Babesia Duncani Are the Main Causative Agents of Human Babesiosis

22 ABSTRACT 23 Babesia microti and Babesia duncani are the main causative agents of human babesiosis 24 in the United States. While significant knowledge about B. microti has been gained over the past 25 few years, nothing is known about B. duncani biology, pathogenesis, mode of transmission or 26 sensitivity to currently recommended therapies. Studies in immunocompetent wild type mice and 27 hamsters have shown that unlike B. microti, infection with B. duncani results in severe pathology 28 and ultimately death. The parasite factors involved in B. duncani virulence remain unknown. 29 Here we report the first known completed sequence and annotation of the apicoplast and 30 mitochondrial genomes of B. duncani. We found that the apicoplast genome of this parasite 31 consists of a 34 kb monocistronic circular molecule encoding functions that are important for 32 apicoplast gene transcription as well as translation and maturation of the organelle’s proteins. 33 The mitochondrial genome of B. duncani consists of a 5.9 kb monocistronic linear molecule with 34 two inverted repeats of 48 bp at both ends. Using the conserved cytochrome b (Lemieux) and 35 cytochrome c oxidase subunit I (coxI) proteins encoded by the mitochondrial genome, 36 phylogenetic analysis revealed that B. duncani defines a new lineage among apicomplexan 37 parasites distinct from B. microti, Babesia bovis, Theileria spp. and Plasmodium spp. Annotation 38 of the apicoplast and mitochondrial genomes of B. duncani identified targets for development of 39 effective therapies. Our studies set the stage for evaluation of the efficacy of these drugs alone or 40 in combination against B. duncani in culture as well as in animal models. 41 Keywords: Babesiosis, Genome, Mitochondria, Apicoplast, Sequencing, Annotation, Drug 42 targets 43 44 2 45 1. Introduction 46 Human babesiosis is a global infectious disease transmitted by ticks, blood transfusion or 47 congenitally (reviewed in Vannier and Krause (2012); Vannier et al. (2015)). The disease is 48 caused by intraerythocytic parasites of the genus Babesia. Immunocompromised and asplenic 49 individuals are at greatest risk of developing babesiosis symptoms, which include fever and 50 headache and can advance to multi-system organ failure and death (Vannier et al., 2015). Most 51 cases of babesiosis occur in the northeastern and midwestern regions of the United States and are 52 caused by Babesia microti and transmitted by Ixodes scapularis, the same tick species that 53 transmits the agents of Lyme disease, anaplasmosis and Powassan virus disease (Burgdorfer et 54 al., 1982; Spielman et al., 1985). Beside these cases, several clinical reports from Washington 55 State, USA and California, USA were linked to another Babesia sp, Babesia duncani (Kjemtrup 56 and Conrad, 2000). The first case of human babesiosis reported to be caused by a new Babesia 57 sp., named WA1, was of a 41-year-old man from Washington State (Quick et al., 1993). Since 58 then 11 more cases of human babesiosis attributable to WA1 and WA1-like organisms have been 59 reported in California and Washington State, and the etiological agent later identified as B. 60 duncani (reviewed in Kjemtrup and Conrad (2000)). Two other cases preceding these 12 B. 61 duncani cases were reported in California and were presumed to be caused by this pathogen 62 (Scholtens et al., 1968; Bredt et al., 1981). Studies in hamsters and mice following infection with 63 the parasite showed that B. duncani pathogenesis is different from that caused by B. microti. 64 Infection with B. microti is characterized by an initial phase of high parasitemia, anemia and 65 splenomegaly followed by rapid decline in parasitemia to undetectable levels, leaving animals 66 surviving infection with little to no detectable symptoms (Cullen and Levine, 1987; Wozniak et 67 al., 1996). In contrast, B. duncani infection of mice and hamsters results in both a rapid increase 3 68 in parasitemia and severe pathology with mortality rates of more than 95% in C3H, A/J, AKR/N 69 and DBA/1J mice, between 40 to 50% in BALB/cJ, CBAJ and 129/J mice and less than10% in 70 C57BL/6 and C57BL/10 mice (Dao and Eberhard, 1996; Moro et al., 1998). 71 While significant knowledge has been gained over the past several years about B. microti, 72 nothing is known about the biology, genome composition and structure, or pathogenesis of B. 73 duncani. Furthermore, recommended therapies have not been evaluated directly against this 74 parasite in vitro or in animal models. Recent studies by Swei and colleagues (in press) suggest 75 that the enzootic tick vector of B. duncani is Dermacentor albipictus and the reservoir host is 76 likely the mule deer. 77 The data described in this study represent, to our knowledge, the first report of the 78 completed sequence, assembly, and annotation of the apicoplast and mitochondrial genomes of 79 B. duncani. Phylogenetic analysis using mitochondrial genes shows that B. duncani defines a 80 new lineage in the Apicomplexa phylum distinct from B. microti. Our analysis further predicts 81 potential therapeutic targets and new strategies to develop effective strategies to treat babesiosis 82 resulting from infection by B. duncani. 83 84 2. Materials and methods 85 2.1. Animal ethics statement 86 All animal experimental protocols followed Yale University institutional guidelines for care and 87 use of laboratory animals and were approved by the Institutional Animal Care and Use 88 Committees (IACUC) at Yale University. Rules for ending experiments were to be enacted if 89 animals showed any signs of distress or appeared moribund. 90 4 91 2.2. Sequencing assembly 92 The B. duncani WA1 isolate was obtained from BEI Resources (www.beiresources.org; 93 Number: NR-12311). This parasite was isolated from the blood of the first reported case of 94 babesiosis acquired in Washington State (Quick et al., 1993). The strain was injected into 95 hamsters and infected red blood cells (RBCs) were purified and used to isolate B. duncani total 96 DNA. This DNA was sequenced using both Illumina Hi-Seq 2500 paired-end 75 bp short-read 97 sequencing and PacBio single molecule long-reads (>10kb) sequencing. Assembly was 98 performed using the following steps: i) raw sequence reads were mapped using BWA-MEM (Li 99 and Durbin, 2009) and BLASR (Chaisson and Tesler, 2012) against the host genome sequence 100 (Golden Hamster MesAur1.0 genome, 101 https://www.ncbi.nlm.nih.gov/assembly/GCF_000349665.1/) to remove host DNA 102 contamination; ii) potential sequencing errors in PacBio reads were corrected using ectools 103 (preprint at http://www.biorxiv.org/content/early/2014/06/18/006395) and then the corrected 104 long reads were assembled using Celera Assembler v7 (Berlin et al., 2015); iii) samtools (Li et 105 al., 2009) was used to extract only unmapped reads, and bedtools (Quinlan and Hall, 2010) 106 bam2fastq to convert those back to fastq; iv) spades (Bankevich et al., 2012), with default 107 parameters, was used to assemble the remaining reads into scaffolds. Two of the assembled 108 contigs encompassed the entire apicoplast and mitochondrial genomes. The sequence of the 109 mitochondrial genome was further validated using long-range PCRs and Sanger sequencing. 110 111 2.3. Annotation of the apicoplast and mitochondrial genomes 112 Annotation of the apicoplast and mitochondrial genomes was performed using Artemis 113 (http://www.sanger.ac.uk/science/tools/artemis) (Rutherford et al., 2000) to identify all open 5 114 reading frames, and BLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi) (Altschul et al., 1990) to 115 identify homologous proteins in other organisms in the GenBank database. The prediction of 116 tRNAs in both genomes was accomplished using tRNAscanSE version 1.21 and version 2.0 117 (http://lowelab.ucsc.edu/tRNAscan-SE/) with the search mode set to “default” and the source set 118 to “Mito/Chloroplast” (Lowe and Eddy, 1997; Schattner et al., 2005; Lowe and Chan, 2016). 119 Only tRNAs with a score above 30% were annotated on the genome. rRNA genes, large subunit 120 (LSU) and small subunit (SSU), were determined by searching for their counterparts in Babesia 121 bovis, Babesia orientalis, B. microti and Theileria parva. The circular genetic map for the 122 apicoplast genome was designed using CGView (http://stothard.afns.ualberta.ca/cgview_server/) 123 (Stothard and Wishart, 2005). 124 125 2.4. Phylogenetic analysis 126 A phylogenetic relationship was established using concatenated sequences of the cob and 127 cox1 mitochondrial genes. The sequences were first added consecutively in MEGA7 (Molecular 128 Evolutionary Genetics Analysis version 7.0) (Kumar et al., 2016) before they were aligned using 129 MUSCLE (Edgar, 2004). A Neighbor-Joining tree (Saitou and Nei, 1987) was then constructed 130 using bootstrap analysis inferred from 1000 replicates (Felsenstein, 1985). The bootstrap 131 consensus tree inferred from 1000 replicates was taken to represent the evolutionary history of 132 the taxa analyzed (Felsenstein, 1985). The evolutionary distances were computed using the JTT 133 matrix-based method and are in the units of the number of amino acid substitutions per site 134 (Jones et al., 1992). Evolutionary analyses were conducted in MEGA7 (Kumar et al., 2016). 135 136 3. Results and Discussion 137 3.1. Identification of the B. duncani apicoplast and mitochondrial genomes 6 138 In order to obtain the genome sequence of the apicoplast and mitochondrial genomes of 139 B. duncani, the WA1 clinical isolate was first propagated in hamsters and DNA isolated from 140 infected RBCs. Search for coding sequences known to be encoded by the apicoplast and 141 mitochondrial genomes of other apicomplexan parasites identified two contigs of ~34 kb 142 (GenBank Accession no. MH107388) and ~6 kb (GenBank Accession no. MH107387) in 143 length, respectively.

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