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Article

Blood Parasites (Haemosporida, Trypanosomatida) in Culex pipiens: A Study and Review of Hibernating and Active Mosquitoes

by
Kristina Valavičiūtė-Pocienė
1,*,
Margarita Kazak
1,
Tatjana Iezhova
2,
Gabrielė Kalinauskaitė
1 and
Rasa Bernotienė
1,*
1
Laboratory of Entomology, Nature Research Centre, Akademijos 2, LT-08412 Vilnius, Lithuania
2
P. B. Šivickis Laboratory of Parasitology, Nature Research Centre, Akademijos 2, LT-08412 Vilnius, Lithuania
*
Authors to whom correspondence should be addressed.
Microbiol. Res. 2024, 15(4), 2184-2198; https://doi.org/10.3390/microbiolres15040146
Submission received: 3 September 2024 / Revised: 21 October 2024 / Accepted: 23 October 2024 / Published: 25 October 2024
(This article belongs to the Special Issue Veterinary Microbiology and Diagnostics)
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Abstract

:
Culex pipiens mosquitoes (Diptera: Culicidae) are widespread during warm periods and actively feed on blood while serving as vectors for various human and animal pathogens. Culex mosquitoes overwinter as adults in temperate zones, raising the question of whether hibernating Cx. pipiens can act as pathogen reservoirs. In this study, hibernating mosquitoes and mosquitoes collected during the warm season were tested for the presence of trypanosomatids and avian haemosporidian parasites using PCR. Midgut preparations were made from Cx. pipiens females in order to search for trypanosomatids morphologically. In total, 1037 Cx. pipiens mosquitoes (556 collected during the warm season and 481 overwintering mosquitos) were investigated. The parasite prevalence differed for mosquitoes collected during the warm season and hibernating ones for both Haemosporida (2.9% in warm-season and no infections in overwintering mosquitoes) and Trypanosomatida (1.6% and 0.4%, respectively) parasites. A phylogenetic analysis confirmed that the trypanosomatids found in hibernating mosquitoes were monoxenous and were not parasites of vertebrates. The peak prevalence of Haemosporida parasites was detected in July (4.9%) and August (2.8%), and for Trypanosomatida, it was detected in May (3.5%). The results of the present study show that overwintering Cx. pipiens mosquitoes are questionable reservoirs for avian haemosporidian parasites, but some monoxenous trypanosomatids can be found in overwintering females.

1. Introduction

Even though mosquitoes have been researched for over a century (ever since Sir Patrick Manson demonstrated in 1877 that they can transmit filarial worms [1] and Sir Ronald Ross discovered in 1897 that Culicinae mosquitoes are a part of the avian malaria life cycle [2]), there is still a significant gap in our knowledge regarding the vector competence of various mosquito species in Europe [3]. The mosquito species Culex pipiens is the most frequently encountered when searching for information on the transmission of various vertebrate pathogens [4,5,6,7]. West Nile (WNV), Ockelbo, Usutu, Sindbis and Equine encephalitis viruses are spread by Culex mosquitoes [6,8,9,10,11]. These mosquitoes are also known as vectors of several Trypanosoma species [4,5,12], avian haemosporidian parasites belonging to the genus Plasmodium [3,7], as well as pathogens causing lymphatic filariasis [13,14]. This is not a complete list of the various pathogens that are known to be spread by Culex mosquitoes [8,9,10,11,12,13,14]. Avian haemosporidian and Trypanosoma parasites are widespread and infect birds from a broad variety of avian families with diverse consequences, ranging from subclinical infections to severe and fatal disease [15]. The prevalence of Plasmodium and Trypanosoma parasites in passerine birds in Lithuania is about 5% and 26% respectively [16], so these parasites can serve as model organisms to study the biology of vector-borne pathogens.
In a temperate climate, the females of Culex, Anopheles and Culiseta can hibernate as adults in various shelters (cellars, bunkers, abandoned buildings, tree holes) [11], and within Culex genera in our study area, Cx. pipiens, Cx. torrentium and Cx. territans hibernate as adult females. Culex pipiens f. pipiens females are anautogenous (they require a blood meal for their egg laying) and, depending on the climatic conditions, can develop several generations per year [11]. Males of this species die during winter, while females search for above-ground overwintering sites [17,18]. Hibernating mosquito females do not express host-seeking behavior and use the accumulated fat body as an energy source. Even though there are studies about Cx. pipiens’s inability to use blood as a source for lipid reserves for overwintering [19], other studies suggest that 40% of female Cx. pipiens who were subjected to warmer temperature periods took a full blood meal and did not undergo ovarian maturation, which gives them an equal chance of surviving [20]. There is evidence of blood-feeding in diapausing Cx. pipiens females in the wild [21]. Because of the fact that Culex females overwinter as adults, it raises the question if (1) overwintering mosquitoes can be winter reservoirs for some pathogens or if (2) mosquitoes are newly reinfected each spring by feeding on the blood of infected hosts. The answer to this question is especially important when studying the biology of pathogens that are transmitted by Culex mosquitoes and cannot be transmitted via vertical transmission.
There are studies molecularly confirming the presence of viruses in overwintering mosquitoes—WNV RNA was found in overwintering Cx. tarsalis [22] and Cx. p. pipiens females [23]. Unlike some arboviruses, Protozoa parasites transmitted by Culex mosquitoes cannot be transmitted via transovarial transmission. Many studies provide information on the prevalence of avian haemosporidian parasites in wild-caught mosquitoes collected during the warm season [3,7,24], but there is almost no knowledge of whether the parasites persist in hibernating mosquitoes. We tested hibernating Cx. pipiens females for the presence of avian Trypanosoma and haemosporidian parasite DNA and compared our results with results obtained from active Cx. pipiens mosquitoes caught during the warm period at similar localities. For this purpose, we employed microscopy and PCR-based methods. We aimed to understand if hibernating Cx. pipiens mosquitoes can be infected with avian Trypanosoma and haemosporidian parasites and in this way act as reservoirs for these parasites.

2. Materials and Methods

2.1. Collection of Mosquitoes

Hibernating mosquitoes were collected in December and January 2023–2024 using mouth aspirators in damp cellars in South Lithuania, Norkūnai village (54.500694, 23.940056), Panemuninkėliai village (54.431389, 24.068250), West Lithuania, Palanga (55.918278, 21.055639) and Vilnius (bunkers in Vilnius (54.700444, 25.332583 and 54.688764, 25.362232) and in the cellar of Verkiai water tower (54.749139, 25.291444)).
Culex pipiens mosquitoes were caught regularly during the warm period (when the average daily temperature was over 10 °C) from May to September using a sweeping net (2021) in Vilnius environs (Dvarčionys 54.736569, 25.384285, Belmontas 54.688764, 25.362232) and CDC trap-baited with CO2 (2022–2023) in South Lithuania Puvočiai (54.112762, 24.302303) and in Vilnius environs (Kairėnai 54.733956, 25.403669, Verkiai 54.748405, 25.289150) and Brinkiškės (54.798850, 25.059928). All investigated localities were mixed forests with small water bodies (ponds, puddles, marshes). Hibernating mosquitoes were collected at similar habitats with cellars as their hibernating places. The material was transported to the laboratory the same day after collection for further processing.

2.2. Mosquito Identification Dissection and Microscopic Examination of Samples

All collected mosquitoes were placed in a freezer for 10–15 min to make them inactive; after that, mosquito females were identified according to Becker et al. and Gunay et al.’s identification keys [11,25] under a stereomicroscope, MOTIC SMZ-171 (Motic, Hong Kong, China). Only Culex mosquito females were further investigated.
Females were dissected immediately after identification for (a) salivary gland [24] and (b) midgut preparations [15,26]. Salivary glands were extracted from the anterior part of the thorax and were used to search for invasive stages of haemosporidian parasites (sporozoites) as described in [24]. Midgut preparations were used to search for Trypanosoma parasites [15,26]: the last two–three segments of a mosquito’s abdomen were gently pulled together with the digestive tract. Half of the midgut was separated from the rest of the organs, pulled into pieces, spread in a thin layer on a microscopic slide, and set aside to dry at room temperature. For each dissection, a fire-sterilized pair of needles was used. Slides were then fixed with a drop of absolute methanol and stained for one hour with a 4% Giemsa solution [15]. The remaining parts of the mosquitoes were placed individually in an SET buffer (solution of Sodium, Ethylenediaminetetraacetic acid (EDTA) and Tris-HCl) for further DNA extraction (see Section 2.3). Data on the prevalence of haemosporidian parasites from the same mosquitoes collected during the warm period have been published as a part of another investigation [24].
The midgut preparations of mosquitoes that were determined to be PCR-positive for trypanosomatid DNA (see Section 2.3) were later examined microscopically. Microscopy was performed using an Olympus BX-43 microscope (Olympus, Tokyo, Japan) equipped with an Olympus DP12 digital camera and image software, Olympus DP-SOFT v.3.2 (Olympus, Tokyo, Japan). Entire samples were screened at ×1000 magnification. Positive slides (49805 NS–49807 NS) were deposited at the Nature Research Centre (Akademijos str. 2, Vilnius, Lithuania).
Culex pipiens and Cx. torrentium mosquito females can only be distinguished by molecular methods. To separate these two species, we used a method which was based on the use of restriction enzymes [27]. After DNA extraction (see Section 2.3) and PCR, the COI region of mitochondrial DNA is cleaved differently by two restriction enzymes, and the results are visualized on the agarose gel. The SspI enzyme cuts a COI fragment of Cx. pipiens into two (~620 pb and 210 bp), and the COI fragment of Cx. torrentium remains uncut. With the enzyme FspBI, this is the opposite of a previously described one [27]. This method enables the distinction of two sibling species, Cx. pipiens and Cx. torrentium, without sequencing.

2.3. DNA Extraction, PCR and Sequencing

We expected a lower prevalence of parasites in hibernating mosquitoes, so they were pooled (up to 10 individuals in one pool collected at the same locality on the same day) for molecular analysis, and active mosquitoes were investigated individually. The total DNA from pools of hibernating Culex females was extracted using an ammonium acetate protocol [28], and DNA from active mosquitoes was extracted using an ammonium acetate protocol or GeneJET Genomic DNA Purification Kit (Thermo Fisher Scientific, Vilnius, Lithuania) as described in [24]. Both DNA extraction methods have been used before and are equally effective.
For the detection of haemosporidian parasites, a nested PCR protocol with two pairs of primers (outer primers: HaemNFI (5′-CATATATTAAGAGAAITATGGAG-3′) and HaemNR3 (5′-ATAGAAAGATAAGAAATACCATTC-3′); inner primers: HaemF (5′-ATGGTGCTTTCGATATATGCATG-3′) and HaemR2 (5′-GCATTATCTGGATGTGATAATGGT-3′)) was used to amplify a fragment of the cytochrome b gene (cytb) [29,30]. For trypanosomatid detection, a nested PCR protocol with two pairs of primers (outer primers: Tryp763 (5′-180 CATATGCTTGTTCAAGGAC-3′) and Tryp1016 (5′-CCCCATAATCTCCAATGGAC-3′); inner primers: Tryp99 (5′-TCAATCAGACGTAATCTGCC-3′) and Tryp957 (5′-182 CTGCTCCTTTGTTATCCCAT-3′)) was used to amplify a DNA fragment encoding SSU 18S rRNA [31,32]. For both types of PCR, a negative control (nuclease-free water) and a positive control (extracted DNA from bird blood, infected with Plasmodium (in the investigation of haemosporidian parasites) or Trypanosoma (in the search for tripanodomatids) parasites confirmed both by PCR and microscopy) were included in every 46 samples. The PCR products were visualized in a 2% agarose gel using MidoriGreen dye (NIPPON Genetics Europe, Düren, Germany). Samples with visible bands (~500 bp for haemosporidians and ~750 bp for trypanosomatids) were considered positive, and the PCR product was precipitated using an ammonium acetate protocol [28] and sent for sequencing. Purified PCR-positive samples were sequenced using an Applied Biosystems Genetic Analyzer 3500 in both strands with inner primers. The obtained sequences were deposited in GenBank (PP946099-PP946107; PP948731, PP948732).

2.4. Analysis of Sequences and Statistics

The obtained chromatograms were analyzed with Geneious Prime (2023.0.4), both strands were aligned, and contig sequences were formed. The analysis of the haemosporidian sequences was described in a previous article [24]. For the identification of trypanosomatid sequences, GenBank BLAST (National Centre of Biotechnology Information website: https://www.ncbi.nlm.nih.gov/BLAST, accessed on 4 June 2024) was used.
A phylogenetic tree was prepared using Geneious Prime (2023.0.4) software, while jModeltest-2.1.10 software suggested the best fit model (GTR+I+G) for further phylogenetic analysis. The phylogenetic tree was constructed using the MrBayes plugin v3.2.6. The analysis was run for 6 million generations (sampling frequency: every 100th generation), and 25% of the initial trees were discarded as a ‘burn-in’ period before the construction of the consensus tree.
Fisher’s exact test was used to compare the prevalences of haemosporidian and Trypanosoma parasites between hibernating and active mosquitoes.

3. Results

Overall, 1037 Cx. pipiens females were processed: 556 individuals were collected during the warm season and 481 hibernating females from wintering places (Table S1). The peak month with the highest numbers of caught Cx. pipiens mosquitoes was July (39.5%), while in August and in June, this amount was 31.1% and 23.9%, respectively (Figure 1). May and September were the months with the least numbers of Cx. pipiens caught, with the percentages being 5.0% and 0.3%, respectively. Based on the results of the restriction enzyme tests [27], we did not find Cx. torrentium, and all investigated mosquitoes were Cx. pipiens. While collecting hibernating Culex, we also found hibernating Culiseta and Anopheles females, but they were not investigated, as the focus of the study was Culex mosquitoes.

3.1. Infections with Trypanosomatida Parasites

Overall, DNA of trypanosomatids was found in 11 (1.1%) Cx. pipiens females: 9 infected individuals were detected in the warm period (1.6%), and 2 infected individuals were collected from overwintering sites (0.4%). The difference between the prevalence of Trypanosoma parasites in hibernating and active mosquitoes was statistically significant (p = 0.005). Trypanosomatids were identified based on the similarity of obtained sequences. Trypanosoma culicavium (Figure 2A) (obtained sequence shows 99.14% similarity to a sequence deposited in GenBank—OM509727) DNA was found in five mosquitoes, Trypanosoma theileri was detected in three individuals, Trypanosoma trinaperronei (Figure 2B) (99.93% similarity to a sequence from GenBank—MN752212) was found in one mosquito, and two hibernating mosquito females were infected with unidentified monoxenous trypanosomatids (Figure 2C), as determined by the phylogenetic analysis and the similarity of obtained sequences (99.51% similarity to a sequence deposited in GenBank—OP748978; Table 1).
The phylogenetic analysis (Figure 3) was used to determine the phylogenetic position of unidentified trypanosomatids, and it showed that all detected trypanosomatids formed two clades: the Trypanosoma species clustered into clade A, and the monoxenous trypanosomatids (Crithidia and unidentified Trypanosomatidae) clustered into clade B. Within clade A, avian Trypanosoma (T. culicavium, T. bennetti, T. everetti, T. thomasbancrofti, T. avium) subclades stand apart from the Trypanosoma of mammals (T. theileri, T. trinaperronei).
All Trypanosoma species detected by PCR were obtained microscopically in the midgut preparations of investigated mosquitoes (Figure 2), except for T. theileri. The Trypanosomatida infection prevalence was the highest in May (3.5%), while during summer, it varied between 1.4% (July) and 1.7% (August).

3.2. Infections with Haemosporidian Parasites

No hibernating mosquitoes were detected to be infected with haemosporidian parasites. There were 16 infected individuals of Cx. pipiens (2.9%) collected during the warm period (Figure 1). The difference between the prevalence of haemosporidian parasites in hibernating and active mosquitoes was statistically significant (p = 0.0002). Three Plasmodium species (Plasmodium homonucleophilum (pSW2), P. matutinum (pLINN1), P. vaughani (pSYAT05)), two Haemoproteus species (Haemoproteus brachiatus (hLK03), H. asymmetricus (hTUPHI01)) and one mixed infection of Leucocytozoon/Plasmodium were identified in the collected mosquitoes.
The highest prevalence of haemosporidian parasites in Cx. pipiens mosquitoes was detected in July (4.9%) and August (2.8%).

4. Discussion

The main finding of this research was that when screening a similar number of Cx. pipiens mosquito females collected during the warm season and in winter for the presence of Trypanosoma and avian haemosporidian parasites, we found significantly different prevalences. Haemosporidian and Trypanosoma parasites were not detected in hibernating mosquitoes, while the prevalence of the same parasites in Cx. pipiens females collected during warm periods was 1.6% (nine positive mosquitoes) for Trypanosoma and 2.9% (16 positive mosquitoes) for haemosporidian parasites.
We detected three Trypanosoma species in the mosquitoes collected during the warm period (Table 1). Until now, quite a high diversity of trypanosomatids has been found in Cx. pipiens mosquitoes worldwide (Table 2). Two Trypanosoma species are known to be transmitted by these mosquitoes: T. culicavium, originally found and described from the gut of Cx. pipiens mosquitoes [5,33], and T. thomasbancrofti [4,12,33], for which Cx. pipiens was confirmed as a vector [4]. Other trypanosomatids have been periodically detected in individuals of this mosquito species, including T. avium [34,35,36] and T. theileri [37]. It was reported that among the avian trypanosomes transmitted by mosquitoes, the prevalence of T. thomasbancrofti in wild-caught mosquitoes, as well as in birds, was lower (0.13%) compared with the prevalence of T. culicavium (4.5%) [4]. The overall prevalence of T. thomasbancrofti in wild-caught mosquitoes in Europe is low [4,35,36], and it is not surprising that during our study, we did not detect this species in Culex mosquitoes.
Trypanosoma theileri was one of the first mammalian trypanosomatids described, and vectors of these parasites are known to be deer keds [39] and tabanids [40]. Information on the prevalence of this parasite in mosquitoes is scarce, with recent studies [4] indicating a prevalence of only 0.05% in Culex, while in Aedes species, it can reach nearly 22%. Our findings demonstrate a higher prevalence of T. theileri in Culex mosquitoes (0.72%), which was confirmed by PCR, although not microscopically (Table 1). Despite dissecting and preparing midgut preparations, it is possible that the preference of the T. theileri group trypanosomes to develop in the hindgut [37] may explain this discrepancy.
Currently, there are several species of Megatrypanum trypanosomes parasitizing deers: Trypanosoma mazamarum [41], T. cervi [42], T. stefanskii [43] and T. trinaperronei [39]. Together with T. theileri, T. melophagium, T. cervi and T. trinaperronei compose a Trypanosoma theileri complex [39,44,45]. Deer keds (Lipoptena cervi and L. mazamae) are known vectors of T. trinaperronei [39], and previously, this parasite has not been found in mosquitoes. We found one Cx. pipiens mosquito infected with T. trinaperronei (Table 1, Figure 2B), and the type of host of this trypanosome species is known to be Odocoileus virginianus (Ruminantia, Cervidae), white-tailed deer [39].
Various monoxenous trypanosomatids, such as several Crithidia species (C. brevicula [36], C. dedva [34], C. fasciculata [35] and C. dobrovolskii), Strigomonas cf. oncopelti, Paratrypanosoma cf. confusum and an unknown lineage of Trypanosomatidae [22], have also been detected. Monoxenous trypanosomatids are generally considered non-pathogenic; however, there is some evidence suggesting negative effects on insect fitness, although this has only been investigated in a few insect species [46,47]. These insect parasites are not limited to blood-feeding habits and can infect hosts through numerous means, such as feeding on infected prey or ingesting contaminated substances like fresh feces (i.e., coprophagy) or contaminated substrates, such as sugar meals [46,47]. Therefore, their detection in overwintering mosquitoes does not indicate previous blood-feeding before the wintering period. However, reports on monoxenous parasites such as Crithidia spp. infecting humans and non-human mammalian hosts alone and in co-infections with Leishmania parasites are increasing in number and consistency each year [48]. Together with the reported cases of non-human trypanosome infections in humans [49], these findings underline the importance of investigating the transmission of trypanosomatids between hosts. Recently, Kostygov et al. [50] discovered an unknown lineage representing a new monoxenous trypanosomatid that has not previously been documented, detected in overwintering Cx. torrentium mosquitoes. Both Cx. torrentium and Cx. pipiens mosquitoes feed on birds and mammals, including humans, making them potential bridge vectors for the transmission of zoonotic pathogens from birds to humans [51]. During our study, two overwintering Cx. pipiens females were found to be infected with an unknown trypanosomatid showing a 98.16% and 99.51% similarity to a parasite found in Cx. torrentium mosquitoes [50].
There seem to be no data on the seasonality of avian trypanosomes, especially in vectors. In 1947, Vanderplank described the seasonal variation in the numbers of mammals with Trypanosoma infections in their blood: infections seem to be the highest during the raining periods, March–April and November–December [52]. Therefore, Pori et al. found no seasonal variations in the avian trypanosome infection prevalence from a total of 685 birds of 87 species [53].
There are at least 16 species and 50 genetic lineages of Plasmodium detected in Cx. pipiens mosquitoes, whereas sporozoites of only 10 Plasmodium species, as well as one genetic lineage not assigned to a species (pCXPIP09), have been found in the salivary glands of Cx. pipiens (Table 3). Sporozoites are the invasive stage of haemosporidian parasites, and the finding of sporozoites in the salivary glands of mosquitos shows that the parasite has finished its development and can be transmitted to another host [54]. Solely PCR-based detection of parasite DNA shows that the mosquito had a blood meal on an infected host but does not show that the mosquito is a vector of a parasite, as parasite DNA can be detected from infected blood meal, indicating the abortive development of a parasite [54]. This is the case when Haemoproteus or Leucocytozoon parasites are detected in Culex mosquitoes [24,54], as they were in this research. Even though mosquitoes are a dead end for parasites of other haemosporidian genera (Haemoproteus and Leucocytozoon) [15], the collection of such data provides new information on the mosquitos’ feeding preferences, as Haemoproteus parasites are specific to vertebrate hosts [15].
There are almost no data on the detection of haemosporidian parasites in hibernating mosquitoes. Plasmodium matutinum (pLINN1) was found in two pools of mosquitoes collected in their hibernation sites in November using PCR [100]. November is just the beginning of the hibernation period for Culex mosquitoes in the temperate zone of the northern hemisphere, and mosquitoes harboring parasites would need to survive at least until April to be able to transmit the parasite to an avian host. We collected hibernating mosquitoes in December and January and collected them in similar parts (south and east) of the country as the active mosquitoes were collected in, except for one locality in western Lithuania. Avian haemosporidian parasites are investigated in the western part of the country, showing high prevalences in different bird species [15,26,46], so we expected to find at least some infected hibernating mosquitoes there. However, we did not find haemosporidian and Trypanosoma parasites in the hibernating mosquitoes. According to different authors, the prevalence of haemosporidian parasites in active Cx. pipiens mosquitoes varies between 0.04% and 6.6% in different studies (0.04%—[88]; 0.52%—[56]; 0.61%—[98]; 3.08%—[60]; 6.6%—[64]), and a 2.9% prevalence was detected in Cx. pipiens active mosquitoes in Lithuania, which falls in the middle [24]. Although all sites of this investigation were similar habitats, the prevalence of haemosporidian parasites differed between study sites and was high in Verkiai (7%) during the warm period. Both active and hibernating mosquitoes were tested in this location, with no infected hibernating mosquitoes being found. The relationship between the study area and parasite prevalence in mosquitoes should be the focus of further research.
Culex pipiens mosquitoes are considered to be mainly ornithophilic [79,101]. The presence of avian haemosporidian DNA in the tested Cx. pipiens females show that these mosquitoes had a blood meal on birds, as this is the only way for mosquitoes to be infected with these parasites [54]. Trypanosoma culicavium is an avian trypanosomatid [5], further proving their ornithophilic tendencies. On the other hand, T. theileri [37] and T. trinaperronei [39] can only develop in mammals, indicating the mammalophilic or opportunistic behavior of Cx. pipiens mosquitoes. Such blood-sucking insects can serve as bridge vectors between different groups of animals, like Cx. pipiens being a bridge vector for WNV from birds to humans [102].
In summary, our investigation did not detect avian haemosporidian or avian Trypanosoma parasites in hibernating Cx. pipiens mosquitoes. In contrast, during the warm season, the prevalence of these parasites was 2.9% for haemosporidian parasites [20] and 1.6% for avian trypanosomatid parasites. Only monoxenous trypanosomatids were detected in overwintering Cx. pipiens mosquitoes. Studies conducted with a larger number of mosquitoes over several years may help to better assess the role of hibernating mosquitoes in avian blood parasite transmission. Given the potential for these parasites to survive in wintering mosquitoes, experimental studies are needed to determine whether the parasites fail to survive in mosquitoes or whether mosquitoes infected with parasites do not survive until spring.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/microbiolres15040146/s1, Table S1: Investigated Culex pipiens mosquitoes with collecting time and location.

Author Contributions

Conceptualization, K.V.-P. and R.B.; methodology, K.V.-P. and R.B.; formal analysis, K.V.-P., G.K., M.K. and R.B.; investigation, K.V.-P., G.K., M.K. and T.J.; resources, R.B.; data curation, K.V.-P., G.K. and M.K.; writing—original draft preparation, K.V.-P., M.K. and R.B.; writing—review and editing, K.V.-P., G.K., M.K., T.J. and R.B.; visualization, K.V.-P., M.K. and T.J.; supervision, R.B.; project administration, R.B.; funding acquisition, R.B. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the grant No. S-MIP-22-50 by the Research Council of Lithuania.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data is contained within the article or Supplementary Materials.

Acknowledgments

The authors would like to thank Miglė Kazlauskaitė for help in collecting hibernating mosquitoes, Vida Petiukonienė for help with hibernating mosquito collection in Verkiai water tower, Alfonso Marzal for providing some of the mosquito traps for this research, and the staff of the Vilnius University Botanical Garden for giving us the possibility to set the traps. The procedures described herein comply with the current laws of Lithuania.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Manson, P. On the development of Filaria sanguinis hominis, and on the mosquito considered as a nurse. J. Linn. Soc. Lond. Zool. 1878, 14, 304–311. [Google Scholar] [CrossRef]
  2. Cox, F.E. History of the Discovery of the Malaria parasites and Their Vectors. Parasit. Vectors 2010, 3, 5. [Google Scholar] [CrossRef] [PubMed]
  3. Santiago-Alarcon, D.; Palinauskas, V.; Schaefer, H.M. Diptera Vectors of Avian Haemosporidian Parasites: Untangling Parasite Life Cycles and Their Taxonomy. Biol. Rev. 2012, 87, 928–964. [Google Scholar] [CrossRef] [PubMed]
  4. Fialová, M.; Santolíková, A.; Brotánková, A.; Brzoňová, J.; Svobodová, M. Complete Life Cycle of Trypanosoma thomasbancrofti, an Avian Trypanosome Transmitted by Culicine Mosquitoes. Microorganisms 2021, 9, 2101. [Google Scholar] [CrossRef] [PubMed]
  5. Votýpka, J.; Szabová, J.; Rádrová, J.; Zídková, L.; Svobodová, M. Trypanosoma culicavium sp. nov., an Avian Trypanosome Transmitted by Culex Mosquitoes. Int. J. Syst. Evol. Microbiol. 2012, 62, 745–754. [Google Scholar] [CrossRef]
  6. Brugman, V.A.; Hernández-Triana, L.M.; Medlock, J.M.; Fooks, A.R.; Carpenter, S.; Johnson, N. The Role of Culex pipiens L. (Diptera: Culicidae) in Virus Transmission in Europe. Int. J. Environ. Res. Public Health 2018, 15, 389. [Google Scholar] [CrossRef]
  7. Bensch, S.; Hellgren, O.; PÉrez-Tris, J. MalAvi: A Public Database of Malaria Parasites and Related Haemosporidians in Avian Hosts Based on Mitochondrial Cytochrome b Lineages. Mol. Ecol. Resour. 2009, 9, 1353–1358. [Google Scholar] [CrossRef]
  8. Nicolescu, G.A. General Characterisation of the Mosquito Fauna (Diptera: Culicidae) in the Epidemic Area for West Nile Virus in the South of Romania. Eur. Mosq. Bull. 1998, 2, 13–18. [Google Scholar]
  9. Lundstrom, J.O. Vector Competence of Western European Mosquitoes for Arboviruses: A Review of Field and Experimental Studies. J. Vector. Ecol. 1994, 19, 23–36. [Google Scholar]
  10. Engel, D.; Jöst, H.; Wink, M.; Börstler, J.; Bosch, S.; Garigliany, M.M.; Jöst, A.; Czajka, C.; Lühken, R.; Ziegler, U. Reconstruction of the Evolutionary History and Dispersal of Usutu Virus, a Neglected Emerging Arbovirus in Europe and Africa. mBio 2016, 7, 10–1128. [Google Scholar] [CrossRef]
  11. Becker, N.; Petrić, D.; Boase, C.; Lane, J.; Zgomba, M.; Dahl, C.; Kaiser, A. Mosquitoes and Their Control, 1st ed.; Springer: Berlin/Heidelberg, Germany, 2003. [Google Scholar]
  12. Šlapeta, J.; Morin-Adeline, V.; Thompson, P.; McDonell, D.; Shiels, M.; Gilchrist, K.; Votýpka, J.; Vogelnest, L. Intercontinental Distribution of a New Trypanosome Species from Australian Endemic Regent Honeyeater (Anthochaera phrygia). Parasitology 2016, 143, 1012–1025. [Google Scholar] [CrossRef]
  13. Shaikevich, E.; Bogacheva, A.; Ganushkina, L. Dirofilaria and Wolbachia in Mosquitoes (Diptera: Culicidae) in Central European Russia and on the Black Sea Coast. Parasite 2019, 26, 2. [Google Scholar] [CrossRef] [PubMed]
  14. Čabanová, V.; Miterpáková, M.; Valentová, D.; Blažejová, H.; Rudolf, I.; Stloukal, E.; Hurníková, Z.; Dzidová, M. Urbanization Impact on Mosquito Community and the Transmission Potential of Filarial Infection in Central Europe. Parasit. Vectors 2018, 11, 261. [Google Scholar] [CrossRef] [PubMed]
  15. Valkiūnas, G. Avian Malaria Parasites and Other Haemosporidia; CRC Press: Boca Raton, FL, USA, 2005; ISBN 9780874216561. [Google Scholar]
  16. Chagas, C.R.F.; Binkienė, R.; Ilgūnas, M.; Iezhova, T.; Valkiūnas, G. The Buffy Coat Method: A Tool for Detection of Blood Parasites without Staining Procedures. Parasit. Vectors 2020, 13, 104. [Google Scholar] [CrossRef]
  17. Becker, N.; Jöst, A.; Weitzel, T. The Culex pipiens Complex in Europe. J. Am. Mosq. Control Assoc. 2012, 28, 53–67. [Google Scholar] [CrossRef]
  18. Byrne, K.; Nichols, R.A. Culex pipiens in London Underground Tunnels: Differentiation between Surface and Subterranean Populations. Heredity 1999, 82, 7–15. [Google Scholar] [CrossRef]
  19. Mitchell, C.J.; Briegel, H. Inability of Diapausing Culex pipiens (Diptera: Culicidae) to Use Blood for Producing Lipid Reserves for Overwinter Survival. J. Med. Entomol. 1989, 26, 318–326. [Google Scholar] [CrossRef]
  20. Eldridge, B.F.; Baileyl, C.L. Experimental Hibernation Studies in Culex pipiens (Diptera: Culicidae): Reactivation of Ovarian Development and Blood-Feeding in Prehibernating Females. J. Med. Entomol. 1979, 15, 462–467. [Google Scholar] [CrossRef]
  21. Bailey, C.L.; Faran, M.E.; Gargan, T.P.; Hayes, D.E. Winter Survival of Blood-Fed and Nonblood-Fed Culex pipiens L. Am. J. Trop. Med. Hyg. 1982, 31, 1054–1061. [Google Scholar] [CrossRef]
  22. Nelms, B.M.; MacEdo, P.A.; Kothera, L.; Savage, H.M.; Reisen, W.K. Overwintering Biology of Culex (Diptera: Culicidae) Mosquitoes in the Sacramento Valley of California. J. Med. Entomol. 2013, 50, 773–790. [Google Scholar] [CrossRef]
  23. Farajollahi, A.; Crans, W.J.; Bryant, P.; Wolf, B.; Burkhalter, K.L.; Godsey, M.S.; Aspen, S.E.; Nasci, R.S. Detection of West Nile Viral RNA from an Overwintering Pool of Culex pipens pipiens (Diptera: Culicidae) in New Jersey. J. Med. Entomol. 2003, 42, 490–494. [Google Scholar] [CrossRef]
  24. Valavičiūtė-Pocienė, K.; Kalinauskaitė, G.; Chagas, C.R.F.; Bernotienė, R. Avian Haemosporidian Parasites from Wild-Caught Mosquitoes with New Evidence on Vectors of Plasmodium matutinum. Acta. Trop. 2024, 256, 107260. [Google Scholar] [CrossRef]
  25. Gunay, F.; Picard, M.; Robert, V. MosKeyTool, an Interactive Identification Key for Mosquitoes of Euro-Mediterranean, Version 2.1; Last Update: 1 August 2018. 2018. Available online: https://www.medilabsecure.com/entomology-tools-0/moskeytool (accessed on 10 March 2024). (In English).
  26. Žiegytė, R.; Markovets, M.Y.; Bernotienė, R.; Mukhin, A.; Iezhova, T.A.; Valkiūnas, G.; Palinauskas, V. The Widespread Biting Midge Culicoides impunctatus (Ceratopogonidae) Is Susceptible to Infection with Numerous Haemoproteus (Haemoproteidae) Species. Parasit. Vectors 2017, 10, 397. [Google Scholar] [CrossRef]
  27. Hesson, J.C.; LundströM, J.O.; Halvarsson, P.; Erixon, P.; Collado, A. A Sensitive and Reliable Restriction Enzyme Assay to Distinguish between the Mosquitoes Culex torrentium and Culex pipiens. Med. Vet. Entomol. 2010, 24, 142–149. [Google Scholar] [CrossRef]
  28. Sambrook, J.; Green, M.R. Molecular Cloning: A Laboratory Manual; Cold Spring Harbor Laboratory: Cold Spring Harbor, NY, USA, 2012; Volume 1, ISBN 9781936113415. [Google Scholar]
  29. Bensch, S.; Stjernman, M.; Hasselquist, D.; Ostman, O.; Hansson, B.; Westerdahl, H.; Pinheiro, R.T. Host Specificity in Avian Blood Parasites: A Study of Plasmodium and Haemoproteus Mitochondrial DNA Amplified from Birds. Proc. R. Soc. Lond. B Biol. Sci. 2000, 267, 1583–1589. [Google Scholar] [CrossRef]
  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. [Google Scholar] [CrossRef]
  31. Sehgal, R.N.M.; Iezhova, T.A.; Marzec, T.; Valkiūnas, G. Trypanosoma naviformis Sp. Nov. (Kinetoplastidae: Trypanosomatidae) from Widespread African Songbirds, the Olive Sunbird (Cyanomitra olivacea) and Yellow-Whiskered Greenbul (Andropadus latirostris). Zootaxa 2015, 4034, 342–350. [Google Scholar] [CrossRef]
  32. Valkiūnas, G.; Iezhova, T.A.; Carlson, J.S.; Sehgal, R.N.M. Two New Trypanosoma Species from African Birds, with Notes on the Taxonomy of Avian Trypanosomes. J. Parasitol. 2011, 97, 924–930. [Google Scholar] [CrossRef]
  33. Zídková, L.; Cepicka, I.; Szabová, J.; Svobodová, M. Biodiversity of Avian Trypanosomes. Infect. Genet. Evol. 2012, 12, 102–112. [Google Scholar] [CrossRef] [PubMed]
  34. Schoener, E.R.; Harl, J.; Himmel, T.; Fragner, K.; Weissenböck, H.; Fuehrer, H.P. Protozoan Parasites in Culex pipiens Mosquitoes in Vienna. Parasitol. Res. 2019, 118, 1261–1269. [Google Scholar] [CrossRef]
  35. Schoener, E.; Uebleis, S.S.; Cuk, C.; Nawratil, M.; Obwaller, A.G.; Zechmeister, T.; Lebl, K.; Rádrová, J.; Zittra, C.; Votýpka, J. Trypanosomatid Parasites in Austrian Mosquitoes. PLoS ONE 2018, 13, e0196052. [Google Scholar] [CrossRef]
  36. Svobodová, M.; Volf, P.; Votýpka, J. Trypanosomatids in Ornithophilic Bloodsucking Diptera. Med. Vet. Entomol. 2015, 29, 444–447. [Google Scholar] [CrossRef]
  37. Brotánková, A.; Fialová, M.; Čepička, I.; Brzoňová, J.; Svobodová, M. Trypanosomes of the Trypanosoma theileri Group: Phylogeny and New Potential Vectors. Microorganisms 2022, 10, 294. [Google Scholar] [CrossRef]
  38. Van Dyken, M.; Bolling, B.G.; Moore, C.G.; Blair, C.D.; Beaty, B.J.; Black IV, W.C.; Foy, B.D. Molecular Evidence for Trypanosomatids in Culex Mosquitoes Collected during a West Nile Virus Survey. Int. J. Parasitol. 2006, 36, 1015–1023. [Google Scholar] [CrossRef]
  39. Garcia, H.A.; Blanco, P.A.; Rodrigues, A.C.; Rodrigues, C.M.F.; Takata, C.S.A.; Campaner, M.; Camargo, E.P.; Teixeira, M.M.G. Pan-American Trypanosoma (Megatrypanum) trinaperronei n. sp. In the White-Tailed Deer Odocoileus virginianus Zimmermann and Its Deer Ked Lipoptena mazamae Rondani, 1878: Morphological, Developmental and Phylogeographical Characterisation. Parasit. Vectors 2020, 13, 308. [Google Scholar] [CrossRef]
  40. Böse, R.; Friedhoff, K.T.; Olbrich, S. Transmission of Megatrypanum Trypanosomes to Cervus dama by Tabanidae. J. Protozool. 1987, 34, 110–113. [Google Scholar] [CrossRef]
  41. Mazza, S.; Romana, C.; Fiora, A. Algunos Hemoparásitos de Mamíferos Del Norte. VII Reun. Soc. Argent. Patol. Reg. Norte 1932, 2, 990–997. [Google Scholar]
  42. Kingston, N.; Morton, J.K. Trypanosoma cervi sp. n. from Elk (Cervus canadensis) in Wyoming. J. Parasitol. 1975, 61, 17–23. [Google Scholar] [CrossRef]
  43. Kingston, N.; Bobek, B.; Perzanowski, K.; Wita, I.; Makis, L. Description of Trypanosoma (Megatrypanum) stefanskii sp. n. from Roe Deer (Capreolus capreolus) in Poland. J. Helminthol. Soc. Wash. 1992, 59, 89–95. [Google Scholar]
  44. Garcia, H.A.; Rodrigues, A.C.; Martinkovic, F.; Minervino, A.H.H.; Campaner, M.; Nunes, V.L.B.; Paiva, F.; Hamilton, P.B.; Teixeira, M.M.G. Multilocus Phylogeographical Analysis of Trypanosoma (Megatrypanum) Genotypes from Sympatric Cattle and Water Buffalo Populations Supports Evolutionary Host Constraint and Close Phylogenetic Relationships with Genotypes Found in Other Ruminants. Int. J. Parasitol. 2011, 41, 1385–1396. [Google Scholar] [CrossRef]
  45. Rodrigues, A.C.; Paiva, F.; Campaner, M.; Stevens, J.R.; Noyes, H.A.; Teixeira, M.M.G. Phylogeny of Trypanosoma (Megatrypanum) theileri and Related Trypanosomes Reveals Lineages of Isolates Associated with Artiodactyl Hosts Diverging on SSU and ITS Ribosomal Sequences. Parasitology 2006, 132, 215–224. [Google Scholar] [CrossRef] [PubMed]
  46. Frolov, A.O.; Kostygov, A.Y.; Yurchenko, V. Development of Monoxenous Trypanosomatids and Phytomonads in Insects. Trends Parasitol. 2021, 37, 538–551. [Google Scholar] [CrossRef] [PubMed]
  47. Votýpka, J.; Petrželková, K.J.; Brzoňová, J.; Jirků, M.; Modrý, D.; Lukeš, J. How Monoxenous Trypanosomatids Revealed Hidden Feeding Habits of Their Tsetse Fly Hosts. Folia Parasitol. 2021, 68, 1–6. [Google Scholar] [CrossRef]
  48. Boucinha, C.; Andrade-Neto, V.V.; Ennes-Vidal, V.; Branquinha, M.H.; dos Santos, A.L.S.; Torres-Santos, E.C.; d’Avila-Levy, C.M. A Stroll Through the History of Monoxenous Trypanosomatids Infection in Vertebrate Hosts. Front. Cell Infect. Microbiol. 2022, 12, 804707. [Google Scholar] [CrossRef] [PubMed]
  49. Truc, P.; Büscher, P.; Cuny, G.; Gonzatti, M.I.; Jannin, J.; Joshi, P.; Juyal, P.; Lun, Z.R.; Mattioli, R.; Pays, E.; et al. Atypical Human Infections by Animal Trypanosomes. PLoS Negl. Trop. Dis. 2013, 7, e2256. [Google Scholar] [CrossRef] [PubMed]
  50. Kostygov, A.Y.; Malysheva, M.N.; Ganyukova, A.I.; Razygraev, A.V.; Drachko, D.O.; Yurchenko, V.; Agasoi, V.V.; Frolov, A.O. The Roles of Mosquitoes in the Circulation of Monoxenous Trypanosomatids in Temperate Climates. Pathogens 2022, 11, 1326. [Google Scholar] [CrossRef]
  51. Jansen, S.; Heitmann, A.; Lühken, R.; Leggewie, M.; Helms, M.; Badusche, M.; Rossini, G.; Schmidt-Chanasit, J.; Tannich, E. Culex torrentium: A Potent Vector for the Transmission of West Nile Virus in Central Europe. Viruses 2019, 11, 492. [Google Scholar] [CrossRef]
  52. Vanderplank, F.L. Seasonal and Annual Variation in the Incidence of Trypanosomiasis in Game. Ann. Trop. Med. Parasitol. 1947, 41, 365–374. [Google Scholar] [CrossRef]
  53. Pori, T.; Ndlovu, M.; Markus, M.B. Influence of Season and Other Factors on Avian Trypanosoma spp. and Microfilarial Prevalence in the Lowveld, South Africa. S. Afr. J. Sci. 2023, 119, 1–6. [Google Scholar] [CrossRef]
  54. Valkiūnas, G.; Kazlauskienė, R.; Bernotienė, R.; Palinauskas, V.; Iezhova, T.A. 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, 2159–2169. [Google Scholar] [CrossRef]
  55. Nourani, L.; Zakeri, S.; Dinparast Djadid, N. Dynamics of Prevalence and Distribution Pattern of Avian Plasmodium Species and Its Vectors in Diverse Zoogeographical Areas—A Review. Infect. Genet. Evol. 2020, 81, 104244. [Google Scholar] [CrossRef]
  56. Ejiri, H.; Sato, Y.; Sawai, R.; Sasaki, E.; Matsumoto, R.; Ueda, M.; Higa, Y.; Tsuda, Y.; Omori, S.; Murata, K.; et al. Prevalence of Avian Malaria Parasite in Mosquitoes Collected at a Zoological Garden in Japan. Parasitol. Res. 2009, 105, 629–633. [Google Scholar] [CrossRef]
  57. Shirotani, A.; Shibata, A.; Ejiri, H.; Sato, Y.; Tsuda, Y.; Hatakeyama, Y.; Iwano, H.; Murata, K.; Yukawa, M. Detection of Avian Malaria DNA from Mosquitoes in Kanagawa in Japan. J. Jpn. Vet. Med. Assoc. 2009, 62, 73–79. [Google Scholar] [CrossRef]
  58. Kim, K.; Tsuda, Y.; Yamada, A. Bloodmeal Identification and Detection of Avian Malaria Parasite from Mosquitoes (Diptera: Culicidae) Inhabiting Coastal Areas of Tokyo Bay, Japan. J. Med. Entomol. 2009, 46, 1230–1234. [Google Scholar] [CrossRef]
  59. Inumaru, M.; Yamada, A.; Shimizu, M.; Ono, A.; Horinouchi, M.; Shimamoto, T.; Tsuda, Y.; Murata, K.; Sato, Y. Vector Incrimination and Transmission of Avian Malaria at an Aquarium in Japan: Mismatch in Parasite Composition between Mosquitoes and Penguins. Malar. J. 2021, 20, 136. [Google Scholar] [CrossRef]
  60. Kim, K.S.; Tsuda, Y. Seasonal Changes in the Feeding Pattern of Culex pipiens pallens Govern the Transmission Dynamics of Multiple Lineages of Avian Malaria Parasites in Japanese Wild Bird Community. Mol. Ecol. 2010, 19, 5545–5554. [Google Scholar] [CrossRef]
  61. Ejiri, H.; Sato, Y.; Kim, K.S.; Hara, T.; Tsuda, Y.; Imura, T.; Murata, K.; Yukawa, M. Entomological Study on Transmission of Avian Malaria Parasites in a Zoological Garden in Japan: Bloodmeal Identification and Detection of Avian Malaria Parasite DNA from Blood-Fed Mosquitoes. J. Med. Entomol. 2011, 48, 600–607. [Google Scholar] [CrossRef]
  62. Odagawa, T.; Inumaru, M.; Sato, Y.; Murata, K.; Higa, Y.; Tsuda, Y.A. Long-Term Field Study on Mosquito Vectors of Avian Malaria Parasites in Japan. J. Vet. Med. Sci. 2022, 84, 1391–1398. [Google Scholar] [CrossRef]
  63. Kim, K.S.; 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, 5374–5385. [Google Scholar] [CrossRef]
  64. Glaizot, O.; Fumagalli, L.; Iritano, K.; Lalubin, F.; van Rooyen, J.; Christe, P. 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. [Google Scholar] [CrossRef]
  65. Lalubin, F.; Delédevant, A.; Glaizot, O.; Christe, P. Temporal Changes in Mosquito Abundance (Culex pipiens), Avian Malaria Prevalence and Lineage Composition. Parasit. Vectors 2013, 6, 307. [Google Scholar] [CrossRef]
  66. Kimura, M.; Darbro, J.M.; Harrington, L.C. Avian Malaria Parasites Share Congeneric Mosquito Vectors. J. Parasitol. 2010, 96, 144–151. [Google Scholar] [CrossRef]
  67. Huff, C.G. Studies on the Infectivity of Plasmodia of Birds for Mosquitoes, with Special Reference to the Problem of Immunity in the Mosquito. Am. J. Hyg. 1927, 7, 706–734. [Google Scholar] [CrossRef]
  68. Herman, G.M. Mosquito Transmission of Avian Malaria Parasites (Plasmodium circumflexum and P. cathemerium). Am. J. Hyg. 1938, 27, 345–350. [Google Scholar] [CrossRef]
  69. Zélé, F.; Vézilier, J.; L’Ambert, G.; Nicot, A.; Gandon, S.; Rivero, A.; Duron, O. Dynamics of Prevalence and Diversity of Avian Malaria Infections in Wild Culex pipiens Mosquitoes: The Effects of Wolbachia, Filarial Nematodes and Insecticide Resistance. Parasit. Vectors 2014, 7, 437. [Google Scholar] [CrossRef]
  70. Schoener, E.; Uebleis, S.S.; Butter, J.; Nawratil, M.; Cuk, C.; Flechl, E.; Kothmayer, M.; Obwaller, A.G.; Zechmeister, T.; Rubel, F.; et al. Avian Plasmodium in Eastern Austrian Mosquitoes. Malar. J. 2017, 16, 389. [Google Scholar] [CrossRef]
  71. Raffaele, G. Un Ceppo Italiano Di Plasmodium elongatum. Riv. Malariol. 1934, 13, 3–8. [Google Scholar]
  72. Micks, D.W. Investigations on the Mosquito Transmission of Plasmodium elongatum Huff, 1930. J. Natl. Malar. Soc. 1949, 8, 206–218. [Google Scholar]
  73. Garnham, P.C.C. Malaria Parasites and Other Haemosporidia; Blackwell Scientific Publications: Oxford, UK, 1966. [Google Scholar]
  74. Corradetti, A.; Garnham, P.C.C.; Laird, M. New Classification of the Avian Malaria Parasites. Parassitologia 1963, 5, 1–4. [Google Scholar]
  75. Corradetti, A.; Verolini, F.; Neri, L. Plasmodium (Haemamoeba) giovannolai n. Sp. Parassita Di Turdus merula. Parassitologia 1963, 5, 11–18. [Google Scholar]
  76. Akiba, K. Studies on Avian Malaria II. On the Species of Plasmodium from Chicken in Japan (P. japonicum Ishiguro, 1957), a Synonym for P. juxtanucleare Versiani and Gomes, 1941. Jpn. J. Vet. Sci. 1959, 21, 18. [Google Scholar]
  77. Christensen, B.M.; Barnes, H.J.; Rowley, W.A. Vertebrate Host Specificity and Experimental Vectors of Plasmodium (Novyella) kempi Sp. n. from the Eastern Wild Turkey in Iowa. J. Wildl. Dis. 1983, 19, 204–213. [Google Scholar] [CrossRef] [PubMed]
  78. Coggeshall, L.T. The Infection of Anopheles quadrimaculatus with a Monkey Malaria Parasite, Plasmodium cynomolgi and with an Avian Parasite, Plasmodium lophurae. J. Parasitol. 1940, 26, S44–S45. [Google Scholar] [CrossRef]
  79. Martinez-de la Puente, J.; Muñoz, J.; Capelli, G.; Montarsi, F.; Soriguer, R.; Arnoldi, D.; Rizzoli, A.; Figuerola, J. 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. [Google Scholar] [CrossRef]
  80. Ferraguti, M.; Martínez-de la Puente, J.; Muñoz, J.; Roiz, D.; Ruiz, S.; Soriguer, R.; Figuerola, J. Avian Plasmodium in Culex and Ochlerotatus Mosquitoes from Southern Spain: Effects of Season and Host-Feeding Source on Parasite Dynamics. PLoS ONE 2013, 8, e66237. [Google Scholar] [CrossRef]
  81. Huff, C.G.A. New Variety of Plasmodium relictum from the Robin. J. Parasitol. 1937, 23, 400–404. [Google Scholar] [CrossRef]
  82. Manwell, R.D. Life-Cycle Of Plasmodium relictum var. matutinum. Am. J. Trop. Med. Hyg. 1940, 20, 859–866. [Google Scholar] [CrossRef]
  83. Manwell, R.D. Failure of Aedes aegypti and Culex pipiens to Transmit Plasmodium vaughani. J. Parasitol. 1947, 33, 167–169. [Google Scholar] [CrossRef]
  84. Kim, K.; Tsuda, Y. Sporogony and Sporozoite Rates of Avian Malaria Parasites in Wild Culex pipiens pallens and C. inatomii in Japan. Parasit. Vectors 2015, 8, 633. [Google Scholar] [CrossRef]
  85. Schmid, S.; Dinkel, A.; Mackenstedt, U.; Tantely, M.L.; Randrianambinintsoa, F.J.; Boyer, S.; Woog, F. Avian Malaria on Madagascar: Bird Hosts and Putative Vector Mosquitoes of Different Plasmodium Lineages. Parasit. Vectors 2017, 10, 6. [Google Scholar] [CrossRef]
  86. Kazlauskienė, R.; Bernotienė, R.; Palinauskas, V.; Iezhova, T.A.; Valkiūnas, G. Plasmodium relictum (Lineages pSGS1 and pGRW11): Complete Synchronous Sporogony in Mosquitoes Culex pipiens pipiens. Exp. Parasitol. 2013, 133, 454–461. [Google Scholar] [CrossRef] [PubMed]
  87. Ionicǎ, A.M.; Zittra, C.; Wimmer, V.; Leitner, N.; Votýpka, J.; Modrý, D.; Mihalca, A.D.; Fuehrer, H.P. Mosquitoes in the Danube Delta: Searching for Vectors of Filarioid Helminths and Avian Malaria. Parasit. Vectors 2017, 10, 324. [Google Scholar] [CrossRef] [PubMed]
  88. Ventim, R.; Ramos, J.A.; Osório, H.; Lopes, R.J.; Pérez-Tris, J.; Mendes, L. Avian Malaria Infections in Western European Mosquitoes. Parasitol. Res. 2012, 111, 637–645. [Google Scholar] [CrossRef] [PubMed]
  89. Inci, A.; Yildirim, A.; Njabo, K.Y.; Duzlu, O.; Biskin, Z.; Ciloglu, A. Detection and Molecular Characterization of Avian Plasmodium from Mosquitoes in Central Turkey. Vet. Parasitol. 2012, 188, 179–184. [Google Scholar] [CrossRef]
  90. Ruge, R. Untersuchungen über das Deutsche Proteosoma. Zentralbl. Bakteriol. Parasitenkd. Infekt. Hyg. 1901, 29, 187–191. [Google Scholar]
  91. Neumann, R.O. Die übertragung von Plasmodium praecox auf Kanarievögel Durch Stegmyia fasciata und die Entwicklung der Parasiten im Magen und der Speicheldr üsen Dieser Stechmucke. Arch. Protistenkd. 1908, 13, 23–69. [Google Scholar]
  92. Sergent, E.D.; Sergent, E.T. Etudes Sur Les Hematozoaires d’oiseaux. Ann. Inst. Pasteur 1907, 21, 251–280. [Google Scholar]
  93. Tate, P.; Vincent, M. The Susceptibility of Autogenous and Anautogenous Races of Culex pipiens to Infection with Avian Malaria (Plasmodium relictum). Parasitology 1934, 26, 512–522. [Google Scholar] [CrossRef]
  94. Hunninen, A.V. Comparative Susceptibility of Four Anopheline Mosquitoes to Plasmodium relictum. J. Natl. Malar. Soc. 1951, 10, 216–223. [Google Scholar]
  95. Hunninen, A.V. Comparative Development of Plasmodium relictum Oocysts in Anopheles quadrimaculatus, A. albimanus, and Culex pipiens. J. Parasitol. 1953, 39, 28–32. [Google Scholar] [CrossRef]
  96. Ejiri, H.; Sato, Y.; Kim, K.S.; Tsuda, Y.; Murata, K.; Saito, K.; Watanabe, Y.; Shimura, Y.; Yukawa, M. Blood Meal Identification and Prevalence of Avian Malaria Parasite in Mosquitoes Collected at Kushiro Wetland, A Subarctic Zone of Japan. J. Med. Entomol. 2011, 48, 904–908. [Google Scholar] [CrossRef]
  97. Raffaele, G. II Plasmodium Della Civetta (Athene noctua). Riv. Malariol. 1932, 9, 3–12. [Google Scholar]
  98. Synek, P.; Munclinger, P.; Albrecht, T.; Votýpka, J. Avian Haemosporidians in Haematophagous Insects in the Czech Republic. Parasitol. Res. 2013, 112, 839–845. [Google Scholar] [CrossRef] [PubMed]
  99. Corradetti, A.; Scanga, M. Plasmodium (Novyella) vaughani Subsp. merulae, n Subsp., Parassita Di Turdus merula, Con Descrizione Del Ciclo Pre-Eritrocitivo. Parassitologia 1972, 14, 85–93. [Google Scholar]
  100. Köchling, K.; Schaub, G.A.; Werner, D.; Kampen, H. Avian Plasmodium spp. and Haemoproteus spp. Parasites in Mosquitoes in Germany. Parasit. Vectors 2023, 16, 369. [Google Scholar] [CrossRef]
  101. Farajollahi, A.; Fonseca, D.M.; Kramer, L.D.; Marm Kilpatrick, A. “Bird Biting” Mosquitoes and Human Disease: A Review of the Role of Culex pipiens Complex Mosquitoes in Epidemiology. Infect. Genet. Evol. 2011, 11, 1577–1585. [Google Scholar] [CrossRef]
  102. Hamer, G.L.; Kitron, U.D.; Brawn, J.D.; Loss, S.R.; Ruiz, M.O.; Goldberg, T.L.; Walker, E.D. Culex pipiens (Diptera: Culicidae): A Bridge Vector of West Nile Virus to Humans. J. Med. Entomol. 2008, 45, 125–128. [Google Scholar] [CrossRef]
Microbiolres 15 00146 g001
Figure 1. Average number of collected Culex pipiens females (left axis), with the average number of mosquitoes infected with Haemosporida and Trypanosomatida parasites (right axis) during the warm season in 2021–2023. Average ± standard error.
Figure 1. Average number of collected Culex pipiens females (left axis), with the average number of mosquitoes infected with Haemosporida and Trypanosomatida parasites (right axis) during the warm season in 2021–2023. Average ± standard error.
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Figure 2. Trypanosomatids found in Culex pipiens midgut preparations: (A)—Trypanosoma culicavium (GenBank No. PP946100); (B)—Trypanosoma trinaperronei (PP946104); (C)—monoxenous trypanosomatid (PP948731). Long arrows—parasite nuclei; arrowheads—kinetoplast; short arrows—free flagellum. Scale bar = 10 µm.
Figure 2. Trypanosomatids found in Culex pipiens midgut preparations: (A)—Trypanosoma culicavium (GenBank No. PP946100); (B)—Trypanosoma trinaperronei (PP946104); (C)—monoxenous trypanosomatid (PP948731). Long arrows—parasite nuclei; arrowheads—kinetoplast; short arrows—free flagellum. Scale bar = 10 µm.
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Microbiolres 15 00146 g003
Figure 3. Bayesian phylogenetic tree of Trypanosoma (A) and other Trypanosomatidae (B) using fragments of 18S rRNA. The tree was rooted using Cryptobia catostomi. The Genbank accession numbers of samples obtained during this investigation are written in bold text.
Figure 3. Bayesian phylogenetic tree of Trypanosoma (A) and other Trypanosomatidae (B) using fragments of 18S rRNA. The tree was rooted using Cryptobia catostomi. The Genbank accession numbers of samples obtained during this investigation are written in bold text.
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Table 1. Trypanosomatidae found in Culex pipiens mosquitoes, with collection date and site given.
Table 1. Trypanosomatidae found in Culex pipiens mosquitoes, with collection date and site given.
Collection DateCollection SiteMicroscopy PositiveGenBank
No		Trypanosomatid SpeciesVertebrate Host
25 May 2022.Verkiai−PP946099Trypanosoma culicaviumBirds
29 June 2022.Kairėnai+PP946101T. culicaviumBirds
29 June 2022.Kairėnai−PP946100T. culicaviumBirds
14 July 2022.Kairėnai−PP946102T. theileriMammals
3 August 2022.Kairėnai+PP946104T. trinaperroneiMammals
3 August 2022.Kairėnai−PP946105T. theileriMammals
30 August 2022.Kairėnai+PP946107T. culicaviumBirds
18 July 2023.Kairėnai+PP946103T. culicaviumBirds
18 July 2023.Kairėnai−PP946106T. theileriMammals
7 December 2023.Verkiai 1+PP948731Monoxenous
trypanosomatid		No vertebrate host
8 December 2023.Vilnius−PP948732Monoxenous
trypanosomatid		No vertebrate host
1 Cellar of water tower.
Table 2. Trypanosoma spp. found in Culex mosquitoes in various studies. EI—experimental infections of mosquitoes; WM—wild-caught mosquitoes.
Table 2. Trypanosoma spp. found in Culex mosquitoes in various studies. EI—experimental infections of mosquitoes; WM—wild-caught mosquitoes.
Trypanosomatid SpeciesIsolateOrigin of IsolateEIWMCountry, Source
Trypanosoma thomasbancroftiCUL15, CUL98Culex pipiens+ Czech Republic [4,5]
OF19Ornithomya fringilline *+ Czech Republic [4]
PAS343Phylloscopus sibilatrix *+ Czech Republic [4]
T. theileri−Cx. pipiens +Czech Republic [4,37]
T. culicaviumCUL1, CUL6, CUL24, CUL28, CUL31Cx. pipiens +Czech Republic [5]
−Cx. pipiens s.l./torrentium, Cx. modestus, Cx. spp +Austria [35]
Trypanosoma sp.CUL5, CUL2Cx. pipiens +Czech Republic [33]
T. avium Cx. pipiens s.l./torrentium +Austria [35]
Cx. pipiens, Cx. tarsalis +USA [38]
*—Culex mosquitoes were experimentally infected using these isolates; +—indication of research method (experimental infection (EI) or study of wild-caught mosquitoes (WM)); −—the isolate was not determined or named.
Table 3. Experimental infections of mosquitoes (EI) and molecular studies (PCR) of wild-caught mosquitoes (WM) showing what haemosporidian parasite DNA or parasites were detected in Culex pipiens mosquitoes. Information acquired from the MalAvi database and [3,55]. Cases with sporozoites detected in mosquito salivary glands are marked in bold and underlined.
Table 3. Experimental infections of mosquitoes (EI) and molecular studies (PCR) of wild-caught mosquitoes (WM) showing what haemosporidian parasite DNA or parasites were detected in Culex pipiens mosquitoes. Information acquired from the MalAvi database and [3,55]. Cases with sporozoites detected in mosquito salivary glands are marked in bold and underlined.
Plasmodium SpeciesLineagesEIWMPCRCountry/References
P. cathemeriumpPADOM02 +Japan [56,57,58,59,60,61,62,63], Switzerland [64,65], USA [66]
unknown lineage USA [67,68]
P. circumflexumpTURDUS1 +Switzerland [64]
P. duraeunknown lineage Africa [15]
P. elongatumpGRW06 +France [69] Austria [34,70]
unknown lineage++ USA [67], Italy [71,72]
P. gallinaceumpGALLUS01 +Japan [57,58,63]
P. garnhamiunknown lineage+ Egypt [73]
P. giovannolaiunknown lineage+ Italy [74,75]
P. homonucleophilumpSW2 +Lithuania [24], Switzerland [64]
P. juxtanuclearepGALLUS02 +Japan [63]
unknown lineage+ Japan [76]
P. kempiunknown lineage+ USA [77]
P. lophuraeunknown lineage USA [78]
P. matutinumpLINN1 +France [69], USA [66], Italy [79], Spain [80], Austria [34]
++Lithuania [24]
unknown lineage USA [81,82,83]
P. relictumpGRW04 +Japan [62,63,84], Madagascar [85]
pGRW11 +Italy [79], Japan [58,60,84], Switzerland [64,65], France [69]
+ Lithuania[86]
pSGS1 +Romania [87], Japan [58,60,61,62,84], Austria [34,70], France [69], Portugal [88], Italy [79], Switzerland [64,65], Spain [80], Turkey [89]
+ Lithuania [86]
unknown lineage +Romania [87]
+ Germany [90,91], Algeria [92], USA [93], Columbia [94,95]
P. rouxiunknown lineage + USA [81,83]
Plasmodium sp.pAFTRU5 +Switzerland [65], Italy [79]
pCOLL1 +France [69], Switzerland [65], Spain [80]
pCXPIP01, CXPIP02, pCXPIP03, pCXPIP04, pCXPIP05, pCXPIP06, pCXPIP07 +USA [66]
pCXPIP09 +Japan [56,57,58,59,60,61,62,63,84]
pCXPIP10, pCXPIP11, pCXPIP12, pCXPIP13, pCXPIP14 +Japan [58,60,63]
pCXPIP15 +Japan [61]
pCXPIP20, pCXPIP21, pCXPIP22, pCXPIP23 +Turkey [89]
pCXPIP24, pCXPIP25, pCXPIP26 +France [69]
pCXPIP30 +Madagascar [85]
pCXPIP31 +Japan [96]
pCXPIP32, pCXPIP33 Italy (Iurescia et al., unpubl)
pCXQUI01 +Japan [63]
pDELURB4 +Italy [79], France [69], Austria [34]
pDELURB5, pDONANA03, pDONANA05 +Austria [34], France [69]
pPADOM01 +France [69], Switzerland [65]
pSPHUM05 +Japan [59]
pSPMAG10, pZOCAP03, pSYCON02 +Japan [62]
P. subpraecoxunknown lineage+ Italy [97]
P. unalispTUMIG03 +USA [66]
P. vaughanipSYAT05 +France [69], USA [66], Lithuania [24], Italy [79], Japan [60], Switzerland [64,65], Turkey [89], Austria [34,70], Czech Republic [98]
unknown lineage+ Italy [99]
+—indication of research method (experimental infection (EI) or study of wild-caught mosquitoes (WM), data obtained using PCR methods (PCR)).
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Valavičiūtė-Pocienė, K.; Kazak, M.; Iezhova, T.; Kalinauskaitė, G.; Bernotienė, R. Blood Parasites (Haemosporida, Trypanosomatida) in Culex pipiens: A Study and Review of Hibernating and Active Mosquitoes. Microbiol. Res. 2024, 15, 2184-2198. https://doi.org/10.3390/microbiolres15040146

AMA Style

Valavičiūtė-Pocienė K, Kazak M, Iezhova T, Kalinauskaitė G, Bernotienė R. Blood Parasites (Haemosporida, Trypanosomatida) in Culex pipiens: A Study and Review of Hibernating and Active Mosquitoes. Microbiology Research. 2024; 15(4):2184-2198. https://doi.org/10.3390/microbiolres15040146

Chicago/Turabian Style

Valavičiūtė-Pocienė, Kristina, Margarita Kazak, Tatjana Iezhova, Gabrielė Kalinauskaitė, and Rasa Bernotienė. 2024. "Blood Parasites (Haemosporida, Trypanosomatida) in Culex pipiens: A Study and Review of Hibernating and Active Mosquitoes" Microbiology Research 15, no. 4: 2184-2198. https://doi.org/10.3390/microbiolres15040146

APA Style

Valavičiūtė-Pocienė, K., Kazak, M., Iezhova, T., Kalinauskaitė, G., & Bernotienė, R. (2024). Blood Parasites (Haemosporida, Trypanosomatida) in Culex pipiens: A Study and Review of Hibernating and Active Mosquitoes. Microbiology Research, 15(4), 2184-2198. https://doi.org/10.3390/microbiolres15040146

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