Sensory Protein Kinase Signaling in Schistosoma mansoni Cercariae: Host Location and Invasion

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Abstract

Schistosoma mansoni cercariae display specific behavioral responses to abiotic/biotic stimuli enabling them to locate and infect the definitive human host. Here we report the effect of such stimulants on signaling pathways of cercariae in relation to host finding and invasion. Cercariae exposed to various light/temperature regimens displayed modulated protein kinase C (PKC), extracellular signal–regulated kinase (ERK) and p38 mitogen-activated protein kinase (p38 MAPK) activities, with distinct responses at 37°C and intense light/dark, when compared to 24°C under normal light. Kinase activities were localized to regions including the oral sensory papillae, acetabular ducts, tegument, acetabular glands, and nervous system. Furthermore, linoleic acid modulated PKC and ERK activities concurrent with the temporal release of acetabular gland components. Attenuation of PKC, ERK, and p38 MAPK activities significantly reduced gland component release, particularly in response to linoleic acid, demonstrating the importance of these signaling pathways to host penetration mechanisms.

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A Systematically Improved High Quality Genome and Transcriptome of the Human Blood Fluke Schistosoma mansoni

Anna V Protasio, Isheng J. Tsai, Anne Babbage … (2012)

Introduction Schistosoma spp. are platyhelminth (flatworm) parasites responsible for schistosomiasis, a tropical disease endemic in sub-tropical regions of Africa, Brazil, Central America, regions of China and Southeast Asia, which causes serious morbidity, mortality and economic loss. An estimated 779 million people are at risk of infection and more than 200 million are infected [1]. The paired adult males and females of S. mansoni reside in the hepatic portal vasculature, each female depositing 200–300 eggs per day near the intestinal wall. These eggs either pass into the gut lumen to be voided in the faeces and continue the life cycle or pass up the mesenteric veins and lodge in the liver, where they can cause serious pathology including granulomatous inflammation response and fibrosis. On contact with fresh water, free-living motile miracidia hatch from the eggs to infect aquatic snails (Biomphalaria spp.), where parasites undergo two rounds of asexual multiplication and are released as infective cercariae into water. Cercariae infect the human host, by penetrating unbroken skin, and transform into schistosomula. After several days the parasites exit the cutaneous tissue via blood (or lymphatic) vessels and travel first to the lungs and onward into the systemic vasculature. They may make multiple circuits before arriving in the hepatic portal system; only then do they start to feed on blood, mature and pair up, the whole process taking approximately five weeks [2]. Two Schistosoma draft genomes (S. mansoni and S. japonicum) were recently published [3], [4] and represent the only described genomes amongst parasitic flatworms to date. Their assemblies were generated by conventional capillary sequencing but are highly fragmented (S. mansoni, 19,022 scaffolds; S. japonicum, 25,048 scaffolds) and severely compromise gene prediction, as well as comparative and functional genomics analyses. The transcriptome has similarly only been partially characterised by large-scale expressed sequence tag (EST) sequencing and low-resolution cDNA-based microarrays. Second-generation sequencing technologies provide new opportunities to characterise both genomes and transcriptomes in depth. In addition to whole genome de novo sequencing [5], [6] and genome improvement [7], massively parallel cDNA sequencing (RNA-seq) can identify transcriptionally active regions at base-pair resolution [8]–[11] and accurately define the exon coordinates of genes [12]. In addition, the quantitative nature and high dynamic range of RNA-seq allows gene expression to be scrutinised [11], [13], [14] in a more sensitive and accurate way than other previous high-throughput methods [15], [16]. In this study we systematically improved the draft genome of S. mansoni, using a combination of traditional Sanger capillary sequencing, second generation DNA sequencing from clonal parasites and reanalysis of existing genetic markers [17]. This allowed us to assemble 81% of the genome sequence into chromosomes. We have also used RNA-seq data from several life-cycle stages to refine the structures of 45% of existing genes as well as to identify new genes and alternatively spliced transcripts. In addition to cis splicing, our data highlight extensive trans-splicing and provide clear evidence that the latter can be used to resolve polycistronic transcripts. With RNA-seq we profiled the parasite's transcriptome during its transformation from the free-living, human-infectious cercariae to the early stages of infection and in the mature adult. As the infective form transforms into a mammalian-adapted parasite, the relative abundance of transcripts shifts markedly during a 24-hour period, from those involved in glycolysis, translation and transcription to those required for complex developmental and signalling pathways. The improved sequence and new transcriptome data are available to the community in a user-friendly and easy to query format via both the GeneDB (www.genedb.org) and SchistoDB (www.schistodb.net) databases. These data demonstrate that revisiting a previously published draft genome, to upgrade its quality, is an option that should not just be reserved for model organisms. Materials and Methods The full description of materials and methods is presented in Supplementary Materials (Text S1). A synopsis of the methods used in this paper is presented below. Parasite material, library preparation and sequencing S. mansoni clonal DNA was obtained from single miracidium infections of Biomphalaria snails. Male and female adults (NMRI strain, Puerto Rican origin) were obtained from infected C57Bl/6 mice. DNA extraction was performed and sequencing libraries were prepared as previously described [18]. Eight and lanes were sequenced for the male samples and one lane for the female sample, both as 108-base paired reads. For RNA-seq samples, total RNA samples were obtained from cercariae, 3 hours and 24 hours post-infection schistosomula, and 7-week old mixed sex adult worms. Schistosomula samples were obtained using mechanical transformation [19]. RNA-seq libraries were prepared using a modified version of the protocol described in [8] and sequenced as 76-base paired reads. All samples were sequenced using the Illumina Genome Analyzer IIx platform. Raw sequence data were submitted to public data repositories; DNA reads were submitted to ENA http://www.ebi.ac.uk/ena/ under accession number ERP000385 and RNA-seq reads were submitted to ArrayExpress http://www.ebi.ac.uk/arrayexpress/ under accession number E-MTAB-451). Generating a new assembly and transferring previous gene annotation The Arachne assembler (version 3.2, [20]) was used to produce a new assembly using the existing capillary reads from the previously published draft assembly [3], supplemented with an additional ∼90,000 fosmid and BAC end sequences. FISH-mapped BACs [3] were also end-sequenced generating 438 reads that were incorporated into the assembly. Illumina reads were used to close gaps with the IMAGE pipeline [7]. The sequences of 243 published linkage markers [17] of S. mansoni were retrieved and used as anchors within the assembly by incorporating them as faux capillary reads. Scaffolds containing these reads were ordered, orientated and merged into chromosomes. Except where indicated, all analyses reported in the present study refer to a frozen dataset taken at this stage of the assembly process (S. mansoni genome v5.0, available at http://www.sanger.ac.uk/resources/downloads/helminths/schistosoma-mansoni.html). All comparisons were made against the previously published draft genome (v4.0). As part of the active finishing process, we randomly checked ∼20% (2,062) of the gaps automatically closed by IMAGE and found 90% of these could be verified by visual inspection. Contigs containing telomeric repeat sequences (TTAGGG) [21] were extended by oligo-walking pUC clones until a unique sequence was identified. Where the unique sequence was linked to a known marker, the telomere could be placed onto a chromosome. All manual improvement changes were included in a subsequent snapshot of the data (v6.0). To transfer the existing annotation to the latest reference we used RATT [22] (with the old assembly split into four parts and using options –q and –r) to define regions with synteny between both assemblies and transform the annotation coordinates onto the new assembly. The annotated genome sequence was submitted to EMBL http://www.ebi.ac.uk/embl/ under the accession numbers HE601624 to HE601631 (nuclear chromosomes); HE601612 (mitochondrial genome); and CABG01000001 to CABG01000876 (unassigned scaffolds). Gene finding using RNA-seq Each lane of RNA-seq reads was independently aligned to the genome using TopHat [23] and the resulting alignment files used as the input for the gene finder Cufflinks [12]. Transcript fragments with less than 10× average read depth coverage and fewer than 50 codons were excluded from subsequent analyses. JIGSAW [24] was used to combine existing models with Cufflinks' transcript fragments. The final set of gene models can be accessed through GeneDB http://www.genedb.org/Homepage/Smansoni and SchistoDB http://www.schistodb.net. Trans-splicing and polycistronic transcription RNA-seq read pairs that contained the splice leader (SL) sequence [25] were used to find trans-splicing sites; where a gene was found within 500 bases from a trans-splice site its transcript was tagged as putative trans-spliced. By looking for genes whose 3′ end was located within 2,000 bp upstream of a putative trans-spliced acceptor site, putative polycistronic units were identified. RT-qPCR was performed to validate both trans-spliced and polycistronic transcripts. Quantification of RNA-seq and differential expression RNA-seq reads were aligned to the reference genome using SSAHA2 [26]. A minimum mapping score 10 was applied to filter aligned reads. Reads per gene and RPKMs (reads per Kilobase per million mapped reads [8]) were calculated using only coding regions coordinates. We also estimated the background signal for non-coding regions (RPKM background RPKM) in pair wise comparisons (adjusted p-value 100 kb) and a further 114 unplaced scaffolds (∼1.1 Mb) that were W-specific. Repeats comprise 90% of the latter, and include previously identified female-specific repeat [32] as well as 0.1 Mb of previously uncharacterised female-specific sequences. These scaffolds usually have female reads mapped many fold higher than the average coverage of the assembly, for example scaffold 1570 has 26 times higher coverage than the average, suggesting that the heterochromatin portion of the W chromosome have been collapsed into these scaffolds. Based on the differences between the genome-wide assembly coverage and the coverage of these scaffolds, we estimate these heterochromatin portions of the W chromosome to comprise ∼3.3 Mb collapsed into the 1 Mb of consensus. Interestingly, the W-specific scaffolds appear to contain no coding genes whereas the Z-specific portion of Z/W sequence contains 782 genes, ∼95% of which exist as single-copies within the assembly. The mitochondrial genome Amongst the unassembled reads there were 5,647 that originated from mitochondrial DNA. An independent assembly of these reads using CAP3 [33] generated a single contig of 21 kb (to which 15 scaffolds from the previous genome assembly could be aligned). The first 14 kb of the contig was 99.9% identical to the published coding portion of the S. mansoni mitochondrial genome [34]. Based on restriction fragment analysis, a long non-coding region that is repetitive and highly variable between individuals has previously been partially characterised [35]. In our data, the additional 9 kb non-coding portion of the mitochondria genome is now complete and comprises known 62 bp repeats [35], plus additional 558 bp repeats and long tracts of low complexity sequence. Improvements to gene models using RNA-seq We obtained total RNA from four time points of the life cycle of S. mansoni: 1) free-living mammalian-infectious cercariae, mechanically transformed schistosomula at 2) three hours and 3) twenty hours post infection, and 4) seven-week old mixed-sex adults recovered from hamster host. The 183 million 76-base RNA-seq read pairs were mapped to the new reference genome using SSAHA2 alignment tool [26]. An average 70% of the RNA-seq reads generated in each sequenced library aligned as proper pairs to the genome (Table 2), an improvement over the previous version of the genome. Less than 6% of reads mapped to the mitochondrial genome in each sample; the lowest (0.5%) corresponding to the schistosomula stages. 10.1371/journal.pntd.0001455.t002 Table 2 Summary of RNA-seq mapping. Cerc 3 h Som 24 h Som Adult Total read pairs sequenced (out of 183,590,080) 69,498,003 53,041,873 50,528,949 10,521,255 Properly mapped read pairsa (%) 70.7 68.6 69.8 72.3 Additional properly mapped read pairs in new assemblyb (%) 2.0 0.2 0.4 2.8 Pairs mapped to repeats (%) 23.8 14.0 16.2 19.7 Pairs mapped to different scaffolds (%) 0.2 2.1 3.0 0.3 One mate mapped or mapped in wrong orientation (%) 4.3 12.2 9.7 6.1 Unmapped (%) 1.0 3.2 1.4 1.6 Proportion of reads mapped to mitochondria 5.1 0.6 0.4 3.7 Number of RNA-seq reads mapped using SSAHA2 to the genome from libraries prepared from cercariae (Cerc); 3-hour post-infection schistosomula (3 h Som); 24-hour post-infection schistosomula (24 h Som); and mixed male and female adult worms (Adult). a reads mapped within expected distance apart and in the correct orientation. b reads that were properly mapped to the new assembly but not in the previous. The majority (91%) of the 11,799 gene models from the previous version of the genome could unambiguously be transferred onto the new assembly. Splitting gene models from the previous assembly increased the gene count by 307; however, the coalescence of genes previously located on multiple different scaffolds caused some redundancy (an example is shown in Figure 2), removal of which reduced the number of transferred genes to 10,123. Of the 1,065 genes that could not be transferred to the new assembly, at least 83% were presumed to represent incorrect annotations due to a lack of sequence similarity and their short lengths, 1- or 2-exon structures (Figure S4) or a lack of start or stop codons. 10.1371/journal.pntd.0001455.g002 Figure 2 Removal of assembly redundancies produces a more reliable set of gene models. Gene models were migrated from previous version using RATT [22]. Repeats and sequencing errors in the old assembly resulted in ambiguities and sequences being represented more than once. In the new version, many scaffolds coalesced into one region and hence the gene models contained in them overlap each other. In this example, four supercontigs from the previous version collapsed on an unplaced region of Chromosome 3 in the new assembly. The smaller gene models are now obsolete as they were clearly incomplete annotations and their coding region are part of the exons of the larger gene model. RNA-seq data has been used to refine and improve gene model predictions in various organisms [10], [36], [37]. In the first draft of the S. mansoni genome, gene models were generated using a combination of ab initio gene predictions and EST evidence [38], with only a few hundred manually curated genes. To systematically upgrade the quality of annotations, we aligned pooled RNA-seq reads using TopHat [23], which allows gaps in the read-to-reference alignment at putative splice sites. Using the upgraded genome sequence 30% more RNA-seq reads with putative splice junctions aligned, highlighting putative new genes or structural refinements that could be made to existing genes. Cufflinks [12] was used to aid the refinement of gene structures by creating transcript “fragments” with sharply defined exon boundaries [23]. Using transcript fragments with at least 10 reads coverage at each base we found 78% of previous gene models had evidence of transcriptional activity within the sampled life cycle stages. Of these models, 3,604 (45%) were modified to include new exons derived from RNA-seq data, hence generating alternative gene predictions (Table 3). Using the transcript data as a guide, 236 genes were merged and 26 split into two or more gene models. 10.1371/journal.pntd.0001455.t003 Table 3 Fate of gene models. Number Total gene models in old genome version a 11,719 Not transferred 1,088 Deleted models 545 Split or merged models 731 Models with additional exons 3,438 Models that have been automatically replaced 1,116 New genes 504 Genes in new version b 10,852 The criteria for including genes into each category are described in the main text. a Version 4.0. b Version 5.0. To assess the accuracy of gene models, we calculated two metrics: the proportion of intron-exon junctions found in previous models that matched to the same intron-exon junction in a transcript fragment, and the proportion of the coding sequence in previous models that overlapped with the transcript fragments. Figure 3A is a heatmap showing these two metrics; existing models are clustered around top right of the plot, which indicates that RNA-seq evidence-based transcript fragments are similar to the existing models. Sixteen percent of gene models were perfectly reproduced by the transcript fragments (Figure 3B), while 90% of gene models with transcriptional evidence have at least 70% of the coding region covered by the transcript fragments. 10.1371/journal.pntd.0001455.g003 Figure 3 Improvement of gene annotation using RNA-seq. (A) Heatmap displaying comparisons between previous gene models and transcript fragments generated from Cufflinks. For each model, the extent of coding region that overlaps with a Cufflinks' model and the proportion of correctly predicted exon boundaries was calculated and categorised into bins of 70–100%. Models in this plot were excluded with less than 70% of their exon boundaries or coding regions predicted. (B), (C) and (D) Example scenarios of Cufflinks' models compared with previous gene models where (B) the Cufflinks prediction is identical to the 1,239 existing models; (C) Cufflinks fails to identify small introns; (D) Cufflinks removes incorrect introns present in the previous gene model, probably due to the improved assembly which, by correcting gaps, produced a longer single exon while the reading frame is preserved. In the new dataset, only 53% of gene models have at least 70% of their exon boundaries preserved. There are two main reasons for this low specificity in predicting exon boundaries. First, Cufflinks was unable to successfully predict the small introns typically observed in the 5′ end of many S. mansoni genes (Figure 3C and [3]). Consistently, when the first four exons of the old gene models were excluded, we found that transcript fragments could perfectly predict 90% of exon boundaries. Second, sequencing errors in the previous assembly resulted in introns being falsely incorporated into gene models during prediction to compensate for apparent frameshifts. These “intron” sequences are no longer necessary to preserve the reading frame and were identified as part of exons by Cufflinks in the new assembly (Figure 3D). For the two reasons above, we used JIGSAW [39] to combine existing models with those produced from RNA-seq data, resulting in 1,264 exon coordinates being changed. We identified 1,370 transcripts corresponding to putative full length coding sequences but which did not overlap with existing gene models. To check whether they indeed represented novel genes, we first screened them against known repeats and transposable elements. The 36 previously published transposable element sequences in S. mansoni matched 866 of the transcribed fragments, the longest of which (5,061 bp) was 99% identical to the coding portion of the LTR retrotransposon Saci-1 [40]. Of the remaining 504 complete transcript fragments we found sequence similarity for 231 in the NCBI nr protein database, mostly to other genes already annotated in S. mansoni (presumably representing gene duplications or members of multi-gene families) or S. japonicum. However, seven out of the remaining 273 full-length transcript fragments did show at least one conserved domain: a putative Tpx-1/SCP related allergen, a rhodopsin-like GPCR domain, a DNA-protein interaction domain, a epidermal growth factor-like (EGF-like) domain, and a polypeptide encoding a fascicline-like domain (FAS1) domain), and two transcripts with ArsR transcriptional regulator sequences. The new transcript fragments were on average shorter (261 bp) and exhibited unusual codon usage (Wilcoxon rank sum test, p<0.01, Figure S5) compared with a typical schistosome gene. Although we cannot rule out at this stage that the small set of atypical genes are non-coding RNA species, they are included in the total number of putative protein coding genes, which stands at 10,852. Trans-splicing Both cis and trans-splicing are used to produce mature transcripts in S. mansoni. By filtering for RNA-seq reads containing the spliced leader (SL) sequence [25], strongly supported trans-splicing events could be mapped on a genome-wide scale and highlighted 1,178 (∼11%) genes (an example is shown in Figure 4A), a figure in close agreement with a previous prediction [41]. For validation, we randomly chose ten putative trans-spliced gene models and could verify the existence of their trans-spliced transcripts by RT-PCR (Figure 4B, Table S1). In many cases, mapping information suggests a second trans-splicing acceptor site, usually within 20–50 bases from the primary acceptor site, indicating that alternative splicing operates at the trans as well as cis levels. Using Gene Ontology enrichment [30], we could find no particular functions or processes enriched within the trans-spliced genes, agreeing with the previous report [41]. 10.1371/journal.pntd.0001455.g004 Figure 4 RNA-seq reveals trans-spliced transcripts. (A) Schematic view of the 5′ end of trans-spliced gene Smp_176420. Shaded coverage plots represent non-normalized RNA-seq reads still containing the spliced-leader (SL) sequence (green – unclipped reads) and reads previously found to contain the SL sequence (orange - clipped). In the latter, the SL sequence was removed prior to aligning the reads to the genome; which improved the reads mapability (lower in the unclipped reads than in the orange reads). (B) RT-PCR validation of 10 putative trans-spliced genes with SL1 as forward primer and a gene-specific reverse primer. Smp_024110.1, previously described as trans-spliced [41], was included as a positive control (indicated with ‘+’) while Smp_045200.1 was included as a negative control (‘−’). All PCRs but one (Smp_176590.1) show bands corresponding to expected PCR product size. (C) Schematic view of the putative polycistron Smp_079750-Smp_079760. PCR1 represents the amplicon obtained from the unprocessed polycistronic transcript containing the intergenic region while PCR2 the trans-spliced form of Smp_079760. (D) RT-PCR validation of 5 putative polycistrons and a positive control (Smp_024110-Smp_024120; lane 9) previously reported in [45]. Each putative polycistron was subjected to two PCRs that correspond to PCR1 (e.g lane 1) and PCR2 (e.g lane 2) in panel C. Polycistronic transcripts originate from a single promoter but are later processed to generate two or more individual mRNAs. This type of transcriptional regulation is characteristic of trypanosomatids [42] and is present in C. elegans [43] and other organisms [44]. It has been suggested [45] that the S. mansoni Ubiquinol-cytochrome-c-reductase (UbCRBP) and phosphopyruvate hydratase (Smp_024120 and Smp_024110 respectively) genes might be transcribed as a polycistronic unit and that trans-splicing of the phosphopyruvate hydratase might resolve the polycistron into individual transcripts. In our study we provide strong evidence that this is indeed the case. One of the characteristics of polycistronic transcripts is a short intergenic distance (<200 bp) between individual “monocistrons”. We identified a total of 46 trans-splicing acceptor sites that fall between gene models that have a maximum intergenic distance of 200 bp, and 115 cases (Figure 4C, Table S2) where the intergenic regions expands up to 2 kb (maximum reported for C. elegans). We validated four of these polycistrons by RT-PCR (Figure 4D, Table S1) and Sanger sequencing (data not shown). Unlike C. elegans, which uses a second spliced leader (SL2) to resolve polycistrons [43], S. mansoni seems to use the same SL for both polycistronic- and non-polycistronic- trans-spliced transcripts. The role of polycistrons in schistosome gene expression remains to be determined but no pattern could be discerned between the ascribed functions of genes within each polycistron. Transcriptome analysis and differentially expressed genes In order to profile the transcriptional landscape of the parasite establishing in the mammalian host, the RNA-seq data from four key time points in the parasite's life cycle were analysed independently. Consistent with RNA-seq experiments elsewhere [16], we found good reproducibility between biological replicates, indicated by high correlation coefficients (average Pearson correlation of log RPKM values, across five pairs of biological replicates, was 0.95; Figure S6). A total of 9,535 (88%) genes were expressed (above an empirically determined background RPKM cut-off of 2 – Text S1 and Figure S7) in at least one surveyed time point and the remaining 12% were regarded as genes with expression too low to be detected or expressed during life stages not surveyed in this study (e.g. intra-molluscan stages) and therefore were excluded from further analysis. Of the excluded genes, 65% are annotated as hypothetical proteins (higher than the genome-wide figure of 44%). To gain better insight into the resolution of the RNA-seq approach in S. mansoni, we compared our results with a few example genes that have been described to undergo pronounce changes in their expression along the parasite's life cycle: an 8 kDa calcium binding protein, associated with tegument remodelling during cercariae transformation into schistosomula [46], [47]; a heat shock protein 70 (HSP70), active in schistosomula after penetration through mammalian host skin [48]–[50]; and the tegument antigen Sm22.6 [51], associated with resistance to re-infection in adult patients of endemic areas [52]. Our RNA-seq results broadly agree (Figure 5) with relative gene expression measurements obtained through other approaches. We also investigated how well the RNA-seq data correlate with previous microarray studies [53], [54]. Comparing normalised intensity values of the array features against the RNA-seq read depth for each microarray probe location in the genome (Figure S8) suggests that these data broadly correlate (Pearson's correlation of the log values 0.67). 10.1371/journal.pntd.0001455.g005 Figure 5 Comparison of expression of genes previously identified to be developmentally regulated. Barplots represent relative normalized reads (from RNA-seq data) for 3 transcripts, asterisks represent comparisons where differential expression is significant (adjusted p-value<0.01). Relative expression reported in the literature [46], [49], [51] is shown at the bottom (+++, high expression, ++ medium expression, + some expression, − not expressed, NA no information available). C = cercariae, 3S = 3-hour schistosomula, 24S = 24-hour schistosomula, A = adult. A total of 2,194 genes had detectable expression in at least one stage but not another and were therefore differentially expressed. We also used a pair-wise approach to analyse genes differentially expressed between the following life cycle stages: cercariae vs. 3-hour schistosomula, 3-hour schistosomula vs 24-hour schistosomula, and 24-hour schistosomula vs. adult. A total of 3,396 non-redundant transcripts (excluding alternative spliced forms) were differentially expressed (adjusted p-value<0.01) within the three pair wise comparisons (Table 4 and Table S3). An example showing differential expression between cercariae and 3-hour post-infection schistosomula is presented in Figure 6. To obtain a broad overview of the biological changes occurring at the gene expression level, we used Gene Ontology term enrichment to identify annotated functions and processes that were overrepresented in genes that were statistically (adjusted p-value<0.01) up- or down- regulated. Aerobic energy metabolism pathways were down regulated in schistosomules compared to cercariae and antioxidant enzymes were overrepresented in transcripts from adults. Three-hour post-infection schistosomula showed enrichment of transcripts involved in transcriptional regulation, G-protein coupled receptor (GPCR) and Wnt signalling pathways, cell adhesion and a considerable number of genes involved in potassium/sodium transport (Table S4). Most of the categories enriched at 3 hours post transformation persist through to 24 hours (e.g. GPCR signalling pathways). A total of 165 proteins are found to be associated with GPCR signalling pathways (annotated via GO). Of these, 30 and 18 were up regulated in 3 and 24 hours post-infection schistosomula, respectively, compared with cercariae. 10.1371/journal.pntd.0001455.g006 Figure 6 Detection of differentially expressed genes. The plot (left) shows the log fold change (y-axis) vs. log relative concentration (x-axis) for the cercariae – 3-hour schistosomula comparison. A total of 1,518 genes are differentially expressed between these two life cycles stages (adjusted p-value<0.01). On the right, example coverage plots for differentially and non-differentially expressed genes. Of particular interest, genes up regulated in the 3-hour schistosomula stage are enriched in G-protein coupled receptors and integrins, suggesting that signalling is a key process in this life-cycle transition. 10.1371/journal.pntd.0001455.t004 Table 4 Number of differentially expressed genes. Stage comparison Up regulated Down regulated Total Cercariae - 3 hour schistosomula 1,002 516 1,518 3 hour schistosomula - 24 hour schistosomula 433 595 1,028 24 hour schistosomula - adult 1,141 935 2,076 Figures refer to those genes with significant differential expression (adjusted p-value<0.01). NB the v5.0 assembly contains 10,852 genes. In order to investigate major processes occurring individually in each life cycle stage, we studied genes with expression above the 95 percentile in cercariae, 24-hour schistosomula and adults (Figure 7). Across the life cycle stages studied, some core cellular processes are consistently highly expressed, including glycolytic enzymes and protein translation but other broad changes are also apparent. Free-living cercariae utilise internal glycogen stores; accordingly genes involved in glycolysis and the tricarboxylic acid cycle (TCA) are highly expressed. After penetrating the skin and transforming into obligate endoparasites, the schistosomula switch to anaerobic metabolism [55], [56] before aerobic metabolism partly resumes in the adult. These events are also reflected in the transcriptome. At the schistosomulum stage there is a switch to high expression of L-lactate dehydrogenase, while TCA cycle transcription markedly decreases. As noted above, the cercariae and adult samples have relatively high contributions from the mitochondrial transcriptome (Figure S9) reflecting the high energy-demands of these two stages. 10.1371/journal.pntd.0001455.g007 Figure 7 Genes with expression above the 95 percentile different in cercariae and intra-mammalian stages. Venn diagram represents the distribution of genes above 95 percentile of expression in 3 different life cycle stages of the parasite. Examples of the genes/processes found within these groups are discussed in the main text. Other genes highly expressed in the schistosomula are involved in protein re-folding and chaperone function: 5 heat shock proteins (Smp_008545, Smp_035200, Smp_062420, Smp_072330, HSP70/Smp_106930) are among the top 50 most expressed genes at this stage and may reflect a response to the rapid temperature rise between fresh-water (∼28°C), in which the cercariae are found, and the warmer mammalian host (∼37°C). Within the host, schistosomes are exposed to potentially damaging reactive oxygen species produced during metabolism. Consistent with previous work [57] we found that antioxidant enzymes - particularly the peroxiredoxins (Prx1, Smp_059480 and Prx2, Smp_158110) - are highly expressed in adults, 24 hours after transformation and for Prx1, as early as 3 hours after transformation. Our results highlight the advantages of RNA-seq transcriptome profiling, especially its ability to dramatically improve the gene annotation alongside accurately recording changes in gene expression. Discussion In 2009 a draft genome of S. mansoni was published and provided a major resource for gene discovery and data mining. Our motivation for this study was to take S. mansoni's genome to the next level, to systematically upgrade its draft sequence so that gene structures can be more accurately predicted and the genomic context of genes can be better explored. Although systematic manual finishing has occurred for some parasite genomes, it is not an economically viable option for most non-model organisms. The genome of S. mansoni is approximately 10 times larger that the genomes of protozoan parasites and is set in the context of a field that attracts less funding. Although additional “traditional” targeted, long-range capillary sequence was introduced, more than 40,000 gaps were closed simply by re-sequencing at deep coverage, from a low-polymorphic population of adult worms. Further substantial changes were made from re-evaluating existing genetic marker information. As a result, the genome is measurably more accurate and its continuity has been transformed; 81% of the data is now assembled into chromosomes. We have also upgraded the annotation using deep coverage RNA-seq. Compared with the 2009 draft genome, the net change in the gene content is that there are now ∼900 fewer genes. However, 500 genes are new and more than 1600 low confidence or erroneous predictions have been removed. Across the genome, more than one third of genes now have new sequences. The value of the genome resource will therefore be tangibly improved: data mining approaches to identify genes will be more sensitive and trawling through kilobases of sequence for missing exons will be come less common. Our results also highlight the major benefit of using RNA-seq for transcriptome profiling - its ability to dramatically improve the gene annotation, whilst accurately recording changes in gene expression. We see major expected changes, for example, the well described metabolic switch on host penetration, plus some previously overlooked ones, such as a battery of receptors up regulated at the onset of infection in the mammalian host. Our data also define with high resolution some of the important building blocks of the schistosome transcriptome – long transcripts, cis and trans-splicing, and for the first time, clear evidence of the trans-splicing being used to resolve polycistrons. By increasing the quality of the genome, we have increased the utility of our RNA-seq data and taken it well beyond the levels attainable by previous microarray approaches. Although only a broad view of gene expression changes are presented herein, the resolution of our analyses reflects the functional annotation that has been previously ascribed. The true value of these data will arise from their use within the context of genome databases such as GeneDB and SchistoDB to query the behaviour of specific genes or groups of genes. The quality of a genome directly influences the uses to which it can be put and with many more, low-cost, draft-genome sequencing projects underway, the requirement for higher quality reference material, is increasing. Chain et al. 2009 recently defined several levels or standards for genome assemblies [58]. In the present study, we have taken an existing draft genome and demonstrated that in relatively modest period of time it can be upgraded to annotation-directed grade using second generation sequencing technology without the need for extensive manual finishing. The much improved genome assembly and gene structures, along with the expression data, are available at GeneDB and SchistoDB and will be an excellent resource not only for the helminth research community but also for in depth comparative genomics studies across metazoa. Supporting Information Figure S1 The frequency and length of newly inserted sequences at gaps. (PDF) Click here for additional data file. Figure S2 The S. mansoni v5.0 genome assembly superimposed over a genetic linkage map [17] . The numbers on the left of chromosomes are map distances in centimorgans, and the identifiers on the right of each chromosome denote contigs and scaffolds of assembly v5.0 (e.g. 6569_28 is contig 6569, which is assembled into scaffold 28). Lines connecting chromosomes indicate where an assembly scaffold contains contigs from two different chromosomes. There are multiple possible reasons for such occurrences, including repetitive sequences, assembly errors. All assembly ambiguities of this kind have been manually inspected and cannot be resolved using the current data. (PDF) Click here for additional data file. Figure S3 Analysis of male and female specific sequences. Sequence data from both Z and W chromosomes assembled together but was resolved by aligning male (blue) and female (red) genome sequence reads. The arrowheads indicate Z-specific genetic linkage markers. (PDF) Click here for additional data file. Figure S4 Plot showing (A) transcript length and (B) number of exons for the three different categories of gene models transfered using the Rapid Annnotation Transfer Tool (RATT). Outliers were not drawn in the boxplot. (PDF) Click here for additional data file. Figure S5 Codon usage of the (manually) curated genes and the 466 novel genes. (PDF) Click here for additional data file. Figure S6 Correlation between replicate experiments. Biological replicates are evaluated by calculating the Pearson's correlation for each pair of samples. (PDF) Click here for additional data file. Figure S7 Cumulative distribution of RNA-seq coverage (expressed as RPKM values, see Methods) for exons, introns, intergenic sequences and untranslated regions. (PDF) Click here for additional data file. Figure S8 Correlation of RNA-seq data and microarray data. The scatter plots show the coverage (Log2-transformed) of reads per probe location compared with normalized microarray intensities (Log2-transformed) from (A) Fitzpatrick et al. 2009 [54] and (B) Parker-Manuel et al. 2011 [53]. The graphs was generated using the smoothScatter function from the R software package [31]. (PDF) Click here for additional data file. Figure S9 Relative gene expression levels for mitochondrial genes. C = cercariae; 3S = 3 hour schistosomula; 24S = 24 hour schistosomula; A = adult. (PDF) Click here for additional data file. Table S1 Primers used for validation of trans-spliced (top) and polycistronic (bottom) transcripts. (XLS) Click here for additional data file. Table S2 Putative polycistrons with a maximum intergenic distance of 200 bp and 2000 bp. (XLS) Click here for additional data file. Table S3 Differentially expressed genes in the cercariae vs. 3 hr schistosomula comparison, 3 hr vs. 24 hr schistosomula comparison and 24 hr schistosomula vs. adult comparison. Only significantly differentially expressed transcripts (adjusted p.value<0.01 – BH correction) are listed. (XLS) Click here for additional data file. Table S4 Gene Ontology (Biological Processes) enrichment for differentially expressed genes in the cercariae vs. 3 hr schistosomula comparison, 3 hr vs. 24 hr schistosomula comparison and 24 hr schistosomula vs. adult comparison. The top 20 hits are shown. (XLS) Click here for additional data file. Text S1 Supplementary Materials and Methods. (DOC) Click here for additional data file.

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Protein kinase C: poised to signal.

Alexandra C Newton (2010)

Nestled at the tip of a branch of the kinome, protein kinase C (PKC) family members are poised to transduce signals emanating from the cell surface. Cell membranes provide the platform for PKC function, supporting the maturation of PKC through phosphorylation, its allosteric activation by binding specific lipids, and, ultimately, promoting the downregulation of the enzyme. These regulatory mechanisms precisely control the level of signaling-competent PKC in the cell. Disruption of this regulation results in pathophysiological states, most notably cancer, where PKC levels are often grossly altered. This review introduces the PKC family and then focuses on recent advances in understanding the cellular regulation of its diacylglycerol-regulated members.

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An Atlas for Schistosoma mansoni Organs and Life-Cycle Stages Using Cell Type-Specific Markers and Confocal Microscopy

James J. Collins, Ryan S King, Alexis Cogswell … (2011)

Introduction Flatworms of the genus Schistosoma are parasites (Phylum Platyhelmithes) that currently infect over 200 million people worldwide [1]. Similar to other trematodes, Schistosoma have complex life cycles consisting of both free-living and parasitic forms [2]. Upon passage from a vertebrate, schistosome eggs that reach freshwater will hatch and produce miracidia, that swim by ciliary movement to locate and penetrate a suitable snail host. Following entry into the snail, miracidia undergo a dramatic developmental conversion, resulting in the production of primary and then secondary sporocysts, which have the capacity to generate thousands of cercariae. Cercariae are then liberated from the snail into freshwater where they can penetrate the epidermis of a vertebrate host. These animals (now called schistosomula) enter the host's circulatory system, eventually migrating to the liver where they feed on blood and develop to adulthood as either male or female worms. Separate sexed worms then pair, begin reproducing, and complete the life cycle by laying eggs that are passed via the urine or feces depending on the schistosome species. Despite the daunting complexity of the schistosome life cycle, great strides have been made in developing modern tools to study these parasites. RNA interference has been used to disrupt gene function in eggs [3], [4], transforming cercariae [5], schistosomula [6], [7], [8], and adults [3], [9]. Such breakthroughs have opened the possibility for large-scale RNAi-based screens [8], [10]. Furthermore, transgenic approaches [11], and whole-mount in situ hybridization [15] techniques have been described, providing additional avenues to analyze gene function. These tools, combined with access to complete genome sequences for S. mansoni [16] and S. japonicum [17], will facilitate a greater understanding of these parasites, making this an exciting time for schistosome research. To fully realize the potential of these newly developed functional genomic tools, however, methods for analyzing changes in cell/tissue morphology following experimental perturbations will need to be developed. Furthermore, a collection of reagents to label distinct organ systems in a variety of life-cycle stages will facilitate a greater understanding of the complex developmental transitions these animals experience. One successful methodology for examining schistosome anatomy has been the combination of fluorescence microscopy with histological stains, such as carmine red or phalloidin. For example, carmine staining has been used to describe the reproductive anatomy of paired vs. unpaired adult females [18], [19] and phalloidin has been utilized to describe the musculature of a number of life-cycle stages [20], [21], [22]. Although these reagents will continue to be valuable tools, they are limited by their individual specificities; thus, there is a need to identify additional markers that will allow high-resolution analyses of distinct schistosome cell types and tissues. Immunofluorescence microscopy has become an indispensable tool for detailed examination of tissue morphology during development and following experimental perturbation. Although species-specific antibodies are widely available for classic model organisms (e.g. Drosophila and mouse), only a limited set of these reagents have been generated for organisms with smaller research communities (e.g. Schistosoma). In emerging model systems such as planarians, a free-living relative of parasitic trematodes, systematic examination of cross-reactivity with commercially available antibodies from other species [23], [24] has been used to overcome the limited access to species-specific immunological resources. Such analyses identified markers for the planarian nervous system, stem cells, protonephridia, intestine, and reproductive system. A similar approach has been used to identify antibodies that label the schistosome nervous system (e.g. [25], [26], [27]); however, this approach has not yielded useful markers for other cell and tissue types. Lectins are proteins that recognize specific carbohydrate moieties, and when conjugated to reporter molecules (e.g. enzymes or fluorophores) can be useful for labeling specific vertebrate [27] and invertebrate [28], [29] tissues. Although some lectins have been used previously to label schistosome tissues [30], [31], [32], [33], [34], detailed descriptions of their labeling have not been reported using modern methodologies such as confocal microscopy. Here we identify and characterize a collection of antibodies and lectins that label specific tissues of S. mansoni cercariae and use these stains to provide a detailed description of cercarial anatomy. We also examined both adults and miracidia and present a description of the protonephridial, reproductive and nervous systems in these stages using these tools. Together, these studies provide new tools and methods for studying these important parasites. Materials and Methods Obtaining free-living stages of S. mansoni Schistosome-infected (Puerto Rican strain NMRI) Swiss-Webster mice were provided by the Schistosome Resource Center. S. mansoni eggs were obtained from mouse livers essentially as previously described [35]. To obtain miracidia, eggs were resuspended in artificial pond water (0.46 µM FeCl3 ⋅6 H2O, 220 µM CaCl2 ⋅2 H2O, 100 µM MgS04 ⋅7 H2O, phosphate buffer [313 µM KH2PO4, 14 µM (NH4)2SO4] pH 7.2) and exposed to light either in a darkened side-arm flask [35] or a 24-well tissue culture plate [36]. S. mansoni cercariae were obtained from Biomphalaria glabrata snails (Schistosome Resource Center) by exposing snails to direct light at 28°C for ∼1–2 hours. B. glabrata snails were maintained in artificial pond water and fed Layer Crumbles (chicken feed) (Rural King, Mattoon, IL). In adherence to the Animal Welfare Act and the Public Health Service Policy on Humane Care and Use of Laboratory Animals, all experiments with and care of vertebrate animals were performed in accordance with protocols approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Illinois at Urbana-Champaign (protocol approval number 10035). Fixation and staining of miracidia and cercariae For fixation of miracidia and cercariae, an 8% formaldehyde solution was prepared by diluting 36% formaldehyde (EMD Chemical, Darmstadt, Germany) in artificial pond water. This solution was added to an equal volume of miracidia or cercariae (also in artificial pond water), agitated vigorously, and incubated for 20–25 m, except for staining with anti-synapsin where samples were fixed for 2 h. Typical volumes for fixation were >20 ml. Fixed animals either settled by gravity or were pelleted by a brief spin (∼5–10 s) in an Eppendorf 5810R centrifuge (Hamburg, Germany) with brake settings on 0 or 1. Following fixation, samples were rinsed in PBSTx (PBS + 0.3% Triton X-100). For subsequent steps, we found two convenient methods for specimen handing. First, samples were aliquoted to a 96-well microtiter plates and allowed to settle by gravity between liquid exchange steps. Alternatively, samples could be processed in 1.7 ml microcentifuge tubes and liquid exchanges could be performed either after gravity settling of the samples or following a brief spin ( 6 h at 4°C. Following >2 h of washing in PBSTx, samples were incubated in goat anti-mouse Alexa fluor 488 (Invitrogen, Carlsbad, Ca; 1∶400–1∶500) for 6 hours to overnight at 4°C. Occasionally, DAPI (1 µg/mL final in PBSTx) and phalloidin conjugated to either rhodamine or Alexa fluor 633 (66 nM final in PBSTx) were added during secondary antibody incubations or wash steps. As an alternative, we experimented with tyramide signal amplification (TSA) to detect antibody staining. For TSA detection, a goat anti-mouse HRP-conjugated secondary antibody (Invitrogen, Carlsbad, CA) was diluted (1∶100) in blocking solution and incubated with samples for 2 hr at room temperature. Samples were washed for 2 hrs in PBSTx and incubated in Amplification Diluent containing fluorescein tyramide (1∶50; TSA-Plus, Perkin Elmer, Waltham, MA). For anti-synapsin staining, TSA resulted in superior signal-to-noise ratios when compared to detection with Alexa Fluor-conjugated secondary antibodies (data not shown). Although this methodology was optimized for anti-synapsin staining, it is anticipated that similar results would be obtained with other murine antibodies. For lectin staining, animals were fixed and blocked as described above and then incubated overnight at 4°C with a fluorophore-conjugated lectin diluted 1∶500 in blocking solution. Lectins were obtained from Vector Laboratories (Burlingame, CA) and diluted from 2 mg/mL stocks. Samples were washed in PBSTx and stained with a combination of DAPI and Alexa fluor 633 phalloidin. Fixation and staining of adult S. mansoni Adult S. mansoni perfused from mice [35] were fixed for 30 minutes at room temperature in 4% formaldehyde diluted in PBSTx. Following brief rinses in 1x PBSTx samples were dehydrated through a methanol/PBSTx series (25%, 50%, 75%, and 100% MeOH) and stored at −20°C until use. Samples were rehydrated by incubation in 1∶1 MeOH:PBSTx followed by incubation in PBSTx. Rehydrated samples were treated with Proteinase K (2 µg/mL) for 10 minutes at room temperature and then post-fixed for 10 minutes in 4% formaldehyde in PBSTx. Samples were processed for immunohistochemistry and lectin staining similar to cercariae and miracidia. For staining with phalloidin, methanol dehydration and Proteinase K treatments were omitted. Imaging Samples were mounted in Vectashield (Vector Laboratories, Burlingame, CA) or a mixture of Vectashield and 80% glycerol and imaged on a Zeiss LSM 710 confocal microscope (Carl Zeiss, Germany) (Plan-Apochromat 63x/1.4 Oil DIC objective). Alexa 488/FITC, Alexa 568/Rhodamine, and Alexa 633 flours were excited with 488 nm, 561 nm, and 633 nm lasers, respectively. Images were processed either using Zen 2009 (Carl Zeiss, Germany), ImageJ [37], or Imaris (Bitplane AG Zurich, Switzerland). Movies were made using iMovie '09 (Apple, Cupertino, CA). Results To elucidate cell- and tissue-specific markers useful for anatomical analyses of Schistosoma, we examined a panel of antibodies, lectins and histological stains in S. mansoni cercariae. We chose cercariae over other stages because these animals are easily maintained in the laboratory and can be obtained in large numbers, facilitating the optimization of fixation and permeabilization conditions. Furthermore, since the cercariae possess many tissues present in other life-cycle stages (e.g. protonephridial and nervous systems), we predicted that cercarial markers would have utility in other stages. Below we report the results of these studies with our observations reported by organ system. Surface structures and musculature Cercaria are covered by a sugar-rich layer, called the glycocalyx, that sits upon the tegument, a modified syncytial epidermis characterized by a trilaminate membrane system [2], [38], [39]. In S. mansoni cercariae spines that face posterior and cover nearly the entire surface of the animal are prominent features of the tegument [20], [38]. Similar to previous reports [20], we find that these spines can be labeled with phalloidin (Figure 1A), consistent with these spines being rich in filamentous actin. Because the goal of this life-cycle stage is to locate and penetrate a suitable host, cercariae also possess a variety of ciliated sensory papillae [38]. We observed ciliated regions of unsheathed papillae scattered along the entire length of the animal by staining with antibodies specific to β-tubulin (Figure 1A, B) and acetylated α-tubulin (Table S1). Additionally, the openings of these ciliated papillae could be visualized as phalloidin-stained discs (Figure 1 A, B) that encircle cilia projecting through the tegument(Figure 1A, B). 10.1371/journal.pntd.0001009.g001 Figure 1 Superficial structures and musculature of cercariae. (A and B) Actin-rich spines and sensory cilia. (A) Anterior region of the head as seen by Differential Interference Contrast (DIC) optics, phalloidin staining to visualize actin, and immunofluorescence with an anti-β-tubulin antibody to label cilia. Images are maximum confocal projections. Bottom, overlay showing distribution of actin and β-tubulin. Arrow indicates an actin-rich channel though which a sensory cilium projects. (B) Maximum confocal projection of a cross section though the tail showing a sensory cilium (green) crossing the musculature (magenta) to project to the outside (right). (C and D) Single confocal sections depicting staining with lectin PSA that labels the basement membrane below the tegument. Panels C and D represent distinct confocal sections of the same animal. (E) Single confocal section through the head showing staining with lectins PSA and PNA and with phalloidin to visualize actin. PSA labels the basement membrane between the actin-rich surface spines and the muscle layer (yellow arrow). PNA marks a layer of material at the level of the actin spines which may represent the tegument or the associated glycocalyx (red arrow). (F) Phalloidin staining in various anatomical regions. Magenta box, actin spines and anterior sensory structures. Green box, longitudinal (white arrowhead), circular (yellow arrowhead), and diagonal muscle fibers (magenta arrowhead). Yellow box, acetabulum and the interface between the head and the tail. Note the intense radially symmetric spheres of phalloidin staining (cyan arrowhead); because of the proximity of these structures to longitudinal muscles in the head and tail, we suggest this staining may represent sites of muscle attachment. White box, longitudinal and helical muscles (green arrowhead) of the tail. Whole cercaria image represents a maximum projection derived from tiled stacks. Insets are magnified views of the indicated regions. All images are maximum projections generated from a Z-stack though an entire animal, except for the green inset that was derived from a subset of optical sections. (G and H) Depth projections showing phalloidin staining of the acetabulum and associated musculature. Scale shown below indicates the color-coding of distances from the ventral surface (i.e. the colors transition from red (ventral) to blue (dorsal) moving deeper into the animal). Panel G represents a dorsal view whereas panel H depicts a transverse section with the ventral surface towards the top. (I) Immunofluorescence with an anti-phospho S/T antibody that labels the longitudinal muscles of the tail. Scale bars, 10 µm. Anterior faces up in panel A and to the left in panels C, D, and F. The cercarial basement membrane serves as the interface for the attachment of the exterior tegument and the underlying body wall musculature. In cercariae this structure stained with a variety of lectins such as Pisum sativum lectin (PSA) (Figure 1C, D, E and Table S1). Double labeling with PSA and Peanut lectin agglutinin (PNA), revealed that PNA marked a layer of material outside the basement membrane at the level of the actin-rich spines embedded in the tegument. This PNA labeling could represent the tegument itself or the sugar-rich glycocalyx coating the outside of the cercarial tegument [38]. Similar to free-living flatworms, the body wall musculature of the cercariae consists of an external layer of circular muscles situated above longitudinal muscles that are interweaved with diagonal fibers [39]. Phalloidin stained all three muscle layers within the animal (Figure 1F and Movie S1) as well as other structures such as the acetabulum (ventral sucker) (Figure 1G and H), mouth, esophagus, and flame cells of the protonephridial system (described below). Since a previous study used phalloidin staining to describe the cercarial musculature [20], we refer readers to this reference for a detailed description of phalloidin staining. In addition to phalloidin staining, we were able to visualize the longitudinal muscles of the tail with an antibody that recognizes phosphorylated serine and threonine (anti-phospho S/T) residues (Figure 1I). Since we failed to observe anti-phospho S/T labeling in other cercarial muscle fibers, it is possible that these muscles are physiologically distinct from other muscles in the animal. Cercarial glands The five pairs of acetabular glands (three pairs of post-acetabular and two pairs of pre-acetabular) are prominent features of the cercarial anatomy, occupying nearly two-thirds of the volume of the cercarial head [38], [40]. The fundi of these unicellular glands fill the body cavity beneath the acetabulum and project anteriorly through the muscle cone, terminating at the anterior apex of the animal. Although the precise function of these glands has yet to be resolved, the secretion of proteinases rom the pre-acetabular glands [41] and mucus-like substances from the post-acetabular glands [42] suggests roles in host penetration and adhesion. Consistent with previous observations of live cercariae [34], a number of fluorescently labeled lectins stained the acetabular glands (Table S1, Figure 2A, Movie S2), both at their base and along their projections. While most of these lectins labeled the post-acetabular fundi, a subset stained both the pre- and post-acetabular glands. Among these was PNA, which labeled the pre- and post-acetabular glands as well as all ten of their projections (Figure 2A, B, C). Co-labeling with lectins (e.g. PSA or PNA) and phalloidin revealed a ring of musculature surrounding the acetabular ducts just posterior to the muscle cone (Figure 2A and Movie S2). Given the position of these muscles, they may act to initiate contractile forces that expel secretions from the ducts or to control secretion release. 10.1371/journal.pntd.0001009.g002 Figure 2 The secretory glands of cercariae. (A) Maximum confocal projections showing labeling of the entire pre- and post- acetabular gland system with lectins PNA and PSA. Yellow arrowheads indicate musculature surrounding the anterior ducts of the acetabular glands and white arrowheads indicate the muscle cone. (B and C) 3D-renderings showing magnified views of the acetabular gland ducts anterior to the muscle cone. The six post-acetabular gland ducts (yellow arrowheads) labeled with both PNA and PSA. PNA labeled the four pre-acetabular ducts (white arrowheads) whereas PSA did not. Panel B represents a ventral view; panel C represents a dorsal view. (D) Secretory globules within the post-acetabular glands labeled with PNA and PSA. (E) Immunofluorescence with an anti-acetylated α-tubulin antibody showing the microtubule-rich periphery of the pre- and post- acetabular glands. Arrowheads indicate putative sensory papillae. (F) sWGA labels the head gland. Left, DIC and DAPI staining. Head gland is pseudocolored orange and acetabular gland ducts are pseudocolored cyan. Center and right most panels show staining with sWGA. Scale bars, 10 µm. Anterior faces up in panels A, B, C and F and to the left in panel E. Double labeling with lectins PSA and PNA showed that these individual lectins label unique sub-cellular regions of the post-acetabular glands (Figure 2A). This was most obvious in the ducts, in which PNA staining was concentrated towards the periphery and PSA staining was detected throughout the duct (Movie S2). Furthermore, secretory globules [38], [40] scattered throughout the posterior regions of the glands were labeled to varying extents with either PSA and/or PNA (Figure 2D). This heterogeneous labeling might reflect individual differences in the content of these secretory globules. Ultrastructural studies describe a microtubule network surrounding the entire acetabular gland system [38], [40]. Consistent with these studies we were able to label the fundi and ducts of these glands with anti-acetylated α-tubulin (Figure 2E). In addition to the acetabular glands, cercariae possess a unicellular gland at the anterior end called the "head gland". This gland has been suggested to provide membrane material required during the cercariae-schistosomula transition following host penetration [43]. Similar to other cercarial glands, the head gland was readily visualized with a variety of lectins (Table S1, Figure 2F). The protonephridial system After being shed from their snail host and before locating and penetrating a mammal, cercariae spend a substantial portion of their short lives in freshwater. Cercaria may regulate water balance by controlling permeability through their outer surface (e.g. the tegument and/or glycocalyx) and by excretion of excess fluid by a network of osmoregulatory tubules called protonephridia [44], [45]. Proximally the protonephridia begin as a heavily ciliated cell called a flame cell. Staining of cercariae with antibodies that recognize various tubulin isoforms and modifications showed labeling of the ciliary tufts of flame cells (Figure 3A and C). Anti-tubulin antibodies also labeled several other structures, including portions of the nervous system (see below); proteinase K treatment often abolished this labeling, leaving predominantly protonephridial labeling (Table S1). For example, staining with the anti-β tubulin antibody following Proteinase K treatment provided robust and specific labeling of the flame cells (Figure 3A arrowheads) and ciliated secondary protonephridial tubules (Figure 3A arrows), allowing for easy identification of these cells even by epifluorescence. In contrast to a previous report stating that cercariae have six pairs of flame cells [38], we find that cercariae typically have 5 pairs of flame cells: an anterior-body dorsal pair, a mid-body ventral pair, posterior-body dorsal and ventral pairs, and a pair in the anterior tail (Figure 3A) similar to that described by Skelly and Shoemaker [46]. The base of the flame cell contains a region of tightly packed ciliary rootlets, which label strongly with an anti-phospho S/T antibody, and a nucleus, which is readily identified by DIC microscopy or DAPI staining (Figure 3C). The ciliary tuft of the flame cells sits in a barrel or basket-like structure formed by interdigitation between the flame cell and first tubule cell [38] and labels strongly with an anti-phospho tyrosine (anti-phospho Y) antibody (Figure 3B and 3C). In addition to labeling the flame cell and first tubule cell, anti-phospho Y also strongly labels the protonephridial tubule extending from the bladder to the nephridiopores located at the tips of the tail furci (Figure 3B). The protonephridial tubule splits anterior to the bifurcation of the tail (Figure 3B; blue inset). Although flame cells in the head were labeled by the anti-phospho Y antibody, it was unclear whether the protonephridial tubules were also labeled, since this antibody marked a variety of structures in the head (Table S1 and Movie S3). 10.1371/journal.pntd.0001009.g003 Figure 3 The protonephridial system of cercariae. (A) Immunofluorescence with an anti-β-tubulin antibody to label the ciliated tufts of the flame cells (yellow arrowheads) and ciliated regions of the protonephridial tubules (white arrows) within the head and tail. Cercariae typically have 10 flame cells, an anterior dorsal pair, a mid-head ventral pair, posterior head dorsal and ventral pairs, and a pair at the anterior tail. Dorsal view is shown to the left and a lateral view is shown to the right. (B) Immunofluorescence with an anti-phospho Y antibody that labels the barrel of the flame cells and the excretory duct. Inserts are magnified views of the indicated regions. Yellow box, flame cells of the tail. Blue box, the protonephridial tubule splits anterior to the point of tail bifurcation. Green box, the protonephridial tubule extends to the nephridiopore at the tip of the tail. Mid-level DIC optical section showing nephridiopore and tegumental structure at tip of tail. (C) Numerous reagents utilized in this study labeled portions of the flame cells providing useful tools for analyzing flame cell morphology. The ciliated tuft stained with various anti-tubulin antibodies (top row). Several lectins showed different staining patterns of the extracellular matrix surrounding the flame cell (middle row). Phospho-specific antibodies (bottom row) also labeled the flame cells: anti-phospho S/T labeled the ciliary rootlet and anti-phosopho Y labeled the flame cell barrel. Differential interference contrast optics also permit observation of flame cell morphology. (D and E) Portions of the protonephridial tubules in the head label with lectin sWGA. (D) Maximum intensity projection of dorsal focal planes showing sWGA labeling of protonephridial tubules (white arrows) leading from the anterior flame cell pair (yellow arrowheads). (E) Cross-section of a maximum intensity projection from animal in D showing the sWGA-labeled protonephridial tubules (arrows) positioned dorsal to the sWGA-labeled acetabular ducts. Abbreviations: Lens culinaris agglutinin (LCA), peanut agglutinin (PNA), Pisum sativum agglutinin (PSA), succinylated wheat germ agglutinin (sWGA). Scale bars, 10 µm. Anterior faces up in all panels. Both the flame cells and first tubule cells contain an actin-rich network that is easily visualized using fluorescently labeled phalloidin (Figure 3C). Interestingly, several fluorescently labeled lectins showed differential staining of the extracellular matrix (ECM) surrounding the flame cells (Figure 3C). Lens culinaris agglutinin (LCA) labeled the ECM surrounding the first tubule cell down to the level where this cell interdigitates with the flame cell (Figure 3C). PNA showed labeling of the entire ECM surrounding the first tubule cell and flame cell. PSA labeling was restricted to the ECM surrounding the junction between the first tubule cell and flame cell. It has long been appreciated from ultrastructural observations that platyhelminth flame cells are surrounded by a significant amount of ECM [44]; our findings indicate surprising complexity in the composition of this material. We also observed lectin staining of portions of the protonephridial ducts in the cercarial head (Figure 3D and E and Table S1). The nervous system The stereotypical flatworm central nervous system consists of paired cephalic ganglia that connect to nerve cords which extend through the body longitudinally [39], [47]. These nerve cords are often connected by transverse commissures, giving the flatworm nervous system a "ladder" or "orthogonal" appearance. Antibodies that recognize various neurochemical signaling molecules (e.g. neuropeptides) have been invaluable reagents for describing the neural anatomy of a variety of free-living and parasitic flatworms [47]. However, since the neurosignaling molecules recognized by these antibodies are often expressed in restricted cell types in flatworms [48], we explored additional markers that recognize features found more generally in neurons. A monoclonal antibody raised against the D. melanogaster Synapsin-1 protein, that is concentrated in nerve terminals of the fly [49], has been used to label the neuropil of the cephalic ganglia, the nerve cords, the transverse commissures, and the sub-muscular plexus of the planarian S. mediterranea [24]. We find this antibody labels similar structures in the central nervous system of cercariae. Specifically, anti-synapsin staining allowed visualization of the neuropil of the cephalic ganglia (Figure 4A, B, C, D and Movie S4), the six-pairs of longitudinal nerve cords (Figure 4A, B, C), and transverse commissures in the head (Figure 4A, B, C). Additionally, anti-synapsin labeled anterior sensory structures, nerves surrounding the acetabulum, and a pair of nerve cords below the longitudinal muscles of the tail. Of particular interest, we observed projections from the cephalic ganglia innervating muscles surrounding the ducts of the acetabular glands (Figure 4E and Movie S4). This innervation suggests that signals directly from the cephalic ganglia may be involved in controlling secretion from the acetabular glands during penetration. 10.1371/journal.pntd.0001009.g004 Figure 4 The central nervous system of cercariae. (A) Immunofluorescence with an anti-synapsin antibody that labels the cephalic ganglia and many peripheral neural structures. Below, synapsin labeling is shown together with phalloidin staining. (B) Maximum confocal projections generated from ventral, medial, or dorsal focal planes. Anatomical regions are given above and staining reagents are listed to the left. (C) Depth projection showing synapsin staining in the CNS. Ventral view is shown above and a lateral view is shown below. Scale shown below indicates color-coding of distances from the ventral surface. Abbreviations: Dorsal nerve cords (DNC), cephalic ganglia (CG), ventral nerve cords (VNC), lateral nerve cords (LNC). For the sake of simplicity we have not distinguished anteriorly projecting cords from dorsally projecting cords. (D) Single confocal section though the cephalic ganglia, showing the central neuropil surrounded by neuronal nuclei. (E) Confocal projection showing innervation between the cephalic ganglia and the musculature surrounding the anterior ducts of the acetabular glands (magenta arrowheads). Scale bars, 10 µm. Anterior faces left in panels A, B, and C and up in panels D and E. In addition to anti-synapsin staining, we found that anti-β-tubulin was useful for detecting the nervous system within the tail (Figure 5A, B, C). However, unlike anti-synapsin that labels synaptic regions, anti-β-tubulin stained microtubule-rich neural projections throughout the tail. These fine projections could be observed transversely projecting through the interior of the tail (Figure 5A), superficially between the tegument and the muscles (Figure 5B), and within a pair of cords beneath the longitudinal muscle fibers (Figure 5C). 10.1371/journal.pntd.0001009.g005 Figure 5 The nervous system of the cercarial tail. (A) Extensive neural projections within the tail visualized by β-tubulin immunostaining. Overlay with phalloidin and DAPI show the position of the nerves relative to the tail musculature and nuclei, respectively. (B) Superficial neural projections (green) laying outside muscle layer (magenta). Arrowhead indicates a sensory papilla. (C) Longitudinal nerve cord (white arrowhead) running along a longitudinal muscle within the tail. Scale bars, 10 µm. Anterior faces left in all panels. Reagents useful for analyzing cercaria anatomy can also be used in adults and miracidia Following host penetration, S. mansoni cercariae (now schistosomula) enter the circulatory system and develop within the lungs before ultimately residing in the vessels of the hepatic portal system. Because cercariae are essentially the template from which the adult animals develop, we examined markers particularly useful for labeling cercariae (Lectins sWGA and PNA and the anti-acetylated α-tubulin antibody) in adult S. mansoni. Consistent with PNA labeling of secretory glands in cercariae (Figure 2A), we observed intense PNA staining of secretory glands surrounding the esophagus in the anterior of male and female worms (Table S1 and Figure 6A). Also similar to cercariae (Table S1), we observed sWGA labeling in the neuropil of the cephalic ganglia (Figure 6A, B) and the extracellular matrix/basement membrane of both sexes. Although adult schistosomes likely rely on the host for regulating fluid balance [44], these animals possess an extensive protonephridial system that consists of flame cells and an elaborate system of ciliated and unciliated ducts [50]. Like cercariae, the ciliated flame cells were detected with anti-acetylated α-tubulin and the ECM of the unciliated ducts stained with sWGA (Figure 6A–D, and Movie S5). Furthermore, anti-acetylated α-tubulin labeled the extensive network of ciliated collecting ducts (Figure 6B–D). Double labeling with sWGA and anti-acetylated α-tubulin allowed for staining of the complete protonephridial system (Figure 6B and D, and Movie S5). 10.1371/journal.pntd.0001009.g006 Figure 6 Labeling of non-reproductive tissues in adult S. mansoni. (A) Lectin PNA staining of the esophageal gland cells. Lectin sWGA labeling of cephalic ganglia and protonephridial ducts in a female. Image is a maximum intensity projection. (B) Lectin sWGA labeling of cephalic ganglia and non-ciliated protonephridial ducts of a male. Also shown is anti-acetylated α-tubulin staining the ciliated regions of the protonephridial system. Image is a maximum intensity projection. (C) Maximum intensity projection depicting anti-acetylated α-tubulin staining of the ciliated regions of the male protonephridial system, including the flame cells and the collecting ducts. Note that many secondary and tertiary ciliated ducts funnel into one of the two main collecting ducts that run along the longitudinal axis. (D) Maximum intensity projection of a protonephridial unit labeled with anti-acetylated α-tubulin and lectin sWGA. Anti-acetylated α-tubulin marks the ciliated flame cells and ducts, whereas sWGA labels non-ciliated tubules. Scale bars, 10 µm. Anterior faces to the upper left in panels A and B. We also found sWGA, PNA and anti-acetylated α-tubulin to be useful for visualizing the male and female reproductive organs. The S. mansoni male reproductive system consists of four to nine testes [2], [51]; the sperm they produce are passed to the female via a cirrus at the anterior end of the gynecophoral canal (Figure 7A). Cortical microtubules surrounding the sperm head and within the sperm flagella were visualized within testes lobes by staining with anti-acetylated α-tubulin (Figure 7B, C). Additionally, we observed sWGA labeling tubular (doughnut shaped) structures between a subset of nuclei in the testes (Figure 7B, C). Because developing germ cells in other animals often remain attached to one another by cytoplasmic bridges (e.g. ring canals [52]), we speculate that these structures could represent canals connecting the cytoplasm of germ cells undergoing spermatogenesis. Spermatozoa passed to a female migrate up the female reproductive tract and are stored in the seminal receptacle located posterior to the ovary; tightly packed sperm were observed in this receptacle by staining with DAPI and anti-acetylated α-tubulin (Figure 8A, B). Occasionally, we also observed “packets” of sperm within the oviduct en route to the seminal receptacle (data not shown). From the seminal receptacle sperm are able to begin the fertilization process as oocytes emerge from the ovary. Oocytes then pass anteriorly though the oviduct (Figure 8A), which merges with the vitelline duct (now the ovo-vitelline duct), before reaching a mass of secretory cells collectively referred to as Mehlis' gland. A variety of functions have been proposed for Mehlis' gland, including providing lubrication for the reproductive tract, activating sperm, and providing aterials for egg shell biosynthesis [53], [54]. Similar to histological [51] and ultrastructural descriptions [55], [56], we observe this gland directly posterior to the ootype (Figure 8C). This observation contrasts that of Neves et al. (2005) [18] who describe the columnar epithelium of the ootype (described by Erasmus [55] and Gönnert [51]) as Mehlis' gland. We found the projections of Mehils' gland cells to label with both sWGA and PNA (Figure 8C, D and Movie S6). Furthermore, we observed anti-acetylated α-tubulin to label microtubules (described previously by Erasmus [55]) at the distal projections of these cells before they enter the ootype (Figure 8C and Movie S6). In cross section the cells of Mehlis' Gland labeled with sWGA, PNA, and anti-acetylated α-tubulin can be seen projecting radially into the posterior ootype (Movie S6). Within the ootype the eggshell coalesces to surround a single fertilized egg and 30–40 vitelline cells [57] (Shown in Figure 8C and Movie S6). This egg is passed anteriorly through the uterus and out the female genital pore. 10.1371/journal.pntd.0001009.g007 Figure 7 The male reproductive system. (A) Staining with DAPI and phalloidin showing the male head and various parts of the male reproductive system. Inset, magnified view of male cirrus. Image represents a maximum intensity projection derived from tiled stacks. Anterior to left, dorsal towards top. (B) Single confocal plane showing a testes lobe stained with anti-acetylated α-tubulin, sWGA and DAPI. DAPI staining shows the nuclear morphology of cells in different stages of spermatogenesis and anti-acetylated α-tubulin (green) stains the flagella and cortical microtubules of sperm. sWGA stains tubular structures that could represent cytoplasmic bridges between cells in mitotic and/or meiotic stages of spermatogenesis. (C) Magnified view of panel B. Scale Bars, 10 µm. 10.1371/journal.pntd.0001009.g008 Figure 8 The female reproductive system. (A) Top, female labeled with DAPI and phalloidin to show various regions of the female reproductive system. Autofluorescence from the vitelline cells is shown in green; this autofluorescence served as a useful marker for the vitelline duct and its contents. Image represents tiled images of confocal sections from a single animal. Yellow and cyan boxes correspond to the regions of the seminal receptacle and the ootype/Mehlis' gland complex, respectively. These boxes are color coded to indicate the relative positions of the structures shown in either panels B (yellow bar in upper left) or C (cyan bar in upper left). (B) Left, anti-acetylated α-tubulin labeling microtubules of sperm present in the seminal receptacle. Right, large nuclei of oocytes in the ovary can be seen stained with DAPI to the left. Dashed line represents the position of the of muscle layer surrounding the ovary. Images are from mid-level maximum intensity projections. (C) Maximum intensity projections showing staining of the ootype (top, shown surrounding an egg) with sWGA and Mehlis' gland with PNA, sWGA and acetylated α-tubulin (bottom). Arrowheads indicate cytoplasmic projections of Mehlis' gland cells. (D) Magnified view of Mehlis' gland from panel C. Scale Bars, 10 µm. Anterior faces left in panel A and up in panels B, C and D. Upon release into freshwater, eggs generated by the adult schistosome hatch, giving rise to free-living miracidia. Following hatching, miracidia must locate and infect an appropriate snail host. Like the cercariae, miracidia contain a number of ciliated sensory structures on their surface [58]; these can also be labeled with anti-tubulin antibodies (Figure 9A, arrowheads). In addition to labeling the multiciliated sensory papilla, anti-tubulin antibodies also label the numerous motile cilia of the epidermal plates that cover the animal. Interestingly, anti-β-tubulin also showed strong labeling of a cytoplasmic meshwork of microtubules in the germ cells (Figure 9B). Previous ultrastructural analysis of miracidial germ cells suggest that these cells may be anchored through several small projections that extend into the intercellular space [58]. It is possible that the microtubule meshwork we observed functions to generate or maintain these cellular projections. 10.1371/journal.pntd.0001009.g009 Figure 9 The organ systems of miracidia. (A) The miracidial surface is covered by motile cilia on the epidermal plates as well as multiciliated sensory papilla (arrowheads) visualized by staining for β-tubulin and DIC optics. (B) Mid-level confocal section showing microtubule meshwork of germ cells (asterisks). (C) Anti-phospho S/T strongly labels the terebratorium and epidermal ridges surrounding the first tier of epidermal plates. (D) Anti-phospho S/T also displayed a weaker circumferential banding pattern similar to circumferential muscle (top). Optical crossections indicate that the weak, superficial anti-phospho S/T labeling is at the level of circumferential muscles (arrowhead) but not longitudinal muscles (arrows). (E) Similar to cercariae, anti-phospho S/T strongly labels the base of flame cells (arrowheads) in miracidia. (F and G) Immunofluorescence with an anti-synapsin antibody labels the cephalic ganglia and peripheral nerve structures. (F) Mid-level confocal section showing labeling of neuropil of the cephalic ganglia with anti-synapsin (green). The neuronal cell bodies of the cephalic ganglia contain small nuclei that stain intensely with DAPI (grey) and surround the neuropil. Phalloidin staining (magenta) labels the muscle as well as flame cells. (G) Maximum intensity projection of a miracidium stained with anti-synapsin antibody. Abbreviations: Cephalic ganglia (CG), nerve cords (NC). (H) Volume rendering of miracidium stained with PNA showing labeling of lateral glands and ducts. Scale Bars, 10 µm. Anterior faces left in all panels except C that represents a view from the anterior surface. Phalloidin staining of miracidia facilitates analysis of the elaborate body wall musculature and flame cells (Figure 9C–E, and Movie S7). A detailed description of phalloidin staining has previously been reported, and we refer readers to that work [59]. In cercariae we observed anti-phospho S/T staining of the longitudinal muscles in the tail (Figure 1I). Similarly we observed anti-phospho S/T staining in the miracidia in a circumferential banding pattern similar to that observed for phalloidin staining of the muscle (Figure 9D). It was difficult to ascertain whether this labeling was in the muscle layer itself or in the overlying epidermal plates and ridges. In addition to the superficial banding pattern, we observed anti-phospho S/T labeling of the base of flame cells (Figure 9E bottom arrowheads). The central nervous system of miracidia resembles that observed in cercariae. The neural mass is centrally located, stains with the anti-synapsin antibody, and is surrounded by neuronal cell bodies that are identifiable by their small ovoid nuclei which stain intensely with DAPI (Figure 9F and Movie S7). Several nerve cords project anteriorly while only a dorsal and ventral nerve cord project posteriorly (Figure 9F, G). Miracidia contain three glands, an apical gland and two lateral glands [58]. Although we observed staining of the lateral glands with a number of the lectins (Figure 9H and Table S1), only a few lectins were observed staining the apical gland (Table S1). This suggests that the products of the apical gland are likely different than those of the lateral glands, an observation consistent with ultrastructural studies demonstrating differences in size of secretory vesicles in these glands [58]. While several lectins stained ECM and other non-secretory cells in cercariae and adults, we failed to observe significant staining of other cell types in miracidia. While this may reflect real biological differences in miracidial tissues compared to cercarial or adult tissues, it is also possible that our fixation protocol for miracidia either blocked epitopes or failed to fix them sufficiently. Discussion These studies describe a number of robust cell-specific labels for three life-cycle stages of S. mansoni. One interesting observation from these studies was that a diverse collection of fluorescently labeled lectins stained secretory glands in all life-cycle stages observed. Specifically, lectins stained the acetabular and head glands of cercariae, the esophageal and Mehlis' gland of adults, and the apical and lateral glands of miracidia. These observations are consistent with previous studies describing lectins, including PNA and WGA, labeling glands, or their secretions [31], in live miracidia and cercariae [34]. Because many of these same lectins label secretory cell types in planarians [29], it is possible these types of cells share a common evolutionary origin and their functions have diverged to accommodate the free-living or parasitic lifestyles of these animals. Many of the reagents we examined labeled portions of the protonephridial system at a number of life-cycle stages. While one of the major functions ascribed to protonephridia is osmoregulation, the system is not only maintained but becomes elaborated in adults when the parasite conforms to the osmolarity of its host [44]. Recent studies demonstrating uptake of molecules including fluorescent analogs of Pgp/MRP substrates and a fluorescent derivative of praziquantel [60], [61], [62], [63] suggests that the protonephridial system is likely important in drug metabolism and excretion. The combination of morphological markers we have identified, along with fluorescent dye labeling experiments, will be instrumental in analyzing protonephridial phenotypes in future drug and RNA interference studies. Since egg-induced granuloma formation is the principle cause of the pathology of schistosomiasis, understanding reproductive processes in schistosomes could illuminate new therapeutic opportunities. Using lectins, an antibody against acetylated α-tubulin, and two common fluorescent stains (DAPI and phalloidin, which has been described previously [21]) we were able to clearly visualize the gonads (testes and ovaries) and most accessory reproductive structures (e.g. ootype and Mehlis' gland) in adult schistosomes. The combination of these reagents provides a step forward from more general fluorescence-based approaches previously used to describe the schistosome reproductive system (e.g. carmine red staining [18]), and should provide a useful tool for accessing reproductive development in these parasites. Collectively, these studies show the utility of this approach for identifying robust markers of schistosome tissues. We anticipate that similar studies focused on other life-cycle stages (e.g. schistosomula and sporocysts) will yield comparable results and will provide additional tools that will enable researchers to dissect the many fascinating biological questions raised by these important parasites. Supporting Information Movie S1 Phalloidin staining of S. mansoni cercariae. Shown are Z-stacks through the head of a cercariae stained with phalloidin. DIC optics and DAPI staining are also shown overlaid with phalloidin staining. (MP4) Click here for additional data file. Movie S2 Pre- and post-acetabular glands. First, Z-stacks though the head of a cercariae labeled with lectins PNA and PSA. Overlay with phalloidin staining is also shown. Second, 3D rendering showing the distal projections of the acetabular glands at the anterior of the animal. (MP4) Click here for additional data file. Movie S3 Whole cercaria view of anti-phospho Y immunofluorescence. First, a Z-stack showing immunofluorescence with the anti-phospho Y antibody. Second, a 3D rendering of anti-phospho Y immunostaining. (MP4) Click here for additional data file. Movie S4 Visualization of the cercariae CNS with anti-synapsin. First, Z-stacks showing immunofluorescence staining with anti-synapsin in a cercarial head. Overlay with DAPI, DIC and phalloidin are also shown. Second, 3D renderings of anti-synapsin immunostaining. (MP4) Click here for additional data file. Movie S5 The protonephridial system of adult S. mansoni . First, 360° rotation of a 3D rendering with depth coding showing ciliated regions of the protonephridial system visualized by immunostaining with anti-acetylated α-tubulin. In this projection warmer colors (e.g. reds) represent more superficial structures, whereas cooler colors (e.g. blues) correspond to deeper structures. Second, a 360° rotation of a protonephridial unit and its associated ducts visualized by staining with sWGA and anti-acetylated α-tubulin. (MP4) Click here for additional data file. Movie S6 The ootype/Mehlis' gland complex. First, Z-stacks showing various regions of the ootype and Mehlis' gland visualized with DIC optics and DAPI, PNA, sWGA, and anti-acetylated α-tubulin stainings. Second, a movie of optically derived cross-sections through Mehlis' gland and the ootype. Sections are shown moving from the posterior end of Mehlis' gland towards the anterior of the ootype. (MP4) Click here for additional data file. Movie S7 3D renderings of miracidia. 3D renderings of miracidia stained with phalloidin, anti-synapsin and DAPI. (MP4) Click here for additional data file. Table S1 Reagents used in this study and their labeling patterns in Schistosoma mansoni. (PDF) Click here for additional data file.

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Journal ID (nlm-ta): J Infect Dis

Journal ID (iso-abbrev): J. Infect. Dis

Journal ID (publisher-id): jid

Journal ID (hwp): jinfdis

Title: The Journal of Infectious Diseases

Publisher: Oxford University Press

ISSN (Print): 0022-1899

ISSN (Electronic): 1537-6613

Publication date (Print): 01 December 2015

Publication date (Electronic): 23 September 2015

Publication date PMC-release: 23 September 2015

Volume: 212

Issue: 11

Pages: 1787-1797

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[1 ]Molecular Parasitology Laboratory, School of Life Sciences, Kingston University , Kingston Upon Thames

[2 ]Department of Life Sciences, The Natural History Museum , London, United Kingdom

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Correspondence: Anthony J. Walker, PhD, School of Life Sciences, Kingston University, Kingston Upon Thames, Surrey KT1 2EE, UK ( t.walker@ 123456kingston.ac.uk ).

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Publisher ID: jiv464

DOI: 10.1093/infdis/jiv464

PMC ID: 4633769

PubMed ID: 26401028

SO-VID: f1af01b0-b3f1-4c72-8a1b-09dcb8ebbb97

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This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial-NoDerivs licence ( http://creativecommons.org/licenses/by-nc-nd/4.0/), which permits non-commercial reproduction and distribution of the work, in any medium, provided the original work is not altered or transformed in any way, and that the work is properly cited. For commercial re-use, please contact journals.permissions@ 123456oup.com .

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Date received : 3 June 2015

Date accepted : 11 September 2015

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Funded by: National Institutes of Allergy and Infectious Diseases (NIAID)

Funded by: NIAID

Funded by: National Institutes of Health http://dx.doi.org/10.13039/100000002

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