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Additional file 3: Figure S17. DBP1 alignment. Complete alignment of DBP1 protein sequences. The presence of the DBL domain (Pfam: PF0311) and the trans-membrane domain is indicated.

Additional file 4: Figure S25. RBP2a alignment. Complete alignment of RBP2a protein sequences.

Additional file 2: Figure S1. Apicoplast and mitochondrial genomes. Circos plots of the assembled apicoplast (top) and mitochondrial (bottom) genomes. The presence of protein-coding genes (grey), ribosomal-RNAs (green), and transfer-RNAs (red) is denoted by bars. For each gene category, outermost circle denotes genes on the forward strand, and the innermost circle denotes genes on the reverse strand. The two circular representations nearest to the center represent GC-skew and GC-content, respectively. Figure S2. Busco assembly assessment. Gene content in Plasmodium simium and other Plasmodium genome assemblies. BUSCO was run using the eukaryota odb9 data set containing 303 BUSCO groups. Figure S3. Protein phylogeny. Maximum likelihood phylogeny based on 3204 concatenated Plasmodium simium protein-coding genes with 1:1 orthologs across a selection of Plasmodium species (see Methods). Tree was constructed using RaxML with the GTRGAMMA model and Plasmodium gallinaceum as outgroup. Branch support from 100 bootstrap replicates. Figure S4. Gene families. A) The number of gene family members among Plasmodium genomes. Diameters of circles are proportional to the log10-transformed number of family members, as shown on the right. B) Bar plot showing the distribution of average read coverage across all annotated P. simium genes. The 5th and 95th percentiles (genes with the highest and lowest read coverage, respectively) are highlighted in dark. Above the plots the proportion of genes belonging to the PIR gene family is shown as pie charts for the two extreme percentiles and the remaining genes ('Rest'). Using a Fisher's exact two-sided test, the proportion of PIR genes in both the 95th and the 5th percentile are significantly higher than in the 'Rest' genes (p = 5.8e-80 and p = 5.1e-67, respectively). Figure S5. Pir/Vir gene clustering. Clustering of PIR protein sequences based on BLASTP similarity. Network threshold is a bit-score of 40 and visualized using the edge-weighted spring embedded layout in cytoscape 3. Figure S6. SNP counts. Top: Number of high-confidence SNPs detected in each sample. Plasmodium simium samples shown in dark red, and Plasmodium vivax samples in orange (see also Additional file 1: Tables S3 & S4). Bottom: For a given SNP loci data is not available from all samples. The bar chart shows how many SNP loci (left y-axis) that have data (coverage) in a given number of samples (x-axis). Only SNPs with data from at least 55 samples were used in this study, as indicated by the blue bars. Purple bars represent discarded SNPs. The cumulative fraction of SNPs is shown as red line (right x-axis). For example, slightly more than half of the SNP loci have data from at least 55 samples, whereas app. 75% of samples have data from at least 25 samples. Figure S7. PCA plot. A) Plot of the first two dimensions from a principal component analysis of 124,968 SNPs. Plasmodium vivax samples are denoted by their geographic origin. B) Magnification of American vivax samples. C) Cumulative contribution (in percent) of each eigenvector. The separate clustering of Plasmodium simium and American P. vivax samples was also observed when plotting 2nd versus 3rd dimensions, and 3rd versus 4th dimensions (not shown). Figure S8. MDS plot. Plot of Multidimensional Scaling of Plasmodium simium and Plasmodium vivax samples. Figure S9. SNP phylogeny. Phylogenetic tree constructed from SNP sites. Identical to Fig. 1, but with sample IDs shown at terminal branches. Figure S10. Phylogenetic network. Network from nucleotide diversity distances produced in SplitsTree using the NeighborNet network. A magnification of the central hubs (red rectangle) is shown at the bottom right. Figure S11. Admixture. Q-estimates from unsupervised ADMIXTURE clustering analysis at K from 2 to 10. Plasmodium vivax samples are ordered by geographic origin. The lowest cross-validation error was observed for K = 3. Figure S12. Gene DXY diversity. A) For different intervals of average DXY values (x-axis), the number of genes with thee values are plotted (y-axis, log10-scaled). B) Box plot showing the distributions of DXY values for members of selected gene families. The right-most, yellow box shows the DXY values for all genes that not member of a gene family. Figure S13. DBP phylogeny. Neighbor-Joining tree of Plasmodium DBP protein sequences. Sequences were aligned using mafft and tree produced with CLUSTALW. Support from 1000 bootstrap replicates. Tree visualized with FigTree, genetic distance shown below tree. Plasmodium simium and Plasmodium vivax sequences derived from this study are highlighted in red and blue, respectively. Remaining sequences are suffixed by their genome of origin (PvivP; P. vivax P01, PvivS; P. vivax SalI, PcynM; P. cynomolgi M, PcynB; P. cynomolgi B, PknoH; P. knowlesi H). Figure S14. RBP phylogeny. Neighbor-Joining tree of Plasmodium RBP protein sequences. Sequences were aligned using mafft and tree produced with CLUSTALW. Support from 1000 bootstrap replicates. Tree visualized with FigTree, genetic distance shown below tree. Plasmodium simium and Plasmodium vivax sequences derived from this study are highlighted in red and blue, respectively. Remaining sequences are suffixed by their genome of origin (PvivP; P. vivax P01, PvivS; P. vivax SalI, PcynM; P. cynomolgi M, PcynB; P. cynomolgi B, PknoH; P. knowlesi H). Figure S15. Coverage across RBP gene loci. Average read coverage across gene loci when mapping human Plasmodium simium (top) and Plasmodium vivax (bottom) reads onto the P. vivax P01 genome. The RBPs genes are shown above plots with the P. vivax P01 gene identifiers below plots. Genes absent in P. simium are highlighted in grey. Note that the P. vivax P01 genome contains two annotated RBP1a and RBP2d genes, respectively. The average read coverage per gene is shown as dots for each individual sample, and the combined distributions are outlined by boxes. A single P. simium sample, AF22, has a typical coverage of around 200X but is omitted from this representation for clarity. The coverage of the P. simium CDC strain is highlighted as red dots. Figure S16. coverage across flanking regions. The average read coverage across Plasmodium vivax DBP and RBP genes is compared to the average read coverage across flanking genomic regions. The log2 ratio between coverage at flanking region and coverage at gene was calculated for four regions: 10 kb-5 kb upstream of gene (UP2), 5 kb-0 kb upstream of gene (UP1), 0 kb-5 kb downstream of gene (DOWN1), 5 kb-10 kb downstream of gene (DOWN2). Boxplot denotes the range of ratios for Plasmodium simium samples (red boxes), American Plasmodium vivax samples (light blue), and remaining Plasmodium vivax samples (blue). Up- and downstream are defined based on genome coordinates irrespective of gene orientation. After Bonferroni correction, only the 'UP1' region at RBP3 for P. simium samples (p = 0.011, denoted by asterisk) had a probability below 0.05 of the mean being above zero assuming a normal distribution. Figure S18. Haplotype network of DBP1 sequences. Haplotype network (minimum spanning network) produced using PopART. Numbers of mutations are indicated by hatch marks on edges. Plasmodium simium samples are shown in red, Plasmodium vivax in green, and P. vivax-like in purple. The previously published P. simium CDC strain sequence (ACB42432) is shown in black and bold. Figure S19. Read support for DBP1 deletion patterns. Among Plasmodium vivax and Plasmodium simium samples multiple deletion patterns in the DBP1 gene are observed (top). Deletions found in P. vivax samples are arbitrarily denoted vivax 1-4. For each sample, the number of reads supporting a given deletion is shown (bottom). Samples with reads supporting multiple deletion forms are indicated by asterisks. Figure S20. Dotplot. Similarity DNA dot plots of DBP1 genes. Top plots show Plasmodium simium and bottom plots show Plasmodium vivax DBP1 genes. Introns are denoted by grey rectangles and the deleted region (or site of deleted region in P. simium) is highlighted in red. Figure S21. Read coverage across the DBP1 deletion in Plasmodium simium samples. Artemis representation of read coverage across the Plasmodium simium DBP1 deletion. Reads from Plasmodium vivax samples AM01 & AM02 (two top panels) and P. simium AF22 & AF36 samples (two bottom panels) were mapped onto the P. simium AF22 assembly. The site of deletion is indicated by vertical red arrows. Note the lack of P. vivax reads spanning the deletion site. Figure S22. read coverage across the DBP1 deletion in Plasmodium vivax samples. Artemis representation of read coverage across the Plasmodium simium DBP1 deletion. Reads from P. simium AF22 & AF36 samples were mapped onto the Plasmodium vivax P01 genome. The site of deletion is indicated by red squares. Note the lack of P. simium coverage across the deleted region. Figure S23. PacBio read coverage across the DBP1 deletion in Plasmodium simium. Schematic depiction of PacBio reads mapping across the deletion in DBP1 gene. X-axis denotes genomic positions. Black line at the centre of plot shows the extent of exons. Reads mapping on the forward strand are shown above exons, reverse strand reads underneath exons. Reads are colored according to the sample from which they are derived. The site of deletion is indicated by arrows and thin vertical orange lines. Figure S24. DBP1 PCR. Top: Schematic overview of DBP1 deletion PCR approach. Gel images shown for human Plasmodium vivax samples (top gel image), human Plasmodium simium samples (middle image), and non-human primate (NHP) P. simium samples (bottom image). Expected band sizes with and without deletion event are indicated by red triangles. Bottom: Primer sequences. Figure S26. Read coverage across the RBP2a deletion in Plasmodium simium samples. Artemis representation of read coverage across the Plasmodium simium RBP2a deletion. Reads from vivax samples AM01 & AM02 (two top panels) and P. simium AF22 & AF36 samples (two bottom panels) were mapped onto the P. simium AF22 assembly. The site of deletion is indicated by vertical red arrows. Note the lack of Plasmodium vivax reads spanning the deletion site. Figure S27. Read coverage across the RBP2a deletion in Plasmodium vivax samples. Artemis representation of read coverage across the Plasmodium simium RBP2a deletion. Reads from P. simium AF22 & AF36 samples were mapped onto the P. vivax P01 genome. The site of deletion is indicated by red squares. Note the lack of P. simium coverage across the deleted region. Figure S28. PacBio read coverage across RBP2a deletion in Plasmodium simium. Schematic depiction of PacBio reads mapping across deletions in the RBP2a gene. X-axis denotes genomic positions. Black line at the centre of plots shows the extent of exons. Reads mapping on the forward strand are shown above exons, reverse strand reads underneath exons. Reads are colored according to the sample from which they are derived. Sites of deletions are indicated by arrows and thin vertical orange lines. Figure S29. RBP2a PCR. Top: Schematic overview of RBP2a deletion PCR approach. Gel images shown for Plasmodium vivax and Plasmodium simium samples. Expected band sizes with and without deletion event are indicated by red triangles. Bottom: Primer sequences. Figure S30. Short indels. A) Pie chart showing the percentage of indels being integers of 1-3 base pairs. B) The percentage of insertions (left bar) and deletions (middle) overlapping low-complexity regions in proteins. The percentage of all proteins consisting of low-complexity sequences is shown on the right. Low-complexity annotation downloaded from PlasmoDB. C) Size distributions of genes with and without indels. Genes with indels are listed in Additional file 1: Table S7. Figure S31. DBP1 protein structures. Plasmodium simium DBP1 is predicted to associate with human DARC. Left: Top-view of modelled DBP1 from Plasmodium vivax strain P01, human-infecting P. simium AF22 (PsDBP1) and monkey-infecting P. simium (CDC PsDBP1) sequences. Models were established based on the crystal structure of the P. vivax DBP1, strain Salvador 1, in complex with human DARC (PDB 4nuv). Individual chains of the DBP1 dimer are shown in light and dark blue. DARC (residues 19-30) is coloured in grey. Residue substitutions between the models and the crystal structure are highlighted in magenta. Right: close-up view of the DARC-binding site, as delimited by a dashed square on the left. Colours as in the left panel. The rearrangement of hydrogen-bonds (black dotted lines) is shown for the Lys-Asn substitution. Figure S32. Mitochondrial haplotype network. Haplotype network (minimum spanning network) produced using PopART (see Methods). Numbers of mutations are shown on edges. The two identical Plasmodium simium samples (AF22 and GenBank accession AY722798) are shown in black. Plasmodium vivax haplogroups coloured according to country of origin. Circle sizes indicate the number of sequences in haplogroups. Figure S33. Protein orthology. Venn diagram showing the number of shared gene orthogroups between Plasmodium simium and three other Plasmodium genomes. Figure S34. SNP allele frequencies & coverage. Top: Allele frequencies for SNPs with available data from at least 37 Plasmodium simium or Plasmodium vivax samples. Bottom: Mean read coverage at SNP sites shown for each samples.

MicroRNAs (miRNAs) are small non-coding RNAs that can play critical roles in regulating various cellular processes including during many parasitic infections. Here, we report a regulatory role for miR-34c-3p in cAMP-independent regulation of PKA activity in Theileria annulata and Plasmodium falciparum infections of bovine leukocytes and human erythrocytes, respectively. We identified prkar2b (cAMP-dependent protein kinase A type II-beta regulatory subunit), as a novel miR-34c-3p target gene and demonstrated how infection-induced up-regulation of miR-34c-3p in leukocytes repressed PRKAR2B expression to increase PKA activity and promote the virulent disseminating tumour phenotype of T. annulata -transformed macrophages. When human erythrocytes are infected by P. falciparum they accumulate miR-34c-3p that ablates both prkar2b and parasite Pfpkar mRNA. However, erythrocytes lack protein translation machinery so only miR-34c-3p-mediated loss of Pfpkar transcripts results in an increase in PfPKA kinase activity. Inhibition of miR-34c-3p increases Pfpkar expression to reduce PfPKA activity leading to slowing of intra-erythrocyte parasite development and a reduction in invasion of fresh red blood cells. Finally, we demonstrate that miR-34c-3p regulation of prkar2b expression is generalizable, by showing that it can negatively regulate prkar2b expression and PRKAR2B protein levels in human cancer cell lines and that brown adipose tissue displays high levels of miR-34c-3p and corresponding low levels of prkar2b mRNA compared to white adipose tissue. Induction of miR-34c-3p therefore, represents a novel cAMP-independent way of regulating PKA activity in a range of cell types associated with cancer, diabetes and parasitic diseases.

DNA sequencing technologies provide unprecedented opportunities to analyze within-host evolution of microorganism populations. Often, within-host populations are analyzed via pooled sequencing of the population, which contains multiple individuals or ‘haplotypes’. However, current next-generation sequencing instruments, in conjunction with single-molecule barcoded linked-reads, cannot distinguish long haplotypes directly. Computational reconstruction of haplotypes from pooled sequencing has been attempted in virology, bacterial genomics, metagenomics and human genetics, using algorithms based on either cross-host genetic sharing or within-host genomic reads. Here we describe PoolHapX , a flexible computational approach that integrates information from both genetic sharing and genomic sequencing. We demonstrated that PoolHapX outperforms state-of-the-art tools in the above four fields, and is robust to within-host evolution. Importantly, together with barcoded linked-reads, PoolHapX can infer whole-chromosome-scale haplotypes from 50 pools each containing 12 different haplotypes. By analyzing real data, we uncovered dynamic variations in the evolutionary processes of HIV previously unobserved in single position-based analysis.

Summary Cell cycle transitions are generally triggered by variation in the activity of cyclin-dependent kinases (CDKs) bound to cyclins. Malaria-causing parasites have a life cycle with unique cell-division cycles, and a repertoire of divergent CDKs and cyclins of poorly understood function and interdependency. We show that Plasmodium berghei CDK-related kinase 5 (CRK5), is a critical regulator of atypical mitosis in the gametogony and is required for mosquito transmission. It phosphorylates canonical CDK motifs of components in the pre-replicative complex and is essential for DNA replication. We also provide evidence for indirect regulation of the concomitant M-phase progression. During a replicative cycle, CRK5 stably interacts with a single Plasmodium -specific cyclin (SOC2), although we obtained no evidence of SOC2 cycling by transcription, translation or degradation. Our results provide evidence that during Plasmodium male gametogony, this unique cyclin/CDK pair fills the functional space of multiple eukaryotic cell-cycle kinases controlling DNA replication and M-phase progression.

Miniopterus nimbae sp. nov. Nimba long-fingered bat Holotype DM 12621 (field no. AM2010_12_18_1), an adult male, was collected by Ara Monadjem. The specimen was fixed in formalin and then transferred to 70% alcohol. The skull has been extracted and cleaned. Photograph of the skull and drawing of the tragus of the holotype are illustrated in Figs. 6 and 7. Type locality Liberia, Nimba Province, Mount Gangra, 10 km to the west of Mount Nimba (Fig. 1). The bat was netted on 18 December 2010 exiting from a mine adit mid-way up Mount Gangra (7.55434°N, 8.62902°W) at 720 m a.s.l, in secondary forest. Paratypes No other specimens of this species were captured or collected on the same day at the same site. However, the previous night an adult female (DM 12614) was collected at the Yiti River 9 km to the south-east of Mount Gangra. Photograph of the paratype is illustrated in Fig. 8. Etymology This species is named after Mount Nimba, one of just three localities from which it is known, further highlighting the critical importance of this mountain for bat conservation in Africa (Monadjem et al., 2016). Diagnosis This is a large-sized Miniopterus from Mount Nimba, Liberia, with a mean forearm length of 47.4 mm (n = 26 individuals). The large size of this bat (particularly its forearm length) readily distinguishes M. nimbae from all other African Miniopterus taxa except the M. inflatus / M. africanus group. In external and craniodental measurements, M. nimbae is similar in size to other members of the M. inflatus group (M. inflatus s.s., M. cf. inflatus and M. africanus) (Tables 3–5); however, in multi-dimensional morphospace based on craniodental measurements, it overlaps only with M. inflatus s.s. The taxon M. cf. inflatus from eastern and southern Africa tends to have a light pelage, being more reddish-brown in colour (compared with a deep chocolate brown in M. nimbae). It is not possible, at present to distinguish M. nimbae from M. inflatus s.s. on external characters. However, they can be readily distinguished by cranial features. In particular, the 1st upper premolar has an additional lingual cusp, posterior to the main cusp, that is present in M. nimbae but absent in M. inflatus s.s. (Fig. 9); this feature being clearly visible, even with a low magnification hand lens, and consistently present in all the specimens examined in this study. The lower tooth row (i 1 –m 3) and lower molar (LWMOLS) lengths are also slightly larger in M. nimbae than M. inflatus s.s. In terms of cranial geometry, M. nimbae differs from M. inflatus s.s. as it bears a slightly more gracile skull, less ‘inflated’ braincase, and the point of maximum curvature along the occiput is more elevated in the newly described taxon than in the nominate form. Furthermore, these two taxa, are also distinguishable on molecular grounds (K2P pairwise genetic distance = 1.6%). Additionally, the ranges of the two taxa do not appear to overlap. Miniopterus africanus appears to be restricted to north-eastern Africa and is genetically distinct (K2P pairwise distance = 16.7%) from M. nimbae. Description External characters.— Miniopterus nimbae is large-sized for the genus, but showing typical generic features including a rounded head, an elongated second phalanx of the third digit, rounded ears, and a relatively long and straight tragus. The tail is slightly less than half that of the total length. The pelage is dark chocolate brown above and slightly paler below. Individual hairs are unicoloured. The mass and standard external measurements of the holotype compared with a small sample of other individuals are shown in Table 3. Craniodental characters.—The skull is robust for a Miniopterus species. The rostrum is broad and the braincase is rounded and high, typical for the genus of Miniopterus. The dentition of M. nimbae is I 2/3, C 1/1, P 2/3, M 3/3, which is typical of the genus. In the upper tooth row, the inner incisor is larger than the outer one, and the anterior premolar is relatively well developed. The cranial and dental measurements of the holotype compared with a small sample of other individuals are shown in Tables 4 and 5. Distribution Analysis of genetic samples taken from five specimens with large forearm lengths (range 46.9– 48.2 mm) at Mount Gangra (Nimba, Liberia), group them all as M. nimbae (Figs. 2 and 3). Although not genetically analysed, it is likely that specimens previously collected from northern Liberia or southeastern Guinea and identified as M. inflatus are instead referable to this new species, M. nimbae. Based on this assumption, this taxon is known from just three localities: Mount Nimba (and surrounding uplands), Liberia; Wonegizi Mountains, Liberia; and Mount Béro, Guinea (Wolton et al., 1982; Koopman et al., 1995; Fahr et al., 2006; Monadjem et al., 2016). The closest locality for M. inflatus s.s. is at least 2,000 km away in eastern Nigeria (Happold, 2013 b), and no other large Miniopterus specimens have been collected in the intermediate area, despite extensive surveys in a number of localities e.g., Thaï and Comoé National Parks in Côte d’Ivoire (Fahr and Kalko, 2011), the Simandou Range in eastern Guinea (Decher et al., 2015), the Fouta Djallon mountains in central Guinea (Weber and Fahr, 2007) or anywhere in Ghana or Sierra Leone (Grubb et al., 1998; Happold, 2013 b, 2013 c). Therefore, M. nimbae is probably an endemic to the upland areas of northern Liberia and south-eastern Guinea, and may be shown to also occur in the upland area of western Côte d’Ivoire at and around Mount Nimba. Biology: Practically nothing is known about the biology of this Upper Guinea forest endemic. It has been recorded roosting in mine adits at 720– 970 m above sea level (a.s.l.) at Mount Gangra and Mount Yuelliton (both within 10 km of Mount Nimba). The size of one roosting colony at Mount Gangra was estimated at between 20–30 individuals; the site was shared with large numbers of Myonycteris angolensis and Hipposideros cf. ruber, and a few H. marisae. A second roosting colony was estimated to include> 200 individuals of this species at Mount Yuelliton (which was not inhabited by any other bat species — A. Monadjem, personal observation). It has been netted at various localities in the foothills of Mount Nimba and in the low-lying rainforest region between these three upland areas. This suggests that this species roosts in upland areas (700 m a.s.l.), and then descends to forage in lower lying forested areas (about 500 m a.s.l.). In a sample of 13 females captured at Nimba between late December and end-March, five were pregnant. During the same period, three out of 10 males had scrotal testes. The mean frequency of the knee of handreleased M. nimbae captured at Mount Nimba was 48.4 kHz (range: 47.2–48.9 kHz, n = 4). Other Taxonomic Considerations In addition to the description of the new species, M. nimbae, the phylogeny presented here (Figs. 2 and 3) also identifies two other distinct taxa. The first is the taxon M. villiersi, which does not appear to be closely related to M. schreibersii s.s. and should therefore not be considered a subspecies of the latter mentioned taxon. In fact, M. villiersi is sister to M. nimbae (Figs. 2 and 3), from which it is readily distinguishable based on genetics and size (see Tables 2–5); the pairwise genetic distance between the two species is 9.0%, and there is no overlap between these two species in forearm length or any of the craniodental measurements presented here. The echolocation calls also differ, with the frequency of the knee of M. villiersi calls (based on hand-released individuals that were captured at Mount Nimba — x = 51.6 kHz, range 51.1–52.4 kHz, n = 5) being distinctly higher than that of M. nimbae (x = 48.4 kHz, range 47.2–48.9 kHz, n = 4) with no overlap between the two species. The second taxon refers to the M. inflatus group, which appears to be paraphyletic. Miniopterus inflatus s.s. (based on sequenced specimens from Gabon, and close to the type locality in southern Cameroon) is sister to the taxon M. nimbae and these two are sister to M. villiersi (Figs. 2 and 3). By contrast, the M. cf. inflatus specimens (from Malawi and Mozambique) are sister to M. fraterculus and M. minor. This suggests that the taxon M. inflatus s.l. comprises, in addition to the newly recognised M. nimbae, two distinct and not closely related taxa which we refer to as M. inflatus s.s. (from Gabon), and M. cf. inflatus (from Malawi and Mozambique)., Published as part of Monadjem, Ara, Shapiro, Julie T., Richards, Leigh R., Karabulut, Hatice, Crawley, Wing, Nielsen, Ida Broman, Hansen, Anders, Bohmann, Kristine & Mourier, Tobias, 2019, Systematics of West African Miniopterus with the description of a new species, pp. 237-256 in Acta Chiropterologica 21 (2) on pages 245-248, DOI: 10.3161/15081109ACC2019.21.2.001, http://zenodo.org/record/3944920, {"references":["MONADJEM, A., L. RICHARDS, and C. DENYS. 2016. An African bat hotspot: the exceptional importance of Mount Nimba for bat diversity. Acta Chiropterologica, 18: 359 - 375.","WOLTON, R. J., P. A. ARAK, H. C. J. GODFRAY, and R. P. WILSON. 1982. Ecological and behavioural studies of the Megachiroptera at Mount Nimba, Liberia, with notes on Microchiroptera. Mammalia, 18: 359 - 375.","KOOPMAN, K. F., C. P. KOFRON, and A. CHAPMAN. 1995. The bats of Liberia: systematics, ecology, and distribution. American Museum Novitates, 3148: 1 - 24.","FAHR, J., B. A. DJOSSA, and H. VIERHAUS. 2006. Rapid assessment of bats (Chiroptera) in Dere, Diecke and Mt. Bero classified forests, southeastern Guinea; including a review of the distribution of bats in Guinee Forestiere. Pp. 168 - 180, 245 - 247, in A rapid biological assessment of three classified forests in Southeastern Guinea (H. E. WRIGHT, J. MCCULLOUGH, L. E. ALONSO, and M. S. DIALLO, eds.). RAP Bulletin of Biological Assessment, 40: 1 - 248.","HAPPOLD, M. 2013 b. Miniopterus inflatus Greater long-fingered bat. Pp. 714 - 716, in The mammals of Africa. Volume IV: Hedgehogs, shrews and bats (M. HAPPOLD and D. C. HAP- POLD, eds.). Bloomsbury Publishing, London, 800 pp.","FAHR, J., and E. K. V. KALKO. 2011. Biome transitions as centres of diversity: habitat heterogeneity and diversity patterns of West African bat assemblages across spatial scales. Ecography, 34: 177 - 195.","DECHER, J., A. HOFFMANN, J. SCHAER, R. W. NORRIS, B. KADJO, J. ASTRIN, A. MONADJEM, and R. HUTTERER. 2015. Bat diversity in the Simandou Mountain Range of Guinea, with the description of a new white-winged vespertilionid. Acta Chiropterologica, 17: 255 - 282.","WEBER, N., and J. FAHR. 2007. Survey of endemic and globally threatened bat species in the Fouta Djallon Plateau for","GRUBB, P., T. S. JONES, A. G. DAVIES, E. EDBERG, E. D. STARIN, and J. E. HILL. 1998. Mammals of Ghana, Sierra Leone and The Gambia. Trendine Press, Cornwall, UK, 265 pp.","HAPPOLD, M. 2013 c. Miniopterus schreibersii Schreibers's long-fingered bat. Pp. 721 - 722, in The mammals of Africa. Volume IV: Hedgehogs, shrews and bats (M. HAPPOLD and D. C. D. HAPPOLD, eds.). Bloomsbury Publishing, London, 800 pp."]}

Kinesin-8 proteins are microtubule motors that are often involved in regulation of mitotic spindle length and chromosome alignment. They move towards the plus ends of spindle microtubules and regulate the dynamics of these ends due, at least in some species, to their microtubule depolymerization activity. Plasmodium spp. exhibit an atypical endomitotic cell division in which chromosome condensation and spindle dynamics in the different proliferative stages are not well understood. Genome-wide shared orthology analysis of Plasmodium spp. revealed the presence of two kinesin-8 motor proteins, kinesin-8X and kinesin-8B. Here we studied the biochemical properties of kinesin-8X and its role in parasite proliferation. In vitro, kinesin-8X has motility and depolymerization activities like other kinesin-8 motors. To understand the role of Plasmodium kinesin-8X in cell division, we used fluorescence-tagging and live cell imaging to define its location, and gene targeting to analyse its function, during all proliferative stages of the rodent malaria parasite P. berghei life cycle. The results revealed a spatio-temporal involvement of kinesin-8X in spindle dynamics and an association with both mitotic and meiotic spindles and the putative microtubule organising centre (MTOC). Deletion of the kinesin-8X gene revealed a defect in oocyst development, confirmed by ultrastructural studies, suggesting that this protein is required for oocyst development and sporogony. Transcriptome analysis of ?kinesin-8X gametocytes revealed modulated expression of genes involved mainly in microtubule-based processes, chromosome organisation and the regulation of gene expression, supporting a role for kinesin-8X in cell division. Kinesin-8X is thus required for parasite proliferation within the mosquito and for transmission to the vertebrate host.

Oral Sessions

The list contains the 65 genes coding for extracellular matrix components or enzymes involved in matrix metabolism that were found upregulated in BCC. (DOCX 18Â kb)

The number of reads in the mRNA sequencing analysis and the following reads that were actually mapped. Whereas approximately 97% of all reads mapped to the human genome, duplicate reads constituted a significant fraction leaving only around 48 million unique reads. (DOCX 33Â kb)