Open Access

PCR detection of malaria parasites and related haemosporidians: the sensitive methodology in determining bird-biting insects

Malaria Journal201615:283

https://doi.org/10.1186/s12936-016-1338-y

Received: 21 January 2016

Accepted: 11 May 2016

Published: 20 May 2016

Abstract

Background

Knowledge about feeding preference of blood-sucking insects is important for the better understanding epidemiology of vector-borne parasitic diseases. Extraction of DNA from blood present in abdomens of engorged insects provides opportunities to identify species of their vertebrate hosts. However, this approach often is insufficiently sensitive due to rapid degeneration of host DNA in midguts. Recent studies indicate that avian malaria parasites (Plasmodium spp.) and related haemosporidians (Haemosporida) belonging to Haemoproteus can persist both in vectors and resistant blood-sucking insects for several weeks after initial blood meals, and these parasites can be readily detected by polymerase chain reaction (PCR)—based methods. Because avian haemosporidians are cosmopolitan, prevalent and strictly specific to birds, the determination of haemosporidian DNA in blood-sucking dipterans can be used as molecular tags in determining bird-biting insects. This hypothesis was tested by investigation of prevalence of natural haemosporidian infections in wild-caught mosquitoes (Culicidae) and biting midges (Ceratopogonidae: Culicoides).

Results

Females of mosquitoes (1072 individuals of three species) and biting midges (300 individuals of three species) were collected in wildlife using simple netting. They were identified and tested individually for the presence of both the haemosporidian parasites and the bird blood using PCR-based methods. Seven different Haemoproteus and two Plasmodium lineages were detected, with overall infection prevalence of 1.12 and 1.67 % in mosquitoes and biting midges, respectively. In all, the detection rate of avian haemosporidian parasites was three fold higher compared with the detection of avian blood.

Conclusions

Molecular markers of avian malaria parasites and other haemosporidians are recommended for getting additional knowledge about blood-sucking dipterans feeding on bird blood. Many genetic lineages of avian haemosporidians are specific to avian hosts, therefore, the detection of these parasite lineages in blood-sucking insects can indicate their feeding preferences on the level of species or groups of related bird species.

Keywords

PCR Blood-sucking insects Plasmodium Haemoproteus Host preference Birds

Background

Many species of blood-sucking insects are involved in transmission of human and animal pathogens [1]. Knowledge on the host preference of vectors is important for better understanding patterns of epidemiology of vector-borne diseases and evolution of host-parasite interactions [2, 3], but remains insufficient, particularly in wildlife. Many species of blood-sucking insects are known to be either ornithophilic or mammalophilic, however, some vector species have broad feeding preference, which often is an adjustment to the availability of their host species [46].

Studies on host preferences of blood-sucking insects in wildlife usually are carried out by (1) the polymerase chain reaction (PCR)-based determination of origin of blood meals reported in females [711] and (2) the observation insect behaviour in choice situations [1214]. The latter method is time-consuming. PCR-based methods are relatively easy to use in research if fresh fully-engorged dipteran insects are available for testing. However, the density of resting fully-engorged mosquito females usually is low [15], and the resting sites of Culicoides biting midges remain insufficiently described in many species [14]. The percentage of engorged mosquitoes and Culicoides spp. females in samples collected by various traps usually is about 0.2–12.5 % [8, 9, 15, 16]. In different studies, the maximum success of PCR-based determination of blood meals in wild-caught fully-engorged insects varied between 17.5 [6] and 92.2 % [10] using vertebrate specific primers. Numerous negative amplifications in these studies likely were due to the DNA degeneration in insect midgut [11]. Kim et al. [17] reported that blood meal identification was successful by PCR-based methods as long as the blood meal residue was distinguishable by eye in the abdomen. Oshaghi et al. [18] determined that the extracted from the blood meal host DNA can be used as a proper template for PCR amplification only 33 h after ingestion.

Mosquitoes, biting midges and blackflies are main vectors of avian haemosporidians, which are obligate heteroxenous blood parasites belonging to the genera Haemoproteus, Plasmodium, Leucocytozoon and some others. Haemosporidian parasites complete sporogony in susceptible dipteran insects after infected blood meals [1921]. These parasites are cosmopolitan in birds [20]. Additionally, birds are the most diverse and common vertebrate hosts of haemosporidians in all ecosystems. Over 250 species of these parasites have been described and named [20], and many more genetic lineages have been identified and deposited in GenBank. Recent molecular studies indicate that avian haemosporidians can persist both in competent vectors and resistant blood-sucking insects for several weeks after initial infected blood meals. These parasites can be readily detected by PCR-based methods up to 17 days after infected blood meals [22]. That provides opportunities to determine bird-biting insects using haemosporidian molecular markers. Because avian haemosporidians are widespread, prevalent, diverse and strictly specific to birds, we predicted that determination of their DNA lineages can be used as molecular tags in determining bird-biting insects. This hypothesis was tested here by investigation of natural haemosporidian infection in wild-caught mosquitoes (Culicidae) and biting midges (Ceratopogonidae: Culicoides) using PCR-based methods.

Methods

Study site and dipteran insect collection

Field-work was carried out at the Biological Station of the Zoological Institute of the Russian Academy of Sciences on the Curonian Spit in the Baltic Sea (55º15′ N, 20º86′ E) between 10th and 20th June 2013. Blood-sucking insects were sampled using an entomological net near the Lake Chaika, where density of mosquitoes and biting midges was high. Insects were collected randomly by simple netting from vegetation; they were collected from the net using a pooter and transported to the laboratory within 3 h post sampling. Collected insects were anaesthetized by placing them into a tube covered with a cotton pad wetted with 96 % ethanol. Anaesthetized mosquitoes were sexed, and females were identified using morphological characters [23]. Biting midges were identified according to their wing patterns and other morphological features [24]; their heads were removed to prepare permanent preparations mounted in Euparal for the further confirmation of the identification.

All collected individual insects were fixed separately in tubes containing 96 % ethanol; they were used for PCR analysis. Before DNA extraction all insect females were checked for blood-meal status dividing them into three classes: fed, gravid and empty. Heads of mosquitoes were removed to eliminate the inhibitory effects on PCRs, as recommended in [25]. New needles were used for dissection of each insect to avoid DNA contamination.

Polymerase chain reaction and sequencing

Total DNA was extracted from each individual insect female using ammonium acetate extraction method [26]. All reactions were performed in 25 µl total volumes, including 50 ng of total genomic DNA template (2 µL), 12.5 µL of DreamTag Master Mix (Thermo Fisher Scientific, Lithuania), 8.5 μL nuclease-free water and 1 μL of each primer.

The quality of extracted DNA was tested using the insect specific primers LCO149 and HCO2198, which amplify a fragment of cytochrome oxidase subunit I (COI) gene of insects’ mitochondrial DNA [27]. The obtained sequences were also used to confirm insect species identification; these sequences were deposited to GenBank. All PCRs with COI primers were positive indicating good quality of extracted DNA.

To determine bird species based on DNA extracted from mosquito blood meals, we applied bird specific primers Avian-3 and Avian-8, which amplify the 473 bp fragment of bird cytochrome b (cyt b) gene [28]. In parallel, we also used recently developed [29] bird specific primers cytbPF1 and cytbBirdR, which amplify 219 bp fragments of cyt b gene. Additional forward primer (cytbBirdF2) was used for semi-nested PCR to amplify the 169 bp fragment of bird cyt b gene.

For genetic analysis of haemosporidian parasites, a nested PCR protocol was applied [30]. We amplified a segment of mitochondrial cyt b gene using primers HaemNFI and HaemNR3, which are specific to haemosporidians belonging to Haemoproteus, Plasmodium and Leucocytozoon parasites. For the second PCR, we used primers HAEMF and HAEMR2, which are specific to Haemoproteus and Plasmodium spp. and amplify 479 bp fragments of cyt b gene. All samples were tested for bird and parasite DNA in parallel. In all PCRs, one negative control (nuclease-free water) and one positive control (one thorax of Ochlerotatus cantans mosquito 7 days after the experimental infection with Haemoproteus tartakovskyi, in the case of parasite testing and a siskin’s (Carduelis spinus) blood sample in case of avian blood testing) were used per every 24 samples to control for false amplifications. No case of false positive or negative amplification was found. Thermal conditions of COI PCR were as described by Day et al. [31], conditions of other PCR followed the protocols mentioned in the references cited.

All positive amplifications were evaluated by running 2 μl of the final PCR product on a 2 % agarose gel; they were sequenced in order to determine cyt b lineages of the detected parasites or to detect bird species from blood meals. DNA fragments were sequenced with corresponding primers twice for both strands. Dye terminator cycling sequencing (BigDye) was used and loaded on an ABI PRISM™ 3100 capillary sequencing robot (Applied Biosystems, Foster City, California). Good quality sequences were obtained; they were edited and aligned using BioEdit (Version 7.0.9.0).

Data analysis

The “Basic Local Alignment Search Tool” (National Centre for Biotechnology Information) was used to determine genetic lineages of detected avian blood parasites or to determine species of birds. The analyses were carried out using “Statistica 7” package. Percentages of infected insects were compared by Yates corrected χ2 test. A P value of 0.05 or less was considered significant.

Results

In all, 1372 females (1072 mosquitoes and 300 Culicoides biting midges) were sampled and investigated for the presence of both bird blood and avian haemosporidian parasites using PCR-based methods (Table 1). Three mosquito species were sampled. Ochlerotatus cantans was the most abundant mosquito species (69.2 % of all collected mosquitoes); Ochlerotatus cataphylla and Ochlerotatus sticticus represented 22.4 and 8.4 % of all collected mosquitoes, respectively. Three Culicoides species were collected and examined. These were Culicoides impunctatus (89.0 % of all sampled midges), Culicoides punctatus (9.3 %) and Culicoides pictipennis (1.7 %). Obtained COI sequences of investigated insects conform for 99–100 % with sequences of corresponding species from GenBank [KJ627800, KX064673–KX064677]. The great majority of collected insects were classified as empty (96.4 %). Gravid and fed females represented 3.1 and 0.5 %, respectively.
Table 1

Sensitivity of PCR-based detection of bird blood and haemosporidian lineages in the same wild-caught blood-sucking dipteran insects

Dipteran insect family and species

No. examined

No. positive for bird blood

No. positive for parasites

Difference, P value

Parasite lineage

Mosquitoes Culicidae

 Ochlerotatus cantans

742

2 (0.27)a

8 (1.08)

0.113

pSGS1, pGRW11, hSISKIN1, hDELURB1, hSW1, hRB1, hRBS2

 Oc. cataphyla

240

1 (0.42)

3 (1.25)

0.616

hRBS2, hSFC1

 Oc. sticticus

90

0 (0)

1 (1.11)

0.316

hSISKIN1

 Total mosquitoes

1072

3 (0.28)

12 (1.12)

0.038

Eight lineages reported

Biting midges Ceratopogonidae

 Culicoides impunctatus

267

2 (0.75)

3 (1.12)

0.472

pSGS1, hTURDUS2, hSISKIN1

 C. punctatus

28

1 (3.57)

1 (3.57)

1.000

hTURDUS2

 C. pictipennis

5

0 (0)

1 (20.0)

0.292

hTURDUS2

 Total biting midges

300

3 (1.00)

5 (1.67)

0.722

Three lineages reported

Grand total

1372

6 (0.44)

17 (1.24)

0.036

Nine lineages reported

aPercentage is given in parenthesis. Significant differences are given in italic font

Seven different Haemoproteus spp. (hSISKIN1, hDELURB1, hSW1, hRB1, hRBS2, hSFC1, hTURDUS2 [GenBank accessions KC559436, KC818455, KJ559439, KJ627802, KR049256–KR049262]) and two Plasmodium spp. (pSGS1, pGRW11 [GenBank accessions KR049253–KR049255]) cyt b genetic lineages were detected, with overall infection prevalence of 1.12 % in mosquitoes and 1.67 % in biting midges (Table 1).

Blood of three bird species (Ficedula hypoleuca, Carduelis spinus and Parus major) was detected in 0.44 % of examined insects; it was identified in three individual mosquitoes and three individual biting midges using primers cytbBirdR/cytbBirdF2 (Table 1). Obtained DNA sequences conform for 98–100 % with the sequences of corresponding bird species from GenBank [accessions KP794613–KP794615]. The efficacy of primers Avian-3/Avian-8 was similar: bird blood was detected in 0.36 % of examined insects (in three individual mosquitoes and two individual biting midges). Both sets of bird specific primers amplified DNA from the same samples, except one sample of C. impunctatus, in which one positive amplification was obtained using cytbBirdR/cytbBirdF2 primers, but not with Avian-3/Avian-8 primers. Differences between sensitivity of different primers in detection of bird blood were insignificant (P = 0.76).

Overall, the detection rate of avian haemosporidian parasites was three-fold higher (P < 0.05) comparing with the detection rate of avian blood in same samples of investigated blood-sucking insects (Table 1).

Discussion

The key result of this study is that molecular markers of avian haemosporidians can be readily used for determining bird-biting dipteran insects. Application of this methodology in parallel with detection of bird blood in engorged insects provides much additional information about feeding preference of blood-sucking dipterans in wildlife.

It is difficult to collect large samples of engorged blood-sucking dipterans for determination of origin of their blood meals [9, 15, 16]. Simple netting from low vegetation was used to collect mosquitoes and biting midges in this study. Blood-sucking insects are also often collected using animal baits and light or ultraviolet traps in wildlife [13]. These sampling methods provide opportunities to collect the actively flying part of dipteran populations, particularly females, which are host seeking insects. Selective collection of engorged females directly in their resting sites is time consuming, particularly because of insufficient knowledge about their resting places, which might differ even for the same insect species at different study sites [14]. Proportion of engorged mosquitoes and Culicoides spp. usually varies markedly in different studies, and it has been reported not to exceed 12.5 % of all sampled females [810, 15]. Even if sufficient number of engorged insects is collected, the probability to obtain positive PCR signals usually is not high using primers specific for vertebrate host DNA. The success of PCR amplification of host DNA in engorged mosquitoes using vertebrate specific primers was 17.5 % in Portugal [6]. Bartsch et al. [8], Lassen et al. [9] and Ninio et al. [32] reported 65, 76 and 91 % of positive PCR amplifications of host DNA in fully engorged biting midges in Germany, Denmark and France, respectively. Negative PCR-based results are usually explained by the low blood meal volumes or the degeneration of host DNA due to blood digestion in the midguts [11]. Additionally, there is a significant negative relationship between the time after blood ingestion and the success of PCR-based analysis [18, 33]. According to Ejiri et al. [28] the success of blood meal identification using PCR is 100 % in fresh full-fed mosquitoes, 75 %—in partially fed, 56 %—in half-gravid and only 38 %—in gravid females.

Molecular markers of avian haemosporidian parasites are suggested to use for determining bird-biting insects (Table 1). The prevalence of haemosporidian infections is high in many bird species worldwide, with the overall prevalence exceeding 40 % and reaching even 80–100 % in many bird populations during the breeding season in Europe [20, 34]. Blood-sucking insects gain these parasites only during infected blood meals on birds. Alive sporozoites of avian malaria parasites and other haemosporidians have been reported in Culex spp. and Culicoides spp. for approximately one month after initial infected blood meal [19, 20]. Additionally, haemosporidian parasites can persist even in resistant blood-sucking insects without DNA degeneration for several weeks after infected blood meals [22]. That provides opportunities to identify bird-biting insects using molecular markers, which target haemosporidian parasite DNA. For example, haemosporidian parasites have been reported between 0.4 and 33.7 % of pools of wild-caught dipteran females tested by PCR at different study sites [6, 35]. We determined avian haemosporidian parasites in 1.24 % of all tested insects (Table 1). Martínez-de la Puente et al. [36] determined 6.4 % of collected Culicoides spp. to be positive for haemosporidian infection in Spain. Ejiri et al. [28] reported 14.6 % of tested Culex pipiens mosquitoes infected by these parasites in Japan. In Switzerland, 6.6 % of Culex pipiens females were PCR positive for avian malaria infection [37]. Inci et al. [38] found DNA of avian haemosporidian parasites in 9.9 % of mosquito pools in Turkey. Thus, it is relatively easy to detect haemosporidian parasites in blood-sucking insects using PCR-based diagnostics. Additionally, it is known that chronically infected vertebrate hosts attract significantly more vectors than uninfected ones [39, 40], and this fact further increases the efficiency of parasite detection compared with the host blood detection in insect. Because avian haemosporidians are strictly specific to birds [2, 20], the reports of their lineages in dipterans indicate their ornithophily (Table 1).

This study detected the widespread bird malaria parasite Plasmodium relictum (lineages pSGS1 and pGRW11) in Ochlerotatus cantans mosquitoes, and the lineage pSGS1 was also detected in Culicoides impunctatus, indicating that these insects naturally feed on birds. Formerly, both insect species were considered to be mainly mammalophilic [4, 41]. Interestingly, Santiago–Alarcon et al. [3], Ferraguti et al. [42], Martínez-de la Puente et al. [36] and this study detected avian Plasmodium lineages in biting midges, the insects which do not transmit these parasites [2], indicating feeding preference of Culicoides midges and possible abortive sporogonic development in non-susceptible insects [22]. In this study, six different Haemoproteus lineages were detected in mosquitoes, which also do not transmit these parasites. These lineages are: hSISKIN1 (Haemoproteus tartakovskyi), hRB1 and hRBS2 (Haemoproteus lanii), hSW1 (Haemoproteus belopolskyi), hDELURB1 (Haemoproteus hirundinis), and hSFC1 (Haemoproteus balmorali) (Table 1). Mosquitoes are non-competent hosts of Haemoproteus spp., but are exposed to these parasites during infected blood meals, and the parasite’s DNA can be detected in mosquitoes even several weeks after initial blood meals [22] providing opportunities to identify bird-biting insects.

It worth noting that some lineages of haemosporidian species are known to complete life cycle and produce gametocytes (the only infective stage for vectors) in certain avian hosts. That is particularly true for some Haemoproteus parasites [20, 43]. Detection of such parasite lineages in blood-sucking insects might indicate their feeding preference on the level of species or groups of relative bird species. For example, the lineage hSISKIN1 of H. tartakovskyi has been reported to complete development and produce gametocytes only in siskins and crossbills (Loxia curvirostra); the lineage hSW1 of H. belopolskyi and the lineages hRB1 and hRBS2 of H. lanii complete development only in Acrocephalus and Lanius birds, respectively (see Table 2). Gametocytes of the lineage hTURDUS2 of H. minutus and the lineage hALARV of H. tartakovskyi were reported only in blackbirds (Turdus merula) and skylarks (Alauda arvensis), respectively [50, 70, 71]. The reports of these Haemoproteus lineages in blood-sucking insects (Table 1) indicate that they likely naturally take blood meals on corresponding bird species.
Table 2

Occurrence of detected Haemoproteus parasites and their cytochrome b lineages in European birds

Parasite species

Genetic lineage

Vertebrate hosts positive by

References

PCR-based detection

Microscopic examination

H balmorali

hSFC1

Muscicapa striata; Cyanistes caerulens

Muscicapa striata

[4449, 50 a]

H. belopolskyi

hSW1

Acrocephalus schoenobaenus; Acrocephalus palustris; Acrocephalus scirpaceus

Acrocephalus schoenobaenus; Acrocephalus palustris

[45, 5153, 54 a55]

H. hirundinis

hDELURB1

Delichon urbicum; Hirundo rustica

Delichon urbicum

[45, 565859 a]

H. lanii

hRB1

Emberiza schoeniclus; Lanius collurio

Lanius collurio

[45, 53, 60 a, 61 a]

hRBS2

Lanius collurio

Lanius collurio

[45, 46, 62 a]

H. minutus

hTURDUS2

Acrocephalus scirpaceus; Bolborhynchus lineola; Cyanistes caeruleus; Emberiza schoeniclus; Eritacus rubecula; Garulus glandarius; Hippolais icterina; Lanius collurio; Panurus biarmicus; Turdus merula; Turdus torquatus

Turdus merula

[4446, 49, 50 a, 53, 60 a, 6367]

H. tartakovskyi

hSISKIN1

Carduelis spinus; Loxia curvirostra; Carpodacus erythrinus

Carduelis spinus; Loxia curvirostra

[22 a, 50 a, 53, 60, 61 a, 68, 69 a]

aStudies reporting presence of gametocytes in the blood of birds

However, the information about feeding preference of blood-sucking insects on the level of bird species should be analysed with caution using parasitology data. Public databases [72] and the GenBank, contain information about reports of some parasite lineages in unusual avian hosts, in which gametocytes of corresponding lineages have not been reported by microscopic methods [20], and solely PCR-based diagnostic data are available (Table 2). The number of such reports is increasing in MalAvi database [72], and they likely are due to abortive development of parasites, which do not produce gametocytes and cannot infect blood-sucking insects [59, 73]. It is important to note that PCR-based method is a sensitive tool for detection of haemosporidian parasites however, it only requires presence of parasite DNA, so can overestimate host range when such DNA is detected from non-competent host species. In other words, solely PCR-based record of a lineage in a host species is not enough evidence of the species being a competent host, i.e. the host in which haemosporidian gametocytes develop. Table 2 provides references of studies, which detected gametocytes in birds for the lineages reported in this study. Vectors can be infected and parasite can develop (completely or abortively) in insects only after blood meal on birds with gametocytes in the blood [20]. Other haemosporidian stages, which might be present in the circulation (remnants of tissue meronts or sporozoites) and provide templates for PCR amplification, are digested in blood-sucking insects [19, 20]. There are increasing evidences that haemosporidian infections in non-competent hosts result in abortive development, and gametocytes are not produced [22, 67, 74]. Because such DNA may leak into the blood, it is likely that the host range across bird families or genera for many parasite lineages reported by PCR is broader than the ranges of their competent hosts (Table 2). That should be taken in consideration when using the methodology described in this study: positive PCR signals in blood-sucking insects likely indicate that they were fed on competent hosts of corresponding parasite lineages. Plasmodium parasites are less specific for vertebrate hosts [19], and reports of their lineages in blood-sucking insects can be used mainly for indication of feeding on the level of class Aves.

Due to complex patterns of transmission of pathogens [75], the determination of ornithophilic blood-sucking insects is important epidemiologically, including the epidemiology of human vector-borne diseases, because the same vector can take blood meal from different hosts and can serve as bridge vector of the disease from birds to humans. Some mosquitoes and biting midges are opportunistic in trait of host choice [9], but others are host-specific and prefer to take blood meals either on mammals or birds [15]. Temporal and spatial variation in host availability might be important in host selection by blood-sucking insects [14]. For example, many Culex species prefer to feed on birds [14], however, this host-insect interaction is not strict. The abundance of birds often changes due to seasonal and other migrations at many sites throughout the year, and Culex species can readily switch from birds to humans when the availability of birds declines [76]. For example, many Culex species prefer to take blood meals on the American robin (Turdus migratorius) in the Nearctic. However, when the density of this bird population decreases during migrations, Culex mosquitoes shift from this preferred avian host to humans, thereby increasing a probability of West Nile virus transmission to humans [77]. More information is needed about feeding specialization of blood-sucking dipterans. Detection of haemosporidian lineages in dipterans contributes to accumulation of such knowledge.

It worth mentioning that the same insect can be infected with several species of haemosporidian parasites due to the blood meal on birds with co-infection or the “contamination” by any subsequent blood meal on different species of avian hosts. This situation makes analysis of possible host species of blood-sucking insects more difficult. Recent experimental study [78] shows that application of several primers in parallel markedly increase detectability of haemosporidian parasite co-infections, and this method worth using in vector research as well.

Haemosporidian parasites are particularly diverse in countries with warm climates [19]. In Europe and other countries with temperate or cold climates, haemosporidian parasites are widespread in birds, but usually are rare or absent in mammals and reptiles. However, many groups of terrestrial vertebrates are parasitized by haemosporidians in tropical countries [19, 20, 79]. It is probable that the methodology developed in this study can be applied for determining reptile, bird and mammal feeding blood-sucking insects in countries where active transmission of these parasites occurs. Wildlife haemosporidians can be used as biological markers for determining feeding preference of blood-sucking insects even more effectively in tropical and subtropical regions where these pathogens inhabit not only birds, as mainly is the case in northern Holarctic [2, 20], but also many species of reptiles and mammals [19, 79].

Conclusions

This study shows that PCR-based detection of lineages of avian haemosporidian parasites is useful methodology for determining ornithophilic blood-sucking insects. If applied together with molecular markers targeting bird DNA, this methodology increases probability of determining host preference of dipteran blood-sucking insects. Molecular markers of avian malaria parasites and other haemosporidians as additional biological tags in determining links between blood-sucking insects and their avian hosts in wildlife are recommended.

Declarations

Authors’ contributions

Both authors contributed to this research and manuscript preparation equally. Both authors read and approved the final manuscript.

Acknowledgements

We thank the staff of the Biological Station “Rybachy”, for assistance in the field. The procedures described herein comply with the current laws of Lithuania and Russia. This research was supported by the Open Access to research infrastructure of the Nature Research Centre under Lithuanian open access network initiative.

Competing interests

The authors declare that they have no competing interests.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

(1)
Nature Research Centre

References

  1. Bush AO, Fernández JC, Esch GW, Seed JR. Parasitism: the diversity and ecology of animal parasites. 1st ed. Cambridge: University Press; 2001.Google Scholar
  2. Atkinson CT, Thomas NJ, Hunter DB. Parasitic diseases of wild birds. 1st ed. Ames: Blackwell Publishing; 2008.View ArticleGoogle Scholar
  3. Santiago-Alarcon D, Havelka P, Schaefer HM, Segelbacher G. Bloodmeal analysis reveals avian Plasmodium infections and broad host preferences of Culicoides (Diptera: Ceratopogonidae) vectors. PLoS ONE. 2012;7:e31098.View ArticlePubMedPubMed CentralGoogle Scholar
  4. Blackwell A, Luntz AJM, Mordue W. Identification of bloodmeals of the Scottish biting midge Culicoides impunctatus by indirect enzyme-linked immunosorbent-assay (ELISA). Med Vet Entomol. 1994;8:20–4.View ArticlePubMedGoogle Scholar
  5. Santiago-Alarcon D, Havelka P, Pineda E, Segelbacher G, Schaefer HM. Urban forests as hubs for novel zoonosis: blood meal analysis, seasonal variation in Culicoides (Diptera: Ceratopogonidae) vectors, and avian haemosporidians. Parasitol. 2013;140:1799–810.View ArticleGoogle Scholar
  6. Ventim R, Ramos JA, Osorio H, Lopes RJ, Perez-Tris J, Mendes L. Avian malaria infections in western Europe mosquitoes. Parasitol Res. 2012;111:637–45.View ArticlePubMedGoogle Scholar
  7. Alcaide M, Rico C, Ruiz S, Soriguer R, Muñoz J, Figuerola J. Disentangling vector-borne transmission networks: a universal DNA barcoding method to identify vertebrate hosts from arthropod bloodmeals. PLoS ONE. 2009;4:e7092.View ArticlePubMedPubMed CentralGoogle Scholar
  8. Bartsch S, Bauer B, Wiemann A, Clausen PH, Steuber S. Feeding patterns of biting midges of the Culicoides obsoletus and Culicoides pulicaris groups on selected farms in Brandenburg, Germany. Parasitol Res. 2009;105:373–80.View ArticlePubMedGoogle Scholar
  9. Lassen SB, Nielsen SA, Kristensen M. Identity and diversity of blood meal hosts of biting midges (Diptera: Ceratopogonidae: Culicoides Latreille) in Denmark. Parasit Vectors. 2012;5:143.View ArticlePubMedPubMed CentralGoogle Scholar
  10. Garros C, Gardès LT, Allène X, Rakotoarivony I, Viennet E, Rossi S, Balenghien T. Adaptation of a species-specific multiplex PCR assay for the identification of blood meal source in Culicoides (Ceratopogonidae: Diptera): applications on Palaearctic biting midge species, vectors of Orbiviruses. Inf Gen Evol. 2011;11:1103–10.View ArticleGoogle Scholar
  11. Martínez-de la Puente J, Ruiz S, Soriguer R, Figuerola J. Effect of blood meal digestion and DNA extraction protocol on the success of blood meal source determination in the malaria vector Anopheles atroparvus. Malar J. 2013;12:109.View ArticlePubMedPubMed CentralGoogle Scholar
  12. Mukabana WR, Olanga EA, Knols BGJ. Host-seeking behaviour of Afrotropical anophelines: field and semi-field studies. In: Takken W, Knols BGJ, editors. Ecology and control of vector-borne diseases. Wageningen: Wageningen Academic Publisher; 2010. p. 181–202.Google Scholar
  13. Takken W, Verhulst NO. Host preferences of blood-feeding mosquitoes. Annu Rev Entomol. 2013;58:433–53.View ArticlePubMedGoogle Scholar
  14. Garcia-Saenz A, McCarter P, Baylis M. The influence of host number on the attraction of biting midges, Culicoides spp., to light traps. Med Vet Entomol. 2011;25:113–5.View ArticlePubMedGoogle Scholar
  15. Martínez-de la Puente J, Figuerola J, Soriguer R. Fur or feather? feeding preferences of species of Culicoides biting midges in Europe. Trends Parasitol. 2015;31:16–22.View ArticlePubMedGoogle Scholar
  16. Lassen SB, Nielsen SA, Skovgård H, Kristensen M. Molecular identification of bloodmeals from biting midges (Diptera: Ceratopogonidae: Culicoides Latreille) in Denmark. Parasitol Res. 2011;108:823–9.View ArticlePubMedGoogle Scholar
  17. Kim KS, Tsuda Y, Sasaki T, Kobayashi M, Hirota Y. Mosquito blood-meal analysis for avian malaria studying wild bird communities: laboratory verification and application to Culex sasai (Diptera: Culicidae) collected in Tokyo, Japan. Parasitol Res. 2009;105:1351–7.View ArticlePubMedGoogle Scholar
  18. Oshaghi MA, Chavshin AR, Vatandoost H, Yaaghoobi F, Mohtarami F, Noorjah N. Effects of post-ingestion and physical conditions on PCR amplification of host blood meal DNA in mosquitoes. Exp Parasitol. 2006;112:232–6.View ArticlePubMedGoogle Scholar
  19. Garnham PCC. Malaria parasites and other haemosporidia. 1st ed. Oxford: Blackwell; 1966.Google Scholar
  20. Valkiūnas G. Avian malaria parasites and other haemosporidia. 1st ed. Boca Raton: CRC Press; 2005.Google Scholar
  21. Perkins SL. Malaria’s many mates: past, present and future of the systematics of the order Haemosporida. J Parasitol. 2014;100:11–25.View ArticlePubMedGoogle Scholar
  22. Valkiūnas G, Kazlauskienė R, Bernotienė R, Palinauskas V, Iezhova TA. Abortive long-lasting sporogony of two Haemoproteus species (Haemosporida, Haemoproteidae) in the mosquito Ochlerotatus cantans, with perspectives on haemosporidian vector research. Parasitol Res. 2013;112:2159–69.View ArticlePubMedGoogle Scholar
  23. Becker N, Petric D, Zgomba M, Boase C, Dahl C, Lane J, Kaiser A. Mosquitoes and their control. 1st ed. New York: Kluwer Academic/Plenum Publischers; 2003.View ArticleGoogle Scholar
  24. Gutsevich AV. Blood-sucking midges of the genera Culicoides and Forcipomyia (Ceratopogonidae). Fauna USSR. 1st ed. Leningrad: Nauka Press; 1973.Google Scholar
  25. Arez AP, Lopes D, Pinto J, Franco AS, Snounou G, do Rosario VE. Plasmodium sp.: optimal protocols for PCR detection of low parasite numbers from mosquito (Anopheles sp.) samples. Exp Parasitol. 2000;94:269–72.View ArticlePubMedGoogle Scholar
  26. Richardson DS, Jury FL, Blaakmeer K, Komdeur J, Burke T. Parentage assignment and extra-group paternity in a cooperative breeder: the Seychelles warbler (Acrocephalus sechellensis). Mol Ecol. 2001;10:2263–73.View ArticlePubMedGoogle Scholar
  27. Folmer O, Black M, Hoeh W, Lutz R, Vrijenhoek R. DNA primers for amplification of mitochondrial cytochrome C oxidase subunit I from diverse metazoan invertebrates. Mol Mar Biol Biotechnol. 1994;3:294–9.PubMedGoogle Scholar
  28. Ejiri H, Sato Y, Kim KS, Hara T, Tsuda Y, Imura T, et al. Entomological study on transmission of avian malaria parasites in zoological garden in Japan: bloodmeal identification and detection of avian malaria parasite DNA from blood-fed mosquitoes. J Med Entomol. 2011;48:600–7.View ArticlePubMedGoogle Scholar
  29. Bobeva A, Zehtindjiev P, Ilieva M, Dimitrov D, Mathis A, Bensch S. Host preferences of ornithophilic biting midges of the genus Culicoides in the Eastern Balkans. Med Vet Entomol. 2015;29:290–6.View ArticlePubMedGoogle Scholar
  30. Hellgren O, Waldenström J, Bensch S. A new PCR assay for simultaneous studies of Leucocytozoon, Plasmodium, and Haemoproteus from avian blood. J Parasitol. 2004;90:797–802.View ArticlePubMedGoogle Scholar
  31. Day JC, Goodall TI, Post RJ. Confirmation of the species status of the blackfly Simulium galeratum in Britain using molecular taxonomy. Med Vet Entomol. 2008;22:55–61.View ArticlePubMedGoogle Scholar
  32. Ninio C, Augot D, Delecolle JC, Dufour B, Depaquit J. Contribution to the knowledge of Culicoides (Diptera: Ceratopogonidae) host preferences in France. Parasitol Res. 2010;108:657–63.View ArticlePubMedGoogle Scholar
  33. Mukabana WR, Takken W, Knols BG. Analysis of arthropod bloodmeals using molecular genetic markers. Trends Parasitol. 2002;18:505–9.View ArticlePubMedGoogle Scholar
  34. Marzal A. Recent advances in studies on avian malaria parasites. 1st ed. Rijeka: INTECH; 2012.Google Scholar
  35. Kim KS, Tsuda Y. Avian Plasmodium lineages found in spot surveys of mosquitoes from 2007 to 2010 at Sakata wetland, Japan: do dominant lineages persist for multiple years? Mol Ecol. 2012;21:5374–85.View ArticlePubMedGoogle Scholar
  36. Martínez-de la Puente J, Martínez J, Rivero-de Aguilar J, Herrero J, Merino S. On the specificity of avian blood parasites: revealing specific and generalist relationships between haemosporidians and biting midges. Mol Ecol. 2011;20:3275–87.View ArticlePubMedGoogle Scholar
  37. Glaizot O, Fumagalli L, Iritano K, Lalubin F, Van Rooyen J, Christe Ph. High prevalence and lineage diversity of avian malaria in wild populations of great tits (Parus major) and mosquitoes (Culex pipiens). PLoS ONE. 2012;7:e34964.View ArticlePubMedPubMed CentralGoogle Scholar
  38. Inci A, Yildirim A, Njabo KY, Duzlu O, Biskin Z, Ciloglu A. Detection and molecular characterization of avian Plasmodium from mosquitoes in central Turkey. Vet Parasitol. 2012;188L:179–84.View ArticleGoogle Scholar
  39. Sherman IW. Malaria: parasite biology, pathogenesis, and protection. 1st ed. Washington, DC: American Society for Microbiology Press; 1998.Google Scholar
  40. Cornet S, Nicot A, Rivero A, Gandon S. Evolution of plastic transmission strategies in avian malaria. PLoS Pathog. 2014;10:e1004308.View ArticlePubMedPubMed CentralGoogle Scholar
  41. Silver JB. Mosquito ecology: field sampling methods. 1st ed. New York: Springer; 2007.Google Scholar
  42. Ferraguti M, Martínez-de la Puente J, Ruiz S, Soriguer R, Figuerola J. On the study of the transmission networks of blood parasites from SW Spain: diversity of avian haemosporidians in the biting midge Culicoides circumscriptus and wild birds. Parasit Vectors. 2013;6:208.View ArticlePubMedPubMed CentralGoogle Scholar
  43. Clark NJ, Clegg SM, Lima MR. A review of global diversity in avian haemosporidians (Plasmodium and Haemoproteus: Haemosporida): New insights from molecular data. Int J Parasitol. 2014;44:329–38.View ArticlePubMedGoogle Scholar
  44. Cosgrove CL, Wood MJ, Day KP, Sheldon BC. Seasonal variation in Plasmodium prevalence in a population of blue tits Cyanistes caeruleus. J Anim Ecol. 2008;77:540–8.View ArticlePubMedGoogle Scholar
  45. Dimitrov D, Zehtindjiev P, Bensch S. Genetic diversity of avian blood parasites in SE Europe: Cytochrome b lineages of the genera Plasmodium and Haemoproteus (Haemosporida) from Bulgaria. Acta Parasitol. 2010;55:201–9.View ArticleGoogle Scholar
  46. Hellgren O, Križanauskiene A, Valkiūnas G, Bensch S. Diversity and phylogeny of mitochondrial cytochrome b lineages from six morphospecies of avian Haemoproteus (Haemosporida: Haemoproteidae). J Parasitol. 2007;93:889–96.View ArticlePubMedGoogle Scholar
  47. Knowles SC, Wood MJ, Alves R, Wilkin TA, Bensch S, Sheldon BC. Molecular epidemiology of malaria prevalence and parasitaemia in a wild bird population. Mol Ecol. 2011;20:1062–76.View ArticlePubMedGoogle Scholar
  48. Szöllosi E, Cichoń M, Eens M, Hasselquist D, Kempenaers B, Merino S, et al. Determinants of distribution and prevalence of avian malaria in blue tit populations across Europe: separating host and parasite effects. J Evol Biol. 2011;24:2014–24.View ArticlePubMedGoogle Scholar
  49. Wood MJ, Cosgrove CL, Wilkin TA, Knowles SC, Day KP, Sheldon BC. Within-population variation in prevalence and lineage distribution of avian malaria in blue tits, Cyanistes caeruleus. Mol Ecol. 2007;16:3263–73.View ArticlePubMedGoogle Scholar
  50. Valkiūnas G, Iezhova TA, Križanauskiene A, Palinauskas V, Bensch S. In vitro hybridization of Haemoproteus spp.; an experimental approach for direct investigation of reproductive isolation of parasites. J Parasitol. 2008;94:1385–94.View ArticlePubMedGoogle Scholar
  51. Bensch S, Stjernman M, Hasselquist D, Ostman O, Hansson B, Westerdahl H, Pinheiro RT, et al. Host specificity in avian blood parasites: a study of Plasmodium and Haemoproteus mitochondrial DNA amplifies from birds. Proc Biol Sci. 2000;267:1583–9.View ArticlePubMedPubMed CentralGoogle Scholar
  52. Biedrzycka A, Migalska M, Bielański W. A quantitative PCR protocol for detecting specific Haemoproteus lineages: molecular characterization of blood parasites in a Sedge Warbler population from southern Poland. J Ornithol. 2015;156:201–8.View ArticleGoogle Scholar
  53. Hellgren O, Waldenström J, Peréz-Tris J, Szöll E, Si O, Hasselquist D, Križanauskiene A, et al. Detecting shifts of transmission areas in avian blood parasites - a phylogenetic approach. Mol Ecol. 2007;16:1281–90.View ArticlePubMedGoogle Scholar
  54. Valkiūnas G, Križanauskiene A, Iezhova TA, Hellgren O, Bensch S. Molecular phylogenetic analysis of circumnuclear hemoproteids (Haemosporida: Haemoproteidae) of Sylviid birds, with a description of Haemoproteus parabeloposkyi sp. Nov. J Parasitol. 2007;93:680–7.View ArticlePubMedGoogle Scholar
  55. Ventim R, Morais J, Pardal S, Mendes L, Ramos JA, Pérez-Tris J. Host-parasite associations and host-specificity in haemoparasites of reed bed passerines. Parasitol. 2012;139:310–6.View ArticleGoogle Scholar
  56. Marzal A, Bensch S, Reviriego M, Balbontin J, De Lope F. Effects of malaria double infection in birds: one plus one is not two. J Evol Biol. 2008;21:979–87.View ArticlePubMedGoogle Scholar
  57. Piersma T, van der Velde M. Dutch house martins Delichon urbicum gain blood parasite infections over their lifetime, but do not seem to suffer. J Ornithol. 2012;153:907–12.View ArticleGoogle Scholar
  58. von Rönn JA, Harrod C, Bensch S, Wolf JB. Transcontinental migratory connectivity predicts parasite prevalence in breeding populations of the European barn swallow. J Evol Biol. 2015;28:535–46.View ArticleGoogle Scholar
  59. Valkiūnas G, Palinauskas V, Ilgūnas M, Bukauskaitė D, Dimitrov D, Bernotienė R, et al. Molecular characterization of five widespread avian haemosporidian parasites (Haemosporida), with perspectives on the PCR-based detection of haemosporidians in wildlife. Parasitol Res. 2014;113:2251–63.View ArticlePubMedGoogle Scholar
  60. Valkiūnas G, Palinauskas V, Križanauskienė A, Bernotienė R, Kazlauskienė R, Iezhova TA. Further observations on in vitro hybridization of hemosporidian parasites: patterns of ookinete development in Haemoproteus spp. J Parasitol. 2013;99:124–36.View ArticlePubMedGoogle Scholar
  61. Valkiūnas G, Iezhova TA, Palinauskas V, Ilgūnas M, Bernotienė R. The evidence for rapid gametocyte viability changes in the course of parasitemia in Haemoproteus parasites. Parasitol Res. 2015;114:2903–9.View ArticlePubMedGoogle Scholar
  62. Valkiūnas G, Kazlauskienė R, Bernotienė R, Bukauskaitė D, Palinauskas V, Iezhova TA. Haemoproteus infections (Haemosporida, Haemoproteidae) kill bird-biting mosquitoes. Parasitol Res. 2014;113:1011–8.View ArticlePubMedGoogle Scholar
  63. Bentz S, Rigaud T, Barroca M, Martin-Laurent F, Bru D, Moreau J, et al. Sensitive measure of prevalence and parasitemia of haemosporidia from European blackbird (Turdus merula) populations: value of PCR-RFLP and quantitative PCR. Parasitol. 2006;133:685–92.View ArticleGoogle Scholar
  64. Drovetski SV, Aghayan SA, Mata VA, Lopes RJ, Mode NA, Harvey JA, et al. Does the niche breadth or trade-off hypothesis explain the abundance—occupancy relationship in avian Haemosporidia? Mol Ecol. 2014;23:3322–9.View ArticlePubMedGoogle Scholar
  65. Martínez-de la Puente J, Muñoz J, Capelli G, Montarsi F, Soriguer R, Arnoldi D, et al. Avian malaria parasites in the last supper: identifying encounters between parasites and the invasive Asian mosquito tiger and native mosquito species in Italy. Malar J. 2015;14:32.View ArticlePubMedPubMed CentralGoogle Scholar
  66. Mata VA, da Silva LP, Lopes RJ, Drovetski SV. The Strait of Gibraltar poses an effective barrier to host-specialised but not to host-generalised lineages of avian Haemosporidia. Int J Parasitol. 2015;45:711–9.View ArticlePubMedGoogle Scholar
  67. Olias P, Wegelin M, Zenker W, Freter S, Gruber AD, Klopfleisch R. Avian malaria deaths in parrots, Europe. Emerg Infect Dis. 2011;17:950–2.View ArticlePubMedPubMed CentralGoogle Scholar
  68. Synek P, Albrecht T, Vinkler M, Schnitzer J, Votýpka J, Munclinger P. Haemosporidian parasites of a European passerine wintering in South Asia: diversity, mixed infections and effect on host condition. Parasitol Res. 2013;112:1667–77.View ArticlePubMedGoogle Scholar
  69. Žiegyte R, Bernotiene R, Palinauskas V, Valkiūnas G. Haemoproteus tartakovskyi (Haemoproteidae): complete sporogony in Culicoides nubeculosus (Ceratopogonidae), with implications for avian haemoproteid experimental research. Exp Parasitol. 2016;160:17–22.View ArticlePubMedGoogle Scholar
  70. Palinauskas V, Iezhova TA, Križanauskienė A, Markovets MY, Bensch S, Valkiūnas G. Molecular characterization and distribution of Haemoproteus minutus (Haemosporida, Haemoproteidae): A pathogenic avian parasite. Parasitol Int. 2013;62:358–63.View ArticlePubMedGoogle Scholar
  71. Zehtindjiev P, Križanauskienė A, Scebba S, Dimitrov D, Valkiūnas G, Hegemann A, et al. Haemosporidian infections in skylarks (Alauda arvensis): a comparative PCR-based and microscopy study on the parasite diversity and prevalence in southern Italy and the Netherlands. Eur J Wildlife Res. 2011;58:335–44.View ArticleGoogle Scholar
  72. Bensch S, Hellgren O, Pérez-Tris J. MalAvi: a public database of malaria parasites and related haemosporidians in avian hosts based on mitochondrial cytochrome blineages. Mol Ecol Res. 2009;9:1353–8.View ArticleGoogle Scholar
  73. Palinauskas V, Žiegytė R, Ilgūnas M, Iezhova TA, Bernotienė R, Bolshakov C, et al. Description of the first cryptic avian malaria parasite, Plasmodium homocircumflexum n. sp. with experimental data on its virulence and development in avian hosts and mosquitoes. Int J Parasitol. 2015;45:51–62.View ArticlePubMedGoogle Scholar
  74. Levin II, Zwiers P, Deem SL, Geest EA, Higashiguchi JM, Iezhova TA, et al. Multiple lineages of avian malaria parasites (Plasmodium) in the Galapagos Islands and evidence for arrival via migratory birds. Conserv Biol. 2013;27:1366–77.View ArticlePubMedGoogle Scholar
  75. Farajollahi A, Fonseca DM, Kramer LD, Kilpatrick AM. ’Bird biting‘mosquitoes and human disease: a review of the role of Culex pipiens complex mosquitoes in epidemiology. Infect Genet Evol. 2011;11:1577–85.View ArticlePubMedPubMed CentralGoogle Scholar
  76. Simpson JE, Hurtado PJ, Medlock J, Molaei G, Andreadis TG, Galvani AP, et al. Vector host-feeding preferences drive transmission of multi-host pathogens: West Nile virus as a model system. Proc R Soc. 2012;279:925–33.View ArticleGoogle Scholar
  77. Kilpatrick AM, Kramer LD, Jones MJ, Marra PP, Daszak P. West Nile virus epidemics in North America are driven by shifts in mosquito feeding behaviour. PLoS Biol. 2006;4:e82.View ArticlePubMedPubMed CentralGoogle Scholar
  78. Bernotiene R, Palinauskas V, Iezhova T, Murauskaite D, Valkiūnas G. Avian haemosporidian parasites (Haemosporida): a comparative analysis of different polymerase chain reaction assays in detection of mixed infections. Exp Parasitol. 2016;163:31–7.View ArticlePubMedGoogle Scholar
  79. Telford SR. Hemoparasites of Reptilia: Color Atlas and Text. 1st ed. Boca Raton: CRC Press; 2009.Google Scholar

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