandrium globosum, LM. (12) Cell outline, with the central nucleus arrowed. (13, 14) Two different cells in ventro-lateral and ventral views, respectively, showing some plates of the epitheca and the sulcus. (15) Epitheca with plate tabulation, arrow indicates the location of the ventral pore in the first apical plate (1′). (16) Hypotheca showing plate tabulation. (17) Po plate. (18) Posterior sulcal plate (Sp). (19) Detail of some precingular, cingular and sulcal plates.

Figures 53–66 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figures 53–66: Alexandrium tamiyavanichii, LM and SEM. (53) Chain of 6 cells,LM. (54) Detail of two cells with cellular content of a chain, LM. (55) Cells in ventral view showing the anterior sulcal plate (Sa), LM. (56) Two cells slightly twisted in a chain, SEM. (57) Cell in ventral view showing plates of the ventral area, LM. (58) Empty cell in ventral view showing plate tabulation, the ventral pore is arrowed, LM. (59) Epitheca in ventro-lateral view with plate tabulation, the left sulcal list is arrowed, SEM. (60, 61) Hypotheca with plate tabulation and pore at the posterior sulcal plate (Sp),SEM.(62, 63) Po and plates around it; the ventral pore is arrowed, LM. (64) Posterior sulcal plate (Sp) with pore (arrow), LM. (65, 66) Anterior sulcal plate, LM.

Figure 71 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figure 71: Maximum-likelihood (ML) tree inferred from ITS sequences of Alexandrium. ML bootstrap and Bayesian posterior probabilities values are shown at branches. Bold letters indicate newly generated sequences in this study. Bootstrap values <50 and posterior probabilities <0.50 are not shown.

Figure 70 in Diversity and distribution of species of the planktonic dinoflagellate genus Alexandrium (Dinophyta) from the tropical and subtropical Mexican Pacific Ocean

Figure 70: Maximum-likelihood (ML) tree inferred from D1-D2 LSU rDNA sequences of Alexandrium. ML bootstrap and Bayesian posterior probabilities values are shown at branches. Bold letters indicate newly generated sequences in this study. Bootstrap values <50 and posterior probabilities <0.50 are not shown.

opencc-zeroNov 2021View details →

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Data from: Environmental variation associated with topography explains butterfly diversity along a tropical elevation gradient

Few studies have evaluated the role of topography on the diversity patterns of biological communities along elevation gradients. We evaluated the influence of microclimate and vegetation structure associated with topographic variation on the richness and composition of species of different families of butterflies on a mountain located in a dry enclave (Chicamocha River Canyon) in the northern Andes, Colombia. We captured butterflies over four months at 18 elevations (300 to 1500 m a.s.l.) in two topographic positions (riverbed and hillslope) using an entomological net and traps baited with fermented fruit. In general, butterfly richness increased with elevation in both topographic positions. However, the richness-elevation relationship changed with butterfly family. The riverbed and hillslope sites host different assemblages of butterflies, and this pattern that was consistent for most families. In the riverbed, two sets of species are recognized along the elevation gradient (one below 700 m a.s.l. and the other above 1000 m a.s.l.), mainly owing to species replacement. On the hillslopes there was no clear pattern of grouping associated with elevation. Microclimate differences between the riverbed and hillslope sites along the elevation gradient were related to the vegetation structure and explained the variation in butterfly species composition. Our results highlight the role of topography not only by explaining the response of species richness and composition to environmental variation determined by elevation, but also as a factor that must be considered in the planning and management of biodiversity conservation in the mountains.

opencc-by-4.0Sep 2021View details →

Data for: Trap type affects dung beetle taxonomic and functional diversity in Bornean tropical forests

Dung beetle community composition data. Data was collected using either dung-baited pitfall traps or flight interception traps. Each row represents one trap, with the author/study information, name of study site, sampling period, trap type and habitat type. Dung beetle species and their abundances are listed. See "metadata" tab for more details. Paper abstract: Baited pitfall traps (BPTs) and flight intercept traps (FITs) are the most common methods employed for sampling dung beetle communities. These methods vary in their efficacy and are affected by factors such as the bait types used and the dispersal abilities of different dung beetle species. We present the first quantitative comparison of the taxonomic and functional diversity, and community composition of dung beetles caught in BPTs and FITs in Bornean tropical forests. We show that BPTs and FITs captured complementary communities with different functional traits, and that BPTs captured more functionally diverse communities. We therefore recommend using a combination of both baited BPTs and FITs for studies assessing the composition of dung beetles across habitat types. Our results also highlight that it is important to consider how trap type affects the trait composition of communities when relating dung beetle communities and functional traits to ecological functioning. We suggest modifications to FITs based on the design of harp traps to increase their effectiveness in capturing larger-bodied beetles.

opencc-by-4.0Jul 2021View details →

Multi-taxa environmental DNA inventories reveal distinct taxonomic and functional diversity in urban tropical forest fragments

Urban expansion and associated habitat transformation drives shifts in biodiversity, with declines in taxonomic and functional diversity. Forests fragments within urban landscapes offer a number of ecosystem services, and help to maintain biodiversity and ecosystem functions. Here, we focus on a tropical forest environment, and on the soil biota. Using eDNA metabarcoding, we compare forest fragments within the city of Cayenne, French Guiana, with a neighbouring continuous undisturbed forest. We wished to determine if urban forest fragments conserve high levels of alpha and beta diversity as well as similar functional composition for plants, soil animals, fungi and bacteria. We found that alpha diversity is similar across habitats for plants and fungi, lower in urban forests for metazoans and higher for bacteria. We also found that urban forests communities differ from undisturbed forests in their taxonomic composition, with urban forests exhibiting greater turnover between fragments potentially caused by ecological drift and limited dispersal. However, their functional composition exhibited limited differences, with an enrichment of palms, arbuscular mycorrhizal fungi and bacteria and a depletion of climber plants and termites. Thus, although urban forest fragments do shelter soil biodiversity that differs from native forests, the losses of soil functions may be relatively limited. This study demonstrates the strong potential of a multi-taxa eDNA approach for rapid inventories across taxonomic kingdoms, in particular for cryptic soil diversity. It also demonstrates the key role of urban forest fragments in conserving biodiversity and ecosystem function, and points to a need for more systematic monitoring of these areas in urban management plans. For each of the 16 samples per plot, 15 g of soil was used for eDNA analyses. Extracellular DNA was extracted as described previously (Zinger et al., 2016; 2019), where each soil sample is added to 15ml of saturated phosphate buffer (Na2HPO4; 0.12m; pH ≈8) in 50ml Falcon tubes. This is placed in an agitator for 15 minutes, before a 2ml aliquot of the soil/phosphate buffer mixture is pipetted into an Eppendorf tube and centrifuged for five minutes at 13000 rcf. 500μL of the resulting supernatant is then recovered and used for the next extraction steps that are carried out with a commercial kit for soil DNA (NucleoSpin® Soil; Macherey-Nagel, Düren, Germany), skipping the lysis step and following manufacturer’s instructions. The DNA extract was recovered in 100 μL and diluted 10 times before being used as PCR template.  For each plot one DNA extraction negative control was performed adding up 17 extractions per plot. PCR amplifications were then conducted for four DNA molecular markers, with primers targeting either Viridiplantae (subsequently referred to as plants), Eukaryotes, Fungi or Bacteria (Table 1). For each marker, PCR amplification of samples occurred across 12 plates. Each PCR reaction was performed in a total volume of 20 μl and comprised 10 μl of AmpliTaq Gold Master Mix (Life Technologies, Carlsbad, CA, USA), 5.84 μl of Nuclease-Free Ambion Water (Thermo Fisher Scientific, Massachusetts, USA), 0.25 μM of each primer, 3.2 μg of BSA (Roche Diagnostic, Basel, Switzerland), and 2 μl of DNA template that was before 10-fold diluted to reduce the amounts of PCR inhibitors. Thermocycling conditions for each primer pair are indicated in Table 1.  A negative extraction control per site and a negative PCR control per PCR plate were amplified and sequenced in parallel with the regular samples. Positive controls were also included and consisted of mock communities of plants and fungi DNA (no mock communities were built for bacteria or eukaryotes here), which were used to guide choices in our data curation process. Two PCR replicates were performed for each sample and control. Amplification was conducted using a double indexing system strategy (Binladen et al. 2007) using a system of 32 by 36 octamers with at least five differences between them located at the 5’ end of each primer (Coissac 2012). In doing so, each PCR product had a unique combination of tags for both forward and reverse primers, allowing for the retrieval of sequence data for each sample. Ten wells per PCR plate were left empty to act as sequencing controls (non-used tag combinations) for downstream data curation (see below). PCR products were pooled and sequencing libraries were constructed using the Illumina TruSeq NanoPCRFree kit following the supplier’s instructions (Illumina Inc., San Diego, California, USA), except that the ligation product was not PCR amplified to limit tag-jump biases (Taberlet et al 2018). The libraries were then sequenced on different Illumina platforms (San Diego, CA, USA) depending on the marker considered (Table S1), using the paired-end technology. Bioinformatic analyses were performed on the GenoToul bioinformatics platform (Toulouse, France), with the OBITOOLS package (Boyer et al. 2016). First, ‘illuminapairedend’ was used to assemble paired-end reads. This algorithm is based on an exact alignment algorithm that considers the quality scores at all positions during the assembly process. Subsequently, we used the ‘ngsfilter’ command to identify and remove the primers and tags on each read, and assign reads to their respective samples. This program was used with its default parameters tolerating two mismatches for each of the two primers and no mismatch for the tags. Following this, sequencing reads were dereplicated using the ‘obiuniq’ command. Sequences of low quality (containing Ns or with paired-end alignment scores below 50) were excluded using the ‘obigrep’ command. The same command was used to exclude sequences represented by only one read (singletons) as they are more likely to be molecular artefacts (Taberlet et al. 2018). Sequences outside of the preset range were also discarded (Table 1). To remove PCR/sequencing errors as well as intraspecific variability, we built OTUs (Operational Taxonomic Units) using the ‘sumaclust’ clustering algorithm (Mercier et al. 2013), which considers the most abundant sequence of each cluster as the cluster representative.  OTUs were set at a sequence similarity threshold of 97% for eukaryotes, fungi and bacteria following the standards in microbial ecology, but this was lowered to 95% for plants since the eDNA target region is shorter (typically around 50 base pairs), where one mismatch inherently results in a lower percentage of similarity. To assign a taxon to plant and fungal OTUs, we built two reference sequence databases, one global, using the ecoPCR programme (Ficetola et al. 2010) and the plant / fungi specific markers on the European Molecular Biology Laboratory (EMBL; release 141), a second local, generated from specimens of fungi (Jaouen et al. 2019) and plants (see Zinger et al. 2019) collected in French Guiana. OTUs were then assigned a taxonomy, using OBITOOL’s ecotag programme (Boyer et al. 2016), which performs a global alignment of each OTU sequence (the query) against each reference. The reference taxon assigned to each OTU corresponds to the Last Common Ancestor of all the best-match sequences for the query. For taxonomic assignment of bacteria and eukaryote OTUs, the SILVA taxonomic database was used (version 1.3; Quast et al., 2012). Classification was performed by a local nucleotide BLAST search against the non-redundant version of the SILVA SSU Ref dataset (release 132; http://www.arb-silva.de) using blastn (version 2.2.30+; http://blast.ncbi.nlm.nih.gov/Blast.cgi) with standard settings (Camacho et al., 2009).  Eukaryote derived metazoan OTUs were then further assigned a taxonomy for Phyla identified at the Arthropoda, Annelida and Nematoda level using reference sequence databases built as above for these groups using the ecoPCR programme on EMBL release 141. Datasets were subsequently filtered to remove contaminants as well as artefacts such as PCR chimeras and remaining sequencing errors, following Zinger et al. (2019) and using routines now implemented in the metabaR R package (Zinger et al 2020b), in R version 3.6.1 (R Development Core Team, 2013). The filtering process consisted of four steps: (i) a negative control-based filtering. OTUs whose maximum abundance was found in extraction/PCR negative controls were removed from the dataset, as they were likely to be reagent/aerosol contaminants, better amplified in the absence of competing DNA fragments as it is the case in biological samples. (ii) a reference-based filtering. OTUs which are too dissimilar from sequences available in reference databases are potential chimeras generated during sequencing and amplification. In this study, we chose to set similarity thresholds at 95% for plants, 80% for bacteria and eukaryotes and due to the marker being more polymorphic, 65% for fungi. For plants and fungi, the remaining assignment was then verified with the local database, to confirm if assigned taxa also occurred in the local dataset, with preference given to local assignment. In addition, we removed all taxa that are not targeted by the primer used. (iii) an abundance-based filtering. This procedure targets incorrect assignment of a few numbers of sequences corresponding to true OTUs occurring to the wrong sample, a phenomenon called “tag-switching” (Esling et al. 2015), “tag jumps” (Schnell et al. 2015) or “cross-talk” (Edgar 2018). It consists in setting OTUs abundances to 0 in samples where their abundance represents < 0.03% of the total OTU abundance in the entire dataset. (iv) Finally, we conducted a PCR-based filtering by considering any PCR reaction that yielded less than 100 reads for plants, 1000 reads for fungi, bacteria and eukaryotes as non-functional, and removed them from the dataset. Data provided consists of 4 x OTU tables for each of the markers used to target different components of the soil biota, with rows representing each OTU, and columns the features of the OTU within the dataset, namely their id code, the number of read counts in the analysed dataset, their similarity score against the taxonomic dataset used to identify them, and when possible, a functional group assignment used in the manuscript. Details of these can be found above and in the manuscript and supplementary information. For each of the four datasets, we also provide a .rds file, corresponding to the processed dataset used in manuscript preparation. This is in the format of a metabaR list which includes PCR, Sample, Read count and the seperately provided OTU datasets. To facilitate interpretation, please refer to Zinger, L., Lionnet, C., Benoiston, A.S., Donald, J., Mercier, C. and Boyer, F., 2021. metabaR: an R package for the evaluation and improvement of DNA metabarcoding data quality. Methods in Ecology and Evolution, 12(4), pp.586-592. For the fungal (ITS) data, we also provide :  - the R1/R2 raw fastq files of the samples used in the paper + experimental controls - a tsv file containing the tag combinations corresponding to the samples/PCR replicates, to enable demultiplexing of data. - a csv file containing the description of each sample.

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Data from: Impact of human foraging on tree diversity, composition and abundance in a tropical rainforest

These are summarised plot data from fifteen 40 m by 40 m sample plots established in Oban Division of Cross River National Park, Nigeria, between 23rd August 2019 and 9th September 2019. We have also included data summaries and RStudio codes used for analysis and generating results for the manuscript entitled: "Impact of human foraging on tree diversity, composition and abundance in a tropical rainforest", submitted for publication as an original research article in Biotropica. All data and R code required to generate the results as shown in the manuscript have been included. Complete tree species and plot data can be accessed at https://forestplots.net/.

opencc-zeroOct 2022View details →

Figure 6 in Four new valviferan isopods from diverse tropical Australian habitats (Crustacea: Isopoda: Holognathidae and Idoteidae)

Figure 6. Synidotea innatans sp. nov. Holotype male left, paratype female right. Scale bar = 1 mm (habitus only).

Figure 2 in Four new valviferan isopods from diverse tropical Australian habitats (Crustacea: Isopoda: Holognathidae and Idoteidae)

Figure 2. Cleantioides carpentaria sp. nov. Holotype. Left antennae 1 and 2 detail in ventral view. x = ventral view of left side of pereonite 7 and anterior pleotelson showing pleonal epimera 1–3. Scale bar = 1 mm (habitus only) and 0.2 mm (pereopods).

Figure 9 in Four new valviferan isopods from diverse tropical Australian habitats (Crustacea: Isopoda: Holognathidae and Idoteidae)

Figure 9. Synidotea karumba sp. nov. Holotype male top in dorsal and ventral view, paratype female below, with antennae 2 peduncles.

Figure 3 in Four new valviferan isopods from diverse tropical Australian habitats (Crustacea: Isopoda: Holognathidae and Idoteidae)

Figure 3. Zenobianopsis cidaris sp. nov. Right limbs from holotype. Scale bars = 1 mm (habitus only) and 0.2 mm (pereopods, uropod).

opencc-by-4.0Dec 2018View details →

Figure 8 in Diversity in Australia's tropical savannas: An integrative taxonomic revision of agamid lizards from the genera Amphibolurus and Lophognathus (Lacertilia: Agamidae)

Figure 8. Photos in life of species under revision: a, Lophognathus horneri sp. nov., adult male with breeding colouration, 80 mile beach, Western Australia (photo: R. Glor); b, Lophognathus gilberti, Katherine, Northern Territory (photo: R. Glor); c, Amphibolurus centralis, adult male with breeding colouration, West MacDonnell Ranges, Northern Territory (photo: J. Melville); d, Amphibolurus burnsi, adult male with breeding colouration, Westmar, Queensland (photo: S. Wilson); e, Tropicagama temporalis, Jabiru, Northern Territory (photo: S. Wilson); f, Gowidon longirostris, adult male with breeding colouration, Ormiston Gorge, Northern Territory (photo: R. Glor).

opencc-by-4.0Dec 2018View details →

Figure 2 in Diversity in Australia's tropical savannas: An integrative taxonomic revision of agamid lizards from the genera Amphibolurus and Lophognathus (Lacertilia: Agamidae)

Figure 2. The distribution of Lophognathus sensu lato samples included in this study along the first three morphological principal components axes. Distribution of each taxon is delineated by a 95% confidence ellipse. Lophognathus gilberti has been separated into L. sp. nov., L. gilberti centralis, and L. gilberti.