Open Access

Sporogony and sporozoite rates of avian malaria parasites in wild Culex pipiens pallens and C. inatomii in Japan

Parasites & Vectors20158:633

https://doi.org/10.1186/s13071-015-1251-1

Received: 2 April 2015

Accepted: 10 December 2015

Published: 15 December 2015

Abstract

Background

Malaria infection in mosquitoes is traditionally detected by microscopic examination for Plasmodium oocysts and sporozoites. Although PCR is now widely used, the presence of parasite DNA in a mosquito does not prove that sporogony is achieved. Thus, detection of sporozoites by microscopy is still required to definitively identify vector mosquitoes. The aim of this study was to confirm sporogony of avian Plasmodium spp. in Culex pipiens pallens and C. inatomii caught from the wild.

Findings

Mosquitoes collected at two sites in Japan were dissected and examined by microscopy for Plasmodium oocysts and sporozoites. DNA was extracted from the midgut and salivary gland of infected mosquitoes, and the infecting Plasmodium species was identified by sequencing 478 bp of cytochrome b. Oocysts, or both oocysts and sporozoites, were found in 3.94 and 0.46 % of C. p. pallens and C. inatomii, respectively. Four (CXPIP09, GRW4, GRW11 and SGS1) and three cytochrome b lineages (CXINA01, CXINA02 and CXQUI01) were confirmed to achieve sporogony in C. p. pallens and C. inatomii, respectively. One mosquito each of C. p. pallens and C. inatomii was co-infected with two different Plasmodium lineages.

Conclusions

These findings demonstrate that C. p. pallens and C. inatomii are natural vectors of four and three lineages of avian Plasmodium spp., respectively. The data indicate that a systematic procedure combining microscopy and PCR is a feasible and reliable approach to identify natural vectors of wildlife malaria.

Keywords

Avian malaria Culex inatomii Culex pipiens pallens Natural vector Plasmodium Sporozoite rateVector competence

Findings

Background

Malaria parasitization in mosquitoes is traditionally detected by dissection and microscopic examination for oocysts in the midgut and sporozoites in the salivary gland [13]. However, many recent field studies have relied on PCR instead [46]. Nevertheless, detection of Plasmodium DNA in a blood-sucking insect does not prove that the insect acts as vector [7, 8], as parasites are eliminated in a refractory insect. Thus, while PCR is sensitive enough to detect DNA from degraded parasites, microscopic detection of sporozoites remains necessary to verify sporogony and to identify competent vectors. On the other hand, oocysts and sporozoites vary little in morphology across Plasmodium species, and are impossible to identify to species or lineage by microscopy [9, 10]. Thus, a combination of dissection and PCR is required [5, 10, 11]. Unfortunately, this combined approach has not been adopted, except in studies of human malaria parasites.

The aim of this study was to use this combined approach to definitively establish whether Culex pipiens pallens and C. inatomii are competent vectors for avian malaria. Although these mosquitoes have been suggested in PCR-based studies to be primary natural vectors of avian malaria in Japan [1113], sporogony has not been confirmed. Our results suggest that a systematic procedure combining dissection and PCR is a reliable approach to identify natural vectors of wildlife malaria.

Methods

Mosquitoes were collected in Rinshi-no-mori park (35°37′ N, 139°42′ E) in Tokyo and Sakata wetland (37° 49′ N, 138° 53′ E) in Niigata, Japan, where transmission of multiple avian malaria parasites has been detected by PCR [11, 13]. The study sites and the ecological differences between C. p. pallens and C. inatomii are described in greater detail in our previous publications [11, 13]. In Rinshi-no-mori park, mosquitoes were collected once or twice a week from May to September in 2012 and from May to June in 2013, using a sweep net 36 cm in diameter as previously described [13]. In Sakata wetland, mosquitoes were collected on 2–3 July 2013 and on 30 June and 1 July 2014 using ten battery-operated suction traps (Inokuchi-Tekko, Nagasaki, Japan) baited with dry ice. The traps are similar to devices designed by the Centers for Disease Control and Prevention. Mosquitoes collected from the field were kept alive until dissection at National Institute of Infectious Diseases in Tokyo and Tottori University in Tottori.

Mosquitoes were immobilized by chilling or by chloroform, dissected according to WHO protocols [1], and examined under a microscope. The midgut was first examined for oocysts, and, when oocysts were present, the midgut and a part of the salivary gland were transferred to a 1.5 ml tube for DNA extraction. In addition, a smear of the salivary gland was stained by Giemsa and carefully examined for sporozoites.

DNA was extracted using REDExtract-N-Amp PCR Reaction Kit (Sigma Chemical Co., St. Louis, MO). A 478 bp fragment of Plasmodium cytochrome b was amplified by nested PCR according to Waldenström et al. [14], with slight modification [7]. Amplification products were purified with QIAEX II-Gel Extraction Kit (QIAGEN), and sequenced in both directions on an ABI PRISM 3130 Genetic Analyzer (Applied Biosystems), using ABI PRISM BigDye Terminator Cycle Sequencing Kit version 1.1 (Applied Biosystems, Foster City, CA, USA). Sequences were analyzed in GENETYX-WIN ver. 11, and compared to published sequences by a BLASTn search against the NCBI GenBank database and MalAvi database [15]. Sequences from one specimen each of C. p. pallens and C. inatomii contained a few doublet peaks. The electropherograms of these sequences were carefully inspected by eye, and were unambiguously resolved into known Plasmodium lineages.

Results and discussion

Plasmodium spp. from mosquitoes in Rinshi-no-mori park

Five mosquito species were collected from Rinshi-no-mori park. Culex p. pallens was the most prevalent (n = 533 females), followed by C. sasai (n = 19), Lutzia vorax (n = 16), Orthopodomyia anopheloides (n = 9), and C. rubithoracis (n = 5). All mosquitoes were dissected, and only C. p. pallens was found to be infected with malaria parasites.

Oocysts were observed in the midgut of 21 (3.94 %) C. p. pallens. Motile sporozoites were found in the salivary gland of 11 of these specimens (Table 1 and Fig. 1). The overall sporozoite rate (i. e., the proportion of mosquitoes with sporozoites) was 2.06 %. This rate is significantly lower (Fisher’s exact test, p = 0.001) than the 8.31 % of samples that tested positive for Plasmodium DNA in a previous study [13]. All infected mosquitoes also tested positive by PCR, and four cytochrome b lineages of avian Plasmodium spp. (CXPIP09, GRW4, GRW11 and SGS1) were identified (Table 1). CXPIP09 and SGS1 were the most prevalent, and accounted for >85 % of infections. The dominance of CXPIP09 and SGS1 was consistent with PCR-based studies [13]. However, we did not detect PADOM02 (Table 1), perhaps due to the small sample size or the inability of C. p. pallens to support its development. GRW4 and GRW11 that had been previously absent and detected at low frequency, respectively, were found to complete sporogony in C. p. pallens (Table 1). Notably, one specimen was co-infected with two different lineages of P. relictum, GRW4 and GRW11.
Table 1

Avian malaria parasites found from Culex pipiens pallens in Rinshi-no-mori park, Tokyo

Plasmodium lineage

Previous PCR study [2]

This study: dissection and PCR

Vector status

(N = 1252)

(N = 533)

DNA

Infection rate (%)

Oocysts

Infection rate (%)

Oocysts & sporozoites

Sporozoite rate (%)

CXPIP09

43

3.43

6

1.13

3

0.56

Competent

SGS1-P. relictum

30

2.40

4

0.75

5

0.94

Competent

PADOM02

16

1.28

    

Unknown

GRW11-P. relictum

3

0.24

  

1*

0.19

Competent

CXPIP11

1

0.08

    

Unknown

CXPIP12

4

0.32

    

Unknown

CXPIP13

1

0.08

    

Unknown

CXPIP14

1

0.08

    

Unknown

GALLUS01- P. gallinaceum

4

0.32

    

Not competent [2, 30]

SYAT05-P. vaughani

1

0.08

    

Unknown

GRW4-P. relictum

    

3*

0.56

Competent

Total

104

8.31

10

1.87

11

2.06

 

*One C. p. pallens specimen was co-infected with CXQUI01 and GRW4-P. relictum. GenBank accession numbers: CXPIP09 [AB458850], SGS1-P. relictum [AF495571], PADOM02 [DQ058612], GRW11- P. relictum [AY831748], CXPIP11 [AB477121], CXPIP12 [AB477122], CXPIP13 [AB477126], CXPIP14 [AB477125], GALLUS01-P. gallinaceum [AY099029], SYAT05 [DQ847271] and GRW4-P. relictum [AF254975]

Fig. 1

Oocysts (a, b) and Giemsa-stained sporozoites (c, d) of avian malaria parasites from Culex pipiens pallens and C. inatomii. ac: CXINA02 [GanBank: AB920777] from C. inatomii, d: SGS1-Plasmodium relictum [AF495571] from C. p. pallens. Scale bar,100 μm (a), 20 μm (b), and 10 μm (c, d)

Of the four lineages that achieve sporogony in C. p. pallens, GRW4, GRW11 and SGS1 belong to the same morphological species, Plasmodium relictum [1618], and are the most widely distributed [15, 19]. For example, SGS1 was found in 62 avian species from Africa, Asia, Europe and Oceania. On the other hand, CXPIP09 has been found exclusively in Japan [15], although its avian hosts are widespread in eastern Asia, such as Corvus macrorhynchos, Passer montanus [2], Lanius bucephalus (KS Kim unpublished data), Cyanopica cyana, Parus minor, Treron sieboldii, and Zosterops japonicus (Koichi Murata personal communication). The reason for the limited distribution of CXPIP09 is unknown, as its natural vector, C. p. pallens, is also found in the same geographic range as the hosts [20]. Of note, field populations of a competent vector species may vary significantly in susceptibility to the same parasite species, depending on innate immunity and the microbiota in the midgut [21, 22]. Therefore, specific adaptation to C. p. pallens in Japan might have stringently limited the distribution of CXPIP09.

Plasmodium spp. from mosquitoes in Sakata wetland

In Sakata wetland, 4293 female C. inatomii were collected, along with C. p. pallens (n = 459), C. orientalis (n = 10) and C. tritaeniorhynchus (n = 6). Of 1314 C. inatomii dissected, six specimens had oocysts and sporozoites (Table 2 and Fig. 1). The sporozoite rate (0.46 %) was similar (Fisher’s exact test, p > 0.05) to the frequency of Plasmodium DNA (0.51 %) in a previous study [11]. All six specimens subsequently tested positive by PCR, and three avian Plasmodium lineages (CXINA01, CXINA02 and CXQUI01) were identified, with sporozoite rates ranging from 0.08 to 0.30 %. One specimen was co-infected with CXINA02 and CXQUI01. CXINA02 was a novel lineage, and was deposited in GenBank under accession number AB920777. Two C. tritaeniorhynchus and 33 C. p. pallens were dissected and none were infected.
Table 2

Avian malaria parasites found from Culex inatomii in Sakata wetland, Niigata

Plasmodium lineage

Previous PCR study [3]

This study: dissection and PCR

Vector status

(N = 7519)

(N = 1314)

DNA

Infection rate (%)

Oocysts & sporozoites

Sporozoite rate (%)

CXQUI01

15

0.20

2**

0.15

Competent

CXINA01

9

0.12

1

0.08

Competent

CXPIP10

8

0.11

  

Unknown

PADOM02

3

0.04

  

Unknown

CXPIP09

1

0.01

  

Unknown

SYBOR02

1

0.01

  

Unknown

GALLUS01-P. gallinaceum

1

0.01

  

Unknown

CXINA02

  

4**

0.30

Competent

Total

38

0.51

6

0.46

 

**One C. inatomii specimen was co-infected with CXQUI01 and CXINA02. GenBank accession numbers: CXQUI01 [AB308051], CXINA01 [AB690267], CXPIP10 [AB477128], PADOM02 [DQ058612], CXPIP09 [AB474376], SYBOR02 [DQ368392] and CXINA02 [AB920777]

The current data demonstrates for the first time that C. inatomii is a natural vector of avian malaria. Indeed, DNA from seven Plasmodium lineages was previously detected in C. inatomii from Sakata (Table 2). Of these, three lineages (CXQUI01, CXINA01 and CXPIP10) were the most prevalent, and comprised >86 % of infections in 2007–2010 [11]. Thus, sporogony of CXINA01 and CXQUI01 in C. inatomii was not unexpected, but the dominance of the novel lineage CXINA02 was. The difference in dominant lineages in mosquitoes between now and 2007–2010 may reflect changes in the parasite species circulating among host birds.

Unfortunately, the avian host species for CXINA01, CXINA02 and CXQUI01 are presently unknown. The spatial distribution of adult C. inatomii is restricted to areas near larval habitats [23, 24], with flight range estimated at <200 m in Sakata [25]. Culex inatomii feeds most commonly on Acrocephalus orientalis, a summer migratory bird that breeds in the reed fields at Sakata [11]. Hence, A. orientalis warrants investigation as a candidate natural host of avian malaria parasites found in C. inatomii.

Conclusions

Microscopic confirmation of sporogony, followed by genetic identification of infecting Plasmodium parasites, demonstrated that C. p. pallens and C. inatomii are natural vectors of four (CXPIP09, GRW4, GRW11 and SGS1) and three (CXINA01, CXINA02 and CXQUI01) lineages of avian Plasmodium, respectively. Ideally, transmission to a vertebrate host via a mosquito vector should be experimentally demonstrated [26]. However, such demonstration is difficult for wildlife parasites, usually because of limited availability of natural hosts and vectors. In addition, previous studies have shown that experimental transmission from wild mosquitoes to laboratory hosts (such as poultry in case of avian malaria) is difficult to achieve as well, usually because infected wild mosquitoes are rare and are reluctant to take blood meals under laboratory conditions [2729]. In light of these, we believe that demonstration of sporogony via a systematic procedure combining dissection and PCR is the most feasible approach to identify natural vectors of wildlife malaria.

Declarations

Acknowledgements

This study was supported by a Grant-in-Aid for Young Scientists (15 K18780) and a Grant-in-Aid from the Ministry of Health, Labour and Welfare of Japan (H24-shinko-ippan-007).

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)
Joint Department of Veterinary Medicine, Faculty of Agriculture, Tottori University
(2)
Department of Medical Entomology, National Institute of Infectious Diseases

References

  1. WHO. Division of Malaria and Other Parasitic Diseases. Manual on practical entomology in malaria Part II Methods and techniques. Geneva: World Health Organization; 1975.Google Scholar
  2. Wharton RH, Eyles DE, Warren M, Cheong WH. Studies to determine the vectors of monkey malaria in Malaya. Ann Trop Med Parasitol. 1964;58:56–77.PubMedGoogle Scholar
  3. Wharton RH, Eyles DE, Warren M, Moorhouse DE, Sandosham AA. Investigations leading to the identification of members of the Anopheles umbrosus group as the probable vectors of mouse deer malaria. Bull World Health Organ. 1963;29:357–74.PubMed CentralPubMedGoogle Scholar
  4. 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;9:1353–8.View ArticlePubMedGoogle Scholar
  5. Valkiūnas G. Haemosporidian vector research: marriage of molecular and microscopical approaches is essential. Mol Ecol. 2011;20:3084–6.View ArticlePubMedGoogle Scholar
  6. Outlaw DC, Ricklefs RE. Species limits in avian malaria parasites (Haemosporida): how to move forward in the molecular era. Parasitology. 2014;141:1223–32.View ArticlePubMedGoogle Scholar
  7. Kim KS, Tsuda Y, Sasaki T, Kobayashi M, Hirota Y. Mosquito blood-meal analysis for avian malaria study in wild bird communities: laboratory verification and application to Culex sasai (Diptera: Culicidae) collected in Tokyo. Japan Parasitol Res. 2009;105:1351–7.View ArticleGoogle Scholar
  8. Valkiūnas G, Kazlauskienė R, Bernotienė R, Palinauskas V, Lezhova TA. Abortive long-lasting sporogony of two Haemoproteus species (Haemosporidia, Haemoproteidae) in the mosquito Ochlerotatus cantans, with perspectives on haemosporidian vector research. Parasitol Res. 2013;112:2159–69.View ArticlePubMedGoogle Scholar
  9. Ramsey JM, Beaudoin RL, Bawden MP, Espinal CA. Specific identification of Plasmodium sporozoites using an indirect fluorescent antibody method. Trans R Soc Trop Med Hyg. 1983;77:378–81.View ArticlePubMedGoogle Scholar
  10. Santiago-Alarcon D, Palinauskas V, Schaefer HM. Diptera vectors of avian Haemosporidian parasites: untangling parasite life cycles and their taxonomy. Biol Rev Camb Philos Soc. 2012;87:928–64.View ArticlePubMedGoogle Scholar
  11. 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
  12. Kim KS, 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;46:1230–4.View ArticleGoogle Scholar
  13. Kim KS, 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;19:5545–54.View ArticlePubMedGoogle Scholar
  14. Waldenström J, Bensch S, Hasselquist D, Östman Ö. A new nested polymerase chain reaction method very efficient in detecting Plasmodium and Haemoproteus infections from avian blood. J Parasitol. 2004;90:191–4.View ArticlePubMedGoogle Scholar
  15. MalAvi: A database for avian haemosporidian parasites Version 2.2.1. http://mbio-serv2.mbioekol.lu.se/Malavi/. Accessed 25 Mar 2015.
  16. Palinauskas V, Kosarev V, Shapoval A, Bensch S, Valkiūnas G. Comparison of mitochondrial cytochrome b lineages and morphospecies of two avian malaria parasites of the subgenera Haemamoeba and Giovannolaia (Haemosporida: Plasmodiidae). Zootaxa. 2007;1626:39–50.Google Scholar
  17. Valkiūnas G, Zehtindjiev P, Hellgren O, Ilieva M, Iezhova TA, Bensch S. Linkage between mitochondrial cytochrome b lineages and morphospecies of two avian malaria parasites, with a description of Plasmodium (Novyella) ashfordi sp. nov. Parasitol Res. 2007;100:1311–22.View ArticlePubMedGoogle Scholar
  18. Kazlauskienė R, Bernotienė R, Palinauskas V, Lezhova TA, Valkiūnas G. Plasmodium relictum (lineages pSGS1 and pGRW11): complete synchronous sporogony in mosquitoes Culex pipiens pipiens. Exp Parasitol. 2013;133:454–61.View ArticlePubMedGoogle Scholar
  19. Hellgren O, Atkinson CT, Bensch S, Albayrak T, Dimitrov D, Ewen JG, et al. Global phylogeography of the avian malaria pathogen Plasmodium relictum based on MSP1 allelic diversity. Ecography. 2015;38:842–50.View ArticleGoogle Scholar
  20. Tanaka K, Mizusawa K, Saugstad E. A revision of the adult and larval mosquitoes of Japan (including the Ryukyu Archipelago and the Ogasawara islands) and Korea (Diptera: Culicidae). Contrib Am Entomol Inst. 1979;16:1–987.Google Scholar
  21. Niaré O, Markianos K, Volz J, Oduol F, Touré A, Bagayoko M, et al. Genetic loci affecting resistance to human malaria parasites in a West African mosquito vector population. Science. 2002;298:213–6.View ArticlePubMedGoogle Scholar
  22. Cirimotich CM, Dong Y, Clayton AM, Sandiford SL, Souza-Neto JA, Mulenga M, et al. Natural microbe-mediated refractoriness to Plasmodium infection in Anopheles gambiae. Science. 2011;332:855–8.PubMed CentralView ArticlePubMedGoogle Scholar
  23. Tsuda Y, Sasaki E, Sato Y, Katano R, Komagata O, Isawa H, et al. Mosquito collections from coastal areas of Tokyo Bay receiving migratory birds. Med Entomol Zool. 2009;60:119–24.Google Scholar
  24. Tsuda Y, Haseyama M, Ishida K, Niizuma J, Kim KS, Yanagi D, et al. After-effects of Tsunami on distribution and abundance of mosquitoes in rice-field areas in Miyagi Prefecture, Japan in 2011. Med Entomol Zool. 2012;63:21–30.View ArticleGoogle Scholar
  25. Tsuda Y, Kim KS. Outbreak of Culex inatomii in disaster areas of the Great East Japan earthquake and tsunami in 2011, with ecological notes on their larval habitats, biting behavior, and reproduction. J Am Mosq Control Assoc. 2013;29:19–26.View ArticlePubMedGoogle Scholar
  26. Barnett HC. The incrimination of arthropods as vectors of disease. In: Strouhal H, Beier M, editors. Proceedings of the 11th International Congress on Entomology, Vienna 1960. Vienna: Wien; 1962. p. 341–5.Google Scholar
  27. Reeves WC, Herold RC, Rosen L, Brookman B, Hammon WM. Studies on avian malaria in vectors and hosts of encephalitis in Kern county, California. II. Infections in mosquito vectors. Am J Trop Med Hyg. 1954;3:696–703.PubMedGoogle Scholar
  28. Forrester DJ, Nayar JK, Foster GW. Culex nigripalpus: a natural vector of wild turkey malaria (Plasmodium hermani) in Florida. J Wildl Dis. 1980;16:391–4.View ArticlePubMedGoogle Scholar
  29. Beier JC, Trpis M. Incrimination of natural culicine vectors which transmit Plasmodium elongatum to penguins at the Baltimore Zoo. Can J Zool. 1981;59:470–5.View ArticleGoogle Scholar
  30. Weathersby AB. Susceptibility of certain Japanese mosquitoes to Plasmodium gallinaceum and Plasmodiun berghei. J Parasitol. 1962;48:607–9.View ArticlePubMedGoogle Scholar

Copyright

© Kim and Tsuda. 2015

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