Santiago Alarcon Et Al2012DipteraVectorsOfAvianHaemosporidianParasites

8/13/2019 Santiago Alarcon Et Al2012DipteraVectorsOfAvianHaemosporidianParasites

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Biol. Rev. (2012),87, pp. 928964. 928doi: 10.1111/j.1469-185X.2012.00234.x

Diptera vectors of avian Haemosporidian

parasites: untangling parasite life cycles

and their taxonomy

Diego Santiago-Alarcon1,2,, Vaidas Palinauskas3 and Hinrich Martin Schaefer2

1Biologa y Conservacion de Vertebrados, Instituto de Ecologa, A.C., carretera antigua a Coatepec 351, Xalapa, C.P. 91070, Mexico2Department of Ecology and Evolutionary Biology, Faculty of Biology I, Hauptstr. 1, University of Freiburg, Freiburg 79104, Germany3 Institute of Ecology, Nature Research Center, Akademijos 2, Vilnius 2100, LT-08412, Lithuania

ABSTRACT

Haemosporida is a large group of vector-borne intracellular parasites that infect amphibians, reptiles, birds, andmammals. This group includes the different malaria parasites (Plasmodiumspp.) that infect humans around the world.Our knowledge on the full life cycle of these parasites is most complete for those parasites that infect humans and, to someextent, birds. However, our current knowledge on haemosporidian life cycles is characterized by a paucity of informationconcerning the vector species responsiblefor their transmission among vertebrates. Moreover, our taxonomic and system-atic knowledge of haemosporidians is far from complete, in particular because of insufficient sampling in wild vertebratesand in tropical regions. Detailed experimental studies to identify avian haemosporidian vectors are uncommon, with onlya few published during the last 25 years. As such, little knowledge has accumulated on haemosporidian life cycles duringthe last three decades, hindering progress in ecology, evolution, and systematic studies of these avian parasites. Nonethe-less, recently developedmolecular tools have facilitated advances in haemosporidian research. DNAcan now be extractedfrom vectors blood meals and the vertebrate host identified; if the blood meal is infected by haemosporidians, theparasites genetic lineage can also be identified. While this molecular tool should help to identify putative vector species,

detailed experimental studies on vector competence are still needed. Furthermore, molecular tools have helped to refineour knowledge on Haemosporida taxonomy and systematics. Herein we review studies conducted on Diptera vectorstransmitting avian haemosporidians from the late 1800s to the present. We also review work on Haemosporidataxonomyand systematics since the first application of molecular techniques and provide recommendations and suggest futureresearch directions. Because human encroachment on natural environments brings human populations into contact withnovel parasite sources, we stress that the best wayto avoid emergent andreemergent diseases is through a program encom-passing ecological restoration, environmental education, and enhanced understanding of the value of ecosystem services.

Key words: Haemosporida, malaria, Plasmodium, Haemoproteus, Leucocytozoon, Diptera, Culicidae, Ceratopogonidae,Hippoboscidae, Simuliidae, insect vector.

CONTENTS

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 929(1) Haemosporidian parasites and their vectors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 929(2) Methods for the study of haemosporidian parasites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 930

II. Diptera vectors transmitting avian haemosporidian parasites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 931(1) Family Culicidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 931

(a) Taxonomic issues for parasites transmitted by Culicidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 943(2) Family Hippoboscidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 943

(a) Taxonomic issues for parasites transmitted by Hippoboscidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 945

* Address for correspondence (Tel: +55 228 842 1800 ext. 4135; Fax: +55 228 818 60 09; E-mail: [email protected];

[email protected]).

Biological Reviews87 (2012) 928964 2012 The Authors. Biological Reviews 2012 Cambridge Philosophical Society


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Avian haemosporidian vectors and taxonomy 929

(3) Family Simuliidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 945(a) Taxonomic issues for parasites transmitted by Simuliidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 946

(4) Family Ceratopogonidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 946III. Advances since the first application of molecular methods (PCR) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 953IV. Future directions in understanding the ecology and evolution of haemosporidian parasites . . . . . . . . . . . . . . . . 955

(1) A current limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 955(2) The dynamic nature of haemosporidian transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 956(3) Taxonomy and systematics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 957

V. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 957VI. Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 958

VII. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 958

I. INTRODUCTION

(1) Haemosporidian parasites and their vectors

Vector-borne parasites of the order Haemosporida (PhylumApicomplexa) are commonly found in amphibians, reptiles,birds, and mammals (Valkiunas, 2005). Avian haemosporid-

ian parasites have a cosmopolitan distribution and aredivided into four genera: Plasmodium, Haemoproteus, Fallisiaand Leucocytozoon (Atkinson & van Riper, 1991; Valkiunas,2005). The four genera of haemosporidian parasites havesimilar life cycles with differences during the asexual phase inthe peripheral blood of their vertebrate host.PlasmodiumandFallisia can undergo asexual reproduction in the peripheralblood of the host (i.e. merogony), whereas this is not the casein the other parasite genera (Valkiunas, 2005). Plasmodiumspecies are considered to be highly virulent, particularly inimmunologically nave hosts (van Riper et al., 1986; Atkin-sonet al., 1995, 2000; Yorinks & Atkinson, 2000; Palinauskaset al., 2008, 2009, 2011).Leucocytozoon is known to be highly

pathogenic in poultry such as turkeys and ducks (Valkiunas,2005). Haemoproteus was considered to be relatively benignwithout causing serious harm to their hosts (Atkinson, 1991;Bennett, 1993; Bennett, Peirce & Earle, 1994), but morerecent evidence demonstrates thatHaemoproteuscan have sig-nificant impacts and some species are highly virulent andlethal (Nordlinget al., 1998; Merinoet al., 2000; Marzalet al.,2005; Valkiunas, 2005).

Avian haemosporidians use dipteran insect vectors dur-ing their sexual and sporogonic phases (Garnham, 1966;Valkiunas, 2005; Martinsen, Perkins & Schall, 2008). Plas-modium species are known to be transmitted by severalspecies of mosquitoes (Culicidae) from different genera (Gar-

nham, 1966; Atkinson & van Riper, 1991; Valkiunas, 2005;Kimura, Darbro & Harrington, 2010);Haemoproteusis knownto be transmitted by several species of hippoboscid and cer-atopogonid flies (Baker, 1957; Atkinson, 1991; Valkiunas,Liutkevicius & Iezhova, 2002; Valkiunaset al., 2010a);Leuco-cytozoonspecies are known to be transmitted by simuliid flies(Malmqvistet al., 2004; Valkiunas, 2005; Hellgren, Bensch& Malmqvist, 2008) and ceratopogonid flies for the sub-genus Akiba (Akiba, 1960; Morii, Kitaoka & Akiba, 1965;Morii & Kitaoka, 1968), and there is some evidence thatFallisia parasites are transmitted by mosquitoes (Gabaldon,Ulloa & Zerpa, 1985). However, insect vectors represent

the life-cycle stage of haemosporidian parasites that we least

understand; further studies areneededinto the ecological andevolutionary dynamics of these host-parasite-vector systems

particularly the inclusion of blood-feeding dipterans from thefamilies Ceratopogonidae, Simuliidae, and Hippoboscidae.

There are three taxonomic issues relevant to a discussionof haemosporidian parasite genera and their vectors. First,

the validity of the Plasmodium subgenera Novyella andGiovannolaiawas questioned by Corradetti & Scanga (1965)and recently by Martinsen, Paperna & Schall (2006); there

are several Plasmodium species whose subgeneric positionis difficult to identify, particularly at low parasitemia.

Second, the genus Haemoproteus was divided into twogroups (the H. columbae group and the H. nettionis group)(Baker, 1963); these two groups later were named as

the subgenera Haemoproteus and Parahaemoproteus (Bennett,Garnham & Fallis, 1965; Valkiunas, 2005). They are well

classified in terms of the vectors transmitting them and theirmorphological features, but it is currently unknown if they

are monophyletic or paraphyletic (Outlaw & Ricklefs, 2011).Third, the genus Leucocytozoon was reclassified into two generaLeucocytozoon and Akiba (Bennett et al., 1965) because theywere transmitted by different Diptera families, but recentlythey were reclassified as one genus with two subgenera

(Valkiunas, 2005).Leucocytozoonand Akiba, unlikePlasmodiumand Haemoproteus, are apparentlyable to metabolize the wholehaemoglobin molecule, and no pigment, i.e. haemozoin, is

left undigested. We suggest that when taxonomic descriptionsare expanded to include competent vector species and

sporogonic morphological traits, many of the currenttaxonomic problems will be resolved.

Because parasite species of the genusPlasmodiumrepresent

a constant threat to human health, most studies on haemo-sporidian parasites have been directed at species infecting

humans (Escalante & Ayala, 1994; Escalante, Barrio & Ayala,1995; Escalante et al., 1998; Hughes & Verra, 2001; Joyet al., 2003; Paul, Ariey & Robert, 2003; Rich et al., 2009;Prugnolle et al., 2010). This results in a high number of

vector studies focusing on a few species (52 anopheline

spp. worldwide; Enayati & Hemingway, 2010) of mosquitoes(Diptera: Culicidae) transmitting human malaria parasites

(e.g. Dong, Manfredini & Dimopoulos, 2009; Muenwornet al., 2009; Muturi et al., 2009; Swain et al., 2009). Fur-thermore, there is abundant research on the control and



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lineages when using DNA sequences versusmorphospecieswhen using microscopy), which might lead to different con-clusions. For example, a cophylogenetic analysis is likelyto show a different cospeciation history between parasitesand their hosts when using genetic lineages (which wouldbe more like a cophylogeographic history) compared withmorphological species. In addition, microscopy studies carrya risk of clumping cryptic species together (e.g. Sehgal et al.,2006; Valkiunaset al., 2010b).

Field and laboratory experimental infection studies wereused to identify Diptera vectors transmitting haemosporidianparasites from the end of the 19th Century to the end ofthe 20th Century (e.g. Huff, 1927; Atkinson, Greiner &Forrester, 1983). Emphasis changed dramatically in recent

years, however, and now most studies use the PCR method

to identify vector feeding preferences (e.g. Imuraet al., 2010)and sporogonic stages in salivary glands (e.g. Valkiunaset al., 2010a). Whereas experimental infection studies canprovide definitive statements about vectorial capacity, PCRstudies can only identify natural putative vectors for thehaemosporidian parasites under study, leaving unclear the

vectorial capacity, as PCR methods cannot prove that thelife cycle of the parasite will be completed. Thus, ideallya combination of both experimental infection and PCRmethods should be used in order to determine the natural

vectors of haemosporidian parasites.This review aims (i) to review the work that has been

done on dipteran vectors transmitting avian haemosporidianparasites since the early 1900s; (ii) to review the work doneon the taxonomy and systematics of avian haemosporidianssince the first application of molecular techniques; and (iii)to integrate this knowledge in order to suggest areas where

additional research is needed.

II. DIPTERA VECTORS TRANSMITTING AVIANHAEMOSPORIDIAN PARASITES

(1) Family Culicidae

By the end of the 1800s it was believed that there was onlyone species of avian malaria parasite; this subsequently wasproved incorrect (Huff, 1927; Herman, 1938). R. Ross firstdiscovered the involvement of a mosquito in the transmissionof an avian Plasmodium parasite in 1897 (Ross, 1898; Cox,

2010 reviewed the history of this discovery); the involve-ment of an Aedes mosquito was first described by Koch(1899) (Aedes communis; Huff, 1927, 1932a). Since then, birdmalaria parasites attracted the attention of researchers as anexperimental model for the investigation of human malaria;they were so used by many laboratories until the discoveryof rodent malaria parasites in 1948 (Killick-Kendrick, 1974)and the successful infection of theAotus trivirgatusmonkey withhuman malaria parasites in 1966 (Young, Porter & Johnson,1966). Rodent and monkey malaria parasites are closerto human malaria in numerous ways (Mulligan & Sinton,1933); hence bird haemosporidians became less attractive

for human malaria research. Despite that, they remain con-venient model organisms for investigations into the general

biology ofPlasmodiumspp. and their close relatives, includingquestions of the evolution of haemosporidians.

The list of discoveries and achievements using avianmalarial models includes new antimalarial drugs (Davey,

1951; Coatney et al., 1953), the first cultivation methods oftissue and erythrocytic stages in vitro (Trager, 1950; Ball &Chao, 1961), and the first steps in the development of anantimalarial vaccine (McGhee, Singh & Weathersby, 1977).

Ball (1964) described a technique for the cultivation of thesporogonic cycle ofPlasmodium relictum: it was possible togrowP. relictum in vitrofrom the gametocyte stage to infectiveSporozoites.

At the beginning of the 20th Century many studiesattempted to survey the different genera and species ofmosquitoes in order to identify possible competent vectors for

Plasmodium parasites (Huff, 1927, 1932a, 1965; Reichenow,1932; Raffaele, 1934; Herman, 1937; Coggeshall, 1940;

Laird, 1941; Hurlbut & Hewitt, 1942; Jeffrey, 1944; Man-well, 1947; Micks, 1949; Corradetti & Scanga, 1965; Niles,

Fernando & Dissanaike, 1965; Garnham, 1966). Such exper-imental infections often used canaries as a model system, and

vector competence was tested mostly using mosquitoes fromthe generaCulexandAedes(Huff, 1927; Herman, 1938; Man-well, 1940; Laird, 1941; Jeffrey, 1944; Micks, 1949). Someresearchers used other bird species, such as ducks, pheasants,

pigeons, and chickens, in experimental infections in responseto the high mortality and infection refraction of some bird

species to somePlasmodium parasites (Coatney, 1938; Laird,1941; Jeffrey, 1944; Micks, 1949; Becker, 1961). By 1907 it

was proven that mosquitoes of two genera Culex and Aedes

were vectors of several avian malaria parasites (Ross, 1898;Koch, 1899; Ruge, 1901; Daniels, 1899; Sergent & Sergent,1907). Subsequently, additional species of Culex and Aedesand other genera were recognized as competent vectors inthe transmission of avian malaria. By 1918 it was known

that birdPlasmodiumcould be transmitted by seven mosquitospecies of three genera [Culex,Aedes, and Culiseta(as Theobaldia)(Huff, 1927)]. Huff (1927) investigated vector susceptibility ofanothernine mosquito species andtwo new genera(PsorophoraandAnopheles) infected with three Plasmodium spp. From these,only two new species (Culex territans and Cx. salinarius) wereadded to the list of susceptible vectors of which only Cx. sali-nariuswas competent at transmittingPlasmodium cathemerium

(Huff, 1927). Reichenow (1932) discovered that mosquitoes ofthe genusCulisetatransmitPlasmodium circumflexum.Corradetti& Scanga (1965) successfully transmitted Plasmodium polareusingCuliseta longiareolata. Nileset al. (1965) describedCoquil-lettidia(as Mansonia)crassipesas a natural vector ofPlasmodiumgallinaceum. Mayne (1928) showed for the first time thatmalaria vectors of the genus Anopheles, which were consid-ered only to transmit human malaria, might also transmit

avian malaria. Later Coggeshall (1940) in experiments withPlasmodium lophurae proved that Anopheles quadrimaculatus iscapable of transmitting parasites other than human malarias(i.e. Plasmodium falciparum, P. vivax, and P. malariae). All these



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932 D. Santiago-Alarcon and others

studies showed that Plasmodium parasites are able to use manyvector species during the transmission cycle, and that some

vectors are capable of feeding on different vertebrate hosts,some of which are not closely related.

At present, more than 20 species of anopheline mosquitoesare known in which sporogony of avian malaria parasites

is completed (Valkiunas, 2005). Species of the genusAnopheles susceptible to avian malaria belong to threesubgenera: Anopheles, Nyssorhynchus and Cellia (as Myzomyia)(Huff, 1965). Anopheles mosquitoes are known to be viable

vectors ofP. cathemerium, P. gallinaceum, P. lophurae, P. fallax,and P. relictum (Huff, 1965; Valkiunas, 2005). There wasno evidence until 1937 that mosquitoes ofCulex and Aedescould be vectors of human malaria parasites. Williamson &

Zain (1937) fed Cx. bitaeniorhynchus on patients with mixedinfections of P. vivax and P. falciparum, and of P. malariaeand P. falciparum. Sporozoites of all tested malaria speciesdeveloped in the salivary glands of laboratory-bred Culexmosquitoes, but the infectivity of the sporozoites was nottested on human subjects. Warren & Wharton (1963) showedthat subspecies ofP. cynomolgiform Oocysts in Culex,AedesandMansoniamosquitoes. However, the development of Oocystsdoes not mean that the insects were able to transmit malaria

parasites. Even though there was no confirmation thatP. falciparumandP. vivaxwere able to complete their life cycleafter Culex transmission, it remains possible that culicinesare competent vectors for human Plasmodium species. Inreptiles, Klein, Young & Telford (1987a) demonstrated thatCx. erraticus is a competent vector for P. floridense, a lizardparasite. Ayala (1971) showed that P. mexicanum (a lizardmalaria parasite) completed its sporogonic cycle in two

sandfly species [Lutzomyia vexator(as Lutzomyia vexatrix occidentis,

but subspecies not recognized to date) andLutzomyia stewarti].Later, Lutzomyia vexator was confirmed to be a competentvector for P. mexicanum by Klein et al. (1987b). Petit et al.(1983) showed that Plasmodium agamaedeveloped partially (theOocysts developed sporozoites, but these never reached the

salivary glands) in Culicoides nubeculosus (Ceratopogonidae).Table 1 shows the results of experimental and natural

infections of Culicidae vectors for different avian Plasmodiumspecies (see also Huff, 1965, for a comprehensive review of

mosquitoes susceptibility to species of avian Plasmodium).There are very few studies testing the ability of avian

Plasmodium parasites to develop in bloodsucking vectorsother than Culicidae. There could be other vector families

transmiting Plasmodium parasites, such as in some lizardPlasmodiumspecies as described above.

Species of Culicidae have a heterogeneous specificity formalaria parasites. Some Culicidae species are competent

for only a small number of Plasmodium species, whereasothers seem to be generalists (Huff, 1927, 1932a; Herman,1938; Laird, 1941; Jeffrey, 1944; Manwell, 1947). Forexample, Coggeshall (1940) showed that An. quadrimaculatus(a vector of human Plasmodium parasites in the US) canbe infected by both a monkey parasite (P. cynomolgi) andan avian parasite (P. lophurae). Furthermore, specificity of aparasite can be altered by artificial selection that increases or

decreases susceptibility of the vector to infection (Huff, 1929;Trager, 1942; Micks, 1949), and adaptation of an avian

parasite (P. lophurae) to a mammal host has been observedfollowing a few infectious injections (as few as four infectious

passages) of infected avian cells into infant mice at differenttime steps (McGhee, 1951). Moreover, it is also possible

to modify the host range of species of Culicidae underexperimental conditions; for instance,Culex apicalisis knownnaturally to feed on cold-blooded vertebrates, but is able toutilize canary bloodunder experimental conditions (Herman,

1938). This suggests that it is possible for vectors to feed onalternative suboptimal hosts, providing an opportunity for

parasites to switch hosts across distantly related vertebrates(e.g. Santiago-Alarcon et al., 2012). The time frame andstability of a vector-parasite-vertebrate interaction couldlead to tight coevolution, and thus, to a specific interaction.However, such specific interactions could vary according to

geographical location, producing a mosaic of coevolutionary

interactions, where the parasite, vector, or vertebrate hostmay change depending on local environmental conditions(Huff, 1938; Thompson, 2005). Ecological factors such as

spatial heterogeneity and phenology may be important indetermining which parasite species are transmitted by which

vectors (Huff, 1938; Applegate & Beaudoin, 1969). Evenwhen a vector is experimentally susceptible to a Plasmodiumparasite, if in nature they do not have overlapping niches theywill never meet unless a change in environmental conditions

allow them to come into contact. The genetic variationwithin haemosporidian parasites might be another factor

determining the specificity or generality of an insect vector.A parasite with a larger spectrum of genetic lineages could

be more likely to be transmitted by a larger array of vector

species, whereas a parasite with low genetic variation is likelyto be transmitted only by vectors to which it is alreadyadapted.

Environmental conditions, especially temperature, mayalso be important factors for the development of different

malarial parasites in mosquitoes. Grassi (1900) showed thattemperatures as low as 15.5 and 17.5

C for nine days were

lethal for P. vivaxand P. falciparum, suggesting that mosquitoesexposed to such conditions immediately after feeding would

not develop an infection. By contrast, Sergent (1919) showedthat exposure of P. relictum to 12 C for 6 h immediatelyafter vector feeding did not interfere with the developmentof the parasite and sporozoites were formed. James (1926)

reported that oocysts and sporozoites ofP. vivaxare able toresist temperatures of 45.5

C for three weeks, or below

freezing temperature for six days. Chao & Ball (1961) showedthat temperatures as low as 4

C are not immediately lethal

to P. relictum infecting Culex tarsalis, even if exposure to thecold temperature occured as soon as 15 min after biting and

continued for 48 h. Oocysts can be exposed to temperaturesas low as 4

C for 23 days and will still retain the ability

to develop after the host is returned to beneficial conditions(Ball & Chao, 1961). Taken together, these studies suggest

that there is considerable variation in the ability ofPlasmodiumspecies to withstand low temperatures.



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Table1.Experimentalstudiesdemonstratingfullorpartial(partofthelifecyclehasnotbeendemonstratede

xperimentally)vectorialcompetenceofdifferentspeciesof

CulicidaetransmittingparasitesofthegeneraPlasmodiumandFallisia.InformationnotobtainedfromtheprimaryliteraturewasacquiredfromGarnham(1966)andValkiunas

(2005)

PCRmethod

Parasitespecies

Vector

species

Proven

experimentally

Proven

natura

l

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCR

on

vectorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

Plasmodiumrelictum

Culexquinquefasciatus(asCx.

pipiensfatigansinold

literature)

+

+

Sporozoite

India,USA

Ross(1898),Daniels(1899),

Huff(1927),andRosen&

Reeves(1954)

Cx.p

ipiens

+

NA

Sporozoite

Germany,

Algeria,

USA,

Columbia

Ruge(19

01),Sergent&

Sergent(1907),Neumann

(1908),Huff(1927),Tate&

Vincent(1934),and

Hunninen(1951,1953)

Cx.hortensis

+

NA

Sporozoite

Algeria

Sergent&Sergent(1918)

Cx.territans

Oocyst

USA

Huff(1927)

Cx.salinarius

Oocyst

USA

Huff(1927)

Cx.tarsalis

+

+

Sporozoite

USA

Huff(1932a),Hermanetal.

(1954),andRosen&

Reeves(1954)

Cx.s

tigmatosoma

+

+

Sporozoite

USA

Herman

etal.(1954)and

Rosen

&Reeves(1954)

Cx.b

itaeniorhynchus

+

Sporozoite

India

Russell&

Mohan(1942)and

Singh

&Mohan(1955)

Cx.gelidus

Oocyst

India

Russell&

Mohan(1942)

Cx.t

heileri

Oocyst

India

Russell&

Mohan(1942)

Cx.w

hitmorei

Oocyst

India

Russell&

Mohan(1942)

Lutziafuscanus(asCx.fuscanus

inoldliterature)

+

Sporozoite

Philippines

Nono(1932)

Culisetaannulata

(asTheobaldia

annulatainoldliterature)

Oocyst

Germany

Reichenow(1932)

Cs.longeareolata(asTheobaldia

spathipaoposinoldliterature)

+

NA

NA

Algeria


Aedescommunis(asCx.nemorosus

inoldliterature)

+

NA

Sporozoite

Germany

Koch(18

99)

Ae.aegypti

+

NA

Sporozoite

Algeria,

USA

Sergent&Sergent(1907,

1918)andHuff(1927)

Ae.mariae

+

NA

NA

Algeria


Ae.dorsalis

Oocyst

USA

Rosen&

Reeves(1954)

Ae.vexans

Oocyst

USA

Rosen&

Reeves(1954)

Anophelescrucian

s

+

NA

Sporozoite

USA

Hunnine

n(1951)

An.freeborni

+

NA

Sporozoite

USA

Hunnine

n(1951)

An.quadrimaculatus

+

NA

Sporozoite

USA

Hunnine

n(1951,1953)

An.subpictus

+

Sporozoite

India

Mayne(1928)



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Table1.(Cont.)

PCRmethod

Parasitespecies

Vector

species

Proven

experimentally

Proven

natura

l

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCR

on

vectorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

An.a

lbimanus

+

NA

Sporozoite

Panama

Hunnine

n(1951,1953)

P.cathemerium

Cx.salinarius

NA

Oocyst

USA

Huff(1927)

Cx.territans

Oocyst

USA

Huff(1927)

Cx.p

ipiens

NA

Oocyst

USA

Huff(1927)andHerman

(1938)

Cx.quinquefascia

tus

Oocyst

USA,Japan

Huff(1927),Tanaka(1946),

andM

icks&McCollum

(1953)

Cx.tarsalis

Oocyst

USA

Huff(1932a)

Cx.b

itaeniorhynchus

+

Sporozoite

Japan

Tanaka(1946)

Cx.tritaeniorhynchus

Oocyst

Japan

Tanaka(1946)

Ae.aegypti

Oocyst

USA

Huff(1927)

Ae.sollicitans

+

NA

NA

USA

Herman

(1938)

Cs.melanura(as

Cs.melaneumin

Valkiunas,2

005)

+

NA

NA

USA

Herman

(1938)

An.quadrimaculatus

Oocyst

USA

Micks&

McCollum(1953)

An.norestensis

Oocyst

Brazil

Barreto(1943)

Lutziafuscanus(asCulex

fuscanusinoldliterature)

Oocyst

Philippines

Nono(1932)

P.gallinaceum

Ae.aegypti

+

NA

Sporozoite

France,

India,USA,

Nigeria,

Japan

+

+

+

Brumpt(1935,1936),Russell

&Mohan(1942),Cantrell

&Jord

an(1949),Okpala

(1958),Weathersby(1962),

Huff(1965),andKimetal.

(2009a

)

Ae.a

lbopictus

+

NA

Sporozoite

France,

India,USA,

Japan

Brumpt(1935,1936),Russell

&Mohan(1942),Cantrell

&Jord

an(1945),and

Weath

ersby(1962)

Ae.geniculatus

+

NA

Sporozoite

France

Roubaudetal.(1939)

Ae.lepidus

+

NA

Sporozoite

Brazil

Paraense

(1945)

Ae.pseudotaeniatus

+

Sporozoite

India

Russell&

Mohan(1942)

Ae.pseudalbopictus

+

Sporozoite

India

Russell&

Mohan(1942)

Ae.scutellaris

+

Sporozoite

India

Russell&

Mohan(1942)

Ae.unilineatus

+

Sporozoite

India

Russell&

Mohan(1942)

Ae.v

ittatus

+

Sporozoite

India

Russell&

Mohan(1942)

Ae.c

hrysolineatus

+

Sporozoite

India

Russell&

Mohan(1942)

Armigereskuchingensis

+

Sporozoite

India

Russell&

Mohan(1942)



8/37


Table1.(Cont.)

PCRmethod

Parasitespecies

Vector

species

Proven

experimentally

Proven

natura

l

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCR

on

vectorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

Ar.o

bturbans

+

Sporozoite

India

Russell&

Menon(1942)

Ae.triseriatus

+

Sporozoite

USA

Cantrell

&Jordan(1945)

Ae.campestris

+

NA

Sporozoite

USA

Cantrell

&Jordan(1945)

Ae.cantator

+

NA

Sporozoite

USA

Cantrell

&Jordan(1945)

Ae.s

timulans

+

NA

Sporozoite

USA

Cantrell

&Jordan(1945)

Ae.a

tropalpus

+

NA

Sporozoite

USA

Trembley(1946)

Ae.s

tokesi

+

Sporozoite

Nigeria

Okpala(1958)

Ae.japonicus

+

NA

Sporozoite

Japan

Weathersby(1962)

Ae.togoi

+

NA

Sporozoite

Japan

Weathersby(1962)

Ae.a

lbolatoralis

Oocyst

India

Russell&

Menon(1942)

Ae.canadensis

+

Sporozoite

USA

Cantrell

&Jordan(1949)

Ae.jamesi

Oocyst

India,USA

Russell&

Menon(1942)and

Cantrell&Jordan(1945)

Ae.pallirostris

+

Sporozoite

India

Russell&

Mohan(1942)

Ae.trivittatus

Oocyst

USA

Cantrell

&Jordan(1945)

Ae.vexans

Oocyst

USA

Cantrell

&Jordan(1945)

Cs.inornata(asTheobaldia

innoratainoldliterature)

Oocyst

USA

Cantrell

&Jordan(1945)

An.quadrimaculatus

+

NA

Sporozoite

USA

Haas&Akins(1947)and

Cantrell&Jordan(1949)

An.freeborni

+

NA

Sporozoite

USA

Eyles(19

60)

An.a

lbimanus

Oocyst

USA

Eyles(19

60)

Ar.aureolineatus

+

Sporozoite

India

Russell&

Mohan(1942)

Ar.annulipalpis

+

Sporozoite

India

Russell&

Menon(1942)

Ar.magnus

Oocyst

India

Russell&

Menon(1942)

Ar.subalbatus

+

NA

Sporozoite

Japan

Weathersby(1962)

Cx.quinquefascia

tus

+

Sporozoite

Mexico

Vargas&

Beltran(1941)

Cx.m

imeticus

Oocyst

India

Russell&

Menon(1942)

Cx.salinarius

Oocyst

USA

Cantrell

&Jordan(1945)

Cx.tarsalis

Oocyst

USA

Huff(1965)

Cx.p

ipienspallen

s

NA

NA

Japan

+

+

+

Kimetal.(2009a)

Coquillettidiacrassipes(as

Mansoniacrassipesinold

literature)

+

+

Sporozoite

SriLanka

Nilesetal.(1965)

Cq.perturbans(asM.perturbans

inValkiunas,2005)

Oocyst

USA

Cantrell

&Jordan(1945)

P.matutinum

Cx.p

ipiens[asC

x.fatigans

(quinquefasciatus)inHuff,

1937]

NA

Oocyst

USA

Huff(1937)andManwell

(1940,1947)

Cx.s

tigmatosoma

+

NA

Sporozoite

Italy

Corradettietal.(1962)



9/37


Table1.(Cont.)

PCRmethod

Parasitespecies

Vector

species

Proven

experimentally

Proven

natura

l

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCR

on

vectorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

Cx.tarsalis

+

Sporozoite

Italy


P.subpraecox

Cx.p

ipiens

+

NA

Sporozoite

Italy

Raffaele

(1932)

P.giovannolai

Cx.p

ipiens

+

NA

Sporozoite

Italy

Corradettietal.(1963a,b)

P.fallax

Ae.a

lbopictus

+

Sporozoite

USA

Huffetal.(1950)andHuff

(1965)

Ae.aegypti

+

Sporozoite

USA

Huffetal.(1950)andHuff

(1965)

Ae.a

tropalpus

+

Sporozoite

USA

Huffetal.(1950)

Ae.triseriatus

+

Sporozoite

USA

Huffetal.(1950)

An.quadrimaculatus

+

Sporozoite

USA

Huffetal.(1950)

Cx.quinquefascia

tus

+

Sporozoite

USA

Huffetal.(1950)

Cx.tarsalis

+

Sporozoite

USA

Huff(1965)

P.polare

Cs.longiareolata

+

NA

Sporozoite

Italy

Corradetti&Scanga(1965)

Cs.morsitans

+

+

Sporozoite

Canada

Meyer&

Bennett(1976)

Cq.perturbans(asM.perturbans

inValkiunas,2005)

+

Sporozoite

Canada

Meyer&

Bennett(1976)

P.lophurae

Ae.aegypti

NA

Oocyst

USA

Coggeshall(1940)andJeffrey

(1944)

Ae.a

lbopictus

+

NA

Sporozoite

USA

Laird(19

41),Jeffrey(1944),

andH

uffetal.(1947)

Ae.a

tropalpus

+

Sporozoite

USA

Laird(19

41)

An.quadrimaculatus

+

NA

Sporozoite

USA

Coggeshall(1940),Hurlbut&

Hewitt(1942),andJeffrey

(1944)

Cx.p

ipiens

Oocyst

USA

Coggeshall(1940)

Cx.restuans

+

Sporozoite

USA

Laird(19

41)

P.durae

Cx.antennatus

NA

+

Sporozoite

Africa

Valkiuna

s(2005)

Cx.p

ipiens

NA

NA

Sporozoite

Africa

Valkiuna

s(2005)

Cx.univittatus

NA

NA

Sporozoite

Africa

Valkiuna

s(2005)

P.garnhami

Cx.p

ipiens

+

NA

Sporozoite

Egypt

Garnham

(1966)

P.circumflexum

Cx.tarsalis

Oocyst

USA

Huff(1965)

Cs.annulata

+

Sporozoite

Germany,

Italy

Reichenow(1932)and


Cs.melanura

+

NA

NA

USA

Herman

(1938)

Cs.longiareolata

+

Sporozoite

Italy


Cs.morsitans

+

+

Sporozoite

Canada

Meyeret

al.(1974)andMeyer

&Ben

nett(1976)

Cq.crassipes

+

+

Sporozoite

SriLanka

Nilesetal.(1965)

Cq.perturbans

+

Sporozoite

Canada

Meyer&

Bennett(1976)

Ae.sollicitans

+

NA

USA

Herman

(1938)



10/37


Table1.(Cont.)

PCRmethod

Parasitespecies

Vectorspecies

Proven

experimentally

Proven

natural

vector

M

ostadvanced

developmental

stageobserved

invector

Region

PCRon

vectorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

R

eferences

P.vaughani

Cx.p

ipiens

+

Sporozoite

Italy

Corradetti&

Scanga(1972)

Cs.morsitans

+

+

Sporozoite

Canada

Williams&Bennett(1978)

Cq.perturbans

+

Sporozoite

Canada

Williams&Bennett(1978)

P.rouxi

Cx.p

ipiens

+

Sporozoite

Algeria,

USA

Huff(1932a)andManwell(1947)

Cx.tarsalis

Oocyst

USA

Huff(1932a)

Cx.territans

Oocyst

USA

Huff(1932a)

P.kempi

Cx.p

ipiens

+

NA

Sporozoite

USA

Christensen

etal.(1983)

Cx.restuans

+

Sporozoite

USA

Christensen

etal.(1983)

Cx.tarsalis

+

NA

Sporozoite

USA

Christensen

etal.(1983)

P.elongatum

Ae.triseriatus

+

Sporozoite

USA

Huff(1930)andMicks(1949)

Cx.p

ipiens

+

+

Sporozoite

USA,Italy

Huff(1927),andRaffaele(1934),

andMicks(1949)

Cx.restuans

Oocyst

USA

Micks(1949

)

Cx.tarsalis

+

NA

Sporozoite

USA

Huff(1932a)andHuff&

Shiroishi(1962)

Cx.territans

Oocyst

USA

Huff(1927)

Cx.salinarius

Oocyst

USA

Huff(1927)

Cx.quinquefasciatus

Oocyst

Italy

Raffaele(1934)

P.hermani

Cx.salinarius

+

+

Sporozoite

USA

Nayaretal.(1981b)

Cx.n

igripalpus

+

+

Sporozoite

USA

Youngetal.(1977),Forrester

etal.(1980),andNayaretal.

(1981b,1982)

Cx.restuans

+

NA

Sporozoite

USA

Nayaretal.(1981a)

Wyeomyiavandu

zeei

+

NA

Sporozoite

USA

Nayaretal.(1980,1981b)

P.juxtanucleare

Cx.gelidus

+

NA

Sporozoite

Malaysia

Bennettetal.(1966)

Cx.quinquefasciatus

+

Sporozoite

Brazil

Paraense(1944)

Cx.p

ipiens

+

NA

Sporozoite

Japan

Akiba(1959

)

Cx.pseudovishnu

i

+

NA

Sporozoite

Malaysia

Bennettetal.(1966)

Cx.s

itiens

+

+

Sporozoite

Malaysia

Bennettetal.(1966)andBennett

&Warren

(1966)

Cx.tritaeniorhynchus

+

NA

Sporozoite

Malaysia

Bennettetal.(1966)

Cx.annulus

+

+

Sporozoite

Malaysia

Bennettetal.(1966)

Fallisianeotropicalis

Aedeomyiasquam

ipennis

+

+

NA

Venezuela

Gabaldonet

al.(1985)

Plasmodiumlineage

LIN1p

Cx.s

itiens

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Cx.annulirostris

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Plasmodiumlineage

LIN2

Ae.hebrideus

+

NA

Vanuatu

+

Ishtiaqetal.(2008)



11/37


Table1.(Cont.)

PCRmethod

Parasitespecies

Vecto

rspecies

Proven

experimentally

Proven

natura

l

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCRon

ve

ctorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

PlasmodiumlineageLIN3p

Ae.notoscriptus

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

P.juxtanuclearelineage

LIN4p

Cx.annulirostris

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Haemoproteuslineage

LIN5h

An.farauti

+

NA

Vanuatu

+

Ishtiaqetal.(2008)

Cx.quinq

uefasciatus

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Ae.v

igilax

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Cq.xanth

ogaster

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Haemoproteuslineage

LIN6h

An.farauti

+

NA

Vanuatu

+

Ishtiaqetal.(2008)

Ae.hebrid

eus

+

NA

Vanuatu

+

Ishtiaqetal.(2008)

Ae.a

lternans

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Ae.vexans

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Verrallina

lineata

+

NA

Vanuatu

+

Ishtiaqetal.(2008)

Ae.aegypti

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Cx.s

itiens

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Cx.annulirostris

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Cx.quinq

uefasciatus

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Ae.v

igilax

+

NA

New

Caledonia

+

Ishtiaqetal.(2008)

Plasmodiumlineage

PlasCoq1

Cq.aurites

+

NA

Cameroon

+

Njaboetal.(2009)

Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)

Plasmodiumlineage

PlasCoq2

Cq.aurites

+

NA

Cameroon

+

Njaboetal.(2009)

Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)

Plasmodiumlineage

PlasCoq3

Cq.metallica

+

NA

Cameroon

+

Njaboetal.(2009)

Cx.spp.

+

NA

Cameroon

+

Njaboetal.(2011)



12/37


Table1.(Cont.)

PCRmethod

Parasitespecies

V

ectorspecies

Proven

experimentally

Proven

natural

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCRon

vectorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

PlasmodiumlineagePlasCoq4

Cq.aurites

+

NA

Cameroon

+

Njaboetal.(2009)

Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cq.aurites

+

NA

Cameroon

+

Njaboetal.(2009)

Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)

HaemoproteuslineagePlasCoq6Cq.aurites

+

NA

Cameroon

+

Njaboetal.(2009)

Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)

Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Mansoniauniformis

+

NA

Cameroon

+

Njaboetal.(2011)


Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)

Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)

PlasmodiumlineagePV3

Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)


Cq.aurites

+

NA

Cameroon

+

Njaboetal.(2009,2011)

Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)

Ae.

mcintoshi

+

NA

Cameroon

+

Njaboetal.(2009,2011)


Cq.aurites

+

NA

Cameroon

+

Njaboetal.(2009)

Cq.pseudoconopas

+

NA

Cameroon

+

Njaboetal.(2009)

Cq.metallica

+

NA

Cameroon

+

Njaboetal.(2009)

Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)

HaemoproteuslineageHaemK1Cq.spp.

+

NA

Cameroon

+

Njaboetal.(2011)

HaemoproteuslineageHaemK2Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)

HaemoproteuslineageHaemK3Cx.

spp.

+

NA

Cameroon

+

Njaboetal.(2011)

PlasmodiumlineageRinshi-1

Cx.

pipienspallens

+

NA

Japan

+

Kimetal.(2009b)

Cx.

sasai

+

NA

Japan

+

Kimetal.(2009a)


Cx.

sasai

+

NA

Japan

+

Kimetal.(2009a)


Cx.

pipienspallens

+

NA

Japan

+

+

Kimetal.(2009b)

Cx.

pipiensmolestus

+

NA

Japan

+

Kimetal.(2009b)

Cx.

sasai

+

NA

Japan

+

Kimetal.(2009a)

Lt.vorax

+

NA

Japan

+

Kimetal.(2009b)


Cx.

pipienspallens

+

NA

Japan

+

Kimetal.(2009b)


Cx.

pipienspallens

+

NA

Japan

+

+

Kimetal.(2009b)



13/37


Table1.(Cont.)

PCRmethod

Parasitespecies

Vec

torspecies

Proven

experimentally

Proven

natura

l

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCRon

ve

ctorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

PlasmodiumlineageYacho-1

Cxpipienspallens

+

NA

Japan

+

Kimetal.(2009b)

PlasmodiumlineagePAN1

Ad.squamipennis

+

NA

Panama

+

Gageretal.(2008)


Cx.ocossa

+

NA

Panama

+

Gageretal.(2008)


Cx.ocossa

+

NA

Panama

+

Gageretal.(2008)


Ad.squamipennis

+

NA

Panama

+

Gageretal.(2008)


Ad.squamipennis

+

NA

Panama

+

Gageretal.(2008)


Ad.squamipennis

+

NA

Panama

+

Gageretal.(2008)


Cx.ocossa

+

NA

Panama

+

Gageretal.(2008)


Ad.squamipennis

+

NA

Panama

+

Gageretal.(2008)


Ad.squamipennis

+

NA

Panama

+

Gageretal.(2008)

Plasmodiumlineagemosquito5

Cx. quinquefasciatus

+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineagemosquito9


+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquito13

Lt.fuscanus

+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquito17

Ae.albopictus

+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquito24


+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquito111


+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquito132


+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquito227


+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquito290

Ma.sp.

+

NA

Japan

+

Ejirietal.(2008)

Plasmodiumlineage

mosquitoZ34

Lt.vorax

+

NA

Japan

+

Ejirietal.(2009)

Plasmodiumlineage

mosquitoS33

Cx.pipiens

+

NA

Japan

+

Ejirietal.(2009)

Plasmodiumlineage

mosquitoZ74

Lt.vorax

+

NA

Japan

+

Ejirietal.(2009)

Plasmodiumlineage

mosquitoZ73

Lt.vorax

+

NA

Japan

+

Ejirietal.(2009)

Plasmodiumlineage

mosquitoZ64

Cx.pipiens

+

NA

Japan

+

Ejirietal.(2009)

Plasmodiumlineage

mosquitoZ83

Lt.vorax

+

NA

Japan

+

Ejirietal.(2009)



14/37


Table1.(Cont.)

PCRmethod

Parasitespecies

Vec

torspecies

Proven

experimentally

Proven

natura

l

vector

Mostadvanced

developmental

stageobserved

invector

Region

PCRon

ve

ctorblood

meal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

PlasmodiumlineageCXPIP01

Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)


Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)


Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)


Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)


Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)


Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)


Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)

PlasmodiumlineageCXRES01Cx.restuans

+

NA

USA

+

Kimuraetal.(2010)


+

NA

USA

+

Kimuraetal.(2010)


+

NA

USA

+

Kimuraetal.(2010)


+

NA

USA

+

Kimuraetal.(2010)


+

NA

USA

+

Kimuraetal.(2010)

PlasmodiumlineageE1

Cx.restuans

+

NA

USA

+

Kimuraetal.(2010)

Plasmodiumlineage

SEIAUR01/F1

Ae.ca

nadensis

+

NA

USA

+

Kimuraetal.(2010)

Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)

Cx.restuans

+

NA

USA

+

Kimuraetal.(2010)

PlasmodiumlineageLINN1

Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)

Cx.restuans

+

NA

USA

+

Kimuraetal.(2010)

P.vaughanilineageSYAT05

Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)

Cx.restuans

+

NA

USA

+

Kimuraetal.(2010)

PlasmodiumlineageTUMIG3

Cx.pipiens

+

NA

USA

+

Kimuraetal.(2010)

Cx.restuans

+

NA

USA

+

Kimuraetal.(2010)

Plasmodiumlineage

PADOM11

Cx.restuans

+

NA

USA

+

Kimuraetal.(2010)

TheseparasitelineagesbelongtothegenusHaemoproteus,whicharesuppo

sedtobeexclusivelytransmittedbylouseflies(Diptera:Hippoboscidae)andbitingmidges(Diptera:

Ceratopogonidae)(seeTables2and4).

Provennaturalvector=

sporogonic

cycleiscompletedinvectorandtransmissionofparasitestoavianhostwasconductedeitherbyvectorbites,byinjectionofaslurryoftriturated

infectedvectors,orbysalivarygland

scontainingSporozoites;acompletemerogoniccyclewasobserved.Onlyvectors

naturallypresentinthestudyareawere

usedinexperiments.

Provenexperimentally=

laboratory

[includingpolymerasechainreaction(PCR)studies]and/orfieldexperimentsha

vebeenconductedonvectorcompetenc

e,experimentscould

havebeenconductedusingvectorsp

eciesnotoccurringatthestudysite(i.e.unnaturalvectors),andinsomecasescomp

letelifecyclehasnotbeenproved.NA,n

otavailable.



15/37


Initial studies on vector specificity showed that it was notpossible to identify broad patterns of infection because of

the paucity of knowledge on genetic variation. The fact thatlarge numbers ofPlasmodiumsporozoites develop in an insect

vectors salivary glands, or that large parasite inoculums areused to infect birds, does not mean that the parasite will be

successfully transmitted or that an infection will develop inthe vertebrate host (Huff, 1927, 1929; Laird, 1941; Jeffrey,

1944; Micks, 1949). Vector susceptibility depends on theparasite strain used during experimental infections (Man-

well, 1940; Micks, 1949). Some authors described parasitesof the same species with no difference in morphology as hav-

ing distinct physiological characteristics (Huff, 1927, 1929,1932a, 1938). For example, P. elongatum was not able toinfect Culex pipiens in a study by Reichenow (1932), but itdid develop in 30% ofC. pipiens when an Italian strain ofthis parasite was used (Raffaele, 1934). Furthermore, Huff(1927) demonstrated that some individuals and/or varieties

of the same competent vector species have different efficien-cies at transmittingP. cathemerium (Huff, 1927, 1929, 1938)indicating the presence of intraspecific variation in vectorial

capacity (see also Tate & Vincent, 1934). Similarly, a moresusceptibleAe. aegyptistrain was selected by Trager (1942) forP. lophuraeinfection and aCx. pipiensstrain forP. elongatumbyMicks (1949). Thus, differences in vector susceptibility could

be explained by both vector and parasite genetic diversity.Recent findings using molecular approaches suggest that

there could be more than 1000 avian haemosporidian strains(Bensch, Hellgren & Perez-Tris, 2009), whereas only about

220 morphological species of avian haemosporidians havebeen described (Valkiunas, 2005; Valkiunas et al., 2010a).Clearly, vertebrate host-parasite-vector interactions will have

a geographical component that must be taken into accountwhen making epidemiological and model predictions.Natural immunity of mosquitoes and vertebrate hosts to

malaria has received considerable attention. Manwell (1938)proved that immunity of a susceptible species of mosquito

might be due to its inheritance. Weathersby (1965) describeda parabiotic twinning method to determine immune mech-

anisms of susceptible and refractory mosquitoes to malariainfections. This involved pairing two mosquitoes (one of a

susceptible and one of a refractory species) using a tiny capil-lary glass, so that haemolymph could be shared between the

two species. Using this method it was shown that there wereenough nutritive elements for development of ookinetes and

early oocyst stages ofP. gallinaceum in refractory Cx. pipiens,but due to some antagonistic action oocysts appeared dis-

torted in immune mosquitoes, while inAe. aegyptimosquitoesoocyst development was successful (Weathersby & McCall,

1968). It was proven later that the innate immunity ofCx.pipiensmosquitoes toP. gallinaceuminfection is due to antago-nistic (antiblastic) factors in mosquito haemolymph and notto the lack of metabolites or required substances (Weath-

ersby & McCroddan, 1982). However, it now appears thatthese innate immunity mechanisms are not generalisable

across species. Alaviet al. (2003) injectedP. bergheiookinetesinto Ae. aegyptihaemocoel without successful development

of the initial stages of oocysts. Inhibitory agents such asthe peritrophic membrane barrier, oxygen free radicals,

melanization and others act throughout the developmentof parasites to block their transmission stages (Billingsley &

Rudin, 1992; Sinden, 2002). Sinden, Alavi & Raine (2004)commented that the refractoriness of mosquitoesmay depend

on the level of inhibitory barriers at each developmental stageand that only when all inhibiting mechanisms fail to block

development can the insect be considered a competent orviable vector. Recently, it was discovered that the mid-gutmicrobiota (bacteria) plays a critical role in the vectorial

capacity to sustain Plasmodium infections; mosquitoes with anormal associated microbiota had a lower infection rate com-

pared to microbe-free mosquitoes (Donget al., 2009). Thissuggests that within-vector microbe interactions represent a

natural immunological barrier to infection, resulting from themodulation of the mosquitos immune genes by the mid-gut

microbial flora (Donget al., 2009). Results of studies on para-site specificities and vector and vertebrate immunitiessupport

Huff, Marchbank & Shiroishis (1959) hypothesis that infec-tivity of the parasite is the result of various factors such as indi-

vidual variability of the immune response, behavioral traitsof the host, genetic diversity of the host and parasite popula-

tions,and coevolutionary history between parasites and hosts.To evaluate the effect of malaria parasites on inverte-

brate hosts, studies have measured the longevity of infectedmosquitoes or flight performance variables (flight speed,

length of initial flight or longest flight). Huff (1965) statedthat the age of the adult mosquito and its nutritional state

might affect its susceptibility to parasite infection. Initialstudies showed no apparent reduction in vitality or longevity

of anopheline mosquitoes infected with P. vivaxeven when

females were heavily infected (King, 1929; DeBuck, 1936).These results were corroborated by other studies (Mayne,1920; Wenyon, 1926; Boyd, 1940; Ragab, 1958). Freier &

Friedman (1987) stated that the relationship ofP. gallinaceumtoAe. aegyptiwas that of a commensal rather than parasitic,however this is at odds with evidence cited below reportingnegative fitness consequences of parasite infection in CulexandAnophelesspp. It remains to be investigated whether vari-ation in tolerance of mosquitoes to infection depends on the

specific species and strains used. By contrast, Buxton (1935)showed that infection with P. relictum led to an increase inmortality in Culex fatigans (now Cx. quinquefasciatus) and thiswas corroborated by Maier (1973), who showed a correlation

between increased death rate in Cx. pipiens fatigans (now Cx.quinquefasciatus) and infection with P. cathemerium. The greatestmortality occurred during the period of ookinete penetrationof the midgut (Maier, 1973). Gad, Maier & Piekarski (1979)

observed higher mortality in An. stephensi during the firstthree days after infection with Plasmodium berghei. A detailedstudy by Klein et al. (1982) using Plasmodium cynomolgi andAn. dirusshowed that mortality peaks coincided with oocystrupturing and later with penetration of salivary gland cellsby sporozoites. A positive correlation between the severity

of infection and a reduction in flight capability was reportedby Schiefer, Ward & Eldridge (1977) who suggested that



16/37


negative effects on vectors were due to decreases in levelsof available carbohydrates, which are crucial for flight. Arecent publication by Ferguson & Read (2002) questionedwhy the impact of malarial parasites on their invertebratehost is still unresolved. They used a meta-analysis to showthat malaria parasites reduce vector survival, but showedthis effect to be due to the use of unnatural vector-parasite

combinations in experimental studies, to the length of theexperiment (e.g. experiments that end before the invasion ofsalivary glands by sporozoites are less likely to find fitnesseffects or behavioural changes in infected mosquitoes), andto the lack of standardized experimental conditions.

Despite avianPlasmodiumbeing the best understood genusof haemosporidian parasites in terms of their transmission

vectors, there are still manyPlasmodiumspecies for which thevector species is unknown, and even for well-studied Plasmod-ium parasites new vectors previously believed to be unsuitablearestill found (Huff, 1965; Garnham, 1966; Valkiunas,2005).If we add to the lack of knowledge on competent vectorsthe fact that the outcome depends on local environmental

conditions and geographic origin (i.e. subspecies, strains orlineages) both of the parasites and hosts, then the inherent dif-ficulty in making definitive statements about the specificityof haemosporidian genera to particular Diptera familiesbecomes clear. Studies on haemosporidian parasites and

vector competence and susceptibility would benefit greatlyfrom the inclusion of a geographic coevolutionary framework(Thompson, 2005) in their design (e.g. Kimura, Dhondt &Lovette, 2006).

(a) Taxonomic issues for parasites transmitted by Culicidae

Corradetti & Scanga (1965) stressed difficulties with the

identification of the systematic position ofPlasmodium polare.They described P. polare as falling between the subgeneraGiovannolaia and Novyella; the final reason for placement ofP. polarein the subgenus Giovannolaiawas the quantity of cyto-plasm. There were also difficulties with attributingPlasmodiumoctamerium to the subgenus Giovannolaia (Manwell, 1968). Itwas pointed out(Manwell, 1968) that this parasite is similar tospecies of the subgenusNovyellain themorphology of its bloodstages. P. octamerium was attributed to the subgenus Novyellain the Garnham collection of malaria parasites (Garnham& Duggan, 1986). However, after examination of the typematerial, P. octamerium was placed in the subgenus Giovannolaiaby Valkiunas (2005), reverting to Manwells (1968) original

systematic position. Strictly speaking, P. octamerium belongs toa group of species (together at least with P. dissanaike), whosesubgeneric position is difficult to identify, particularly at lowparasitemia.

(2) Family Hippoboscidae

The oldest record of transmission of a haemosporidian para-site by hippoboscid flies was by Aragao (1908) working withHaemoproteus columbae(Adie, 1915). This parasite infects thecommon pigeonColumba livia. By 1906 a fly from the genusLynchia was implicated in the transmission of H. columbae,

the first studies used the species Lynchia maura (originallydescribed as Olfersia maura, now Pseudolynchia canariensis) andLynchia brunea lividicolor (originally described as Olfersia lividi-color, now Pseudolynchia canariensis) (Sergent & Sergent, 1906;

Adie, 1915). The role of these flies in transmission was con-firmed by later observers, including Wenyon (1926), Huff

(1932b), Coatney (1933, 1935), and Mohammed (1958). By1911 it was believed that H. columbae developed up to theookinete stage in the insect vector and proceeded no further;hence, researchers thought that the ookinete was the stage

inoculated into the pigeon (Minchin, 1912, cited in Adie,1915). A few years later the complete development of the

parasite in Lynchiaflies was demonstrated (Adie, 1915). Unlikemosquitoes and ceratopogonid vectors, both males and

females of hippoboscid flies take blood meals, are susceptibleto infection, and are able to transmit the parasite; moreover,

infection rates of Lynchia flies can be up to 100% (Adie,1915). Rendtorff, Jones & Coatney (1949) conducted experi-

ments onH. columbaeusingP. canariensisas vector; they foundthat vectors feeding on birds with patent infections developed

sporozoites only if the bird was infected for 25 days or longer;birds with patent infections of less than 25 days most likelyhad immature gametocytes, which precluded the develop-

ment of the sexual and sporogonic stages in the vector.Early studies working on host specificity of Haemopro-

teus parasites suffered from taxonomic confusion and lackof knowledge on genetic variability. Baker (1957) noted

that many hippoboscid species used in earlier studies weresynonyms ofP. canariensis, and that this fly was the onlyknown vector of H. columbae at that time. Subsequently,he demonstrated thatOrnithomya aviculariawas also suscepti-ble to infections with a Haemoproteus sp. parasite similar to

H. columbae (Baker, 1957). However, it was not possible toinfect uninfected C. livia pigeons with the Haemoproteus sp.parasites obtained from the wood pigeon Columba palumbusvia infectious bites ofO. avicularia(Baker, 1957, 1963). Thisled to the suggestion that Haemoproteus parasites acquiredfrom C. palumbuswere from a subspecies or lineage unableto develop in C. livia (Baker, 1963). This parasite was ableto develop in P. canariensisbut at a low rate (two out of 73flies were successfully infected) (Baker, 1966b). Baker (1966a)showed that the C. palumbus parasite was a different species(named H. palumbis) unable to develop in C. livia. A subsequentexperiment showed that H. columbaewas unable to developinC. palumbus, and although it was able to infect O. avicularia,

the oocysts appeared degenerated and no sporozoites wereobserved (Baker, 1968). This series of experiments suggests

that there is strong host specificity of these parasites, in partic-ular for the vertebrate host (Baker, 1968). However, Rashdan

(1998a) showed thatH. columbaecould successfully infect twoother dove speciesStreptopelia senegallusand S. turturthrougheither injection of infected salivary glands or infectious bitesofP. canariensis. The course of infection of H. columbae inthe new hosts was the same in S. senegallus but a longerpre-patent period and lower parasitemia was observed inS. turturcompared toC. livia (Rashdan, 1998a). P. canariensiswas able to feed and survive normally on the Streptopelia



17/37


doves (Rashdan, 1998a). Furthermore,P. canariensiswas ableto successfully transmitHaemoproteus turturto bothS. senegallusand S. turtur, but was unable to transmit this parasite toC. livia, even by inoculation of infected tissue (i.e. salivaryglands or macerated lungs; Rashdan, 1998b). These stud-ies suggest that host specificity of haemosporidian parasites

cannot be evaluated based on phylogenetic relationships, atleast not below the family taxonomic rank. Table 2 lists all

experimental vector studies forHaemoproteusparasites.The hippoboscid flyO. aviculariais not only involved in the

transmission ofH. palumbis, but is also a competent vector forTrypanosoma avium(Baker, 1956, 1957). It has been suggestedthat hippoboscid flies can act as a bridge in the transmissionofHaemoproteusspecies across different vertebrate hosts dueto their plastic feeding preferences (when the preferred hostis absent or in low numbers, these flies will use alterna-

tive hosts; Greiner, 1975). Host-generalist insect vectors arepotentially capable of transmitting parasites from different

genera, and can act as reservoirs and vectors of introducedand expanding parasites (Perkins et al., 2008). A study ana-lyzing the ecology of hippoboscid flies feeding on mourning

doves (Zenaida macroura) recordedMicrolynchia pusillaand Stil-bometopa podopostyla flies as natural vectors of H. sacharoviand suggested that someCulicoides(Ceratopogonidae) speciescould act as supplementary vectors in the transmission of this

dove parasite, although no experimental evidence was pro-vided (Greiner, 1975). When a parasite species is transmited

by different vectors, their population may show less fluctu-ations over time within a given area compared to parasites

specializing on a single vector both due to the alternativepathways (i.e. vector species) and because some vectors can

act as reservoirs. On the other hand, use of an abundant

vector species would compensate for the narrow host pref-erences of a specialized parasite. Such patterns are likelyto vary as a geographic mosaic; a parasite species that is a

vector generalist in one location may be a vector specialist inanother. For example,P. relictumis mostly transmitted byCxquinquefasciatusin Hawaii, whereasCx. pipiens,Cx. restuansandAe. canadensisare its vectors in Eastern USA (LaPointe, Goff& Atkinson, 2005; Kimura et al., 2010).

The other bird species used intensively in studies of

Haemoproteusparasites transmitted by hippoboscid flies is theCalifornia quail (Lophortyx californica) (Tarshis, 1955). Tarshis(1952, 1954) developed field methods for the collection andtransport of hippoboscid flies from wild-trapped California

quail. This bird is parasitized by H. lophortyx, which is success-fully transmitted byStilbometopa impressa, and has an averageinfection prevalence of 63% (Tarshis, 1955). The prepatentperiod in naturally infectedS. impressaflies was determinedto be 2144 days and for experimentally infected flies theprepatent period was 44 days or longer, depending on the

temperature at which flies were maintained (Tarshis, 1955).These experiments were later criticized because aviaries

were used with mesh sizes that did not exclude Cerato-pogonid vectors (Valkiunas, 2005; Mullenset al., 2006); thus,Culicoidesflies could not be eliminated as competent vectorsofH. lophortyx(see Section II.4). T

able2.Experimentalstudiesdem

onstratingfullorpartialvectorialcompetenceofdifferentspeciesofHippoboscidaefliestransmittingparasitesofthegenusHaemoproteus

PCRmethod

Parasitespecies

Vectorspecies

Proven

experimentally

Proven

natural

vector

M

ostadvanced

developmental

stageobserved

invector

Region

PC

Ron

vecto

rblood

m

eal

PCRon

whole

unengorged

vectors

PCRon

thoracic

partsof

vectors

References

HaemoproteuscolumbaePseudolynchiacanariensis

+

+

Sporozoite

USA,Egypt,Brazil

Aragao(1908),Rendtorff

etal.(1949),and

Ra

shdan(1998a)

H.palumbis

Ornithomyaavicularia

+

+

Sporozoite

England

Baker(1957,1963,1966a,

b,1968)

P.canariensis

+

Sporozoite

England

Baker(1966a,b)

H.turtur

P.canariensis

+

Sporozoite

Egypt

Rashdan(1998b)

H.sacharovi

P.canariensis

+

Sporozoite

USA

Huff(1932b)

H.multipigmentatus

Microlynchiagalapagoensis

+

None

Ecuador(Galapagos)

+

Valki

unasetal.(2010a)

H.iwa

Olfersiaspinifera

+

None

Ecuador(Galapagos)

+

Levin

etal.(2011)

SeeTable1fordefinitionsofexperim

entalandnaturalvectors.



18/37


Few studies have been conductedon the seasonal dynamicsof hippoboscid vectors, limiting our understanding of thetransmission cycle of the Haemoproteus parasites transmittedby these flies. For example, Klei & DeGiusti (1975) showedthat intensity of infection and prevalence ofH. columbae inpigeon populations from Detroit was higher in the autumnand winter seasons and lower in spring. This was explained

by the higher number of flies in late summer and during theautumn, in contrast to patterns observed in other populationsexposed to less seasonal habitats.

(a) Taxonomic issues for parasites transmitted by Hippoboscidae

The genus Haemoproteus was divided into two groups(H. columbae group and H. nettionis group) (Baker, 1963);later these two groups were named as Haemoproteus andParahaemoproteus (Bennett et al., 1965). Haemoproteus containsH. columbae (type species), H. sacharovi and H. multipigmenta-tus (Valkiunas et al., 2010a), H. lophortyx was considered tobelong to this subgenus, but has recently been placed in the

subgenus Parahaemoproteus; hippoboscid flies transmit theseparasites. The features of the Haemoproteusgroup are: (i) theinvertebrate host is a hippoboscid fly; (ii) sporogony occursrelatively slowly with the production of expanding oocysts,giving rise to several hundred sporozoites, having one endmore pointed than the other; (iii) exoerythrocytic merogonyoccurs in the vascular endothelium without the productionof compartmentalized cytomeres. The second group, Para-haemoproteus, has strikingly different features. It was originallydescribed by Wenyon (1926), who stated that unpigmentedschizonts occurred in masses in the lung, liver, and kidneysof infected birds. Sporogony ofParahaemoproteus has beendescribed in detail by Fallis & Wood (1957) and Fallis &

Bennett (1961a) in Canadian birds and was shown to occurin species of Culicoides (Ceratopogonidae). The Parahaemo-proteusgroup may thus be defined as including parasites ofbirds that (i) undergo sporogony in species ofCulicoides,withthe production of small, non-expanding oocysts containingfew sporozoites, with tapered ends, and (ii) mostly presentexoerythrocytic schizogony in large compartmentalized fociin the organs.Parahaemoproteus danilewskii(Kruse, 1890a,b) isthe type species. Nowadays, these two groups are recognizedas the subgenera H. (Haemoproteus) andH. (Parahaemoproteus)(Valkiunas, 2005), and this division is supported by molec-ular phylogenetic studies (Martinsenet al., 2008; Santiago-

Alarconet al., 2010), but it is still unclear whether they are

sister or paraphyletic subgenera (Outlaw & Ricklefs, 2011).

(3) Family Simuliidae

Before the 1930s only speculations about the life cycle ofLeucocytozoon parasites were available (Schaudinn, 1904),and the vector responsible for transmitting the parasiteLeucocytozoon smithi in turkeys, which caused high mortalitiesin farms across the USA was unknown. During an outbreakofL. smithiin 1930 in Nebraska, healthy turkeys were injectedwith blood preparations derived from heavily infectedturkeys, but the infections could not be maintained beyond

28 days (Skidmore, 1931, see also Johnson et al., 1938).Turkey lice (Eomenacanthus stramineum) and the fly Stomoxyscalcitrans(Muscidae) were suggested as possible vectors, butexperiments did not validate this (Skidmore, 1931). Finally,

Simulium meridionale (referred as S. occidentale) was observedfeeding on infected turkeys; macerated flies were prepared in

saline solution and injected into healthy turkeys, infectionsdeveloped on the 12th day and parasites were observed in

the blood even >70 days after infection (Skidmore, 1931).Future outbreaks that killed large numbers of turkeys in the

USA prompted further studies that concluded that S. jenningsi(asS.nigroparvum) and S. venustum [now a species groupwith some recognized cytospecies (the use of cellular leveltraits such as chromosomal inversions to differentiate closely

realted species)] could vectorL. smithiin turkeys andL. anatis(= L. simondi) in ducks (Johnson et al., 1938). Due to theeconomic losses generated in the poultry industry byL. smithiinfections and to the vectorial capacity of many simuliids

to transmit human and domestic animal parasites, large-scale aerial treatments to control blackfly populations were

conducted in some areas of the United States (Kissam, Noblet& Garris, 1975), and efforts to establish laboratory coloniesof several simuliid species were made (Edman & Simmons,

1985; Lacoursiere & Boisvert, 1987), in order to facilitate in-depth studies ofL. smithi(e.g. Steele, Noblet & Noblet, 1992).

Blackflies were bred in captivity since the early 1900sbecause of their involvement in the transmission ofOnchocercaparasites (Blacklock, 1926), but were difficult to maintainsuccessfully (Johnsonet al., 1938). With the discovery of theirinvolvement in the transmission ofL. smithi, further attemptsto culture these insects in the laboratory were made. Davies

(1953) was able to keep S. venustum and Simulium decorum in

captivity for up to 63 days. During his studies he measuredfly mortality and found that flies with a partial blood mealsurvived better than those that took a full blood meal. Fur-

thermore, flies that fed on ducks infected with Leucocytozoonwere less viable than those that fed on uninfected ducks,

but both survived less than flies that did not take a bloodmeal at all (Davies, 1953). This study suggests that flies pay a

fitness cost even when feeding on uninfected hosts, and suchcosts increase when large blood meals or infected blood is

ingested (see also Desser & Yang, 1973; Allison, Desser &Whitten, 1978).

Once the vector family was identified, work on othersimuliids was conducted. Johnson et al. (1938) described

the different stages of L. smithi in both turkeys and inS. nigroparvum; however, they were not able to identify oocystsin the insects gut wall and some developmental stages inthe internal organs of turkeys. Starting in the 1950s more in

depth studies on the developmental stages, in particular thesporogonic cycle, ofLeucocytozoon parasites were conducted.Desser & Fallis (1967) provided a detailed description of thesporogony ofL. simondi in Simulium rugglesi; this fly specieswas recorded attacking 14 different bird species around LakeMichigan, with a clear preference for ducks (Barrow, Kelker

& Miller, 1968). The complete life cycle ofL. sakharoffiwasrecorded usingSimulium angustitarsefeeding on rooks (Corvus



19/37


frugilegus; Baker, 1970). The parasite L. bonasae(= L. lovati), aparasite infecting ruffed grouse (Bonasa umbellus), was exper-imentally transmitted to grouse chicks obtained from wildbroods viainjection of infected salivary glands of the simuliidsS. latipesand S. aureum; S. venustum flies were unable to produce

viable infections in this model (Fallis & Bennett, 1958). In

this same study, the infected salivary gland suspension usedto inoculate grouse chicks was used to inoculate ducklings

and white-crowned sparrows (Zonotrichia leucophrys), neitherof which developedL. bonasaeinfection. Furthermore, grousechicks andheavily infected ducklings with L. simondiwere keptin proximity, and no cross infection was detected; the flies

feeding on grouse and duckings were different and seemedto have different parasite affinities (Fallis & Bennett, 1958).

Nonetheless, further experimental studies proved that differ-entspecies of simuliids could be competent for more than oneLeucocytozoonspecies, successfully transmitting the parasite todifferent bird species (Fallis & Bennett, 1962). EvenS. venus-tum, which is not ornithophilic, was a suitable vector for thebird parasites L. fringillinarum and L. simondiwhen forced tofeed experimentally on birds (Fallis & Bennett, 1962; Desser

& Yang, 1973), but S. venustum rarely feeds on ducks undernatural conditions (Fallis & Bennett, 1961b). Fallis (1964)provided an extensive list of simuliids and their feedinghabits from which it is clear that many species of blackflies

feed habitually on both mammals and birds (e.g. Simuliumaureosimile, S. latipes, S. mexicanum, S. meridionale). Moreover,many simuliids have flexible host preferences, which mightdepend on host availability (Fallis, 1964) and potentially

provides opportunities for parasites to jump across speciesand vertebrate groups. For example,Simulium veracruzanumisimplicated in the transmission ofOnchocercanematodes from

cows and horses to man (Fallis, 1964). If cross infections arerare, parasite jumps could instantly isolate parasite popu-lations, eventually leading to genetic divergence and/or to

a new emergent disease. Experimental studies of simuliidstransmittingLeucocytozoonparasites are listed in Table 3.

More recently, molecular techniques have been usedto untangle the host-parasite-vector relationships. Using

engorged vectors it is possible to extract the DNA of theblood meal and amplify specific sections of DNA to identify

the vertebrate host, and the haemosporidian parasite generawhen the blood meal is infected. By using this method it

has been possible to determine that black flies have somedegree of host specificity (Hellgren et al., 2008). Moreover,

the Leucocytozoon lineages that specific simuliid species trans-mit are closely related, implying an ecological restriction

in terms of the vertebrate host spectrum that Leucocytozoonparasites are able to infect due to the specific host pref-

erences of simuliids (Malmqvist et al., 2004; Hellgren et al.,2008). Care must be taken in the conclusions drawn from

molecular studies, however, because: (i) vectors can switchtheir feeding

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Santiago Alarcon Et Al2012DipteraVectorsOfAvianHaemosporidianParasites