Ecology

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4.Bongrand, C. & Ruby, E. G. Achieving a multi-strain symbiosis: strain behavior and infection dynamics. ISME J. 13, 698–706 (2019).PubMed 
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7.Graf, J. & Ruby, E. G. Host-derived amino acids support the proliferation of symbiotic bacteria. Proc. Natl Acad. Sci. USA 95, 1818–1822 (1998).CAS 
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8.McFall-Ngai, M. J. & Ruby, E. G. Developmental biology in marine invertebrate symbioses. Curr. Opin. Microbiol. 3, 603–607 (2000).CAS 
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9.Moriano-Gutierrez, S. et al. The noncoding small RNA SsrA is released by Vibrio fischeri and modulates critical host responses. PLoS Biol. 18, e3000934 (2020).CAS 
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10.Schwartzman, J. A. & Ruby, E. G. Stress as a normal cue in the symbiotic environment. Trends Microbiol. 24, 414–424 (2016).CAS 
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11.Nyholm, S. V. & McFall-Ngai, M. J. A lasting symbiosis: how the Hawaiian bobtail squid finds and keeps its bioluminescent bacterial partner. Nat. Rev. Microbiol. https://doi.org/10.1038/s41579-021-00567-y (2021).12.Schwartzman, J. A. et al. The chemistry of negotiation: rhythmic, glycan-driven acidification in a symbiotic conversation. Proc. Natl Acad. Sci. USA 112, 566–571 (2015). In this study, the host’s delivery of chitin-derived N-acetylglucosamine is shown to develop 4 weeks after hatching, and this chitin is apparently delivered by haemocytes that lyse in the crypts only at night. A nocturnal acidification of the crypts results, and a model for how this outcome enhances bioluminescence is described.CAS 
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13.Heath-Heckman, E. A. et al. Bacterial bioluminescence regulates expression of a host cryptochrome gene in the squid-vibrio symbiosis. mBio https://doi.org/10.1128/mBio.00167-13 (2013).Article 
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15.Bouskra, D. et al. Lymphoid tissue genesis induced by commensals through NOD1 regulates intestinal homeostasis. Nature 456, 507–510 (2008).CAS 
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16.Thaiss, C. A. et al. Transkingdom control of microbiota diurnal oscillations promotes metabolic homeostasis. Cell 159, 514–529 (2014).CAS 
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17.Ruby, E. G. Symbiotic conversations are revealed under genetic interrogation. Nat. Rev. Microbiol. 6, 752–762 (2008).CAS 
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18.Bongrand, C. & Ruby, E. G. The impact of Vibrio fischeri strain variation on host colonization. Curr. Opin. Microbiol. 50, 15–19 (2019).CAS 
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19.Colton, D. M. & Stabb, E. V. Rethinking the roles of CRP, cAMP, and sugar-mediated global regulation in the Vibrionaceae. Curr. Genet. 62, 39–45 (2016).CAS 
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20.Mandel, M. J. & Dunn, A. K. Impact and Influence of the natural Vibrio-squid symbiosis in understanding bacterial-animal interactions. Front. Microbiol. 7, 1982 (2016).PubMed 
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21.Aschtgen, M. S. et al. Insights into flagellar function and mechanism from the squid-vibrio symbiosis. NPJ Biofilms Microbiomes 5, 32 (2019).PubMed 
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22.Stabb, E. V. & Visick, K. L. in The Prokaryotes (eds Rosenberg, E., DeLong, E. F., Lory, S., Stackebrandt, E. & Thompson, F.) 497–532 (Springer, 2013).23.Nawroth, J. C. et al. Motile cilia create fluid-mechanical microhabitats for the active recruitment of the host microbiome. Proc. Natl Acad. Sci. USA 114, 9510–9516 (2017). This work provides the first glimpse into the cilium-driven fluid mechanics that position V. fischeri cells to reach and settle in ‘quiet zones’ on the light organ surface, permitting a selective ‘recruitment’ of this microorganism from the planktonic environment.CAS 
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24.Altura, M. A. et al. The first engagement of partners in the Euprymna scolopes-Vibrio fischeri symbiosis is a two-step process initiated by a few environmental symbiont cells. Environ. Microbiol. 15, 2937–2950 (2013). Aggregations of only a few V. fischeri cells are observed to initiate normal host responses, and reveal that aggregation is a two-part process that begins with bacterial attachment to the cilia.PubMed 
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25.Nyholm, S. V., Stabb, E. V., Ruby, E. G. & McFall-Ngai, M. J. Establishment of an animal-bacterial association: recruiting symbiotic vibrios from the environment. Proc. Natl Acad. Sci. USA 97, 10231–10235 (2000).CAS 
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26.Yip, E. S., Geszvain, K., DeLoney-Marino, C. R. & Visick, K. L. The symbiosis regulator RscS controls the syp gene locus, biofilm formation and symbiotic aggregation by Vibrio fischeri. Mol. Microbiol. 62, 1586–1600 (2006).CAS 
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27.Koehler, S. et al. The model squid-vibrio symbiosis provides a window into the impact of strain- and species-level differences during the initial stages of symbiont engagement. Environ. Microbiol. https://doi.org/10.1111/1462-2920.14392 (2018).Article 
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28.Morris, A. R. & Visick, K. L. Control of biofilm formation and colonization in Vibrio fischeri: a role for partner switching? Environ. Microbiol. 12, 2051–2059 (2010).CAS 
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29.Norsworthy, A. N. & Visick, K. L. Gimme shelter: how Vibrio fischeri successfully navigates an animal’s multiple environments. Front. Microbiol. 4, 356 (2013).PubMed 
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30.Shibata, S., Yip, E. S., Quirke, K. P., Ondrey, J. M. & Visick, K. L. Roles of the structural symbiosis polysaccharide (syp) genes in host colonization, biofilm formation, and polysaccharide biosynthesis in Vibrio fischeri. J. Bacteriol. 194, 6736–6747 (2012).CAS 
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31.Yip, E. S., Grublesky, B. T., Hussa, E. A. & Visick, K. L. A novel, conserved cluster of genes promotes symbiotic colonization and sigma-dependent biofilm formation by Vibrio fischeri. Mol. Microbiol. 57, 1485–1498 (2005).CAS 
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32.Bassis, C. M. & Visick, K. L. The cyclic-di-GMP phosphodiesterase BinA negatively regulates cellulose-containing biofilms in Vibrio fischeri. J. Bacteriol. 192, 1269–1278 (2010).CAS 
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33.Chavez-Dozal, A., Hogan, D., Gorman, C., Quintanal-Villalonga, A. & Nishiguchi, M. K. Multiple Vibrio fischeri genes are involved in biofilm formation and host colonization. FEMS Microbiol. Ecol. 81, 562–573 (2012).CAS 
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34.Tischler, A. H., Lie, L., Thompson, C. M. & Visick, K. L. Discovery of calcium as a biofilm-promoting signal for Vibrio fischeri reveals new phenotypes and underlying regulatory complexity. J. Bacteriol. 200, e00016–e00018 (2018). This article expands our understanding of the regulatory controls and signals leading to biofilm formation by identifying calcium as a signal that induces a coordinate upregulation of Syp- and cellulose-dependent biofilm formation and revealing the sensor kinase HahK as a new biofilm regulator.CAS 
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35.Ziemba, C., Shabtai, Y., Piatkovsky, M. & Herzberg, M. Cellulose effects on morphology and elasticity of Vibrio fischeri biofilms. NPJ Biofilms Microbiomes 2, 1 (2016).PubMed 
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36.Ray, V. A., Driks, A. & Visick, K. L. Identification of a novel matrix protein that promotes biofilm maturation in Vibrio fischeri. J. Bacteriol. 197, 518–528 (2015).PubMed 
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37.Shibata, S. & Visick, K. L. Sensor kinase RscS induces the production of antigenically distinct outer membrane vesicles That depend on the symbiosis polysaccharide locus in Vibrio fischeri. J. Bacteriol. 194, 185–194 (2012).CAS 
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38.Hussa, E. A., Darnell, C. L. & Visick, K. L. RscS functions upstream of SypG to control the syp locus and biofilm formation in Vibrio fischeri. J. Bacteriol. 190, 4576–4583 (2008).CAS 
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39.Mandel, M. J., Wollenberg, M. S., Stabb, E. V., Visick, K. L. & Ruby, E. G. A single regulatory gene is sufficient to alter bacterial host range. Nature 458, 215–218 (2009).CAS 
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40.Ray, V. A., Eddy, J. L., Hussa, E. A., Misale, M. & Visick, K. L. The syp enhancer sequence plays a key role in transcriptional activation by the sigma54-dependent response regulator SypG and in biofilm formation and host colonization by Vibrio fischeri. J. Bacteriol. 195, 5402–5412 (2013).CAS 
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41.Visick, K. L. & Skoufos, L. M. Two-component sensor required for normal symbiotic colonization of Euprymna scolopes by Vibrio fischeri. J. Bacteriol. 183, 835–842 (2001).CAS 
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42.Norsworthy, A. N. & Visick, K. L. Signaling between two interacting sensor kinases promotes biofilms and colonization by a bacterial symbiont. Mol. Microbiol. 96, 233–248 (2015).CAS 
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43.Thompson, C. M., Marsden, A. E., Tischler, A. H., Koo, J. & Visick, K. L. Vibrio fischeri biofilm formation prevented by a trio of regulators. Appl. Environ. Microbiol. https://doi.org/10.1128/AEM.01257-18 (2018).Article 
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44.Brooks, J. F. II & Mandel, M. J. The histidine kinase BinK Is a negative regulator of biofilm formation and squid colonization. J. Bacteriol. 198, 2596–2607 (2016).PubMed 
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45.Pankey, M. S. et al. Host-selected mutations converging on a global regulator drive an adaptive leap by bacteria to symbiosis. eLife https://doi.org/10.7554/eLife.24414 (2017). Evolutionary pathways that can lead to symbiotic colonization are revealed in this elegant study that follows the serial passage of a non-colonizing strain through many E. scolopes juveniles, resulting in altered, symbiosis-competent strains.Article 

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46.Morris, A. R., Darnell, C. L. & Visick, K. L. Inactivation of a novel response regulator is necessary for biofilm formation and host colonization by Vibrio fischeri. Mol. Microbiol. 82, 114–130 (2011).CAS 
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47.Morris, A. R. & Visick, K. L. The response regulator SypE controls biofilm formation and colonization through phosphorylation of the syp-encoded regulator SypA in Vibrio fischeri. Mol. Microbiol. 87, 509–525 (2013).CAS 
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48.Brooks, J. F. II et al. Global discovery of colonization determinants in the squid symbiont Vibrio fischeri. Proc. Natl Acad. Sci. USA 111, 17284–17289 (2014). This large-scale investigation of colonization factors provides important information on genetic requirements for symbiosis and provides a wealth of data for hypothesis generation that will foster many subsequent studies.CAS 
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49.Thompson, C. M. & Visick, K. L. Assessing the function of STAS domain protein SypA in Vibrio fischeri using a comparative analysis. Front. Microbiol. 6, 760 (2015).PubMed 
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50.Rotman, E. R. et al. Natural strain variation reveals diverse biofilm regulation in squid-colonizing Vibrio fischeri. J. Bacteriol. https://doi.org/10.1128/JB.00033-19 (2019).Article 
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51.Bongrand, C. et al. A genomic comparison of 13 symbiotic Vibrio fischeri isolates from the perspective of their host source and colonization behavior. ISME J. 10, 2907–2917 (2016). This study of the genomes and behaviours of a collection of a number of squid symbionts propelled the field from the near-exclusive study of a single isolate, ES114, into new and exciting directions with the genomic sequencing of dominant strains that contain numerous additional genetic sequences and factors.CAS 
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52.Newell, P. D., Boyd, C. D., Sondermann, H. & O’Toole, G. A. A c-di-GMP effector system controls cell adhesion by inside-out signaling and surface protein cleavage. PLoS Biol. 9, e1000587 (2011).CAS 
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53.Christensen, D. G., Marsden, A. E., Hodge-Hanson, K., Essock-Burns, T. & Visick, K. L. LapG mediates biofilm dispersal in Vibrio fischeri by controlling maintenance of the VCBS-containing adhesin LapV. Mol. Microbiol. 114, 742–761 (2020). This article addresses a major long-standing question concerning the initiation of the light organ association; specifically, how do aggregated V. fischeri cells release themselves and migrate into host tissue? One factor may be an adhesin-cleaving protease, which is kept in check by a c-di-GMP-responsive protein, and can promote symbiont dispersal from biofilms.CAS 
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54.Fidopiastis, P. M. et al. Characterization of a Vibrio fischeri aminopeptidase and evidence for its influence on an early stage of squid colonization. J. Bacteriol. 194, 3995–4002 (2012).CAS 
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55.Davidson, S. K., Koropatnick, T. A., Kossmehl, R., Sycuro, L. & McFall-Ngai, M. J. No means ‘yes’ in the squid-vibrio symbiosis: nitric oxide (NO) during the initial stages of a beneficial association. Cellul. Microbiol. 6, 1139–1151 (2004).CAS 
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56.Wang, Y. et al. Vibrio fischeri flavohaemoglobin protects against nitric oxide during initiation of the squid-Vibrio symbiosis. Mol. Microbiol. 78, 903–915 (2010).CAS 
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57.Stabb, E. V. Should they stay or should they go? Nitric oxide and the clash of regulators governing Vibrio fischeri biofilm formation. Mol. Microbiol. 111, 1–5 (2019).CAS 
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58.Thompson, C. M., Tischler, A. H., Tarnowski, D. A., Mandel, M. J. & Visick, K. L. Nitric oxide inhibits biofilm formation by Vibrio fischeri via the nitric oxide sensor HnoX. Mol. Microbiol. 111, 187–203 (2019). This publication provides insight into the complex role in symbiosis of the squid-produced defence molecule NO by uncovering its ability to inhibit biofilm formation via the NO sensor HnoX, a finding that suggests that NO may influence the location or timing of biofilm formation and/or promote dispersal during symbiotic initiation.CAS 
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59.Singh, P., Brooks, J. F. II., Ray, V. A., Mandel, M. J. & Visick, K. L. CysK plays a role in Biofilm formation and colonization by Vibrio fischeri. Appl. Environ. Microbiol. 81, 5223–5234 (2015).CAS 
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60.Raina, J. B., Fernandez, V., Lambert, B., Stocker, R. & Seymour, J. R. The role of microbial motility and chemotaxis in symbiosis. Nat. Rev. Microbiol. 17, 284–294 (2019).CAS 
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63.Millikan, D. S. & Ruby, E. G. FlrA, a sigma54-dependent transcriptional activator in Vibrio fischeri, is required for motility and symbiotic light-organ colonization. J. Bacteriol. 185, 3547–3557 (2003).CAS 
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64.Millikan, D. S. & Ruby, E. G. Vibrio fischeri flagellin A is essential for normal motility and for symbiotic competence during initial squid light organ colonization. J. Bacteriol. 186, 4315–4325 (2004).CAS 
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65.Wolfe, A. J., Millikan, D. S., Campbell, J. M. & Visick, K. L. Vibrio fischeri sigma54 controls motility, biofilm formation, luminescence, and colonization. Appl. Environ. Microbiol. 70, 2520–2524 (2004).CAS 
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71.Nikolakakis, K., Monfils, K., Moriano-Gutierrez, S., Brennan, C. A. & Ruby, E. G. Characterization of the Vibrio fischeri fatty acid chemoreceptors, VfcB and VfcB2. Appl. Environ. Microbiol. 82, 696–704 (2015).PubMed 
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72.Mandel, M. J. et al. Squid-derived chitin oligosaccharides are a chemotactic signal during colonization by Vibrio fischeri. Appl. Environ. Microbiol. 78, 4620–4626 (2012). While it was long-expected that V. fischeri might sense and be attracted to squid-produced molecules to facilitate directed migration into the light organ crypts, this work is the first to identify squid-produced molecules, chitin oligosaccharides, that function as a chemotactic signal promoting colonization.CAS 
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73.Bennett, B. D., Essock-Burns, T. & Ruby, E. G. HbtR, a heterofunctional homolog of the virulence regulator TcpP, facilitates the transition between symbiotic and planktonic lifestyles in Vibrio fischeri. mBio https://doi.org/10.1128/mBio.01624-20 (2020). Comparisons of V. fischeri with the related pathogen Vibrio cholerae reveal that a regulator conserved among Vibrio spp. plays very different roles in the interactions of these two microorganisms with their respective hosts.Article 
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77.Visick, K. L., Foster, J., Doino, J., McFall-Ngai, M. & Ruby, E. G. Vibrio fischeri lux genes play an important role in colonization and development of the host light organ. J. Bacteriol. 182, 4578–4586 (2000).CAS 
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This study demonstrated that several anuran genera (Xenopus, Lithobates [Rana], Hyla, and Anaxyrus [Bufo]), as well as Nile monitor lizards, green anoles, and featherfin catfish, are susceptible to infection with D. insignis and/or D. medinensis L3s. We also found that D. insignis and D. medinensis larvae can persist in anuran tissues for at least eight and two months, respectively, although the number of L3s recovered from each infected animal was generally low. Regardless, these data show that these animals could serve as paratenic hosts if they ingest infected copepods in nature and are subsequently ingested by an appropriate definitive host.We exposed animals using two different methods (group batch or by mouth [PO]), but aimed to primarily batch expose animals as that better mimics natural exposure. A few anuran species (i.e., American toads, Cope’s gray treefrogs, and adult African clawed frogs) were exposed to D. medinensis-infected copepods PO, as they had metamorphosed into adults before D. medinensis larvae became available for use and would be unlikely to ingest all copepods autonomously. Our primary goal in this study was to determine susceptibility to Dracunculus infection.Six anurans that were exposed as tadpoles underwent metamorphosis to froglets before being necropsied. Dracunculus L3s were recovered from two of these animals, supporting previous findings that D. insignis larvae can persist in anuran tissues through metamorphosis14. The persistence of larvae in the tissues through metamorphosis may facilitate Dracunculus transmission from aquatic to terrestrial food chains. This could be an important factor in transmission, as the majority of definitive hosts of Dracunculus nematodes are terrestrial. This study found that, in addition to X. laevis and Lithobates spp. (which have previously been infected with Dracunculus spp. larvae), Anaxyrus sp. and Hyla sp. can also become infected with Dracunculus L3s14,21. The infection of Anaxyrus sp. and Hyla sp. is particularly interesting, as members of these genera transition to a terrestrial or arboreal existence as adults, compared to Xenopus sp. and Lithobates spp. which remain completely or predominantly aquatic, even as adults. This transition to a terrestrial habitat could carry infectious larvae further from water sources, making them available to definitive hosts more widely across the landscape. However, the role of these animals in Dracunculus transmission would still depend on many other factors, including the natural history of these amphibian species, diets of definitive hosts, and how long Dracunculus L3s persist in paratenic hosts, as terrestrial anurans would be unlikely to acquire new infections after metamorphosis.During a previous experimental study, D. insignis L3s persisted in amphibian paratenic hosts for up to 37 DPI, at which time the animals were necropsied16. In this long-term infection trial, we found that D. insignis larvae persisted for at least 244 days (approximately eight months), while D. medinensis larvae persisted for at least 58 days (approximately two months). These results demonstrate that infection of a paratenic host can extend the time that L3s may persist in the environment well beyond the lifespan of a copepod21. As we had a limited supply of D. medinensis L3s, we were unable to conduct sufficient trials to determine whether D. insignis may persist longer in paratenic hosts than D. medinensis. If this difference was found to exist, it could contribute to the higher proportion of wild-caught adult frogs found to be infected with D. insignis than with D. medinensis during field surveys18,19. Further testing with an increased sample size would be required to determine whether the persistence of larvae actually differs between Dracunculus species or paratenic host species.No Dracunculus larvae were recovered from the two adult African clawed frogs that were fed D. medinensis L3s that had been recovered from other paratenic hosts. It is likely that our very small sample size (two animals) and the prolonged period before necropsy (4 months) explain these negative results. In our persistence trials, there was attrition over time so these animals should have been examined earlier after exposure. Future efforts to investigate transmission of Dracunculus between different paratenic hosts should use larger sample sizes and shorter infection periods. It would also be interesting to know if predatory animals, such as Nile monitor lizards, which can experimentally become infected with Dracunculus sp. larvae could become infected by ingesting other paratenic hosts.Fish were investigated for their potential role in Dracunculus transmission, as many fish species consume copepods as part of a natural diet25,26. Despite this, Dracunculus larvae have not been recovered during multiple studies screening wild-caught fish17,19. Dracunculus insignis L3s have rarely been recovered from previous experimental trials with fish16. When larvae were recovered from fish, larval recovery rates were very low (0.6–2.0% recovery; 1–2 larvae per fish) and only 3/43 (7.0%) of the fish harbored Dracunculus larvae upon necropsy16. In a separate trial, fish experimentally functioned as short-term transport hosts of D. medinensis and D. insignis to infect domestic ferrets7. Our findings from this trial were surprising, as we recovered up to 6 D. medinensis L3s from the tissues of three out of four (75%) exposed featherfin catfish. This fish species is common in the Chari River Basin area in Chad, Africa where high numbers of D. medinensis infections are reported in domestic dogs living in fishing villages, and is consumed by both people and dogs17. Dogs in these villages often eat discarded small fish or fish viscera4. Although our sample size was small, our current findings are evidence that some fish species may be more capable of serving as paratenic hosts for Dracunculus than those that have been previously tested. This finding further supports the continuation of the screening of wild fish muscle tissues for Dracunculus larvae.Lizards were included in this study because large, subcutaneous nematodes (believed to be Dracunculus sp.) were historically reported from Nile monitor lizards and these lizards are consumed by people20,22. However, a lack of contemporary reports and recent work in Chad, Africa, determining that large, subcutaneous nematodes recovered from wild Nile monitor lizards were not Dracunculus sp. but actually most similar to Ochoterenella sp., suggest that monitor lizards in this region are not definitive hosts for D. medinensis17. This current study confirms that Nile monitor and green anole lizards could become infected with Dracunculus larvae. As the diet of Nile monitor lizards can include amphibians and fish, were those prey to contain Dracunculus larvae, it is possible that monitors could serve as paratenic hosts, either by ingestion of larvae in fish intestines or in tissues of amphibians or fish, although these modes of transmission to paratenic hosts have not been confirmed23,24. It is unlikely that green anoles would become naturally infected with Dracunculus spp. due to their diet and primarily arboreal habitat; however, their infection demonstrates that multiple, distantly related lizard species are susceptible to experimental infection.Although anoles were exposed to both D. insignis and D. medinensis larvae, it is most likely that the recovered larvae were D. insignis, as only two D. medinensis-infected copepods were administered (in addition to 23 D. insignis-infected copepods). Species identity of these larvae could not be confirmed, however, as Dracunculus larvae can only be identified to species using molecular diagnostic techniques, which would destroy the sample, and these larvae were used in an experimental infection trial after recovery. Exposure of a ferret PO to the four larvae recovered from this anole (as part of a separate study) did not yield an infection, which is unsurprising given the low dose of larvae used. A previous study has shown that as few as 10 Dracunculus larvae may lead to infection of a ferret when administered interperitoneally (IP) (which was a more effective infection route than PO inoculation), therefore, four larvae administered PO would be unlikely to yield infection of a ferret27,28.In all trials, infection occurred only in those animals that were inoculated with or exposed to at least 20 copepods per individual, suggesting an impact of parasite dose-dependent infection probability for Dracunculus infection in paratenic hosts. As copepod infection rate during this study was estimated to be (ge) 25%, it is likely that animals ingesting 20 copepods would consume at least 5 Dracunculus sp. larvae. Previous studies demonstrated that 10 larvae (administered IP) were sufficient to infect a ferret, but that percent recovery was higher with IP infection than PO27,28. It is likely that a similar minimum infectious dose also exists for paratenic hosts and may differ by paratenic host species and mode of infection. Parasite dose-dependent infection probability of Dracunculus spp. merits further investigation, as understanding this relationship could help researchers to more effectively study transmission in the laboratory by performing experimental infection trials with greater reliability.Despite the variable sample sizes and exposure routes in this study, we demonstrated that a wide range of animals (anurans, fish, and lizards) were susceptible to infection with D. insignis and/or D. medinensis L3s. Importantly, one exposed fish species (Synodontis eupterus) was susceptible, opening up further concerns that certain fish species could serve as transport and paratenic hosts of Dracunculus species. Nile monitor lizards and anoles were successfully infected with L3s, demonstrating the first experimental infection of lizards with Dracunculus larvae. Dracunculus larvae remained L3s in the tissues of tested anurans for up to 244 days, extending the known persistence time of infectious larvae. Although no larvae were recovered from frogs that were fed L3s recovered from other paratenic hosts, continued investigation into the possibility of paratenic host to paratenic host transmission would be particularly interesting in determining if some predatory frogs (tadpoles or adults), fish, or lizards may concentrate higher numbers of L3s over time through predation of other infected paratenic hosts. Despite this study not determining how infectious larvae recovered from each of these paratenic hosts would be to another host, our findings contribute to a better understanding of the ability of these paratenic hosts to harbor Dracunculus L3s. This information is valuable to understanding how transmission to animal definitive hosts may be occurring, in addition to informing GWEP management decisions aiming to decrease transmission of D. medinensis to humans and animals. More

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