Hosts of Amblyomma dissimile and Amblyomma rotundatum in the Brazilian Amazon.
\r\n\t
",isbn:"978-1-83962-547-3",printIsbn:"978-1-83962-546-6",pdfIsbn:"978-1-83962-548-0",doi:null,price:0,priceEur:0,priceUsd:0,slug:null,numberOfPages:0,isOpenForSubmission:!0,hash:"e5ba02fedd7c87f0ab66414f3b07de0c",bookSignature:" John P. Tiefenbacher",publishedDate:null,coverURL:"https://cdn.intechopen.com/books/images_new/10765.jpg",keywords:"Managing Urbanization, Managing Development, Managing Resource Use, Drought Management, Flood Management, Water Quality Monitoring, Air Quality Monitoring, Ecological Monitoring, Modeling Extreme Natural Events, Ecological Restoration, Restoring Environmental Flows, Environmental Management Perspectives",numberOfDownloads:null,numberOfWosCitations:0,numberOfCrossrefCitations:null,numberOfDimensionsCitations:null,numberOfTotalCitations:null,isAvailableForWebshopOrdering:!0,dateEndFirstStepPublish:"January 12th 2021",dateEndSecondStepPublish:"February 9th 2021",dateEndThirdStepPublish:"April 10th 2021",dateEndFourthStepPublish:"June 29th 2021",dateEndFifthStepPublish:"August 28th 2021",remainingDaysToSecondStep:"20 days",secondStepPassed:!1,currentStepOfPublishingProcess:2,editedByType:null,kuFlag:!1,biosketch:"A geospatial scholar working at the interface of natural and human systems, collaborating internationally on innovative studies about hazards and environmental challenges. Dr. Tiefenbacher has published more than 200 papers on a diverse array of topics that examine perception and behaviors with regards to the application of pesticides, releases of toxic chemicals, environments of the U.S.-Mexico borderlands, wildlife hazards, and the geography of wine.",coeditorOneBiosketch:null,coeditorTwoBiosketch:null,coeditorThreeBiosketch:null,coeditorFourBiosketch:null,coeditorFiveBiosketch:null,editors:[{id:"73876",title:"Dr.",name:"John P.",middleName:null,surname:"Tiefenbacher",slug:"john-p.-tiefenbacher",fullName:"John P. Tiefenbacher",profilePictureURL:"https://mts.intechopen.com/storage/users/73876/images/system/73876.jfif",biography:"Dr. John P. Tiefenbacher (Ph.D., Rutgers, 1992) is a professor of Geography at Texas State University. His research has focused on various aspects of hazards and environmental management. Dr. Tiefenbacher has published on a diverse array of topics that examine perception and behaviors with regards to the application of pesticides, releases of toxic chemicals, environments of the U.S.-Mexico borderlands, wildlife hazards, and the geography of wine. More recently his work pertains to spatial adaptation to climate change, spatial responses in wine growing regions to climate change, the geographies of viticulture and wine, artificial intelligence and machine learning to predict patterns of natural processes and hazards, historical ethnic enclaves in American cities and regions, and environmental adaptations of 19th century European immigrants to North America's landscapes.",institutionString:"Texas State University",position:null,outsideEditionCount:0,totalCites:0,totalAuthoredChapters:"1",totalChapterViews:"0",totalEditedBooks:"6",institution:{name:"Texas State University",institutionURL:null,country:{name:"United States of America"}}}],coeditorOne:null,coeditorTwo:null,coeditorThree:null,coeditorFour:null,coeditorFive:null,topics:[{id:"12",title:"Environmental Sciences",slug:"environmental-sciences"}],chapters:null,productType:{id:"1",title:"Edited Volume",chapterContentType:"chapter",authoredCaption:"Edited by"},personalPublishingAssistant:{id:"194667",firstName:"Marijana",lastName:"Francetic",middleName:null,title:"Ms.",imageUrl:"https://mts.intechopen.com/storage/users/194667/images/4752_n.jpg",email:"marijana@intechopen.com",biography:"As an Author Service Manager my responsibilities include monitoring and facilitating all publishing activities for authors and editors. From chapter submission and review, to approval and revision, copyediting and design, until final publication, I work closely with authors and editors to ensure a simple and easy publishing process. I maintain constant and effective communication with authors, editors and reviewers, which allows for a level of personal support that enables contributors to fully commit and concentrate on the chapters they are writing, editing, or reviewing. I assist authors in the preparation of their full chapter submissions and track important deadlines and ensure they are met. I help to coordinate internal processes such as linguistic review, and monitor the technical aspects of the process. As an ASM I am also involved in the acquisition of editors. 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Venkateswarlu",coverURL:"https://cdn.intechopen.com/books/images_new/371.jpg",editedByType:"Edited by",editors:[{id:"58592",title:"Dr.",name:"Arun",surname:"Shanker",slug:"arun-shanker",fullName:"Arun Shanker"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}},{type:"book",id:"878",title:"Phytochemicals",subtitle:"A Global Perspective of Their Role in Nutrition and Health",isOpenForSubmission:!1,hash:"ec77671f63975ef2d16192897deb6835",slug:"phytochemicals-a-global-perspective-of-their-role-in-nutrition-and-health",bookSignature:"Venketeshwer Rao",coverURL:"https://cdn.intechopen.com/books/images_new/878.jpg",editedByType:"Edited by",editors:[{id:"82663",title:"Dr.",name:"Venketeshwer",surname:"Rao",slug:"venketeshwer-rao",fullName:"Venketeshwer Rao"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}},{type:"book",id:"4816",title:"Face Recognition",subtitle:null,isOpenForSubmission:!1,hash:"146063b5359146b7718ea86bad47c8eb",slug:"face_recognition",bookSignature:"Kresimir Delac and Mislav Grgic",coverURL:"https://cdn.intechopen.com/books/images_new/4816.jpg",editedByType:"Edited by",editors:[{id:"528",title:"Dr.",name:"Kresimir",surname:"Delac",slug:"kresimir-delac",fullName:"Kresimir Delac"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}},{type:"book",id:"3621",title:"Silver Nanoparticles",subtitle:null,isOpenForSubmission:!1,hash:null,slug:"silver-nanoparticles",bookSignature:"David Pozo Perez",coverURL:"https://cdn.intechopen.com/books/images_new/3621.jpg",editedByType:"Edited by",editors:[{id:"6667",title:"Dr.",name:"David",surname:"Pozo",slug:"david-pozo",fullName:"David Pozo"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}}]},chapter:{item:{type:"chapter",id:"57528",title:"Challenging the Paradigms on the Origin, Specification and Development of the Female Germ Line in Placental Mammals",doi:"10.5772/intechopen.71559",slug:"challenging-the-paradigms-on-the-origin-specification-and-development-of-the-female-germ-line-in-pla",body:'\nGerm cells are the only cell types capable of transmitting the genetic traits of an individual. They differentiate into spermatozoa and oocytes in adult testis and ovary, respectively, and give rise to a totipotent zygote after fertilization. Germ cells guarantee the perpetuation and diversification of the genetic information across the generations in most multicellular organisms. The developmental pathways that lead to the formation of a highly specialized germ cell are long and complicated, and the molecules that are involved in this process are still a matter of discussion. One extraordinary feature in the germ cell lineage in mammals is the fact that specification occurs far from the gonads, implying a necessary migratory phase after specification. A second feature is their unique capacity to undergo meiosis, in which chromosome recombination generates genetic variation in the haploid gametes [1, 2, 3, 4].
\nMost of our understanding regarding germ cell specification and differentiation in mammals comes from studies in the laboratory mouse. It is widely accepted that specification of primordial germ cells (PGCs) in mouse takes place at a very early stage in development; more precisely, they are thought to be set apart following blastocyst implantation in the proximal epiblast of the gastrulating embryo [2]. So far, however, no lineage tracing study has shown that those early segregated PGCs finally end up in the gonads [5]. Alternatively, it has been proposed that presumptive early specified PGCs in the proximal epiblast are rather a primordial pool of stem cells from which PGCs can be specified later on in development, probably during migration toward the emerging gonad [5]. Both explanations have been raised from mouse embryo studies. Nevertheless, there are some key embryological differences between the mouse and other mammals, especially at the epiblast stage when PGCs are specified. The epiblast of the murine rodent forms a cup-shaped egg cylinder, but most other mammals have a flat disk-like epiblast. Signals from extraembryonic tissues induce germ cell fate in a subset of epiblast cell at a specific position with optimal concentration and timing of signals. As PGC specification largely depends on signals from surrounding tissues, the morphology of the embryo is crucial for dissecting out the mechanisms of germ line establishment in different mammals since tissues surrounding the epiblast in the egg cylinder are not the same in flat-disk embryos [6].
\nIn mouse, the blastocyst implants in the uterus by E4.5. The inner cell mass (ICM) of blastocyst is the source of epiblast cells. The ICM is segregated into epiblast and hypoblast or the primitive endoderm. Epiblast cells are equipotent and give rise to all the somatic and germ cells. During implantation, when the syncytiotrophoblast starts to penetrate the wall of the uterus, the epiblast and hypoblast are physically constrained and form a bilaminar embryo. The internal epiblast cells reorganize from a ball of cells into a cup-shaped epithelium surrounded by hypoblast. Immediately before gastrulation (E6.0 and E6.5), the mouse embryo can be visualized as a thick-walled cup of tissue (the epiblast or embryonic ectoderm), which gives rise to the entire fetus and some of the placental membranes. A second thick-walled cup of tissue (the extraembryonic ectoderm, ExE) placed overturned on the epiblast will give rise to the main part of the placenta. Both cups are enclosed in a thin bag of primitive endoderm-derived visceral endoderm (VE) [2, 7, 8].
\nThe embryonic disk is forced into a complex shape called the ‘egg cylinder’ in which the anterior and posterior poles of the embryo come in close proximity to each other. Around E4.5 and E5.5, the ExE arises from the polar trophoectoderm (TE) and makes contact with the underlying epiblast. At E6.5, gastrulation starts with the formation of the primitive streak at the posterior region of the embryo. At E7.5, epiblast cells migrating first through this structure include the PGC precursors, which form the extraembryonic mesoderm [9, 10].
\nIn mouse, PGCs originate from the most proximal epiblast cells by induction of the ExE and VE. Both extraembryonic tissues surround the epiblast cell of the postimplantation egg cylinder at around E5.0–E6.0. The ExE and VE release the bone morphogenetic protein (BMP) 4, 8b and 2 to instruct a small number of pluripotent proximal epiblast cells to become competent to be PGCs, suppressing a somatic program that is adopted by neighboring cells [11] (Figure 1).
\nSchematic comparison of the early-implanted embryo in the egg cylinder of mouse and the flat disk embryo of human, rabbit and vizcacha. No extraembryonic ectoderm (ExE) is found in human/rabbit embryo; in the vizcacha, it locates far apart from the epiblast to be responsible for inducing germ cell specification as in mouse. The black circle indicates the presumptive location from which PGCs originate. PGCs: primordial germ cells, pPGC: presumptive PGC, VE: visceral endoderm.
Accordingly, BMP4 released from the ExE activates the expression of B-lymphocyte–induced maturation protein 1 (Blimp1), also known as PR domain-containing protein 1 (Prdm1), at ~E6.25 and PR domain-containing protein 14 (Prdm14) at ~E6.5 in a dose-dependent manner [12, 13, 14, 15, 16]. Bmp2 expressed in the proximal VE enhances the same signaling pathway, ensuring that the highest levels of Bmp signaling occur in the most proximal epiblast [11, 13, 16]. Both Blimp1 and Prdm14 together with Tcfap2c (also known as Ap2γ) [17, 18] are required for PGC specification.
\nBlimp1 protein signal first appears in about 6 cells in the most proximal epiblast at the posterior side of the embryo. Blimp1+ cells initially express the Hox genes as well as many other genes known to be involved in embryonic development and suppress the expression of genes associated with pluripotency, such as Sox2, Nanog, and Zic3. At around E6.75–E7.0, Hox genes are downregulated and Blimp1+ cells regain the expression of pluripotency genes [13, 14, 15]. Therefore, the PGC precursors appear to be initially induced toward a somatic mesodermal fate, but then regain their potentially pluripotent nature.
\nFollowing lineage restriction, PGC precursors initiate germ cell specification by activating Prdm14 and Tcfap2c [15]. Prdm14 plays crucial roles in two successive events characterizing the germ cell program: reacquisition of pluripotent potential and epigenetic reprogramming [12, 18]. Prdm14 is first expressed in Blimp1+ cells and later on in PGCs. Although the activation of Prdm14 is independent of Blimp1, the expression of Tcfap2c at ~E6.75 appears to be dependent on Blimp1 [12]. Indeed, when these factors are coexpressed they can induce PGC-like cells in the absence of cytokines, suggesting that the tripartite gene network Blimp1/Prdm14 /Tcfap2c is sufficient for mouse PGC specification [18]. The first two genes acting in the founder PGC population are Fragilis and Stella. Fragilis (also known as mouse interferon-induced protein like gene-1 [mil-1]/interferon-induced transmembrane protein 3 [Ifitm3]) [19] expression marks the beginning of germ cell competence and it starts expressing at ~E6.25–E6.5 before Blimp1 expression. Fragilis expression intensifies in the posterior extraembryonic mesoderm at ~E7.0–E7.25. In fact, the high levels of BMPs activate the expression of Fragilis and competent cells acquire the ability to form PGCs when they begin to express Blimp1.
\nStella (also known as primordial germ cell 7 [Pgc7]/developmental pluripotency-associated 3 [Dppa3]) [20] begins to express specifically in Fragilis-Blimp1 expressing cells in the extraembryonic mesoderm at ~E7.0–E7.25 and continues to be expressed in migrating PGCs. The function of Stella gene product is uncertain, but it has domains characteristic of proteins involved in RNA splicing. Actually, Stella represses homeobox genes in the nascent germ cell and as such maintains the pluripotency of PGCs during their migration toward the genital ridge. However, gene-knockout studies revealed that neither Fragilis nor Stella is essential for PGC specification [21, 22].
\nBlimp+ PGC precursors proliferate and move into the extraembryonic mesoderm (ExM); they reexpress pluripotency-associated genes (Oct4, Nanog, Sox2 and Klf2) and Stella at around E7–E7.75. While Klf2 and Stella are apparently dispensable proteins for PGC development [2, 23], the three core pluripotency factors Oct4, Nanog and Sox2 are essential for PGC development [24, 25, 26]. Although the specific roles of these factors during germ cell development are unclear, it is thought that their expression confers latent pluripotency to the germ line. In the case of PGCs, this regulatory network is thought to protect them from somatic-inducing signals during the extensive epigenetic reprogramming they undergo [27]. Oct4 is uniformly expressed in postimplantation epiblast and also in nascent PGCs during specification. Oct4 expression remains high until germ cells undergo sexual differentiation in the gonad [28, 29]. It is apparently essential for both germ cell specification [30] and maintenance [31]. Nanog is enriched at the proximal posterior epiblast, the position where PGCs are specified from, in E6.5 and E7.5 embryos [32]. Nanog appears to be dispensable for mouse PGC specification but is essential for germ cell maintenance. Sox2 is active in mouse PGC from E7.5 forward. Conditional knockout of Sox2 shortly after specification caused a dramatic decrease of germ cell numbers by E7.5, being undetectable by E13.5 [32].
\nWith the establishment of germ cell fate, germ cells express factors like alkaline phosphatase (AP), Nanos3, Dazl, mouse vasa homologue (Mvh) and Dnd1 [2]. They increase in number and move out of the embryo by the primitive streak in formation toward the extraembryonic mesoderm at the base of the allantois at E7.25. As mentioned above, PGCs form a cluster of cells 6 to 16 cells at around E6.5; then, they increase to approximately 20–28 cells, move posteriorly and develop into PGCs at E6.75-E7. During early gastrulation, the PGCs form a cluster of 40–50 cells at the base of the incipient allantois in the ExM at around E7.25 [33, 34]. Subsequently, and concomitant with an increase in their number, at around E8, they start to translocate one by one toward the developing hindgut endoderm and move through it. They then leave the endoderm to emerge in the mesentery and at around E10.5 colonize the embryonic gonads, where they proliferate and initiate a differentiation into either oocytes or spermatozoa depending on the embryo sex.
\nPGCs in the mouse may be motile from their onset (E7.25) until they colonize the genital ridge (E11.5). After formation, PGCs move through the posterior primitive streak and invade the definitive endoderm and posterior extraembryonic structures. Following subsequent migration within the hindgut during its anterior extension (E8-E9.5), mouse PGCs migrate through the hindgut tissue to the mesoderm, followed by bilateral migration toward the gonadal ridges (E10.5–11.5). During this pregonadic phase, PGCs can be identified by morphological criteria and surface markers, such TNAP and SSEA-1, and the expression of pluripotent markers like Oct4, Sox2 or Nanog [35].
\nSix distinct stages of PGC behavior in the migratory process were identified, including: (i) invasion of the endoderm, (ii) passive or active migration into the hindgut, (iii) random migration within the hindgut, (iv) migration from the gut to the genital ridges, (v) clustering at the ridges and (vi) cell death within midline structures [36].
\nAt E7.5, PGCs move through the primitive streak and into the definitive endoderm. Some PGCs also end up in the allantois and/or parietal endoderm. The fate of PGCs in extraembryonic structures remains uncertain, but PGCs in the definitive endoderm become incorporated into the hindgut, and at E9.0, they can be found moving within and around the cells of the hindgut epithelium [36]. At E8.5, PGCs on the lip of the hindgut pocket have a rounded nonmotile morphology suggesting that PGCs are passively incorporated into the gut and then, at stage (iii), they reinitiate active motility around the epithelial cells.
\nInteractions between PGCs may also be important for their homing behavior. PGCs emerge from the gut individually, but during migration, they interact with each other forming a migrating network of cells [37]. This network becomes progressively aggregated into clusters of cells toward the end of migration. Antibodies against E-cadherin blocked the process of PGC aggregation in cultured embryo slices and prevented PGCs from forming tight clusters at the genital ridges [38].
\nAt the end of their migration, PGCs presumably lose their motile properties as they associate with somatic cells in the gonad and acquire sex-specific morphologies. There does not seem to be any evidence for sex-specific differences during germ cell migration.
\nThe mouse pathway described above is the classical currently accepted model of PGC formation. This path establishes that PGCs originate and specify as an early lineage-restricted cluster of cells in the base of the allantois soon after implantation. Nevertheless, no definitive proof demonstrating the continuity of those presumptive early-specified PGC and the germ cells, which colonize the genital ridge, has so far been provided. In view of this, and critically reviewing the literature on PGC origin and specification in mouse, Mikedis and Downs [5] advocate in favor of an alternative hypothesis. These authors propose an alternative model in which the presumptive PGCs in the base of the allantois are instead a pool of pluripotent progenitor cells in the posterior end of the primitive streak that builds up the fetal-placental interface. The pluripotent cell pool condenses into a specific area of the proximal epiblast, namely the allantoic core domain (ACD), which extends the body axis posteriorly through the allantoic midline. The pluripotent cells in the ACD express all PGC markers and contribute to both embryonic and extraembryonic tissues. From this pluripotent population, it is suggested that PGC could be segregated later. PGC specification could take place for example during migration toward the genital ridge once evolutionarily conserved genes of germ line development, such as VASA, Dazl and Nanos, begin to be expressed. Although this alternative explanation is proposed for the mouse egg cylinder, it may well apply in flat embryos where the ExE is absent or far apart from the epiblast.
\nThe embryo proper of most gastrulation-stage mammals, including humans, rabbits and pigs among others, has the shape of a flat disk with two cell layers: epiblast and hypoblast (equivalent to VE in mice) [39, 40, 41]. In the flat disk of non-rodent embryos, the epiblast contacts with the VE (hypoblast), and the ExE is absent. In basal rodents of the suborder Hystricognathi such as the guinea pig (Cavia porcellus) and the vizcacha (Lagostomus maximus), the ExE persists in the flat-disk embryo, but it remains far apart from the epiblast at the time of PGC induction [42, 43]. Moreover, murine PGC nest in the growing mesodermal allantois in the proximal/posterior region of the embryo is a precocious structure found in the mouse, but not seen in nonrodent mammals. These differences have a critical effect on PGC specification factors [4].
\nDue to ethical and technical reasons, there is limited information on the origin of human PGCs in postimplantation embryos. PGCs have been described in human embryos at early somite stage in the dorsal wall of the yolk sac near the developing allantois [44, 45, 46]. Decades later, AP activity in presumably PGCs was observed by several groups in human embryos with 5–8 somites at a similar location. Using single cell analysis, human PGCs isolated at 4 weeks of development seem to express PRDM14 and TFAP2C, whereas BLIMP1 and FRAGILIS are not expressed [48], suggesting that the critical molecular network in mouse and human early PGCs is divergent. A recent report showed that BLIMP1 is activated in human PGC-like (hPGCL) cells after specification by SOX17, and it is suggested that its role is to inhibit the potential for somatic differentiation [47, 48]. SOX17 is the earliest marker of hPGCL cells and it is in fact the key regulator of their fate, which is not the case in mice. BLIMP1 is downstream of SOX17, and it represses endodermal and other somatic genes. Furthermore, hPGCL cells arise from precursors expressing high levels of T and low levels of SOX2, resembling posterior primitive streak-derived progenitors [32]. This suggests that human germ cell precursors may arise from a population of posterior primitive streak-derived cells that activate BLIMP1 in response to paracrine signals, a process that occurs during postgastrulation (later than mice) [34]. The precise combination of signals that promote germ line segregation in humans is currently unknown; however, recent studies in emerging models as cynomolgus macaque, together with in vitro studies in human PGC-like cells derived from induced pluripotent stem cells, reveal a different molecular pathway from that evolved in mouse [46].
\nIn pregastrulation rabbit embryos, BMP2 is first expressed from the hypoblast and yolk sac epithelium at the boundary of the embryonic disk, which is equivalent to the proximal VE and extraembryonic VE in mice, respectively. In turn, rabbit BMP4 expression is significantly delayed compared to the mouse. BMP4 marker is first detected during primitive streak formation and it is expressed peripherally in intraembryonic hypoblast and epiblast and in the mesoderm at the posterior pole of the embryonic disk. Interestingly, BLIMP1+ single PGC precursors are detected before primitive streak formation and BLIMP1 mRNA distribution closely follows the expression pattern of BMP2. Thus, it is proposed that BMP2 may play a more essential role in rabbit PGC specification than BMP4 [49].
\nOn the other hand, PG-2 (a germ cell epitope) and BLIMP1-expressing cells have been localized at early gastrulation stage in a region identified in the posterior upper layer (epiblast) and mesoderm [50]. However, BLIMP1 shows a wider expression pattern during these developmental stages, with positive cells in the hypoblast all around the circumference of the embryo, adjacent to the site of BMP4 expression in the extraembryonic cells surrounding the embryo. Nevertheless, from these ‘blimped’ pPGCs, only the posterior ones seem to become PG-2-positive [50].
\nA recent study in the basal Hystricognathi rodent Lagostomus maximus, which develops through a flat-disk epiblast far apart from the ExE (Figure 1), showed that OCT4 protein seems to play an essential role in the establishment and maintenance of the germ line [43]. OCT4 expression in the pregastrulating embryo was observed across all the epiblast cells, but after the primitive streak stage, OCT4 was mostly downregulated, and its expression only persisted in a group of cells that was later restricted to the mesoderm of the posterior end of the embryo. It seems likely that OCT4 expression is required for maintaining pluripotency, helping to epigenetically reprogram cells for PGC development that will be specified at a later stage, probably suppressing expression of genes involved in mesodermal specification [43]. In this model, BLIMP1 expression has not been detected during early gastrulation or later stages of development (migration and colonization of the genital ridges). It seems likely that BLIMP1 would not be necessary for the specification of the germ line in the basal rodent L. maximus.
\nIn an advanced stage of development, at neural plate stage, in the base of the allantois in the ectoderm and mesoderm after gastrulation, OCT4+ cells become restricted in number to a group of 6–8 cells, and they begin to express SOX17, STELLA and FRAGILIS. The temporal colocalization of SOX17 and OCT4 proteins in L. maximus seems to play a major role in inhibiting somatic genes and maintaining pluripotency instead of the mouse alternative SOX2/OCT4 [51]. During migration through the gut, SOX17 is downregulated, and its expression is restored in the oogonia after the colonization of the genital ridges.
\nThen, in the early- and late-head fold stages in mesoderm and endoderm tissues, the expression of OCT4 and SOX17 continues but FRAGILIS and STELLA are downregulated and turned on again during migration. Another notable protein, the germ line marker VASA, was observed early during the translocation of OCT4+ cells to the hindgut. Thereafter, VASA-expressing cells were detected throughout the migration toward the genital ridges. OCT4+/VASA+ cells sequentially turned on STELLA and FRAGILIS during migration. Leopardo and Vitullo [43] suggested that OCT4/STELLA/FRAGILIS-expressing cells are finally restricted and specified to form PGCs during migration when the evolutionarily conserved germ line marker VASA is expressed.
\nThe spatiotemporal pattern of expression of germ line markers found in L. maximus diverges from the currently accepted model on the origin of PGCs as a lineage-restricted cluster of cells in the base of the allantois, specified early just before, or during, gastrulation. In contrast, in this rodent, specification of germ cells seems to occur during migration of a stem cell pool derived from a pluripotent progenitor population within the embryonic axis as proposed by Mikedis and Downs as an alternative pattern of the classical mouse model [5].
\nThe number of PGCs that colonize the genital ridges depends on the species. In mice, beginning with 100–145 PGCs at 8 days postconception (dpc), the number increases exponentially up to 15,000–20,000 oogonia per ovary at 15.5 dpc, the time of entry into meiosis and cessation of mitosis [33, 52, 53, 54, 55, 56]. A similar pattern of germ cell proliferation was described in rats [57]. In the basal rodent L. maximus, approximately 1000 PGCs are detected by the end of migration, rapidly increasing to 55,000 oogonia once fetal ovary colonization is finished; the number continues to increase to reach more than 3×106 germ cells by the end of gestation [43, 58]. The limited human data suggest that 1000–2000 colonizing PGCs reach a maximum of approximately 5–6×106 germ cells per ovary at 20 weeks of gestation [59, 60, 61, 62].
\nAfter a few rounds of mitosis, colonizing PGCs, now referred to as oogonia, cease proliferation and enter a premeiotic phase, with downregulation of pluripotency-associated genes such as Oct4 and Lin28 and upregulation of meiotic genes such as Scp3 [63, 64]. Oogonia entering meiosis, now called oocytes, undergo prophase of the first meiotic division. Just before or early after birth, depending on the species, oocytes in diplotene stage of meiotic prophase I enter a quiescent state known as dictyate, in which they remain arrested, sometimes for years or decades, until just before ovulation [65, 66].
\nIn mice, entry into meiosis seems to be a synchronized event, with no overlapping between mitosis and meiosis. By 17 dpc, mitotic proliferation is finished and all germ cells initiate meiosis [65] entering meiotic prophase in a wave from the anterior to the posterior end of the ovary [64]. However, there is a marked asynchrony of germ cell development in the human ovary. The onset of meiosis occurs by week 11 of gestation [55], but mitosis continues in more peripherally located germ cells for many weeks thereafter, even when primordial follicles begin to form [64, 67]. In the rat, non overlapping mitosis and meiosis of germ cells occurs as in the mouse [68]. However, the basal rodent L. maximus shows asynchrony and overlapping of mitotic and meiotic phases of germ cells in a comparable way as humans [58].
\nThe persistence of PGCs or oogonia in the postnatal ovary has been a matter of discussion throughout the twentieth century since Pearl and Schoppe [69] proposed, in 1921, that postnatal oogenesis might occur in the mammalian adult ovary. Three decades later, in an extensive review of the literature of the time, Zuckerman [70] advocated for the absence of oocyte renewal in the mature mammalian ovary, proposing that mammals are born with a finite nonrenewable oocyte pool, a perspective that was widely accepted for more than 50 years generating a useful framework in advancing our knowledge of ovarian dynamics in placental mammals. Nevertheless, this long-held dogma was challenged in 2004 by Tilly’s team [71] with the description of a small population of germ line stem cells in the adult ovary of the laboratory mouse. This observation refueled the possibility that neo-oogenesis could take place in the adult ovary of mammals and evidence for and against this possibility has accumulated over the recent years [72]. Although it has not been proved yet that ovarian stem cells may contribute to replenishment of the adult ovary if needed, the persistence of germ line stem cells has been independently proven in the human, mouse and rat models, as well as their ability to be manipulated in vitro, and to give rise to offspring following transplantation [73, 74, 75].
\nDeath is a prominent feature of mammalian germ line development, with a predictable temporal and spatial pattern. In fetal life, direct germ cell depletion occurs by means of a constitutive massive germ cell death program, referred to as attrition [59, 60, 76, 77, 78, 79]. In adult life, germ cell demise is mainly the result of death of the supporting follicular cells, a process known as follicular atresia [64, 76, 77, 78].The main mechanism underlying germ cell attrition and follicular atresia requires the activation of a conserved intracellular program of cell death called apoptosis. The execution of the apoptotic program depends on the coordinated action of a group of genes that will activate as a signaling cascade in response to different stimuli. Depending on the source and type of the stimuli, apoptosis can be initiated through an extrinsic pathway, also referred to as the death receptor pathway, which includes the recognition of death ligands to their cell surface receptors [80] or the intrinsic or mitochondrial pathway, which is mainly regulated through the BCL2 protein family whose members are divided into three groups: proapoptotic proteins, antiapoptotic (or prosurvival) proteins and pore-forming proteins [81]. Extrinsic apoptosis molecules are mainly involved in final follicular regression and atresia and corpus luteum regression [82, 83]. BCL2 gene family executing the intrinsic apoptosis path plays an essential role in the death of the germ cell proper in the antenatal ovary and of granulosa cells during follicular atresia in the adult ovary [84].
\nThe analysis of the spatial and temporal expression of members belonging to the BCL2 gene family in the mammalian ovary showed that, in general, the expression of proapoptotic genes is continuous throughout prenatal oogenesis, whereas antiapoptotic members are expressed in a time-restricted pattern associated mainly to differentiation and proliferation of the germ cell [85, 86, 87, 88, 89, 90]. The enhanced expression of proapoptotic genes such as BAX in the face of antiapoptotic members like BCL2 gives support to the high rate of apoptosis characterizing the mammalian ovary. The involvement of this biased gene balance in determining death or survival of the germ cell has been experimentally supported by showing that Bcl2- and Bax-knockout mice have decreased or increased primordial follicle reserve, respectively [91, 92].
\nThe causes that determine massive constitutive death of mammalian female germ cells are poorly understood. This massive elimination may avoid the persistence in the ovary of germ cells exhibiting nuclear or mitochondrial chromosomal/genetic defects [93]. Alternatively, death may relate to the exhaustion of germ cells acting as nurse cells to the surviving oocyte pool [94]. Finally, it has been suggested that massive death may enable the appropriate association between germ cells and pregranulosa cells during ovigerous cords or ovarian cyst breakdown, just before primordial follicles begin to form [95]. In any case, the balance between germ cell death and survival seems to be critical to preclude ovarian dysgenesis or premature ovarian failure and to ensure reproductive success.
\nGerm cell elimination occurs at different points of fetal development. There are three main waves of germ cell death: (i) at prophase and metaphase of proliferating oogonia, (ii) at pachytene of meiotic prophase I oocytes and (iii) at diplotene of meiotic prophase I oocytes [57, 59, 96]. The vast majority of germ cell death occurs during the second and the third waves. Thus, germ cells entering meiosis are particularly susceptible to cell death [55, 60].
\nIn mice, the maximum number of germ cells is registered at the time of entry of primary oocytes into meiotic prophase. However, up to two-thirds of the germ cells are lost before the ovarian reserve is established just after birth [64, 97, 98]. In rats, germ cells proliferate to reach a peak of 64,000 oogonia at 17.5 dpc, but the number of oocytes falls down to about 39,000 at birth and 19,000 at 2 dpc [57, 96]. Humans display a similar dynamics of germ-cell elimination. After the germ cell peak number of 5–6×106 oocytes that occurs at 5 months postconception, there is a dramatic decline in germ cell numbers similar to that seen in mice and rats. By the time of birth, the number of germ cells drops dramatically to 1–2×106 [59, 61, 64, 96] (Figure 2).
\nGerm cell growth curves in mammalian species. Human, mouse and rat share the same growth pattern with maximum germ cell endowment at approximately mid-gestation followed by a massive decline through intraovarian cell death. On the contrary, the vizcacha shows a continuous increase of germ cell population, unaffected by cell death.
Moreover, the process of germ-cell apoptosis continues during postnatal life through follicular atresia. In humans, only 300,000 oocytes survive at 7 years postpartum and fewer than 1000 are present in the years just prior to menopause [59, 61, 96].
\nOnce PGCs have colonized the fetal gonad, the final endowment that will constitute the oocyte reserve seems to depend largely on the balance between cell proliferation and death. Based on the results of germ cell death displayed by mouse, rat and human, it has been widely accepted that massive intraovarian elimination of germ cells is a constitutive attribute of mammalian ovary for the final establishment of the germinal reserve. After a period of high proliferation of colonizing PGCs to reach the maximal oocyte endowment of the species, the activation of the apoptotic pathway generates a point of inflection in the growth curve of the oocyte population that eliminates from 60 to 85% of newly formed oocytes depending on the species [54, 59, 97] (Figure 2). The comparable pattern following the elimination of germ cells quantified in mouse, rat and human, together with the recognition that apoptosis in fetal ovary is active in a few other mammals, proved sufficient to establish massive elimination as a general rule controlling the final oocyte endowment of the ovary in placental mammals.
\nChallenging this established rule, a quantitative estimate based on unbiased stereological methods showed that the mean germ cell number per ovary increases continuously from the early-developing fetal ovary up to 45–60 days after birth in the South American plains vizcacha, L. maximus [58] (Figure 2). Female vizcacha displays a constitutive ovary-specific overexpression of the antiapoptotic BCL2 gene and low to absent expression of proapoptotic BAX gene that leads to a strong suppression of apoptosis-dependent germ cell attrition throughout fetal development [99] and apoptosis-dependent follicular atresia throughout adult life [100]. The detection of germ cells undergoing last steps of apoptosis revealed by TUNEL assay never surpasses 4% of the entire germinal population. Hence, the healthy germ cell population increases continuously from early-developing ovary reaching a 50 times higher population number by the end of gestation. Beginning with an endowment of around 56,000 oogonia at 50 dpc, total germ cell number grows up approximately to 3×106 by the end of gestation [58].
\nWhether the vizcacha is just the exception that confirms the rule or it represents another strategy for establishing the germ cell endowment in mammals, we will have to wait for quantitative studies in a more representative number of placental mammals. Until then, the vizcacha is the first mammal so far described in which female germ line develops in the absence of constitutive massive germ cell elimination since the balance between pro- and anti-apoptotic BCL2 genes is biased in favor of suppressing apoptosis.
\nOur current knowledge regarding the origin and specification of PGCs and the establishment of the ovarian reserve in placental mammals comes by and large from model organisms, notably the mouse. The mouse model has erected as the paradigm for germ line development; however, studies in a few other species unveil differences that challenge the mouse gene network as an established path that may apply to all mammals.
\nThe molecular pathway disclosed for the mouse embryo in the last fifteen years still lacks a final proof showing that the presumptive PGCs, originating early in the proximal epiblast of the egg cylinder, are the same cells that finally colonize the genital ridge later on development. Until this could be traced, alternative hypothesis proposing that PGCs may specify just before colonization from a migrating pluripotent cell population when evolutionarily conserved genes begin to express cannot be ruled out.
\nThe peculiar morphology of the early-implanted mouse embryo, the egg cylinder, sets aside from most mammals that develop through a flat disk embryo. Hence, it is reasonable to suppose that the topographical difference of the gastrulating flat embryo may create a different morphological scenario for signaling and specification of PGCs. The current knowledge in flat embryos, such as those of human and vizcacha, supports a divergent molecular path from that of mouse.
\nOnce the fetal gonad has been colonized by PGCs, it is widely accepted that a balance between proliferation and cell death determines the final oocyte reserve. Massive germ cell death is regarded as an intrinsic shared mechanism in the mammalian ovary regulating the establishment of the final oocyte pool. Nevertheless, only four species have been quantified at the moment and one of these four shows a continuous growth of the germinal population with a minimum cell death. If this is an exception to a general rule or an alternative strategy for establishing the oocyte pool remains unanswered for now.
\nAt this time, we are still far from having a comprehensive knowledge on the possible variety of mechanisms regulating the origin and specification of PGCs and the establishment of the final oocyte reserve in placental mammals. The few species investigated so far seem to indicate that strategies that remain hidden in the great diversity of mammals have not yet been revealed. Comparative studies from different mammalian orders are still lacking and needed.
\nThis study was supported by intramural grant program from Universidad Maimónides-Fundación Científica Felipe Fiorellino and PICT-1281-2014 granted to ADV from the Agencia Nacional de Promoción Científica y Tecnológica, Ministerio de Ciencia, Tecnología e Innovación Productiva, Argentina.
\nPGCs | primordial germ cells |
ICM | inner cell mass |
ExE | Extraembryonic ectoderm |
VE | visceral endoderm |
TE | trophoectoderm |
ExM | extraembryonic mesoderm |
ACD | allantoic core domain |
hPGCL | human primordial germ cell-like |
pPGC | preprimordial germ cell |
The Amazon or Amazon Rainforest is the largest remnant of tropical forest in the world, occupying a region of approximately 6.7 million/km2, covering nine countries in South America: Ecuador (≅ 2%), Suriname (≅ 2%), Bolivia (≅ 4%), Venezuela (≅ 4%), Guyana (≅ 3%), French Guiana (≅ 2%), Colombia (≅ 10%), Peru (≅ 13%), and Brazil (≅ 60%) (Figure 1) [1, 2]. In Brazil, the Amazon biome, also known as “Legal Amazon” occupies approximately 49% of its territory, covering the states of Acre, Amapá, Amazonas, Mato Grosso, Pará, Rondônia, Roraima, Tocantins and Maranhão (Figure 1). The Brazilian Amazon is known for its high richness of landscapes composed of 23 ecoregions, whose main domain is the humid rainforest (≅ 78%). Due to this heterogeneity, the Amazon has an immeasurable amount of essential habitats for the maintenance of flora and fauna, represented with high biological diversity, which is regarded as the largest in the world [3, 4] The Amazon has approximately 45,000 species of flora (39,474 species) and fauna (5,526 species) [2, 5]. However, even with this expressive diversity and the numerous faunal studies carried out in the region, there are still many gaps in the biological diversity of the Amazon. Due to its enormous extension and high degree of preservation (e.g., unexplored areas), new bioecological associations and new species are discovered every year. In the past 20 years, more than 1,200 new species have been described in the Amazon region from which we can highlight 16 birds, 39 mammals, 55 reptiles and ≅ 100 amphibians [2]. However, anthropic action has negatively impacted the animal-forest relationships for decades, resulting in the extinction of ecologically demanding species and, at the same time, the appearance of opportunistic and/or generalist species [6].
\nDiversity of reptiles, amphibians, birds and mammals in Brazil, including the Brazilian Amazon.
Many domestic and wild animals are responsible for the maintenance and dispersion of ticks in nature. In addition, vertebrates act as amplifiers and/or reservoirs for viruses, protozoa and bacteria transmitted by these ectoparasites. The antropic action affects the population dynamics of both ticks and their wild hosts directly or indirectly, and consequently, the epidemiology of tick-borne diseases once restricted to wild fauna can reach domestic animal and humans interfaces [7]. Ticks are responsible for more than 100,000 cases of diseases in humans and animals worldwide [8], therefore a concern for public health professional. In Brazil, ticks are vectors of diseases such as babesiosis, ehrlichiosis, anaplasmosis and rickettsioses, including Brazilian Spotted Fever (BSF) [9, 10].
\nTicks belong to the Sub-Class Acari, Super-Order Parasitiformes, Order Ixodida, and four families: Ixodidae, Argasidae, Nuttallielidae and Deinocrotonidae [11, 12], the latter extinct. Of these, only the Ixodidae and Argasidae families occur in Brazil, with nine genera and 75 species [11, 13, 14, 15, 16]. Ixodidae family is the most diverse with 51 species and five genera: Amblyomma (33 species), Ixodes (12 species), Rhipicephalus (two species), Haemaphysalis (three species) and Dermacentor (one species). Argasidae family is represented by 24 species and four genera: Ornithodoros (18 species), Antricola (three species), Argas (one species) and Nothoaspis (two species). In general, the immature stages (larvae and nymphs) of two or three host ticks feed on small mammals and birds, while adults exploit medium to large-sized hosts [17]. One-host species Dermacentor nitens and Rhipicephalus microplus, complete the life cycle mainly on large animals such as horses and cattle, respectively. Second Esser et al. [18] the diversity of ticks increases with the increase of diversity of hosts, inasmuch as these hematophagous arthropods rely on the hosts to complete their life cycles. Therefore, if we take into account the high diversity of hosts living in this biome and the amount of unexplored ecoregions, it is reasonable to infer that the diversity of ticks in the Amazon biome is insufficiently addressed. Thus, a review on the subject is necessary to point out the existing gaps and encourage new studies on ticks in the Brazilian Amazon.
\nThe class Amphibia includes the orders Anura, Caudata, Gymnophiona, whereas the class Reptilia includes the orders Squamata, Testudines and Crocodylia. There are 331 amphibian and 550 reptile species in the Brazilian Amazon (Figure 1), although that faunal records are far from complete [2, 19]. Brazil has witnessed an increasing number of reports on tick parasitism of amphibians and reptiles over the past few years [20, 21, 22, 23, 24]. However, knowledge of this tick fauna as their hosts remains incomplete. To date, major tick-amphibian associations reported in Brazil are the ixodids Amblyomma dissimile, Amblyomma rotundatum, Amblyomma humerale, Amblyomma fuscum and Amblyomma goeldii [20, 21, 22, 23, 24], and the argasids Ornithodoros saraivai and Ornithodoros faccinii associated with species Cycloramphus boraceiensis and Thoropa miliaris, respectively [25, 26]. Occasional records on reptiles have also been reported for Ornithodoros mimon and Ornithodoros rietcorreai [22]. However, none of these argasids have been reported in the Amazon biome. Although the Brazilian Amazon has a high diversity of amphibians and reptiles, studies on ticks in association with these hosts are still rare, with a dominance of species A. dissimile, A. rotundatum and A. humerale [21, 22, 23, 27, 28].
\nThe tick A. dissimile is constantly misidentified with the morphologically similar A. rotundatum. Recently, the distribution of A. dissimile in the country was reorganized, showing that this tick is restricted to the Pantanal and Amazon biomes [28]. Therefore, reports outside these biomes are considered misidentifications and must be viewed with caution. Most publications on this species are simple records of occurrence with some authors including data on prevalence and intensity of infestation both in the Amazon biome and elsewhere. According to Luz et al. [29] 12 (54.5%) out of 22 Rhinella marina toads captured in Amapá state, were parasitized by a total of 97 ticks (6 males, 39 females, 31 nymphs, 21 larvae) and mean intensity of 8.1 ticks per infested toad. In the Amazonian biome, A. dissimile is common on R. marina (Anura) and Boidae (Squamata) (Table 1). The experimental life cycle including pre-attachment periods for each parasitic stage lasts approximately 350 days, as reported by Schumaker et al. [43] who started a colony from one engorged female collected from the Amazonian biome. Ogrzewalska et al. [37] reported Rickettsia bellii and ‘Candidatus Rickettsia colombianensi’ in ticks collected from Bothrops atrox from Pará state. Luz et al. [29] reported ‘Ca. R. colombianensi’ in ticks collected on R. marina from Amapá state.
\nHost | \nSpecie | \n\nA. dissimile\n | \n\nA. rotundatum\n | \n\n | ||||
---|---|---|---|---|---|---|---|---|
\n | \n | L | \nN | \nA | \nL | \nN | \nA | \n\n |
Amphibian (Anura) | \n\n | \n | \n | \n | \n | \n | \n | \n |
Bufonidae | \n\nRhinella marina\n | \n\n | 37 | \n12M; 16F | \n\n | 164 | \n133F | \n[23, 29, 30, 31, 32, 33] | \n
\n | \n\nRhinella major\n | \n\n | \n | 2F | \n\n | 15 | \n2F | \n[33, 34, 35] | \n
\nRhaebo guttatus\n | \n\n | \n | \n | \n | 1 | \n1F | \n[23] | \n|
\nRhinella margaritifera\n | \n\n | \n | \n | \n | 3 | \n1F | \n[23] | \n|
\nRhinella gildae\n | \n\n | \n | \n | \n | 1 | \n\n | [36] | \n|
Leptodactylidae | \n\nLeptodactylus pentadactylus\n | \n\n | \n | \n | \n | \n | 1F | \n[23] | \n
Reptile | \n\n | \n | \n | \n | \n | \n | \n | \n |
Boidae | \n\nBoa constrictor\n | \n\n | 1703 | \n723M; 1346F | \n\n | 12 | \n1M; 77F | \n[23, 30, 32, 33, 34, 36, 37, 38, 39, 40] | \n
\n | \n\nCorallus hortulanus\n | \n\n | 3 | \n1M | \n\n | \n | \n | [31, 34] | \n
\nEunectes murinus\n | \n\n | 305 | \n21M; 3F | \n\n | \n | \n | [31, 40] | \n|
Viperidae | \n\nBothrops atrox\n | \n4 | \n11 | \n24M; 14F | \n\n | \n | 1M; 2F | \n[32, 33, 36, 37] | \n
\n | \n\nCrotalus durissus\n | \n\n | \n | 1M; 1F | \n\n | \n | \n | [34] | \n
\nLachesis muta\n | \n\n | 1 | \n\n | \n | \n | \n | [34] | \n|
Elapidae | \n\nMicrurus averyi\n | \n\n | \n | 3F | \n\n | \n | \n | [31] | \n
\n | \nMicrurus lemniscatus\n | \n\n | \n | 12M; 9F | \n\n | \n | \n | [31] | \n
Colubridade | \n\nLeptophis ahaetulla\n | \n\n | \n | \n | \n | 3 | \n\n | [33] | \n
\n | \n\nChironius multiventris\n | \n\n | \n | \n | \n | 3 | \n\n | [33] | \n
\nChironius scurrulus\n | \n\n | \n | 1M; 1F | \n\n | \n | \n | [34] | \n|
\nChironius laevicollis\n | \n\n | 12 | \n15M; 6F | \n\n | \n | \n | [31] | \n|
\nMastigodryas boddaerti\n | \n\n | \n | \n | \n | 1 | \n\n | [33] | \n|
\nHydrodynastes gigas\n | \n\n | \n | \n | \n | 1 | \n7F | \n[30] | \n|
\nHelicops polylepis\n | \n\n | 1 | \n\n | \n | \n | \n | [34] | \n|
\nLeptodeira annulata\n | \n\n | 1 | \n\n | \n | \n | \n | [34] | \n|
\nPhimophis guerini\n | \n\n | \n | 1F | \n\n | \n | \n | [34] | \n|
\nErythrolamprus reginae\n | \n\n | 1 | \n\n | \n | \n | \n | [34] | \n|
\nSpilotes pullatus\n | \n\n | 72 | \n42M; 9F | \n\n | \n | \n | [31] | \n|
\nErythrolamprus reginae semilineatus\n | \n\n | 1 | \n\n | \n | \n | \n | [34] | \n|
Dipsadidae | \n\nXenodon severus\n | \n\n | \n | 8 | \n\n | \n | \n | [40] | \n
Testudinidae | \n\nChelonoidis denticulatus\n | \n\n | \n | 3M | \n\n | 3 | \n2M; 42F | \n[31, 41] | \n
\n | \nChelonoidis carbonaria\n | \n\n | 1 | \n1M; 1F | \n\n | 11 | \n\n | [32, 39] | \n
Podocnemididae | \n\nPodocnemis expansa\n | \n\n | \n | \n | \n | \n | 1F | \n[39] | \n
\n | \n\nPodocnemis unifilis\n | \n\n | \n | \n | \n | 1 | \n\n | [39] | \n
\nTrachemys dorbigni\n | \n\n | \n | \n | \n | 2 | \n12F | \n[39] | \n|
Kinosternidae | \n\nKinosternon scorpioides\n | \n\n | \n | \n | \n | \n | 2F | \n[42] | \n
Chelidae | \n\nPlatemys platycephala\n | \n\n | \n | \n | \n | \n | 5F | \n[32] | \n
Tropiduridade | \n\nUranoscodon superciliosus\n | \n\n | \n | \n | \n | 7 | \n6F | \n[33] | \n
\n | \nTropidurus sp. | \n\n | \n | \n | \n | \n | 1M | \n[38] | \n
Teiidae | \n\nAmeiva ameiva\n | \n\n | 4 | \n\n | \n | \n | 1F | \n[31, 34] | \n
Iguanidae | \n\nIguana iguana\n | \n\n | 31 | \n118M; 69F | \n\n | \n | 2F | \n[31, 36, 39, 40] | \n
Dactyoloidae | \n\nNorops auratus\n | \n\n | 1 | \n\n | \n | \n | \n | [34] | \n
Gekkonidae | \n\nHemidactylus mabouia\n | \n\n | 1 | \n\n | \n | \n | \n | [34] | \n
Alligatoridae | \n\nPaleosuchus trigonatus\n | \n\n | \n | 9M; 15F | \n\n | 4 | \n12F | \n[31, 32] | \n
\n | \nCaiman crocodilus\n | \n\n | 3 | \n\n | \n | \n | \n | [31, 39] | \n
Total | \n\n | 4 | \n2189 | \n983M; 1496F | \n\n | 221 | \n307F | \n\n |
Hosts of Amblyomma dissimile and Amblyomma rotundatum in the Brazilian Amazon.
The tick A. rotundatum is an obligate parthenogenetic species, although there are six reports of adult males, two males in the laboratory [44] and six in the Amazon region parasitizing Tropidurus sp., Boa constrictor, Iguana iguana, Chelonoidis denticulatus and B. atrox [30, 36, 44, 45, 46, 47]. Most publications on A. rotundatum are simple records of its occurrence with some additional data on prevalence and intensity of infestation in the Cerrado, Atlantic forest and Amazon biomes [22, 23, 31, 34, 48]. In the Brazilian Amazon, the families Bufonidae (Amphibian) and Boidae (Reptilia) are the most frequently parasitized by A. rotundatum, but there are reports on other species of Amphibian and Reptiles (Table 1). Recently, Gianizella et al. [31] reported A. rotundatum in different municipalities of the Amazonas state, on B. constrictor and several unknown hosts. The bioecology of this species under quasi-natural environment demonstrated a peak of larvae and nymphs in the dry season and females in the rainy season; however, there is a hypothesis of the absence of seasonality in this tick [48, 49]. Although A. rotundatum is a three-host tick on amphibians, it can behave as a two-host tick when feeding on snakes [50]. The complete experimental life cycle, including pre-attachment periods for each parasitic stage, ranged from 126 to 228 days on toads [51] or 56 to 163 days on snakes [50] and ulcerative lesions and hemorrhages after A. rotundatum feeding [48] were reported for ticks collected on toads from the Cerrado biome. Transmission of the hemogregarine Hemolivia stellata by R. marina collected in Belém, state of Pará was reported by Petit et al. [52]. This tick has also been found infected with R. bellii in the state of Amazon in the municipalities of Cacaulândia and Monte Negro, in the state of Rondônia; in the municipalities of Amapá (Ilha de Maracá) and Santana, in the state of Amapá; and in the municipality of Rio Branco, in the state of Acre [29, 53, 54].
\nMost reports of A. humerale mention the adult stage parasitizing species of tortoises, namely the “yellow-footed tortoise” C. denticulatus and the “red-footed tortoise” Chelonoidis carbonarius [27, 55]. There is also a record of A. humerale on Rhinoclemmys punctularia in central Amazon [56]. Labruna et al. [27] collected 215 adult ticks from six C. denticulatus and nine Chelonoidis sp. from state of Rondônia, with mean infestation of 14.3 ± 12.0 ticks per tortoise. In addition, seven engorged nymphs were collected on lizards. Morais et al. [57] collected 120 adult ticks from 18 (75%) out of 24 C. denticulatus captured in a transitional area between the biomes Cerrado and southwestern Amazon rainforest, in the state of Mato Grosso. The mean intensity of infestation was 6.7 ticks/tortoise. In both surveys, male ticks were mostly attached in clusters on the carapace whereas females were found attached to the tortoise skin. The sex ratio (males:females) for A. humerale were different in both surveys, 10.3:1 [27] and 1.1:1 [57]. There is one additional record of two nymphs on Paleosuchus trigonatus (Crocodylia) [32]. The life-cycle in the laboratory, including pre-feeding periods for each of the parasitic stages, could be completed in an average period of ca. 200 days [58]. Rickettsia bellii and Rickettsia amblyommatis have been detected in A. humerale [42, 53]. The records of A. fuscum in association with amphibians and reptiles are rare in Brazil. Dantas-Torres et al. [59] collected one male tick in one out of 490 caimans (Caiman latirostris and Paleosuchus palpebrosus) trapped in the Atlantic rainforest biome in Pernambuco state, north-eastern Brazil. Amblyomma goeldii has been recorded only in the Amazonas state. Martins et al. [60] reported two males collected on B. constrictor and recovery of ≈100 (20%) engorged larvae out of ≈500 unfed larvae experimentally infested on a B. constrictor.\n
\nThe importance of birds to maintain biodiversity and ecological balance of nature is notorious [61]. Due to migration, wild birds are of concern to human and animal health worldwide [62] because they can carry infected ticks over long distances, directly influencing the epidemiology of tick-borne diseases in animals and humans. In addition, wild birds themselves can be reservoirs of Borrelia burgdorferi sensu lato, and potentially to Anaplasma phagocytophilum and Rickettsia spp. [62, 63]. Wild birds play an important role in maintaining and dispersing immatures (larvae and nymphs) of several tick species into new locations [61, 64].
\nOf the total genera of ticks described in Brazil, five have at least one species recorded in association with wild birds. The most common are the hard ticks of the genera Amblyomma, Haemaphysalis and Ixodes [61, 64]. There are also occasional reports of the genera Rhipicephalus and Ornithodoros. Ticks of the genus Amblyomma are the most common on wild birds in the Brazil including the Amazon biome, exclusively for the larvae and nymphs [61, 64]. Adult ticks are only occasionally found on wild birds, with the exception of Ixodes paranaensis and Ixodes auritulus, which have the entire cycle synchronized with birds [65]. In Brazil, there are no reports of wild birds as a source of pathogens transmitted by ticks to humans, but they can serve as disperser hosts for vectors of Brazilian Spotted Fever (BSF) as Amblyomma sculptum, Amblyomma aureolatum and Amblyomma ovale, in the larvae and nymph stages [64]. Therefore, wild birds act indirectly in the epidemiology of BSF by dispersing and maintaining their vectors in nature.
\nOver more than 1,900 birds recorded in Brazil, approximately 1,300 reside in the Brazilian Amazon, with a 20% of endemism [66] (Figure 1). Of these, approximately 7% are migrants from the northern hemisphere and southern South America, including migrations from other Brazilian biomes [67]. To date, 86 bird species of Brazilian Amazonian have been recorded in association with at least one tick species (Table 2). This is equivalent to approximately only 7% of bird species found in this biome and 5% of the total birds recorded in Brazil (Table 2). Similar to other studies regarding tick-bird associations in Brazil [73, 74, 75], Passeriformes birds were the most parasitized in the Amazon biome, including 14 families and 72 species (Figure 2 and Table 2). In this group, the greatest diversity of parasitized birds was Thamnophilidae (20 species) followed by Dendrocolaptidae (16 species) and Tyrannidade (10 species) (Figure 2). The least parasitized families were Conopophagidae, Furnariidae, Xenopidae, Tityridae, Cardinalidae, Columbidae, Cuculidae, Momotidae, Capitonidae, Ramphastidae, Psittacidae, Accipitridae and Falconidae with only one species of parasitized bird each (Figure 2 and Table 2). Non-Passerines were represented by 10 different orders and 11 families, with emphasis on Bucconidae with three species (Figure 2 and Table 2). To date, approximately 1,068 specimens of ticks have been collected from birds in the Brazilian Amazon, in the stages of larvae (884/83%), nymphs (184/17%) and no adults (Figure 3). These are included in the genera Amblyomma and Haemaphysalis. The genus Amblyomma was the dominant with eight species (Table 2). The greatest diversity of ticks was reported for the Thamnophilidae family with seven species: A. longirostre, A. nodosum, A. humerale, A. calcaratum, A. geayi, A. coelebs and Haemaphysalis juxtakochi. The Dendrocolaptidae family was the second with five species: A. longirostre, A. nodosum, A. humerale, A. calcaratum and A. geayi (Figure 2 and Table 2).
\nHosts | \nTicks | \nReferences | \n||||
---|---|---|---|---|---|---|
\n | \n | \n | Species | \nStages | \n\n | |
\n | \n | \n | \n | LL | \nNN | \n\n |
Passeriformes | \nThamnophilidae | \n\nThamnomanes schistogynus\n | \n\nA. humerale\n \nA. nodosum\n | \n\n | 1 1 | \n[68] | \n
\nThamnophilus doliatus\n | \n\nA. nodosum\n | \n\n | 2 | \n[68] | \n||
\nThamnophilus schistaceus\n | \n\nA. nodosum\n \nAmblyomma sp. | \n2 | \n2 | \n[68] | \n||
\nThamnophilus aethiops\n | \n\nA. calcaratum\n \nA. longirostre\n | \n\n | 5 1 | \n[68] | \n||
\nMyrmelastes hyperythrus\n | \n\nA. nodosum\n | \n\n | 9 | \n[68] | \n||
\nSciaphylax hemimelaena\n | \n\nA. nodosum\n | \n\n | 1 | \n[68] | \n||
\nSchistocichla leucostigma\n | \n\nA. humerale\n \nAmblyomma sp. | \n4 5 | \n\n | [69] | \n||
\nPhlegopsis nigromaculata\n | \n\nA.coelebs\n \nA. longirostre\n \nA. nodosum\n \nAmblyomma sp. | \n1 1 24 | \n5 | \n[68, 69] | \n||
\nHypocnemis subflava\n | \n\nAmblyomma sp. | \n3 | \n\n | [70] | \n||
\nHylophylax naevius\n | \n\nH. juxtakochi\n \nA. humerale\n \nA. longirostre\n \nA. calcaratum\n \nAmblyomma sp. | \n1 3 1 8 | \n1 2 1 | \n[69] | \n||
\nHylophylax punctulatus\n | \n\nAmblyomma sp. | \n1 | \n\n | [69] | \n||
\nThamnomanes caesius\n | \n\nA. geayi\n \nA. longirostre\n \nA. humerale\n \nAmblyomma sp | \n1 3 4 | \n2 1 | \n[69, 71] | \n||
\nHylophylax poecilinotus\n | \n\nA. geayi\n \nA. humerale\n \nA. longirostre\n \nAmblyomma sp. \nH. juxtakochi\n | \n1 1 3 22 | \n1 1 1 1 | \n[69] | \n||
\nMyrmornis torquata\n | \n\nAmblyomma sp. | \n3 | \n\n | [69] | \n||
\n | \n\n | \n\nMyrmotherula hauxwelli\n | \n\nAmblyomma sp. | \n1 | \n\n | [69] | \n
\nMyrmotherula longipennis\n | \n\nA. longirostre\n \nAmblyomma sp. | \n3 3 | \n\n | [69] | \n||
\nPyriglena leuconota\n | \n\nA. coelebs\n \nA. humerale\n \nA. longirostre\n \nAmblyomma sp. | \n1 1 9 4 | \n1 | \n[69] | \n||
\nMyrmoborus myotherinus\n | \n\nA. longirostre\n | \n\n | 1 | \n[69] | \n||
\nEpinecrophylla leucophthalma\n | \n\nA. longirostre\n | \n\n | 1 | \n[71] | \n||
\nWillisornis poecilinotus\n | \n\nA. calcaratum\n | \n\n | 1 | \n[71] | \n||
Conopophagidae | \n\nConopophaga aurita\n | \n\nAmblyomma sp. | \n\n | 1 | \n[69] | \n|
Scleruridae | \n\nSclerurus cf. rufigularis\n | \n\nA. humerale\n | \n\n | 1 | \n[68] | \n|
\nSclerurus caudacutus\n | \n\nA. humerale\n | \n\n | 1 | \n[69] | \n||
Furnariidae | \n\nAutomolus paraensis\n | \n\nAmblyomma sp. | \n\n | 1 | \n[69] | \n|
Dendrocolaptidae | \n\nDendrocincla merula\n | \n\nA. longirostre\n \nAmblyomma sp. | \n7 34 | \n1 | \n[68, 69, 71] | \n|
\nDendrocincla fuliginosa\n | \n\nAmblyomma sp. | \n10 | \n\n | [31, 70] | \n||
\nDeconychura longicauda\n | \n\nA. longirostre\n \nAmblyomma sp. | \n4 39 | \n1 | \n[69, 71] | \n||
\nDeconychura stictolaema\n | \n\nA. longirostre\n \nAmblyomma sp. | \n1 1 | \n1 | \n[69] | \n||
\nDendrocolaptes hoffmannsi\n | \n\nA. calcaratum\n \nA. geayi\n \nA. humerale\n \nA. longirostre\n \nAmblyomma sp. | \n1 2 4 39 59 | \n2 1 3 | \n[69] | \n||
\nDendrocolaptes certhia\n | \n\nA. longirostre\n | \n\n | 1 | \n[71] | \n||
\nGlyphorynchus spirurus\n | \n\nA. longirostre\n | \n\n | 1 | \n[71] | \n||
\nGlyphorynchus spirurus\n | \n\nA. geayi\n \nA. longirostre\n \nAmblyomma sp. | \n1 6 16 | \n1 2 | \n[69] | \n||
\nHylexetastes brigidai\n | \n\nA. geayi\n \nA. longirostre\n \nAmblyomma sp. | \n1 6 21 | \n2 | \n[69] | \n||
\nSittasomus griseicapillus\n | \n\nA. humerale\n \nAmblyomma sp. | \n2 | \n1 | \n[70] | \n||
\nXiphorhynchus guttatus\n | \n\nA. longirostre\n \nA. nodosum\n | \n\n | 2 1 | \n[68] | \n||
\nXiphorhynchus guttatoides\n | \n\nA. humerale\n \nAmblyomma sp. | \n1 | \n2 | \n[70] | \n||
\nXiphorhynchus elegans\n | \n\nA. longirostre\n \nAmblyomma sp. | \n3 2 | \n2 | \n[69] | \n||
\n | \n\n | \n\nXiphorhynchus ocellatus\n | \n\nA. calcaratum\n | \n\n | 1 | \n[71] | \n
\nXiphorhynchus pardalotus\n | \n\nAmblyomma sp. | \n3 | \n\n | [31] | \n||
\nDendroplex picus\n | \n\nA. longirostre\n \nA. nodosum\n | \n\n | 3 3 | \n[68, 70] | \n||
Xenopidae | \n\nXenops minutus\n | \n\nA. nodosum\n | \n\n | 2 | \n[69] | \n|
Pipridae | \n\nPipra filicauda\n | \n\nA. geayi\n \nAmblyomma sp. | \n\n | 1 | \n[68, 70] | \n|
\nPipra fasciicauda\n | \n\nA. longirostre\n \nA. nodosum\n \nAmblyomma sp. | \n1 8 | \n7 | \n[68] | \n||
\nMachaeropterus pyrocephalus\n | \n\nA. longirostre\n | \n\n | 1 | \n[68] | \n||
\nDixiphia pipra\n | \n\nA. geayi\n \nA. longirostre\n | \n1 1 | \n1 1 | \n[69] | \n||
\nLepidothrix nattereri\n | \n\nAmblyomma sp. | \n1 | \n\n | [71] | \n||
Rhynchocyclidae | \n\nLeptopogon amaurocephalus\n | \n\nA. geayi\n | \n1 | \n\n | [68] | \n|
\nRhynchocyclus olivaceus\n | \n\nA. geayi\n \nA. humerale\n \nA. longirostre\n \nA. nodosum\n \nAmblyomma sp. | \n1 1 4 | \n2 1 17 | \n[68] | \n||
\nTodirostrum maculatum\n | \n\nA. nodosum\n | \n\n | 1 | \n[68] | \n||
\nPoecilotriccus latirostris\n | \n\nA. nodosum\n \nAmblyomma sp. | \n6 | \n3 | \n[70] | \n||
\nHemitriccus flammulatus\n | \n\nAmblyomma sp. | \n1 | \n\n | [70] | \n||
Tyrannidae | \n\nMyiozetetes similis\n | \n\nA. nodosum\n | \n\n | 1 | \n[68] | \n|
\nEmpidonax alnorum\n | \n\nA. nodosum\n \nA. longirostre\n | \n4 | \n1 | \n[68] | \n||
\nPlatyrinchus saturatus\n | \n\nAmblyomma sp. | \n1 | \n\n | [69] | \n||
\nHemitriccus minor\n | \n\nA. longirostre\n | \n1 | \n\n | [69] | \n||
\nMionectes macconnelli\n | \n\nA. longirostre\n \nAmblyomma sp. | \n4 5 | \n\n | [69] | \n||
\nAttila spadiceus\n | \n\nA. geayi\n \nAmblyomma sp. | \n1 3 | \n\n | [69] | \n||
\nMyiobius barbatus\n | \n\nA. longirostre\n | \n3 | \n\n | [69] | \n||
\nOnychorhynchus coronatus\n | \n\nA. longirostre\n \nAmblyomma sp. | \n3 6 | \n\n | [69] | \n||
\nRamphotrigon megacephalum\n | \n\nA. nodosum\n \nAmblyomma sp. | \n1 | \n3 | \n[70] | \n||
\nKnipolegus poecilocercus\n | \n\nAmblyomma sp. | \n1 | \n\n | [71] | \n||
\n | \nTroglodytidae | \n\nPheugopedius genibarbis\n | \n\nA. nodosum\n \nAmblyomma sp. | \n363 | \n1 | \n[68] | \n
\nMicrocerculus marginatus\n | \n\nAmblyomma sp. | \n3 | \n\n | [69] | \n||
Tityridae | \n\nSchiffornis turdina\n | \n\nA. longirostre\n \nAmblyomma sp. | \n3 14 | \n\n | [69] | \n|
Turdidae | \n\nTurdus hauxwelli\n | \n\nA. longirostre\n | \n\n | 2 | \n[68] | \n|
\nTurdus sanchezorum\n | \n\nA. longirostre\n | \n\n | 1 | \n[68] | \n||
\nTurdus ignobilis\n | \n\nA. geayi\n \nA. nodosum\n \nA. longirostre\n \nAmblyomma sp. | \n2 | \n1 1 1 | \n[68] | \n||
\nTurdus amaurochalinus\n | \n\nAmblyomma sp. | \n1 | \n\n | [70] | \n||
\nTurdus albicollis\n | \n\nA. longirostre\n \nA. coelebs\n \nAmblyomma sp. | \n2 3 | \n3 | \n[31, 69] | \n||
Thraupidae | \n\nRamphocelus carbo\n | \n\nA. nodosum\n \nA. longirostre\n \nAmblyomma sp. | \n44 | \n15 4 | \n[30, 68, 70, 71] | \n|
\nTachyphonus surinamus\n | \n\nA. humerale\n \nA. longirostre\n \nAmblyomma sp. | \n1 1 5 | \n\n | \n | ||
Cardinalidae | \n\nCyanoloxia cyanoides\n | \n\nAmblyomma sp. | \n1 | \n\n | [71] | \n|
Columbiformes | \nColumbidae | \n\nColumbina talpacoti\n | \n\nA. nodosum\n | \n\n | 1 | \n[68] | \n
Cuculiformes | \nCuculidae | \n\nCrotophaga major\n | \n\nA. nodosum\n | \n\n | 1 | \n[68] | \n
Coraciiformes | \nMomotidae | \n\nMomotus momota\n | \n\nA. longirostre\n \nA. humerale\n | \n\n | 1 1 | \n[68, 70] | \n
Piciformes | \nCapitonidae | \n\nCapito auratus insperatus\n | \n\nA. nodosum\n \nA. longirostre\n | \n\n | 2 1 | \n[68] | \n
Ramphastidae | \n\nRamphastos vitellinus\n | \n\nA. geayi\n | \n\n | 3 | \n[31] | \n|
\n | \nPteroglossus bitorquatus\n | \n\nA. longirostre\n | \n\n | 1 | \n[30] | \n|
Galbuliformes | \nBucconidae | \n\nMonasa nigrifrons\n | \n\nAmblyomma sp. | \n1 | \n\n | [70] | \n
\nMalacoptila rufa\n | \n\nAmblyomma sp. | \n1 | \n\n | [69] | \n||
\nGalbula cyanicollis\n | \n\nAmblyomma sp. | \n2 | \n\n | [69] | \n||
Psittaciformes | \nPsittacidae | \n\nPrimolius maracana\n | \n\nA. dissimile\n | \n\n | 1 | \n[72] | \n
Accipitriformes | \nAccipitridae | \n\nHarpia harpyja\n | \n\nA. cajennense\n | \n\n | 16 | \n[41] | \n
\n | \n | \n | \nH. juxtakochi\n | \n\n | 1 | \n[41] | \n
Cariamiformes | \nCariamidae | \n\nCariama cristata\n | \n\nA. cajennense\n | \n\n | 5 | \n[39] | \n
Galliformes | \nCracidae | \n\nPenelope superciliaris\n | \n\nA. cajennense\n | \n1 | \n1 | \n[42] | \n
Falconiformes | \nFalconidae | \n\nMicrastur ruficollis\n | \n\nA. longirostre\n | \n\n | 1 | \n[71] | \n
Total | \n\n | \n | \n | 884 | \n184 | \n\n |
Ticks identified on wild birds in the Brazilian Amazon biome.
Diversity of hard ticks parasitizing wild birds from the Brazilian Amazon.
Distribution of studies reporting the association of hard ticks on wild birds from the Brazilian Amazon.
Overall, A. longirostre and A. nodosum are the two most common species on wild birds in the Brazilian Amazon [68, 69, 70, 71] (Figure 4 and Table 2). The tick A. longirostre was the most common, recorded in 12 families and 40 species of birds (36 Passerines and 4 non-Passerines) (Figure 2). Amblyomma longirostre has been treated as an arboreal tick, with immatures parasitizing birds and adults parasitizing rodents Erethizontidae (e.g., Sphiggurus spp.) [73, 76]. This tick was also the most abundant with 110 larvae and 39 nymphs. Amblyomma longirostre was found in co-infestation with the following species: A. calcaratum, A. nodosum, A. coelebs, A. humerale, A. geayi and H. juxtakochi (Table 2). In addition, as it is frequent on birds, A. longirostre is popularly known as “bird tick” or “bird earring” [77]. Amblyomma nodosum was the second most common species collected on 12 families and 22 species of birds. This tick was recorded in co-infestation with A. longirostre, A. coelebs, A. humerale and A. geayi (Table 2 and Figures 2 and 4). The birds most infested by A. nodosum were Rhynchocyclus olivaceus and Ramphocelus carbo with 17 and 15 nymphs, respectively. These birds inhabit the forest understory and visit the soil occasionally [67]. As Xenartha mammals (Myrmecophaga tridactyla, Tamandua tetradactyla) are the primary hosts of A. nodosum [17], it is believed that the low areas of the understory are a major source of infestation. Interestingly, T. tetradactyla may have arboreal habits [78], which helps to explain the presence of A. nodosum also on birds of different forest strata.
\nHard tick species parasitizing wild birds from the Brazilian Amazon.
To date, of the total of ticks collected, 736 (70%) were larvae identified as Amblyomma sp. due to lack of reliable tools for larval identification, thus, indicating that the diversity of ticks on birds of the Amazon may be underestimated. Luz et al. [75], using molecular biology, identified more than 90% of the larvae collected from birds in the Atlantic Forest biome, reporting the greatest diversity of ticks on birds in a single study in Brazil and description of the new tick Amblyomma romarioi [14]. Therefore, the identification of all larvae by molecular biology in addition to morphological identification is extremely important to ascertain the diversity of ticks in the Amazon biome.
\nThe fauna of wild mammals in Brazil is quite diverse and more than half lives in the Amazon biome [79]. Like birds, amphibians and reptiles, mammals play an important role in preserved or anthropized ecosystems. Therefore, the knowledge of the local diversity of wild mammals, and their relationship with ticks is considered an important tool for public conservation policies and consequently for public health. The alteration of wild habitats can determine changes in the patterns of parasitic specificity, inducing tick species to seek new groups of hosts, increasing the risk of disease transmission [79, 80]. Some wild mammals (small, medium and large) are directly or indirectly involved in the transmission cycles of many tick-borne pathogens worldwide, including Anaplasma spp., Babesia spp., Borrelia spp., and Rickettsia spp. [80, 81]. In Brazil, the main zoonosis transmitted by ticks is Brazilian Spotted Fever caused by the bacterium Rickettsia rickettsii, which has the rodent Hydrochoerus hydrochaeris as its main amplifier. In addition, there is evidence that marsupials and small rodents can serve as amplifier hosts for R. rickettsii in nature [82].
\nIn Brazil there are approximately 755 species of mammals distributed in all its six biomes, including the Amazon biome [78, 79]. Of the total mammal species, 41% (≅ 311 species) occur in the Amazon biome (Figure 1) [2, 76]. These vertebrates are distributed in 11 orders, 51 families and 249 genera [78, 79]. Among the families, Cricetidae is the most diverse, with 144 species [78, 79]. The vast majority of tick species, including all life stages, in Brazil have records on wild mammals of different sizes [17]. In general, small mammals of the orders Rodentia and Didelphimorphia are those that have a greater number of studies in association with ticks, especially the families Cricetidae and Didelphidae [17, 82]. In general, medium and large mammals are parasitized by ticks in all stages (larva, nymph and adult), while in small mammals the stages of larva and nymph are more common. In this last group of hosts we can highlight the cricetids Akodon spp., Calomys spp., Oligoryzomys spp. and Nectomys spp. as the most parasitized by immature ticks in nature.
\nNine orders and 24 families of wild mammals have representatives in association with ticks in the Brazilian Amazon (Figure 5). Rodentia was the most diverse with 16 species of mammals, followed by the orders Carnivora (13 species) and Didelphimorphia (12 species) (Figure 5). However, it was the family Didelphidae that presented a greater number of parasitized species, followed by Dasyproctidae (seven species) and Mustelidae (four species). These records corroborate with numerous studies of tick parasitism on wild animals from South America, with emphasis on the orders Rodentia and Didelphimorphia [17, 82, 83].
\nDiversity of hard ticks parasitizing wild mammals from the Brazilian Amazon.
Hard ticks parasitizing wild mammals in the Brazilian Amazon are represented by five genera: Amblyomma, Ixodes, Haemaphysalis, Dermacentor and Rhipicephalus. Of these, the genus Amblyomma was more frequently recorded with 23 species (Figure 5). The orders Rodentia, Pilosa and Didelphimorphia are hosts for highest diversity to Amblyomma species 17, 15 and eight, respectively. All these species are also recorded on a variety of wild mammals in Brazil [83, 84, 85, 86, 87], except for A. rotundatum and A. dissimile, which are more specific ticks of cold-blooded animals (Amphibians and Reptiles), although there are occasional reports on mammals in South America [20, 83]. Amblyomma humerale, of which the adult stage is more specific to tortoises, immature stages have been found on a variety of small mammals, reptiles and birds [17, 68, 71, 83].
\nThe second most common genus in the Amazon is Ixodes, with six reported species: Ixodes amarali, Ixodes bocatorensis, Ixodes lasallei, Ixodes luciae, Ixodes schulzei and Ixodes spinosus (Table 3). Although there are two exclusive species on birds in Brazil [65], all species recorded in the Brazilian Amazon parasitize mainly wild mammals [13, 16, 31, 32, 88, 99, 102]. Ixodes spp. were found on families Didelphidae, Dasyproctidae, Cricetidae, Myrmecophagidae, Bradypodidae and Cyclopedidae (Figure 5 and Table 3). In general Ixodes ticks mainly parasitize rodents (e.g., Cricetidae) in the larvae and nymph stages, with adults feeding mostly on marsupials (e.g., Didelphis spp., Monodelphis spp.) [17, 83], and agoutis (e.g., Dasyprocta) [13, 16]. Most species were found parasitizing Monodelphis glirina (I. amarali, I. schulzei, I. luciae) and Monodelphis touan (I. amarali, I. schulzei), with some reports of I. amarali on the rodent Hylaeamys megacephalus [30, 38, 92, 99, 100]. Also on rodents the species I. spinosus, I. lasallei and I. bocatorensis have been recorded [30, 31, 38], although these previous records referred to them erroneously as Ixodes fuscipes, which according to more recent data, does not occur in the Amazon biome (Figure 5 and Table 3) [13, 16].
\nTick species | \nDomestic mammals | \nWild mammals | \nStates | \nReferences | \n
---|---|---|---|---|
\nAmblyomma auricularium\n | \n\n | \nCabassous unicinctus\n\n4\n, Dasypus novemcinctus\n\n4\n\n,\n\n5\n\n,\n\n7\n, Dasypus septemcinctus\n\n4\n\n,\n\n9\n\n, Euphractus sexcinctu\n\n4\n\n,\n\n9\n\n, Galactis cuja\n\n4\n\n,\n\n9\n, Galictis vittata\n\n4\n, Lycalopex vetulus\n\n9\n, Myrmecophaga tridactyla\n\n9\n, Pecari tajacu\n\n9\n, Tamandua tetradactyla\n\n4\n, Thrichomys inermis\n\n4\n\n | \nMA\n4\n, MT\n5\n, RO\n7\n, TO\n9\n\n | \n[30, 32, 36, 39] | \n
\nAmblyomma cajennense sensu stricto | \n\nEquus caballus\n\n4\n\n\n,\n\n\n5\n\n\n,\n\n\n6\n\n\n,\n\n\n7\n\n\n,\n\n\n8\n\n\n,\n\n\n9\n\n, Sus scrofa\n\n4\n\n\n,\n\n\n9\n, Canis familiaris\n\n4\n\n\n,\n\n\n9\n, Bubalus bubalis\n\n7\n\n | \n\nCaluromys philander\n\n5\n\n, Cerdocyon thous\n\n5\n, Didelphis marsupialis\n\n5\n\n,\n\n6\n\nDasyprocta azarae\n\n5\n, Hydrochoerus hydrochaeris\n\n5\n, Mazama gouazoubira\n\n4\n, Metachirus nudicaudatus\n\n4\n, Myrmecophaga tridactyla\n\n5\n\n\n,\n\n\n9\n, Nasua nasua\n\n5\n, Panthera onca\n\n5\n\n, Pecari tajacu\n\n5\n, Pteronura brasiliensis\n\n3\n, Tapirus terrestris\n\n5\n, Tayassu pecari\n\n5\n\n | \nAM\n3\n, MA\n4\n, MT\n5\n, PA\n6\n, RO\n7\n, RR\n8\n, TO\n9\n\n | \n[32, 36, 42, 88, 89, 90, 91] | \n
\nAmblyomma calcaratum\n | \n\n | \nMyrmecophaga tridactyla\n\n7\n, Tamandua tetradactyla\n45,6\n | \nMA\n4\n, MT\n5\n, PA\n6\n; RO\n7\n\n | \n[36, 38, 42] | \n
\nAmblyomma coelebs\n | \n\nEquus caballus\n\n7\n\n | \n\nAgouti paca\n\n7\n, Cuniculus paca\n\n6\n, Dasyprocta azarae\n\n5\n, Dasyprocta sp\n7\n\n; Didelphis albiventris\n\n5\n, Didelphis marsupialis\n\n3\n\n,\n\n5\n, Hydrochoerus hydrochaeris\n\n7\n, Metachirus myosuros\n\n5\n, Nasua nasua\n\n5\n, Neacomys spinosus\n\n5\n, Panthera onca\n\n5\n, Sapajus macrocephalus\n\n3\n, Tayassu pecari\n\n6\n, Pecari tajacu\n\n7\n, Tapirus terrestris\n\n3\n\n,\n\n5\n\n,\n\n7\n\n | \nAM\n3\n, MT\n5\n, PA\n6\n, RO\n7\n\n | \n[30, 31, 32, 38, 42, 88] | \n
\nAmblyomma dissimile\n | \n\n | \nBradypus tridactylus\n\n3\n, Choleopus didactylus\n\n3\n, Coendou sp\n3\n, Dasyprocta leporina\n\n3\n\n | \nAM\n3\n\n | \n[31, 36, 40] | \n
\nAmblyomma dubitatum\n | \n\n | \nHydrochoerus hydrochaeris\n\n1\n\n\n,\n\n\n5\n\n\n,\n\n\n7\n\n, Tamandua tetradactyla\n\n9\n\n | \nAC\n1\n, MT\n5\n, RO\n7\n, TO\n9\n\n | \n[30, 32, 39, 54] | \n
\nAmblyomma geayi\n | \n\n | \nAlouatta nigerrima\n\n6\n, Artibeus lituratus\n\n3\n, Bradypus tridactylus\n\n3\n\n\n,\n\n\n6\n, Bradypus variegatus\n\n1\n\n,\n\n3\n\n,\n\n4\n\n,\n\n6\n\n,\n\n9\n, Caluromys lanatus\n\n3\n, Choloepus hoffmanni\n\n7\n, Choloepus didactylus\n\n3\n\n,\n\n7\n, Cyclopes didactylus\n\n3\n, Didelphis marsupialis\n\n6\n, Philander opossum\n\n3\n, Proechimys sp\n3\n, Oecomys sp\n6\n, Saguinus bicolor\n\n3\n, Tamandua tetradactyla\n\n3\n\n | \nAC\n1\n, AM\n3\n, MA\n4\n, PA\n6\n, RO\n7\n, TO\n9\n\n | \n[30, 31, 33, 36, 39, 40, 42, 92] | \n
\nAmblyomma goeldii\n | \n\n | \nTamandua tetradactyla,\n\n3\n\n,\n\n6\n\n, Bradypus tridactylus\n\n3\n\n | \nAM\n3\n, PA\n6\n\n | \n[31, 40, 42] | \n
\nAmblyomma humerale\n | \n\n | \nBradypus tridactylus\n\n3\n, Chrotopterus auritus\n\n3\n, Cyclopes didactylus\n\n7\n, Dasypus novemcinctus\n\n5\n\n,\n\n6\n\n, Didelphis marsupialis\n\n3\n\n,\n\n5\n\n,\n\n6\n\n, Hydrochoerus hydrochaeris\n\n1\n, Marmosa constantiae\n\n5\n, Marmosa sp\n3\n, Metachirus myosuros\n\n5\n\n, Philander opossum\n\n3\n, Proechimys roberti\n\n5\n\n, Tamandua tetradactyla\n\n3\n\n\n,\n\n\n6\n\n | \nAC\n1\n, AM\n3\n, MT\n5\n, PA\n6\n, RO\n7\n\n | \n[30, 31, 32, 33, 38, 39, 40, 42, 54, 88] | \n
\nAmblyomma incisum\n | \n\n | \nTapirus terrestris\n\n7\n\n | \nRO\n7\n\n | \n[38, 93] | \n
\nAmblyomma latepunctatun\n | \n\n | \nTapirus terrestris\n\n3\n\n\n,\n\n\n6\n\n\n,\n\n\n7\n, Tayassu pecari\n\n3\n, Dasyprocta fuliginosa\n\n3\n, Didelphis marsupialis\n\n3\n\n | \nAM\n3\n; PA\n6\n; RO\n7\n\n | \n[31, 38, 93] | \n
\nAmblyomma longirostre\n | \n\n | \nCoendou sp\n3\n, Coendou prehensilis\n\n6\n\n\n,\n\n\n7\n, Coendou nycthemera\n\n6\n\n | \nAM\n3\n, PA\n6\n, RO\n7\n\n | \n[30, 31, 33, 40, 42] | \n
\nAmblyomma naponense\n | \n\n | \nDasyprocta fuliginosa\n\n3\n, Galictis sp\n3\n, Hydrochoerus hydrochaeris\n\n1\n, Mazama gouazoubira\n\n4\n, Myoprocta acouchy\n\n3\n, Tapirus terrestris\n\n7\n, Tayassu pecari\n\n3\n\n,\n\n4\n\n | \nAC\n1\n, AM\n3\n, MA\n4\n, RO\n7\n\n | \n[31, 36, 38, 54] | \n
\nAmblyomma nodosum\n | \n\n | \nTamandua tetradactyla\n\n4\n\n,\n\n5\n\n,\n\n6\n\n,\n\n7\n\n,\n\n9\n; Bradypus variegatus\n\n9\n, Myrmecophaga tridactyla\n\n9\n\n | \nMA\n4\n, MT\n5\n, PA\n6\n, RO\n7\n, TO\n9\n\n | \n[30, 32, 38, 39, 40, 42] | \n
\nAmblyomma oblongoguttatum\n | \n\nCanis familiaris\n\n4\n\n,\n\n6\n\n,\n\n9\n, Sus scrofa\n\n7\n\n | \n\nAgouti paca\n\n7\n\n, Cuniculus paca\n\n9\n, Dasyprocta sp\n7\n, Myrmecophaga tridactyla\n\n9\n, Tayassu tacaju\n\n7\n\n, Tayassu pecari\n\n3\n\n,\n\n5\n\n,\n\n6\n\n,\n\n7\n, Dasyprocta fuliginosa\n\n3\n, Mazama americana\n\n3\n, Tapirus terrestris\n\n5\n\n,\n\n7\n, Panthera onca\n\n7\n\n | \nAM\n3\n, MA\n4\n, MT\n5\n, PA\n6\n, RO\n7\n, TO\n9\n\n | \n[31, 32, 38, 39, 42, 91, 94, 95] | \n
\nAmblyomma ovale\n | \n\nCanis familiaris\n\n3\n\n,\n\n4\n\n,\n\n9\n, Sus scrofa\n\n7\n\n | \n\nPanthera onca,\n\n3\n\n,\n\n5\n\n,\n\n7\n\n,\n\n9\n, Tapirus terrestres\n\n3\n, Leopardus pardalis\n\n3\n, Proechimys quadruplicatus\n\n3\n\n, Proechimys sp\n3\n, Tapirus terrestres\n\n3\n\n, Pauxi tuberosa\n\n5\n, Puma concolor\n\n5\n, Procyon cancrivorus\n\n7\n, Tayassu tacaju\n\n7\n, Tayassu pecari\n\n7\n\n, Agouti paca\n\n7\n, Dasyprocta sp\n7\n\n, Eira barbara\n\n7\n\n, Nasua nasua\n\n7\n, Cerdocyon thous\n\n9\n\n | \nAM\n3\n, MA\n4\n, MT\n5\n, RO\n7\n, TO\n9\n, | \n[30, 31, 32, 38, 39, 42, 91, 94] | \n
\nAmblyomma pacae\n | \n\nCanis familiaris\n\n7\n\n | \n\nHydrochoerus hydrochaeris\n\n1\n\n, Cuniculus paca\n\n5\n, Tamandua tetradactyla\n\n6\n, Monodelphis glirina\n\n6\n\n, Didelphis marsupialis\n\n7\n\n | \nAC\n1\n, MT\n5\n, PA\n6\n, RO\n7\n\n | \n[38, 42, 54, 92] | \n
\nAmblyomma parkeri\n | \n\n | \nDidelphis marsupialis\n\n5\n\n | \nMT\n5\n\n | \n[32] | \n
\nAmblyomma romitii\n | \n\n | \nHydrochoerus hydrochaeris\n\n5\n\n,\n\n6\n\n,\n\n7\n\n | \nMT\n5\n, PA\n6\n, RO\n7\n\n | \n[32, 41, 96] | \n
\nAmblyomma rotundatum\n | \n\n | \nHydrochoerus hydrochaerisi, Xenodon merremii\n\n9\n\n | \nRO7, TO\n9\n\n | \n[30, 31, 32, 33, 36, 38, 39, 41, 42, 54] | \n
\nAmblyomma scalpturatum\n | \n\nCanis familiaris\n\n7\n\n, Sus scrofa\n\n7\n\n | \n\nAgouti paca\n\n7\n\n, Dasyprocta fuliginosa\n\n3\n\n, Didelphis marsupialis\n\n5\n, Pecari tajacu\n\n5\n\n,\n\n7\n, Tapirus terrestris\n\n5\n\n,\n\n7\n, Hydrochoerus hydrochaeris\n\n7\n\n | \nAM3, MT\n5\n, RO\n7\n\n | \n[30, 31, 32, 38, 88] | \n
\nAmblyomma sculptum\n | \n\nEquus caballus\n\n,4,7,9\n | \n\nMyrmecophaga tridactyla\n\n9\n\n | \nMA\n4\n, RO\n7\n, TO\n9\n\n | \n[88] | \n
\nAmblyomma varium\n | \n\n | \nBradypus tridactylus\n\n3\n\n, Bradypus sp\n\n3\n\n, Bradypus variegatus\n\n7\n\nCholeopus didactylus\n\n3\n\n\n,\n\n\n7\n\n, Choleopus hoffmanni\n\n3\n\n\n,\n\n\n7\n\n, Choloepus sp\n7\n\nDasyprocta aguti\n\n3\n\n, Tamandua tetradactyla\n\n3\n\n | \nAM\n3\n, PA\n6\n, RO\n7\n\n | \n[30, 31, 33, 38, 40, 97] | \n
\nDermacentor nitens\n | \n\nBos taurus\n\n4\n, Canis familiaris\n\n4\n\n,\n\n7\n, Equus caballus\n\n,4,7,8,9\nCapra hircus\n\n4\n, Ovis aires\n\n4\n\n | \n\nAgouti paca\n\n7\n, Mazama gouazoubira\n\n9\n\n | \nMA\n4\n, RO\n7\n, RR\n8\n, TO\n9\n\n | \n[36, 38, 39, 95, 98] | \n
\nIxodes amarali\n | \n\n | \nHylaeamys megacephalus\n\n6\n, Monodelphis glirina\n\n6\n, Monodelphis touan\n\n6\n, | \nPA\n6\n\n | \n[99] | \n
\nIxodes bocatorensis\n | \n\n | \nBradypus tridactylus\n\n3\n, Cyclopes didactylus\n\n3\n, Dasyprocta leporina\n\n3\n, Tamandua tetradactyla\n\n3\n\n | \nAM\n3\n\n | \n[16] | \n
\nIxodes spinosus\n | \n\n | \nDasyprocta sp\n7\n, Myoprocta pratti\n\n1\n\n | \nRO7, AC\n1\n\n | \n[13] | \n
\nIxodes lasallei\n | \n\n | \nDasyprocta variegata\n\n7\n\n | \nRO\n7\n\n | \n[16] | \n
\nIxodes luciae\n | \n\n | \nDidelphis marsupialis\n\n1\n\n\n,\n\n\n3\n\n\n,\n\n\n4\n\n\n,\n\n\n7\n, Marmosa murina\n\n6\n, Marmosa demerarae\n\n3\n, Monodelphis sp\n7\n, Oecomys sp.\n7\n, Orysomys sp.\n7\n\n, Oligorysomys sp.\n7\n, Philander opossum\n\n6\n\n | \nAC\n1\n, AM\n3\n, MA\n4\n, PA\n6\n, RO\n7\n\n | \n[31, 36, 38, 100] | \n
\nIxodes schulzei\n | \n\n | \nMonodelphis touan\n\n6\n\n | \nPA\n6\n\n | \n[92] | \n
\nHaemaphysalis juxtakochi\n | \n\nCanis familiaris\n\n4\n\n,\n\n7\n, | \n\nMazama americana\n\n3\n\n,\n\n9\n, Mazama p\n7\n, Dasyprocta azarae\n\n5\n; Dasyprocta fuliginosa\n\n3\n, | \nAM\n3\n, MA\n4\n, MT\n5\n; RO\n7\n, TO\n9\n\n | \n[31, 36, 38, 39, 42] | \n
\nRhipicephalus microplus\n | \n\nBos taurus\n\n4\n\n,\n\n7\n\n,\n\n8\n\n,\n\n9\n\n, Canis familiaris\n\n4\n\n,\n\n7\n\n,\n\n9\n\n, Felis catus\n\n7\n, Equus caballus,\n\n4\n\n,\n\n7\n\n,\n\n8\n\n,\n\n9\n\n, Capra hircus\n\n4\n\n,\n\n7\n\n, Ovis aires\n\n4\n\n | \n\nAlouatta puruensis\n\n7\n, Mazama gouazoubira\n\n5\n\n,\n\n9\n, Mazama sp.\n7\n, Tapirus terrestres\n\n7\n\n,\n\n9\n, Tayassu sp\n7\n\n | \nMA\n4\n, MT\n5\n; RO\n7\n, RR\n8\n; TO\n9\n\n | \n[32, 33, 36, 38, 39, 95, 98] | \n
\nRhipicephalus sanguineus sensu lato | \n\nCanis familiaris\n\n1\n\n,\n\n3\n\n,\n\n4\n\n,\n\n7\n\n,\n\n9\n\n | \n\nBradypus tridactylus\n\n3\n, Galictis cuja\n\n4\n, Saquinus bicolor\n\n3\n\n | \nAC\n1\n, AM\n3\n, MA\n4\n, RO\n7\n\n | \n[30, 31, 32, 33, 36, 38, 39, 95, 98, 101] | \n
Records of ticks in the Amazon biome, Brazil, according to tick species, hosts (domestic and wild animals), states, references.
Acre – AC.
Amapá – AP.
Amazonas – AM.
Maranhão – MA.
Mato Grosso – MT.
Pará – PA.
Rondônia – RO.
Roraima – RR.
Tocantins – TO.
In general, the greatest diversity of ticks was recorded on T. tetradactyla with 10 species, followed by the rodent H. hydrochaeris with nine species. These records indicates the importance of these hosts for maintaining local tick diversity, in addition to act as dispersers of these ectoparasites. Additional hosts have also been shown to be important in maintaining diversity of tick in the Amazon: Tapirus terrestris, Tayassu pecari, Pecari tajacu, Agouti paca, Dasyprocta fuliginosa, Bradypus tridactylus, Bradypus variegatus, Panthera onca, Nasua nasua, Choleopus didactylus, Choloepus hoffmanni, Cyclopes didactylus, Dasyprocta azarae, Dasyprocta leporina, Philander opossum, M. glirina, M. touan, Mazama gouazoubira, Mazama americana and M. tridactyla (Table 3), because they are parasitized by more than one species of ticks. There is also a record of A. cajennense s.s. on Pteronura brasiliensis a semi-aquatic animal [89].
\nInterestingly, three species of ticks commonly found on domestic animals have also been found parasitizing wild animals: Rhipicephalus sanguineus sensu lato, R. microplus and D. nitens. Although occasional, the encounter of these species of ticks on wild animals is possible, especially when wild animals coexist with domestic animals infested by ticks (e.g., pastures, corrals, houses). A total of 11 species of ticks have been recorded parasitizing domestic animals in Amazon (Table 3).
\nSpotted fevers caused by R. rickettsii and Rickettsia parkeri are currently the only confirmed tick-borne disease affecting humans in the country, and A. sculptum, A. aureolatum, A. ovale are the main vectors. The disease is severe and highly lethal when caused by the bacterium R. rickettsii (vectors A. sculptum and A. aureolatum) and moderate, non-lethal, when caused by R. parkeri, vectored mainly by A. ovale [10]. Only A. sculptum and A. ovale have been recorded in the Brazilian Amazon, the first in rare reports and the second frequently recorded and populations established in this biome. In fact, considering only the tick adult stage, A. ovale has been reported as the most common human-biting tick in the Amazon [38].
\nIn the last century, papers about ticks on human beings have been published in the Brazilian Amazon, however these studies are fragmented and scarce. At least 14 species of hard ticks have already been found and documented parasitizing humans within the limits of the Brazilian Amazon. Of these, the vast majority (11 species) belong to the genus Amblyomma: A. cajennense s.s., A. coelebs, A. dissimile, A. latepunctatum, A. naponense, A. oblongoguttatum, A. ovale, A. romitii, A. rotundatum, A. scalpturatum and A. sculptum. The other three species belong to the genus Rhipicephalus, including R. microplus and R. sanguineus s.l., and the genus Haemaphysalis with only H. juxtakochi. The public health importance of human–tick associations in the Amazon biome is unclear and further research are needed to clarify the issue.
\nNymphs of H. juxtakochi have been reported on humans in the states of Rondônia (municipality not precisely indicated) and Amazonas, in the municipality of Santa Isabel do Rio Negro [31, 38]. H. juxtakochi has been found infected with the bacterium Rickettsia rhipicephali in the Amazon region, more precisely in the municipalities of Monte Negro and Confresa, states of Rondônia and Mato Grosso, respectively [42, 93]. Human parasitism by R. microplus was expected, since this species can parasitize humans when it comes into direct contact with domestic cattle, as reported by [38]. Therefore, its importance is much more economical, causing severe losses to livestock in the country than for public health.
\nDespite sporadic records of R. sanguineus s.l. on humans in Brazil, this association in the Brazilian Amazon requires confirmation. Records of approximately 4,020 R. sanguineus s.l. (larva, nymph, and adult), supposedly collected on humans, in the state of Pará by Serra-Freire et al. [103] do not corroborate the common findings in country. The bioecology of this endophilic and introduced tick is well studied. In other countries, R. sanguineus s.l. is vector of some zoonotic agents for humans (Rickettsia conorii, Rickettsia massiliae and R. rickettsii) [83]. Adults of A. cajennense s.s. have been reported on humans in the municipality of Monte Alegre (Pará state) [90]. Reports of Amblyomma cajennense sensu lato parasitizing humans published by Martins et al. [90] in the municipalities of Sinop and Tucuruí in the states of Mato Grosso and Pará, is possibly A. cajennense s.s. taking into account the area of occurrence of this species. This tick is aggressive to humans. In the Amazon biome the R. bellii bacterium was detected in this species in Mato Grosso state and R. amblyommatis in Mato Grosso, Maranhão and Rondônia state [32, 42, 53, 83, 88, 90, 91, 104, 105].
\nNymphs and adult of A. coelebs were found on humans in Rondônia state [38]. An adult of this tick was found on human in Roraima state, municipality of Caroebe [106]. The nymph and adult stages of A. colelebs were also found on humans in Amazonas state, in the municipalities of Coari, Fonte Boa, Presidente Figueiredo and Santa Isabel do Rio Negro [31, 107]. There are reports of infection by R. amblyommatis in A. coelebs in the Amazon region in the states of Rondônia and Mato Grosso, respectively [32, 53, 88]. Adult of A. dissimile was recorded on humans in the Maicurú River/Amazon biome, located in Pará state [106]. Adults of the species A. latepunctatum have been reported on humans in Coari and Santa Isabel do Rio Negro, both municipalities located in the state of Amazonas [31].
\nAdults of the tick A. naponense (published as Amblyomma mantiquirense) were found on humans in the state of Pará (municipality not reported) [108]. Human parasitism also by adults was later reported on the Maicurú River, located in this same state [106]. However, nymphs have been found on humans in the states of Rondônia (municipality not specified precisely) and Amazonas, in the municipalities of Coari and Santa Isabel do Rio Negro [31, 38]. The bacteria R. bellii and Rickettsia sp. strain PA were identified in this species in the Amazon region of the municipalities of Santarém and Rurópolis, in the state of Pará [42]. Rickettsia sp. strain Tapirapé was found in this tick in the municipalities of Confresa and Rio Branco, in the states of Mato Grosso and Acre, respectively [42, 54].
\nAdults of the species A. oblongoguttatum were recorded parasitizing humans in the state of Pará, in the Maicurú River and in the municipality of Uruará [106, 108, 109]. Human parasitism by nymphs and adults of this tick was recorded in the state of Rondônia (municipality not specified with precision) [38]. However, the nymphal stage of this species was later recorded on humans in the municipality of Monte Negro in the same state [106]. Human parasitism by adult of this tick was also recorded in the municipality of Caroebe, state of Roraima [106]. Nymphs and adults of this species have been recorded parasitizing humans in three municipalities (Coari, Manacapuru and Santa Isabel do Rio Negro), all located in the state of Amazonas [31, 106, 107]. On the other hand, the life cycle of this tick has already been studied under laboratory conditions [110]. The bacteria R. bellii and R. amblyommatis were detected in this species in the Amazon in the municipalities of Governador Jorge Teixeira and Pimenta Bueno, in the state of Rondônia [53, 105].
\nAdults of the tick A. ovale have been reported to parasitize humans in several areas of the state of Rondônia [38]. As previously reported in this chapter, adults of A. ovale parasitize mainly domestic and wild carnivores, while immature (larva and nymph) parasitize rodents of the families Cricetidae and Echimyidae, with sporadic reports on wild birds [17, 75, 83]. This preference for such host groups was observed in the laboratory [111]. This tick is a vector of the protozoan Hepatozoon canis and the bacterium R. parkeri, agents of importance in veterinary and human medicine, respectively [83]. In the Amazon region, the bacterium R. bellii was identified in this species in the municipalities of Governador Jorge Teixeira and Monte Negro, in the state of Rondônia; and in the municipality of Cururupu, in the state of Maranhão [53, 91].
\nThe adult stage of the species A. romitii (published as Amblyomma tasquei) was found parasitizing humans in the north of the state of Pará (municipality not reported) [108]. In this same state, the larval and adult stages of this tick were found on humans in the municipality of Rurópolis [96, 112]. The life cycle of this tick was studied under laboratory conditions [113].
\nParasitism by adults of A. rotundatum was recorded in the municipality of Belém in the state of Pará on a human who worked in frequent contact with reptiles in the Zoobotanical Park of the Museu Paraense Emílio Goeldi [114].
\nAdults of the A. scalpturatum have been reported to parasitize humans in the state of Pará (municipality not reported) [108], and in the Maicurú River located in this state [93, 106]. Human parasitism by the nymphal and adult stages of this species has been reported in the states of Mato Grosso (municipality of Jauru) and Rondônia (municipality not precisely indicated), however in this latter state it was not specified whether these adult ticks were fixed or walking on humans [38, 93]. The nymphal stage was also collected from a human who was probably infested with this tick in the municipality of Porto Velho in the state of Rondônia [115]. The nymph and adult stages of this species were also found parasitizing humans in the state of Amazonas, in the municipalities of Coari, Fonte Boa, Jutaí, Presidente Figueiredo and Santa Isabel Rio Negro [31, 107]. In the Amazon biome, two species of rickettsiae (R. bellii and R. amblyommatis) were detected in A. scalpturatum, in the municipalities of Governador Jorge Teixeira and Sinop, states of Rondônia and Mato Grosso, respectively [53, 88].
\nHuman parasitism by adults of A. sculptum was recorded in the municipality of Jauru in the state of Mato Grosso [90]. The Jauru territory comprises 90% of the Amazon biome and 10% of the Cerrado biome, so this human record in this region is in accordance with the transition area of the occurrence of this species in sympatry with A. cajennense s. s. [90]. The bioecology of this native tick has been extensively studied due to its aggressive parasitism towards domestic animals and humans in the Brazilian territory. In the geographical area in which this species occurs, it completes an annual generation, with larvae occurring mostly during autumn, nymphs during winter, and adults during spring and summer, with larval behavioral diapause being the main regulating factor of its life cycle [116, 117]. Besides being considered a pest for domestic animals (dogs, cattle and horses), the species can transmit to humans the Brazilian Spotted Fever caused by the bacterium R. rickettsii, being this the most important zoonotic disease transmitted by ticks within the limits of the National territory. Additionally, A. sculptum has also been identified with the bacterium R. amblyommatis in the municipality of Pium, state of Tocantins [118].
\nThere is a trend of seasonal behavior for some species of Amblyomma in the Amazon biome, with immatures predominating in the dry period (June to October), and adults in the rainy period (October to March) [119]. Similar to the pattern observed for A. sculptum in areas outside the Amazon biome [116].
\nTaxonomy of Argasidae is currently questionable and relies on five schemes that divide the family in up to ten genera [120]. Achieving a consensus between soft tick taxonomists depends now chiefly on the molecular characterization of early collected type specimens for some genera. For instance, elucidating the status of pivotal taxa such as Alectorobius is mandatory if we are to understand the systematics of the Argasidae, particularly in the American Continent. Considering a practical approach, in this chapter we adopt but not necessarily endorse the classification of soft ticks into five genera, namely: Antricola, Argas, Nothoaspis, Ornithodoros, and Otobius [121].
\nThe fauna of argasid ticks in Brazil is currently composed by 24 species [15]. Ticks of this family parasitize terrestrial vertebrates including amphibians in this country [25, 26]. With the exception of larvae from an undetermined Ornithodoros sp. collected on Potus flavus (Procionidae) [84], reports of soft ticks parasitizing mammals in the Brazilian Amazon are few and almost restricted to larvae collected on bats. Adults and nymphs have been collected either inside bat-inhabited caves or over massive rock formations where bats shelter (Table 4).
\nSpecies | \nState | \nLocality | \nMilieu of collection/Host | \nReference | \n
---|---|---|---|---|
\nAntricola delacruzi\n | \nRondônia | \nPorto Velho | \nBat inhabited cave/Pteronotus spp. | \n[122] | \n
\nAntricola guglielmonei\n | \nRondônia | \nPorto Velho | \nBat inhabited cave/Pteronotus spp | \n[122] | \n
\nNothoaspis amazoniensis\n | \nRondônia | \nPorto Velho | \nBat inhabited cave/Pteronotus parnellii\n | \n[123] | \n
\nNothoaspis setosus\n | \nRondônia | \nMonte Negro | \nRock formation/ \nMolossus molossus\n | \n[124] | \n
\nOrnithodoros cavernicolous\n | \nRondônia | \nMonte Negro | \nBat inhabited caves/Unknown | \n[125] | \n
\nOrnithodoros hasei\n | \nAmapá | \nunknown | \nMistneted bat Artibeus planisrostris\n | \n[87] | \n
\n | Pará | \nMarajó Island | \nBat-inhabited three hole/Unknown | \n[126] | \n
\nOrnithodoros kohlsi\n | \nRondônia | \nMonte Negro | \nMistneted bat Molossops mattogrossensis\n | \n[30] | \n
\nOrnithodoros marinekellei\n | \nRondônia | \nPorto Velho | \nBat inhabited cave/Pteronotus spp | \n[127] | \n
\n | Pará | \nCarajás, Paraupebas, Canaã dos Carajás | \nBat inhabited caves/Unknown | \n[128] | \n
\nOrnithodoros peropteryx\n | \nRondônia | \nMonte Negro | \nBat inhabited cave/Unknown | \n[125] | \n
\nOrnithodoros rondoniensis\n | \nRondônia | \nPorto Velho | \nBat inhabited cave/Unknown | \n[122] | \n
\n | Pará | \nCarajás, Paraupebas, Canaã dos Carajás | \nBat inhabited caves/Unknown | \n[128] | \n
\nOrnithodoros stageri\n | \nRondônia | \nUnknown | \nUnknown/Noctilio albiventris, Nyctinomops laticaudatus\n | \n[129] [11] | \n
Soft ticks and their respective hosts occurring in the Brazilian Amazon.
The first record of a soft tick in the Brazilian Amazon was published by North American entomologists Robert A. Cooley and Glen M. Kohls back in 1941. They received a tick collected inside a bat-inhabited three hole at Marajó Island (Pará state) and identified it as female of Ornithodoros hasei (mentioned as Ornithodoros dunni) [126]. Nowadays, we know that O. hasei is a wide spread species in Brazil and that three-roosting bats Artibeus planirostris and Noctilio spp. could act as main hosts [87, 130, 131, 132]. With the exception of recent collection of larvae on A. planirostris [87], knowledge on the distribution of O. hasei along the Brazilian Amazon is still poorly vague.
\n\nTadarida laticaudata were the first bats reported to be parasitized by soft ticks in the Brazilian Amazon [133]. This report referred to Ornithodoros setosus, which was recently reclassified as Nothoaspis setosus [124]. In 1972, the bats Noctilio labialis and T. laticaudata were reported to be parasitized by Ornithodoros stageri in the Brazilian Amazon [129], which was recently confirmed [11]. A report in the Brazilian Amazon is a remarkable fact for O. stageri, since it also has distribution in Southern United States and Mexico [126, 134].
\nBat inhabited caves constitute excellent niches to find argasid ticks. In particular, special, large colonies of insectivorous bats dwelling inside small chambers create high temperature conditions (28-40°C) where hundreds of Antricola, Nothoaspis and Ornithodoros ticks might occur [135]. Between 2004 to 2010 collections of ticks performed in hot caves from Porto Velho (Rondônia) fostered the description of two novel species (e.g., Nothoaspis amazoniensis and Ornithodoros rondoniensis) [122, 123]; the redescription of larvae and description of postlarval stages of Ornithodoros marinkellei [127], and the expansion of geographical distribution of Antricola delacruzi and Antricola guglielmonei into Brazilian Amazon [122]. At least for two species, O. marinkellei and O. rondoniensis, further collections performed in caves from Pará state underpined a larger distribution along the Amazon ecosystems [128].
\nMost amazing feature of soft ticks inhabiting hot caves, is that adaptation to this particular milieu seems to have modified their morphology and biology drastically. For instance, evidence showing that adults of Antricola ticks lack the capacity to digest blood has been gained after transcriptomic analyses of their saliva [136]. In fact, scoop-like short mouth parts suggest that adults of Antricola do not suck blood [137]. Moreover, adults of Antricola, O. marinkellei and O. rondoniensis possess huge spiracular plates [122, 127, 137], perhaps necessary to thrive in such hot and extremely humid environments.
\nExcept for larvae of Ornithodoros kohlsi collected on the bat Molossops mattogrossensis [30], prospections performed by our group between 2016 and 2019 focused mainly in the search for soft ticks inside natural cavities over massive rock formations. During these expeditions, Ornithodoros cavernicolous and Ornithodoros peropteryx were collected for the first time in caves from Monte Negro (Rondônia), extending their distribution of both species to the Brazilian Amazon [125]. Furthermore, analyses performed on large larvae isolated from Molossus molossus bats clarified that O. setosus was incorrectly classified in its original description. Indeed, O. setosus matches morphologically and molecularly within the genus Nothoaspis therefore the statement of N. setosus n. comb. was proposed [124]. It is important to note that our last collections performed at Monte Negro included several morphotypes of Ornithodoros pending formal description. Consequently, the fauna of soft ticks occurring in the Brazilian Amazon is likely to increase soon.
\nTicks parasitize a wide variety of vertebrates around the world such as amphibians, reptiles, mammals and birds, including humans. Although there are a variety of studies of parasitism by ticks on animals and humans in the different ecoregions of Brazil, in the Amazon biome they are scarce and fragmented. Because of this, it is possible to infer that the diversity of ticks in the Brazilian Amazon is underestimated. In the Amazon, amphibians and reptiles were important hosts for A. rotundatum, A. dissimile and A. humerale, hard ticks common in these hosts in other regions of the country. Birds and mammals, on the other hand, proved important for the maintenance and dispersion of over 30 species of hard ticks in the Amazon, but there are still many gaps between hosts and their ticks in this region. Interestingly, immature A. humerale appears to be frequent on wild birds in this biome. No soft ticks have been reported parasitizing amphibians, reptiles, birds and/or non-flying mammals (except for a single record of Ornithodoros sp. on P. flavus), possibly due to scarcity of more studies in this biome. Some studies show that bats seem to have an important role spreading soft tick populations along Amazonian caves and that the diversity described for this group is still very poor. Two vector species of spotted fevers were found in the Amazon biomes parasitizing humans (A. sculptum and A. ovale). However, to date, there are no reports of BSF in humans in the region. Finally, it is of paramount importance that researchers (acarologists and epidemiologists) direct their attention to the Amazon biome, in order to fill the numerous existing gaps in the diversity of ticks in Brazil and prevent possible outbreaks of diseases transmitted by these ectoparasites to animals and humans.
\nNo conflict of interest declared.
Supporting women in scientific research and encouraging more women to pursue careers in STEM fields has been an issue on the global agenda for many years. But there is still much to be done. And IntechOpen wants to help.
",metaTitle:"IntechOpen Women in Science Program",metaDescription:"Supporting women in scientific research and encouraging more women to pursue careers in STEM fields has been an issue on the global agenda for many years. But there is still much to be done. And IntechOpen wants to help.",metaKeywords:null,canonicalURL:null,contentRaw:'[{"type":"htmlEditorComponent","content":"At IntechOpen, we’re laying the foundations for the future by publishing the best research by women in STEM – Open Access and available to all. Our Women in Science program already includes six books in progress by award-winning women scientists on topics ranging from physics to robotics, medicine to environmental science. Our editors come from all over the globe and include L’Oreal–UNESCO For Women in Science award-winners and National Science Foundation and European Commission grant recipients.
\\n\\nWe aim to publish 100 books in our Women in Science program over the next three years. We are looking for books written, edited, or co-edited by women. Contributing chapters by men are welcome. As always, the quality of the research we publish is paramount.
\\n\\nAll project proposals go through a two-stage peer review process and are selected based on the following criteria:
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\\n\\nAdvantages of Publishing with IntechOpen
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\n\nWe aim to publish 100 books in our Women in Science program over the next three years. We are looking for books written, edited, or co-edited by women. Contributing chapters by men are welcome. As always, the quality of the research we publish is paramount.
\n\nAll project proposals go through a two-stage peer review process and are selected based on the following criteria:
\n\nPlus, we want this project to have an impact beyond scientific circles. We will publicize the research in the Women in Science program for a wider general audience through:
\n\nInterested? If you have an idea for an edited volume or a monograph, we’d love to hear from you! Contact Ana Pantar at book.idea@intechopen.com.
\n\n“My scientific path has given me the opportunity to work with colleagues all over Europe, including Germany, France, and Norway. Editing the book Graph Theory: Advanced Algorithms and Applications with IntechOpen emphasized for me the importance of providing valuable, Open Access literature to our scientific colleagues around the world. So I am highly enthusiastic about the Women in Science book collection, which will highlight the outstanding accomplishments of women scientists and encourage others to walk the challenging path to becoming a recognized scientist." Beril Sirmacek, TU Delft, The Netherlands
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USA, CRC Press Taylor & Francis, Asia Pacific, Trans Tech Publications Ltd., Switzerland, and Materials Science Forum, USA. 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Omar obtained\nhis Bachelor degree in electrical and\nelectronics engineering from Universiti\nSains Malaysia in 2002, Master of Science in electronics\nengineering from Open University\nMalaysia in 2008 and PhD in optical physics from Universiti\nSains Malaysia in 2012. His research mainly\nfocuses on the development of optical\nand electronics systems for spectroscopy\napplication in environmental monitoring,\nagriculture and dermatology. He has\nmore than 10 years of teaching\nexperience in subjects related to\nelectronics, mathematics and applied optics for\nuniversity students and industrial engineers.",institutionString:null,institution:{name:"Universiti Sains Malaysia",country:{name:"Malaysia"}}},{id:"191072",title:"Prof.",name:"A. K. M. Aminul",middleName:null,surname:"Islam",slug:"a.-k.-m.-aminul-islam",fullName:"A. K. M. Aminul Islam",position:null,profilePictureURL:"https://mts.intechopen.com/storage/users/191072/images/system/191072.jpg",biography:"Prof. Dr. A. K. M. 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