Filarial and Wolbachia genomics

Review Article

Filarial and Wolbachia genomics

A. L. SCOTT,1 E. GHEDIN,2 T. B. NUTMAN,3 L. A. MCREYNOLDS,4 C. B. POOLE,4 B. E. SLATKO4 & J. M. FOSTER4

1Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore,MA, USA, 2Department of Computational & Systems Biology, Center for Vaccine Research, University of Pittsburgh School of Medicine,Pittsburgh, PA, USA, 3Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, NIH, Bethesda, MD, USA,4New England Biolabs, Ipswich, MA, USA

SUMMARY

Filarial nematode parasites, the causative agents for a spec-trum of acute and chronic diseases including lymphatic fila-riasis and river blindness, threaten the well-being andlivelihood of hundreds of millions of people in the developingregions of the world. The 2007 publication on a draft assem-bly of the 95-Mb genome of the human filarial parasiteBrugia malayi – representing the first helminth parasitegenome to be sequenced – has been followed in rapid succes-sion by projects that have resulted in the genome sequencingof six additional filarial species, seven nonfilarial nematodeparasites of animals and nearly 30 plant parasitic and free-living species. Parallel to the genomic sequencing, tran-scriptomic and proteomic projects have facilitated genomeannotation, expanded our understanding of stage-associatedgene expression and provided a first look at the role of epi-genetic regulation of filarial genomes through microRNAs.The expansion in filarial genomics will also provide a signif-icant enrichment in our knowledge of the diversity and vari-ability in the genomes of the endosymbiotic bacteriumWolbachia leading to a better understanding of the geneticprinciples that govern filarial–Wolbachia mutualism. Thegoal here is to provide an overview of the trends andadvances in filarial and Wolbachia genomics.

Keywords Brugia, filarial, genome, Nematode, noncodingRNA, proteomics, transcriptomics, Wolbachia

INTRODUCTION

Filarial nematode parasites are vector-borne pathogens ofsubstantial medical and veterinary importance. While it isdifficult to estimate the current prevalence of filarial infec-tions in humans owing to ongoing eradication efforts (1),in 2006 it was estimated that well over 1 billon were atrisk and approximately 150 million people were infectedwith the major species of filarial nematodes: Wuchereriabancrofti, Brugia malayi, Onchocerca volvulus and Loa loa(2). In humans, filarial parasites typically result in persis-tent infections that cause some of the most debilitatingdisease states recorded: lymphatic blockage resulting inelephantiasis, ocular pathology leading to river blindnessand severe dermatitis manifesting as sowda. While currentdrugs are effective (predominantly against the larval formsof the parasite), development of resistant strains hasrecently been reported (3). There is no vaccine to protectagainst filarial infection or to modulate disease.

There are eight filarial species of medical importancefor humans and a number of additional species of veteri-nary and scientific interest (Table 1). All of the filarial par-asites have common elements to their life cycle; mostnotably, all species are diecious, undergo four larvalmoults, are transmitted by hematophagus arthropod vec-tors and maintain developmental pauses at the first andthird larval stages. The developmental pauses are criticalfor successful transmission. The nature and severity ofpathology varies across filarial species with most of the tis-sue pathology and immunological changes associated withthe adult parasites and the transmissible larval stage.

The Phylum Nematoda contains five clades (4) and anestimated 1 million species (5). Filarial species reside inclade III (Spiruria), which is comprised of animal para-sites including ascaridid, spirurid and oxyurid parasites ofvertebrates and oxyurid and rhigonematid parasites ofarthropods (6). Since 1998, when it became the first multi-cellular organism to be fully sequenced (7), the terrestrial,

Correspondence: Alan L. Scott, Department of MolecularMicrobiology and Immunology, Bloomberg School of PublicHealth, Johns Hopkins University, 615 North Wolfe, Baltimore,MA 21205, USA (e-mail: [email protected]).Disclosures: None.Received: 3 August 2011Accepted for publication: 4 November 2011

Parasite Immunology, 2012, 34, 121–129 DOI: 10.1111/j.1365-3024.2011.01344.x

� 2011 Blackwell Publishing Ltd 121

bacteriovorus, clade V species Caenorhabditis elegans hasdominated as a prototype system for advancing under-standing in the molecular genetics and developmental biol-ogy of nematodes. It also represents a powerful model thatcontinues to contribute to a variety of important fieldsincluding medical genetics, ageing, cancer and infectiousdiseases [reviewed in (8–10)]. Early on, it was presumed,given the common body plan and developmental progres-sion of all nematode species, that the structure, organiza-tion, gene content and regulation of the C. elegansgenome could serve as an apt general model for nema-todes. With the publication of the genome from the firstparasitic nematode, B. malayi, with its unique genomestructure, large number of unique genes and the discoverythat many filarial species harbour an endosymbiont (11),the idea was sown that no single nematode genome couldserve as a general model. This idea has been supported bysubsequent reports on the composition and structure ofother filarial and nonfilarial nematode genomes (12–14).It appears that diversity in nematode genome structureand content parallels the extensive diversity found in theNematoda itself.

One of the notable advances to emerge from theB. malayi sequencing project was an appreciation thatstudies on the genomics and biology of many filarial spe-cies are incomplete without a consideration of the genomeof their endosymbiotic bacterium Wolbachia (Table 1).The current idea that Wolbachia and filarial parasites haveevolved a mutualistic relationship in which the bacteriumexerts control over the development (moulting), fecundityand viability of its host (15,16) underscores the importanceof integrated ensemble studies of these two organisms.

The advances associated with next-generation deepsequencing have significantly reduced the costs of sequenc-ing, assembling and annotating genomes. These technicaland informatics advances have already made an impact onthe field of nematode genomics through increasing thecorpus of nematode (including filarial) sequenced genomesand by providing support data that refine and expandassembly and annotation. The ability to produce sequencefree of the traditional bias associated with conventionalsequencing library construction has facilitated the fillingof gaps and the generation of higher-order assemblies.The enhanced transcriptomic capabilities conferred by

Table 1 Filarial and Wolbachia genomes

Vertebratehost

Status of genomesequencing

Number ESTs(clusters)a Wolbachia

Status ofgenome

Brugia malayi Human Build 1 complete build 2in progress

25 844 (9961) Yes Completed Draft

Brugia timori Human – – Yes –Wuchereria bancrofti Human Assembly and annotation

in progress4871 (2279) Yes Assembly and annotation

in progressOnchocerca volvulus Human Assembly and annotation

in progress14 791 (5144) Yes Assembly and annotation

in progressLoa loa Human Assembly and annotation

in progress27 (26) No –

Mansonella perstans Human – – Yes –Mansonella

streptocercaHuman – – No –

Mansonella ozzardi Human – – Yes –Brugia pahangi Feline – 28 (28) Yes –Dirofilaria immitis Canine, feline Assembly and annotation


in progressOnchocerca ochengi Bovine Assembly and annotation


in progressSetaria digitata Bovine – – No –Onchocerca cervicalis Equine – – Yes –Onchocerca flexuosa Cervidae Genome surveyb 2063 (1527) No –Litomosoides

sigmodontisRodent Assembly and annotation


in progressAcanthocheilonmea

viteaeRodent Genome surveyb – No –

aNumber of ESTs and clusters compiled in the most recent release of NEMBASE 4 (http://www.nematodes.org/nembase4/); bMcNulty et al.2010 (50). ESTs, expressed sequence tags.

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next-generation sequencing approaches are providingunprecedented insights into the full spectrum of stage-,sex- and tissue-associated gene expression that is refiningand expanding the complement of genes for each speciesand enhancing annotation through the identification ofnew exons, spliced variants and gene boundaries. In addi-tion, the next-generation sequencing platforms have facili-tated the initial studies in defining gene regulation infilarial species through the identification of micro RNAs.The goal of this review is to provide an outline of theadvances that are taking place in filarial nematode andWolbachia genomics and how these advances are expand-ing and refining our understanding of the structure andcomposition of filarial genomes, the regulation of geneexpression and the molecular interactions that defineendosymbiosis between filarial parasites and Wolbachia.

THE BRUGIA MALAYI GENOME

The reported draft of the B. malayi 95-Mb nuclear gen-ome was based on a traditional shotgun sequencingapproach that accounted for an estimated 90% of the gen-ome at approximately nine-fold coverage (11). Thisresulted in an assembly distributed on >8000 scaffolds thatranged in size from <2 kb to more than 6Æ5 Mb. It wasestimated that 14% of the genome was made up ofrepeated DNA comprised mainly of the major A+T-rich322-bp HhaI repeat family, the 62 ⁄ 53-bp MboI repeat fam-ily and large number of simple and low complexity repeats.The number and distribution of the tandemly arrayedrepeat elements presented a significant challenge to the fullassembly of the B. malayi genome into chromosomal units(1N = 5 autosomal and 1X).

Deep sequencing approaches have recently been appliedto extend and enhance the resolution of the B. malayi gen-ome and to resolve some of the content and assemblyissues encountered in the conventional shotgun-basedsequencing. While the analysis of these data is in progress,it is clear that the combined short read (Illumina) andlong read (pyrosequencing, 454 Life Sciences; Roche,Branford, CT, USA) strategy will have a significant impacton the resolution of the B. malayi genome. The combinedIllumina ⁄ 454 sequence data have increased the coverage to20X resulting in the annotation of an additional approxi-mately 6 Mb into scaffolds. In addition, the new sequenc-ing efforts have resulted in a significant increase in theN50 of the scaffolds from 93 to 357 kb and decreased thenumber of scaffolds from 8236 to under 2900 (this work isin progress). The expectation is that this next build of theB. malayi genome will also contain enhanced annotationof genes and regulatory elements provided by recentlycompleted transcriptomic and proteomic studies (17–20).

TRANSCRIPTOMICS

Analyses of the gene complements of the three free-living(C. elegans, Caenorhabditis briggsae and Pristionchus paci-ficus) and the four parasitic (B. malayi, Trichinella spiralis,Meloidogyne incognita and Meloidogyne hapla) publishednematode genomes indicate that as many as 45% of thepredicted protein-coding genes for each organism representnovel, species-specific entries into the major databases(7,12–14,21,22). Given the challenges that this level of pro-tein sequence diversity space presents for informatics-based gene finding and annotation of nematode genomes,transcriptomic and proteomic approaches play especiallyimportant roles in defining and verifying gene content.Indeed, the >25 000 expressed sequence tags (EST)derived from over a dozen larval and adult cDNAlibraries (representing over approximately 10 000 clusters;Table 1) (23) provided critical support data during theannotation of the B. malayi genome (11). Although ESTdatabases have proven to be a rich source for gene discov-ery and genome annotation, the EST approach is relativelyinefficient and fraught with cloning bias and limitationsassociated with single-pass sequence data and dealing withpartial sequences. The recent ability to apply massivelyparallel sequencing strategies provides a more efficient andostensibly less biased approach to define full-length tran-scripts and detail gene expression. The value of this com-plementary approach is exemplified by the modENCODEproject for C. elegans, where RNAseq and proteomics wereemployed along with other methods to identify 1650 here-tofore unrecognized protein-coding genes, redefine thestructure of known genes, reveal the scope of alternativesplicing and increase the estimate of diversity of noncod-ing RNAs 20-fold (24).

The initial use of this RNAseq approach for filarial trans-criptomics is a project in which the Illumina deep sequenc-ing platform was employed to produce over 108 paired endreads to gain a high-resolution expression profile ofB. malayi eggs ⁄ embryos, immature (£3 days of age) andmature microfilariae, third- and fourth-stage larvae andadult male and female worms (in annotation; S. Michalskiand B. Christensen, personal communication). It is antici-pated that this analysis will, in addition to expanding on the11 508 predicted genes (11), provide enhanced detail on thegenomic organization of previously identified B. malayigenes including mispredicted and unpredicted exons, thenature of splice variants and 5¢ and 3¢ UTR sequences.

PROTEOMICS

The identification of a complement of 11 508 protein-cod-ing genes in the initial draft of B. malayi genome has

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provided the baseline data needed to apply high-through-put proteomic approaches to augment genome annotationincluding identification of translational start and stopsites, frame shifts and the verification of the translation ofthose gene models designated as hypothetical ⁄ predicted.Bennuru et al. (18) have recently published a high-densityproteome map of B. malayi using reverse-phase liquidchromatography–tandem mass spectroscopy to define theexcretory ⁄ secretory (ES) and somatic proteins (17) pro-duced by the adults, microfilariae and infective stagelarvae. Taken together, these analyses identified 7103(>60%) of the proteins predicted in the genome including2336 of the 4956 genes that had been assigned hypotheti-cal ⁄ predicted status (11). A noteworthy observation fromthe analyses of the ES and somatic extracts is that a sig-nificant percentage (approximately 30%) of the proteinsare synthesized in a developmentally regulated manner.While many of these stage-associated proteins are uniqueto the databases and have no discernable functionalmotifs, this regulated pattern of translation implicatesthese molecules in processes important for filarial survival.As other filarial species attain the level of annotation thatwill allow the application of similar proteomicsapproaches, it will be fascinating to determine whether thisdevelopmental regulation of protein production holds trueand whether integrated comparative genomic ⁄ transcrip-tomic ⁄ proteomic analysis will provide the power to assignputative roles to the many hypothetical ⁄ predicted filarialgenes.

ADDITIONAL FILARIAL GENOMES

As noted above, next-generation sequencing technologieshave made it possible to rapidly increase the number ofnematode genomes available for analysis of structure andcontent. Recently, sequence data for the genomes of threemajor human filarial pathogens, W. bancrofti, O. volvulusand L. loa, have been made publically available (http://www.broadinstitute.org/annotation/genome/filarial_worms/MultiHome.html). In addition, draft genome sequence willsoon be available for a number of filarial pathogens of ani-mals including Dirofilaria immitis, Litomosoides sigmodontisand O. ochengi (http://www.nematodes.org/nematodegenomes/index.php/Species_in_sequencing).

In the very near future, this rich substrate of filarialgenomes will enable an unprecedented opportunity toidentify the biochemical and regulatory pathways that arecommon and unique to filarial species and thus generaterational models for filarial physiology that point to targetsfor drug- or small molecule-based control measures. Com-parative analysis of closely related species pairs such asB. malayi and W. bancrofti and O. volvulus and O. ochengi

will provide data for better models of filarial evolutionand the adaptations that resulted in certain species beingrestricted to a single vertebrate host. It will be of greatinterest to define the degree of short- and long-range synt-eny between the genomes of filarial species. The genomesmay also provide evidence of the distinct and commonmechanisms required for nematode survival and transmis-sion by mosquitoes and flies.

SMALL NONCODING RNAS

The classical view of RNAs [tRNA, mRNA and rRNA]as molecules restricted to protein synthesis has been sig-nificantly modified over the last decade to include smallnoncoding RNAs that function as vital regulatory mole-cules. Nematodes have played a key role in seminal obser-vations in this arena. The first 22-nucleotide microRNA(miRNA) was discovered by Ambros and Ruvkun in asearch for the cause of a developmental timing defect inC. elegans. They discovered that a mutation in a miRNA,lin-4, altered its ability to bind to the 3¢ untranslatedregion and inhibit the expression of LIN-14, a hetero-chronic regulatory nuclear protein (25,26). The discoveryof RNA interference (RNAi) in C. elegans (27) providedthe basis for defining the mechanisms that govern thegeneration and action of miRNAs. From these initialnematode-derived observations, it has been demonstratedin eukaryotes from sponges to mammals that, in additionto the regulation of gene expression, small noncodingRNAs such as miRNAs (microRNAs), siRNAs (smallinterfering RNAs) and piRNAs (Piwi-interacting RNAs)are critical for the control of cellular metabolism, growthand differentiation, to maintain genome integrity and as adefence mechanism against viruses and mobile genetic ele-ments (28,29).

Recently, noncoding RNAs have emerged as a majorcomponent of the eukaryotic transcriptome. Transcriptionof miRNAs results in the formation of a dsRNA hairpinthat is sequentially cleaved by the double-stranded (ds)RNA-specific endoribonucleases Drosha and Dicer to gen-erate a 22-nt dsRNA. In association with proteins of theArgonaute family, the dsRNA is unwound and directed toits cognate RNA and ⁄ or DNA molecules in the context ofthe RNA-induced silencing complex (RISC) (30). Part ofthe miRNA binding specificity for target mRNAs is deter-mined by a 6- to 7-nt ‘seed’ sequence at the 5¢ end of eachmiRNA (31). The B. malayi genome contains many of thecomponents required for miRNA processing includingorthologues of Drosha, Dicer and Argonaute (11).Recently, deep sequencing has been employed to explorethe diversity of small RNAs expressed by mature male,female and microfilariae of B. malayi (32).



Employing approximately 30 million deep sequencingreads, 145 miRNAs were identified in the adults and lar-vae of B. malayi. The miRNAs represent 99 families eachdefined by a unique seed sequence. Sixty of the miRNAfamilies found in B. malayi are conserved in other species.Examples of conserved miRNAs include the let-7, miR-1and bantam families (32, C. B. Poole, W. Gu, P. Davis,S. Kumar, J. Jin, D. Conte, Mello C. & L. A. McReynolds,unpublished data). These highly conserved miRNAs arelikely involved in the regulation and maintenance ofconserved developmental rather than species-specificfunctions.

Only 11 ⁄ 99 B. malayi miRNA families are restricted tohelminths. Of this group, nine are found only in filarialparasites, including L. loa, W. bancrofti and O. volvulus.The sequences of the filarial-specific miRNAs are quitesimilar in all the filarial species examined, suggesting thatthey have recently evolved by gene-duplication and ⁄ or sin-gle base substitutions from existing miRNA families (C. B.Poole, W. Gu, P. Davis, S. Kumar, J. Jin, D. Conte, MelloC. & L. A. McReynolds, unpublished data). Additionalstudies are needed to identify the mRNA targets of theB. malayi miRNAs and to determine whether they areinvolved in the maintenance of parasitism.

About 30% of the B. malayi miRNAs are preferentiallyexpressed in one stage over another. For example, miR-71and miR-34 are 15-times more highly expressed in micro-filariae compared to adult male and female parasites.Together, miR-71 [27%] and mR-34 [13%] represent 40%of the total miRNAs found in microfilariae. A similar dif-ferential increase in miR-71 and miR-34 has been noted inC. elegans diapause L1s compared to embryos (33). Theprolonged pause in development of blood-stage microfila-riae is thought to be analogous to diapause in C. elegans.The role of miR-71 and miR-34 extends beyond the con-trol of developmental arrest. For both C. elegans adultsand mammalian cells, miR-34 enhances radiation resis-tance (34). In C. elegans, the loss of miR-71 results in atwo-fold reduction in life span possibly through blockingmembers of the DAF-2, insulin-like growth factor pathway(35). It will be of interest to determine whether miR-71and miR-34 govern stress resistance and maintenance ofarrested development in filarial nematodes.

Inspection of the B. malayi miRNA families revealsseveral examples of miRNA evolution. Current models pro-pose that miRNAs evolve by a combination of gene ampli-fication, mutation and arm switching mechanisms (36). TheB. malayi miR-2 family is an example of gene amplifica-tion. This family has 11 members and has expanded com-pared to the five-member C. elegans miR-2 family. Six ofthe B. malayi miR-2 family members are more closelyrelated to each other than to C. elegans miR-2, suggesting

a recent expansion of this family in filarial parasites.Several of the filarial-specific miRs likely arose by mutationfrom conserved miR families. For example, alignment ofB. malayi miR-57 with filarial-specific miR-A identified asingle mutation in the seed sequence of miR-A comparedto miR-57. With the overall identity of the two miRNAs at67%, these two miRNAs likely arose from a commonancestor by duplication of miR-57 followed by nt substitu-tions to create the two different miRNAs (C. B. Poole,W. Gu, P. Davis, S. Kumar, J. Jin, D. Conte, Mello C. &L. A. McReynolds, unpublished data).

As noted above, the repeat content of the B. malayi gen-ome is approximately 14%. The dominant repeat, the 322-bp HhaI DNA repeat family (37), encodes a small RNA.The HhaI RNA is present in both the adult and mf stagesand is transcribed from one strand of the DNA. A com-putational search did not find any sequences that couldform a stable dsRNA hairpin, suggesting that this RNA isnot generated by Drosha or Dicer cleavage. The functionof this small noncoding RNA is not known.

WOLBACHIA

Most species of filarial nematodes harbour within their tis-sues an a-proteobacterial endosymbiont belonging to thegenus Wolbachia. These Rickettsia-like bacteria are presentin all filarial species that infect humans with the exceptionof L. loa and M. streptocerca. The maternally transmittedendosymbionts are found predominantly in the hypoder-mal cells of the lateral cord as well as the ovaries, oocytesand developing embryos within the uteri of adult femaleworms [see (38) for review]. Wolbachia were first describedin insect hosts and are widespread in arthropods withclose to 70% of species believed infected (39). In arthro-pod hosts, the endobacteria are considered reproductiveparasites because they induce a variety of phenotypes suchas cytoplasmic incompatibility, parthenogenesis, feminiza-tion of males and the killing of male embryos, which serveto promote the fitness of infected females and the spreadof endosymbiont through populations (40). Furthermore,Wolbachia can be cleared from their arthropod hosts byantibiotic treatment with little or no consequence (41).

In contrast, the Wolbachia infecting filarial nematodesdisplay the features of an obligate mutualist – neitherorganism can survive long term without the other.Attempts to culture the Wolbachia endosymbiont fromB. malayi (wBm) or to adoptively transfer wBm to analternate host species have not been successful. Treatmentwith tetracycline family antibiotics clear the Wolbachiafrom filarial parasites, and this clearance precipitates ablock in embryogenesis followed eventually by the deathof the worm. Similar treatment of Wolbachia-free filarial



species shows no such effects on worm viability. Clinicaltrials with tetracyclines have validated the Wolbachiaendosymbionts as an anti-filarial target, thus demonstrat-ing a novel effective treatment, which, importantly, hasboth microfilaricidal and macrofilaricidal activity. How-ever, the required long courses of tetracycline treatmentand the contraindications for young children and pregnantwomen preclude its widespread implementation.

The genome sequence of the Wolbachia endosymbiontfrom B. malayi (wBm) was determined to facilitate under-standing of the symbiosis between the bacterium and itsfilarial host and to assist in the identification of Wolba-chia biochemical processes that could serve as novel anti-filarial drug targets. Typical of most endosymbionts, the1Æ08-Mb-circular wBm genome shows loss of many meta-bolic processes and is predicted to encode only 806 pro-tein-coding genes. Mapping the genes to metabolicpathways indicated that the biosynthesis of riboflavin, fla-vin adenine dinucleotide (FAD), haeme and nucleotidesmight be important for the symbiotic relationship withthe B. malayi host (42). These biosynthetic capabilitiesappear not to be encoded by the B. malayi genome,although nucleotide salvage pathways exist (11). Con-versely, nematode-derived amino acids appear importantfor wBm growth (42). Interestingly, among the few wBmgenes showing positive selection in a genome-wide screenfor the presence of diversifying selection were severalgenes involved in biosynthesis of haeme, riboflavin andnucleotide biosynthesis (43). This observation lendsweight to the notion that these pathways may be keycomponents of the mutualistic association.

Additional bioinformatic-based discovery and experi-mentation have predicted the essential gene repertoire ofwBm and identified several Wolbachia biochemical pro-cesses that represent candidate drug targets. These includeenzymes of lipid II biosynthesis, lipoprotein biosynthesis,haeme biosynthesis, and the glycolytic enzymes pyruvatephosphate dikinase and cofactor-independent phospho-glycerate mutase. (44–49). Several of these are undergoingfurther evaluation as potential targets or are already sub-ject to high-throughput inhibitor screens.

A recent proteomic analysis of extracts from wholeadult and larval B. malayi also identified 577 of the 806wBm proteins (17). This analysis demonstrated that manyof the wBm proteins were differentially transcribed in thevarious parasite stages. In addition, 96 of the 166 hypo-thetical ⁄ predicted genes (42) were validated as producing aprotein during one or more stages of filarial development.Proteomic-level identification of all of the proteinsrequired for nucleotide and haeme biosynthesis in wBmsupports the hypothesis that the endosymbiont providescritical factors required for parasite survival.

If Wolbachia provides metabolites to its filarial hosts aspart of the endosymbiotic relationship, an obvious ques-tion is how do those Wolbachia-free filarial species fulfiltheir metabolic needs? To address this question, low-cover-age genomic sequencing of O. flexuosa and Acanthochei-lonmea viteae, filarial species uninfected by Wolbachia, wasundertaken (50). In both species, fragments of DNA origi-nating from Wolbachia were evident in the nucleargenomes, indicating an ancestral Wolbachia infection fol-lowed by secondary loss of endosymbiont. Although somefragments were transcribed, none appeared capable ofencoding a full-length functional protein. The laterallytransferred Wolbachia DNA was present as short degener-ated gene fragments that typically contained frame shiftsand stop codons. Interestingly, this low-coverage sequencedata also indicated that the Wolbachia-free filarial species,like B. malayi, were deficient in the biosynthesis of haeme,nucleotides, riboflavin and FAD [J. M. Foster and B. E.Slatko, unpublished data]. The absence of nematode- orWolbachia-derived genes encoding these biosynthetic capa-bilities raises questions regarding how the Wolbachia-freespecies fulfil these functions and the relevance of theseWolbachia-derived metabolites in species that do harbourWolbachia. It is possible that the low-coverage genomicsequencing of A. viteae and O. flexuosa simply missedfunctional genome- or Wolbachia-encoded versions thesemolecules. An alternative explanation is that filarial nema-todes can meet their requirements for these moleculesfrom exogenous sources and that metabolic provisioningby Wolbachia in infected filarial species is perhaps simplybeneficial under steady state and only essential either atcritical points in the filarial life cycle or under certainstressful conditions. Indeed, although Wolbachia withinarthropods are typically considered ‘parasites’, there isgrowing evidence that they can also confer benefits totheir hosts – including metabolic provisioning of moleculessuch as haeme and vitamin B (e.g. riboflavin) – under cer-tain conditions (51,52).

While Wolbachia-infected filarial species may havebecome dependent on their endosymbiont for metabolitesover evolutionary time, uninfected filarial species may haveevolved to be more proficient at securing essential compo-nents from their surroundings. Despite uncertainties aboutthe contribution of metabolic provisioning to the mutualis-tic association, the Wolbachia within filarial nematodes areclearly essential as demonstrated by the outcome of antibi-otic treatment [see (38) for review].

Sequencing of Wolbachia genomes from additional filar-ial hosts together with more nematode genomes from bothWolbachia-infected and uninfected filarial species will facil-itate a greater understanding of the nature of the symbio-sis. Phylogenetic analyses resolve Wolbachia into about



seven distinct supergroups and a number of additional lin-eages (53). Wolbachia from filarial nematodes are foundalmost exclusively in Supergroups C and D, while theother supergroups are comprised of arthropod Wolbachia.Supergroup F is an interesting case because it containsWolbachia from both arthropods and filariae (Mansonellaperstans) and probably reflects a relatively recent transferof Wolbachia between these invertebrate phyla (54). Apartfrom wBm (supergroup D), only three other completed cir-cular Wolbachia genomes exist – those from Drosophilamelanogaster (wMel) and Drosophila simulans (wRi) fromsupergroup A and Culex quinquefasciatus (wPip) fromsupergroup B (55–57). The Wolbachia genomes from thesedifferent supergroups are highly scrambled relative to eachother and show little synteny (42,55). A few areas ofshort-range synteny exist and probably reflect functionallyrelated gene products such as those comprising the TypeIV secretion system. Comparison of the two supergroup Agenomes also shows a high degree of mosaicism caused byextensive recombination facilitated by the insertion ele-ments and repeat blocks that populate arthropod Wolba-chia genomes, but in this case there is considerably morelong-range synteny (56).

A number of additional Wolbachia genomes have beensequenced to high coverage by next-generation sequencingtechnologies, but they remain incomplete or are under-going finishing. These include Wolbachia genomes fromthe filarial nematodes O. volvulus, O. ochengi, D. immitis(all supergroup C), W. bancrofti and L. sigmodontis(both supergroup D). [see http://www.broadinstitute.org/annotation/genome/filarial_worms/GenomesIndex.html andhttp://xyala.cap.ed.ac.uk/downloads/959nematodegenomes/blast/blast.php for details].

Analysis of a collection of bacterial artificial chromo-somes prepared from genomic DNA from D. immitis iden-tified a subset that was derived from the Wolbachiaendosymbiont (wDi; supergroup C) [J. M. Foster andB. E. Slatko, unpublished data]. DNA sequences weredetermined for both ends of these Wolbachia inserts (typi-cally about 60 kb long) and mapped onto both the wBmgenome (supergroup D) and, where possible, the wOvo(supergroup C) contigs that represent about 10% of theWolbachia genome from O. volvulus (available at NCBI).No syntenic arrangement of adjacent genes from one end

of a wDi BAC or genes from opposite ends of a wDi BAC(about 60 kb apart) were found when mapped to wBm(supergroup D). However, examples of synteny betweenwDi and wOvo (both supergroup C) were apparent. Thisanalysis predicts that future comparisons of new Wolbachi-a genome sequences from various filarial nematode hostswill reveal greatest structural conservation when membersof the same Wolbachia supergroup are compared, similarto the situation observed for Wolbachia genomes fromarthropod hosts. The new insight into the symbiotic rela-tionship between Wolbachia and their filarial nematodehosts that might be afforded by the completion of theongoing Wolbachia genome projects and subsequent com-parative genomics is eagerly awaited.

WHERE DO WE GO FROM HERE?

Filarial nematodes impose an appalling toll on humandevelopment and well-being. While C. elegans will maintainits importance as a model for understanding certain aspectsof nematode biology, the genome-level data from theincreasing number of filarial and nonfilarial nematode spe-cies make it clear that the C. elegans genome cannot be usedas a prototype for gene content or genome structure inNematoda. It will soon be possible to employ large-scalecomparative genomics within Nematoda to aid in identify-ing the core genetic elements that define a nematode. Inaddition, detailed genomics combined with transcriptomics,proteomics, metabolomics, biochemistry, physiology, etc.will serve as the bases for a systems biology approach toestablish a high resolution understanding the cellular andmolecular foundation for filarial nematode development intheir vertebrate and arthropod hosts, the requirements tomaintain the mutualistic relationship with their endos-ymbionts and the immunobiology of filarial infections.

ACKNOWLEDGEMENTS

We thank our colleagues of the Filarial Genome Projectand the filarial research community for their continuedsupport and encouragement. We thank Shelly Michalski(University of Wisconsin, Oshkosh) and Bruce Christensen(University of Wisconsin, Madison) for their input onB. malayi transcriptomics.

REFERENCES

1 Chu BK, Hooper PJ, Bradley MH, McFar-land DA & Ottesen EA. The economic bene-fits resulting from the first 8 years of theGlobal Programme to Eliminate LymphaticFilariasis (2000–2007). PLoS Negl Trop Dis2010; 4: e708.

2 Ottesen EA. Lymphatic filariasis: treatment,control and elimination. Adv Parasitol 2006;61: 395–441.

3 Osei-Atweneboana MY, Awadzi K, AttahSK, Boakye DA, Gyapong JO & PrichardRK. Phenotypic evidence of emerging iver-

mectin resistance in Onchocerca volvulus.PLoS Negl Trop Dis 2011; 5: e998.

4 Blaxter ML, De Ley P, Garey JR, et al. Amolecular evolutionary framework for thephylum Nematoda. Nature 1998; 392: 71–75.



5 Lambshead PJ, Brown CJ, Ferrero TJ, Haw-kins LE, Smith CR & Mitchell NJ. Biodiver-sity of nematode assemblages from theregion of the Clarion-Clipperton FractureZone, an area of commercial mining interest.BMC Ecol 2003; 3: 1.

6 Blaxter ML. Nematoda: genes, genomes andthe evolution of parasitism. Adv Parasitol2003; 54: 101–195.

7 Consortium, C. e. S. Genome sequence of thenematode C. elegans: a platform for investi-gating biology. Science 1998; 282:2012–2018.

8 Olsen A, Vantipalli MC & Lithgow GJ.Using Caenorhabditis elegans as a model foraging and age-related diseases. Ann N YAcad Sci 2006; 1067: 120–128.

9 Poulin G, Nandakumar R & Ahringer J.Genome-wide RNAi screens in Caenorhabd-itis elegans: impact on cancer research. Onco-gene 2004; 23: 8340–8345.

10 Pradel E & Ewbank JJ. Genetic models inpathogenesis. Annu Rev Genet 2004; 38: 347–363.

11 Ghedin E, Wang S, Spiro D, et al. Draftgenome of the filarial nematode parasiteBrugia malayi. Science 2007; 317: 1756–1760.

12 Mitreva M, Jasmer DP, Zarlenga DS, et al.The draft genome of the parasitic nematodeTrichinella spiralis. Nat Genet 2011; 43: 228–235.

13 Opperman CH, Bird DM, Williamson VM,et al. Sequence and genetic map of Meloido-gyne hapla: a compact nematode genome forplant parasitism. Proc Natl Acad Sci U S A2008; 105: 14802–14807.

14 Abad P, Gouzy J, Aury JM, et al. Genomesequence of the metazoan plant-parasiticnematode Meloidogyne incognita. Nat Bio-technol 2008; 26: 909–915.

15 Fenn K & Blaxter M. Are filarial nematodeWolbachia obligate mutualist symbionts?Trends Ecol Evol 2004; 19: 163–166.

16 Taylor MJ, Bandi C & Hoerauf A. Wolbachi-a bacterial endosymbionts of filarial nema-todes. Adv Parasitol 2005; 60: 245–284.

17 Bennuru S, Meng Z, Ribeiro JMC, et al.Stage-specific proteomic expression patternsof the human filarial parasite Brugia malayiand its endosymbiont Wolbachia. Proc NatlAcad Sci U S A 2011; 108: 1234–1235.

18 Bennuru S, Semnani R, Meng Z, RibeiroJM, Veenstra TD & Nutman TB. Brugia ma-layi excreted ⁄ secreted proteins at thehost ⁄ parasite interface: stage- and gender-specific proteomic profiling. PLoS Negl TropDis 2009; 3: e410.

19 Hewitson JP, Harcus YM, Curwen RS, et al.The secretome of the filarial parasite, Brugiamalayi: proteomic profile of adult excretory-secretory products. Mol Biochem Parasitol2008; 160: 8–21.

20 Moreno Y & Geary TG. Stage- and gender-specific proteomic analysis of Brugia malayiexcretory-secretory products. PLoS NeglTrop Dis 2008; 2: e326.

21 Dieterich C, Clifton SW, Schuster LN, et al.The Pristionchus pacificus genome provides aunique perspective on nematode lifestyle andparasitism. Nat Genet 2008; 40: 1193–1198.

22 Stein LD, Bao Z, Blasiar D, et al. The gen-ome sequence of Caenorhabditis briggsae: aplatform for comparative genomics. PLoSBiol 2003; 1:E45.

23 Elsworth B, Wasmuth J & Blaxter M. NEM-BASE4: the nematode transcriptomeresource. Int J Parasitol 2011; 41: 881–894.

24 Gerstein MB, Lu ZJ, Van Nostrand EL,et al. Integrative analysis of the Caenorhabd-itis elegans genome by the modENCODEproject. Science 2010; 330: 1775–1787.

25 Lee RC, Feinbaum RL & Ambros V. TheC. elegans heterochronic gene lin-4 encodessmall RNAs with antisense complementarityto lin-14. Cell 1993; 75: 843–854.

26 Wightman B, Ha I & Ruvkun G. Posttran-scriptional regulation of the heterochronicgene lin-14 by lin-4 mediates temporal pat-tern formation in C. elegans. Cell 1993; 75:855–862.

27 Fire A, Xu S, Montgomery MK, Kostas SA,Driver SE & Mello CC. Potent and specificgenetic interference by double-stranded RNAin Caenorhabditis elegans. Nature 1998; 391:806–811.

28 Wheeler BM, Heimberg AM, Moy VN, et al.The deep evolution of metazoan microR-NAs. Evol Dev 2009; 11: 50–68.

29 Tabara H, Sarkissian M, Kelly WG, et al.The rde-1 gene, RNA interference, andtransposon silencing in C. elegans. Cell 1999;99: 123–132.

30 Ghildiyal M & Zamore PD. Small silencingRNAs: an expanding universe. Nat RevGenet 2009; 10: 94–108.

31 Lewis BP, Burge CB & Bartel DP. Con-served seed pairing, often flanked by ade-nosines, indicates that thousands of humangenes are microRNA targets. Cell 2005;120: 15–20.

32 Poole CB, Davis PJ, Jin J & McReynoldsLA. Cloning and bioinformatic identificationof small RNAs in the filarial nematode, Bru-gia malayi. Mol Biochem Parasitol 2010; 169:87–94.

33 Karp X, Hammell M, Ow MC & Ambros V.Effect of life history on microRNA expres-sion during C. elegans development. RNA2011; 17: 639–651.

34 Kato M, Paranjape T, Muller RU, et al. Themir-34 microRNA is required for the DNAdamage response in vivo in C. elegans and invitro in human breast cancer cells. Oncogene2009; 28: 2419–2424.

35 de Lencastre A, Pincus Z, Zhou K, Kato M,Lee SS & Slack FJ. MicroRNAs both pro-mote and antagonize longevity in C. elegans.Curr Biol 2010; 20: 2159–2168.

36 de Wit E, Linsen SE, Cuppen E & BerezikovE. Repertoire and evolution of miRNAgenes in four divergent nematode species.Genome Res 2009; 19: 2064–2074.

37 McReynolds LA, DeSimone SM & WilliamsSA. Cloning and comparison of repeatedDNA sequences from the human filarial par-asite Brugia malayi and the animal parasiteBrugia pahangi. Proc Natl Acad Sci U S A1986; 83: 797–801.

38 Foster JM, Hoerauf A, Slatko BE & TaylorMJ. The molecular biology, immunology andchemotherapy of Wolbachia bacterial endos-ymbionts of filarial nematodes. In KennedyM, Harnett W (eds): Parasitic Nematodes:Molecular Biology, Biochemistry and Immunol-ogy. Wallingford, UK, CABI, 2011: (in press).

39 Hilgenboecker K, Hammerstein P, Schlatt-mann P, Telschow A & Werren JH. Howmany species are infected with Wolbachia? –A statistical analysis of current data. FEMSMicrobiol Lett 2008;281:215–220.

40 Werren JH, Baldo L & Clark ME. Wolbachi-a: master manipulators of invertebrate biol-ogy. Nat Rev Microbiol 2008; 6: 741–751.

41 Werren JH. Biology of Wolbachia. Annu RevEntomol 1997; 42: 587–609.

42 Foster J, Ganatra M, Kamal I, et al. TheWolbachia genome of Brugia malayi: endo-symbiont evolution within a human patho-genic nematode. PLoS Biol 2005; 3: e121.

43 Brownlie JC, Adamski M, Slatko B &McGraw EA. Diversifying selection and hostadaptation in two endosymbiont genomes.BMC Evol Biol 2007; 7: 68.

44 Foster JM, Raverdy S, Ganatra MB, ColussiPA, Taron CH & Carlow CK. The Wolbachi-a endosymbiont of Brugia malayi has anactive phosphoglycerate mutase: a candidatetarget for anti-filarial therapies. Parasitol Res2009; 104: 1047–1052.

45 Henrichfreise B, Schiefer A, Schneider T,et al. Functional conservation of the lipid IIbiosynthesis pathway in the cell wall-less bac-teria Chlamydia and Wolbachia: why is lipidII needed? Mol Microbiol 2009; 73: 913–923.

46 Holman AG, Davis PJ, Foster JM, CarlowCK & Kumar S. Computational predictionof essential genes in an unculturable endo-symbiotic bacterium, Wolbachia of Brugiamalayi. BMC Microbiol 2009; 9: 243.

47 Johnston KL, Wu B, Guimaraes A, Ford L,Slatko BE & Taylor MJ. Lipoprotein biosyn-thesis as a target for anti-Wolbachia treatmentof filarial nematodes. Parasit Vectors 2010; 3: 99.

48 Raverdy S, Foster JM, Roopenian E & Car-low CK. The Wolbachia endosymbiont ofBrugia malayi has an active pyruvate phos-phate dikinase. Mol Biochem Parasitol 2008;160: 163–166.

49 Wu B, Novelli J, Foster J, et al. The hemebiosynthetic pathway of the obligate Wolba-chia endosymbiont of Brugia malayi as apotential anti-filarial drug target. PLoS NeglTrop Dis 2009; 3: e475.

50 McNulty SN, Foster JM, Mitreva M, et al.Endosymbiont DNA in endobacteria-freefilarial nematodes indicates ancient horizontalgenetic transfer. PLoS ONE 2010; 5: e11029.

51 Brownlie JC, Cass BN, Riegler M, et al. Evi-dence for metabolic provisioning by a com-mon invertebrate endosymbiont, Wolbachiapipientis, during periods of nutritional stress.PLoS Pathog 2009; 5: e1000368.

52 Hosokawa T, Koga R, Kikuchi Y, Meng XY& Fukatsu T. Wolbachia as a bacteriocyte-associated nutritional mutualist. Proc NatlAcad Sci U S A 2010; 107: 769–774.



53 Bordenstein SR, Paraskevopoulos C, HotoppJC, et al. Parasitism and mutualism inWolbachia: what the phylogenomic trees canand cannot say. Mol Biol Evol 2009; 26: 231–241.

54 Casiraghi M, Bordenstein SR, Baldo L, et al.Phylogeny of Wolbachia pipientis based ongltA, groEL and ftsZ gene sequences: clus-tering of arthropod and nematode symbionts

in the F supergroup, and evidence for furtherdiversity in the Wolbachia tree. Microbiology2005; 151: 4015–4022.

55 Klasson L, Walker T, Sebaihia M, et al.Genome evolution of Wolbachia strain wPipfrom the Culex pipiens group. Mol Biol Evol2008; 25: 1877–1887.

56 Klasson L, Westberg J, Sapountzis P, et al.The mosaic genome structure of the Wolba-

chia wRi strain infecting Drosophila simulans.Proc Natl Acad Sci U S A 2009; 106: 5725–5730.

57 Wu M, Sun LV, Vamathevan J, et al. Phylog-enomics of the reproductive parasite Wolba-chia pipientis wMel: a streamlined genomeoverrun by mobile genetic elements. PLoSBiol 2004; 2: E69.



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Filarial and Wolbachia genomics