CC BY
103Russian Journal of Nematology, 2017, 25 (2), 121 - 127
Susceptibility of Anopheles stephensi (Diptera: Culicidae) to Dirofilaria immitis (Spirurida:
Onchocercidae)
Rahmat Solgi1, Seyed Mahmoud Sadjjadi1, 2, Mehdi Mohebali3, 4, Navid Dinparast Djadid5, Abbasali Raz5, Sedigheh Zakeri5 and Zabihollah Zarei3
department of Parasitology and Mycology, School of Medicine, Shiraz University of Medical Sciences,
71345-1735, Shiraz, Iran 2Basic Sciences in Infectious Diseases Research Center, Shiraz University of Medical Sciences,
71348-45794, Shiraz, Iran
3Department of Medical Parasitology and Mycology, School of Public Health, Tehran University of Medical Sciences,
14155-6446, Tehran, Iran 4Centre for Research of Endemic Parasites of Iran (CREPI), Tehran University of Medical Sciences,
14155-6446, Tehran, Iran
5Malaria and Vector Research Group (MVRG), Biotechnology Research Center (BRC), Pasteur Institute of Iran,
13169-43551, Tehran, Iran e-mail: smsadjjadi@sums.ac.ir
Accepted for publication 27 November 2017
Summary. Dirofilaria immitis is endemic in the southern parts of Iran where there is a favourable environment for the development of Anopheles stephensi. The aim of the present study was to evaluate the susceptibility of A. stephensi as D. immitis vector under laboratory conditions. Anticoagulated dog blood containing 1,500 microfilariae ml-1 was artificially administered to 140 mosquitoes of the test group (T group), while 94 mosquitoes were left as a control (C group). Blood-fed mosquitoes of the T group were dissected for morphological and molecular analyses at pre-set time points to observe the developmental stage of D. immitis. The results showed that the average number of the microfilariae ingested per female in T group was 9.6. At the end of the study, 16 mosquitoes developed infective third-stage larvae of D. immitis, and 49 mosquitoes survived at end of incubation period, giving an estimated infection rate of 32.6% and the vector efficiency index of 17%. The infection rate and vector efficiency index suggested that A. stephensi could be considered as a potential vector of D. immitis. Key words: dog heartworm, filariasis, microfilariae, molecular analysis, mosquitoes.
The dog heartworm, Dirofilaria immitis (Leidy), is a cause of the zoonotic filariasis with a cosmopolitan distribution (Simon et al., 2009). This nematode needs a definite vertebrate host and a culicid, as a vector, to complete its life cycle. It primarily infects canines but may accidentally infect both humans and felines (Simon et al., 2012; Khodabakhsh et al., 2016). Human dirofilariasis, as an emerging zoonotic disease, has been reported increasingly from different parts of the world, including Iran (Negahban et al., 2007; Mirahmadi et al., 2017). Canine dirofilariasis, on the other hand, is endemic in Iran, and its prevalence has been reported to be as high as 33%. However, there are few data related to invertebrate hosts of D. immitis in Iran (Sadjjadi et al., 2004; Azari-Hamidian,
2007). The presence of different competent zoophilic vectors in the areas where the prevalence of canine dirofilariasis is high implies that human population is at risk of infection (Simon et al., 2005). Among culicid species, more than 77 species are assumed to be the potential vectors of D. immitis (Azari-Hamidian et al., 2009). Few of them such as Culex pipiens (Linnaeus), Aedes vexans (Meigen), A. albopictus (Skuse) and Anopheles maculipennis (Meigen) are considered to be efficient vectors for transmitting the infective third-stage larvae (L3) in nature (Aranda et al., 1998; Ferreira et al., 2015). Culex theileri (Theobald) is the only reported mosquito in Iran that is able to be infected naturally with infective L3 of D. immitis (Azari-Hamidian et al. , 2009). The Malpighian tubules of some
mosquitoes are capable of inducing a melanotic response to the invasion of D. immitis larvae and providing resistance against this nematode (Bradley & Nayar, 1985). The vector competence is different among mosquito species, strains, or even individuals of the same strain (Serrao et al., 2001). Anopheles stephensi is abundant throughout the southern part of Iran (Fig. 1) where the prevalence of dirofilariasis is high (Jafari et al., 1996; Khedri et al., 2014; Salahi-Moghaddam et al., 2017). Previously, it was shown that A. stephensi could be infected by feeding on a dog infected with D. repens and subsequently, developed the ingested microfilaria (mf) into the infective larvae (Webber & Hawking, 1955). The current study was performed to analyse the susceptibility of the A. stephensi, as the dominant strain in Sistan and Baluchestan Province, for transmission of D. immitis.
MATERIAL AND METHODS
Mosquitoes' colony. The colony of mosquitoes was originated from eggs collected from Sarbaz city of Sistan and Baluchestan Province, the endemic region of D. immitis in the southeast of Iran (Khedri et al. , 2014). The larvae were reared to adults in emergent cages (50^50^50 cm) in an insectarium at 26 ± 1°C, with the relative humidity 80%, and 12L: 12D photoperiod. The emerged adults were maintained at the same conditions on 10% sucrose for 5-6 days for mating. The identity of the species of collected mosquitoes was examined at the National Insectarium, Malaria and Vector Research Group (MVRG), Biotechnology Research Centre (BRC), Pasteur Institute of Iran by means of morphological identification keys and a molecular analysis as described (Subbarao et al., 1987; Azari-Hamidian & Harbach, 2009; Chavshin et al., 2014). Briefly, for molecular identification, genomic DNA was extracted from samples using DNAzol® (Invitrogen, Gaithersburg, USA) according to the procedure supplied by the manufacturer. Amplification of the mtDNA-COII partial gene was performed using primer pairs COII-F (5'-ATGGCAACATGAGCAAATT-3') and COII-R (5' -GTATAAAACTATGATTAGC-3').
Dirofilaria immitis microfilariae. The microfilaremic and amicrofilaremic canine blood samples were obtained by one of the authors from his previous study (Zarei et al., 2016). The infected dog was a naturally infected 11-year-old mix breed. The presence of D. immitis in the samples was identified and confirmed by Knott's concentration technique (KCT) (Knott, 1939) and PCR (Rishniw et al. , 2006), respectively. The concentration of mf
was determined by the mean value of ten counts of 20 ^l blood smear under magnification 40x.
Anopheles stephensi infection. A colony of 5-8-day-old female mosquitoes was selected and starved for 1 d prior to blood feeding. Two groups of female mosquitoes were fed using microfilariaemic blood (n = 140; test group, T) or amicrofilaremic blood (n = 94; control group, C). The mosquitoes from each group (T and C) were placed separately in cages 50x50x50 cm in size. All mosquitoes in test and control groups were fed using an artificial feeding apparatus (Cosgrove et al., 1994) loaded with positive or negative blood samples containing heparin anticoagulant solution, respectively. The time exposure to blood for each group was one hour. Immediately after blood feeding, the unfed mosquitoes were removed from the cages. All engorged mosquitoes were fed with 10% sucrose solution.
Dissection of mosquitoes colonies. To determine the number of ingested mf, three mosquitos from the T group were killed within 24 h post infection and dissected. For a period of 16 d, the members of the engorged group T were examined at scheduled days (Table 1). The groups of mosquitoes, comprising 1-8 killed or naturally dead mosquitoes, were dissected daily to detect different stages of D. immitis larvae. The mosquitoes were killed by putting them in dry ice and dissected in phosphate buffer saline (PBS) solution for detection of D. immitis larvae under a stereo-microscope, and the development stages were determined as described previously (Taylor, 1960). After microscopic studies, the slides were rinsed with a few drops of PBS for subsequent molecular analysis. Molecular analysis was applied for each sample to monitor the presence of D. immitis infective/non-infective larvae using PCR (Ferreira et al., 2015). For this purpose, DNA extraction was performed separately for both head+thorax and abdomen in order to differentiate between D. immitis infective and infected larvae. The amplification of ITS2 region was carried out using DIDR-F and DIDR-R primers.
Susceptibility analysis of A. stephensi for transmission of D. immitis. The mortality rate of the mosquitoes was calculated daily, within 10 days post infection (dpi) and until 16 dpi as the percentage of naturally dead mosquitoes over the total blood-fed mosquitoes in each cage. The significant differences in mortality rates between the two cages were calculated using Fisher's exact test; P values below 0.05 were considered as statistically significant. The infection rate (IR) was defined as the number of blood-fed mosquitoes with infective L3 in their body multiplied by 100 and divided by the number of surviving mosquitoes at the end of
Fig. 1. Map of Iran indicating the distribution of Anopheles stephensi and heartworm disease using various markers.
Table 1. Anopheles stephensi specimens examined microscopically for microfilaria (mf) and larvae stages of Dirofilaria immitis at scheduled days post infection (dpi).
Total mosquitoes
dpi Killed: total larval stage counted/ Naturally dead: total larval stage counted/
Examined mosquito infected; mean mosquito infected; mean
Mf L2 L3 Mf L1 L3
i 3 29/3; 9.6 0 0 - - -
2 5 - - - 25/5; 5 0 0
3 6 - - - 25/2; 12.5 18/3; 6 0
4 7 - - - 20/4; 5 17/3; 5.6 0
5 6 - - - 0 13/3; 4.3 0
6 6 - - - 0 20/4; 5 0
7 8a 0 0 0 0 12/4;3 0
8 4 0 8/3; 2.6 0 - - -
9 7b 0 0 0 5/3; 1.6 0 0
10 6 0 0 8/2; 4 - - -
11 6 0 3/3; 1 0 - - -
12 7 0 0 8/3; 2.6 - - -
13 8c 0 4/1 9/2; 4.5 0 3/1 10/1
14 4d 0 0 0 0 0 8/3; 2.6
15 4 - - - 0 0 9/3; 3
16 3 - - - 0 0 8/2; 4
Total 90 29/3; 9.6 15/7; 2.1 25/7; 3.5 75/14; 5.3 83/18; 4.6 35/9; 3.8
Mosquitoes found dead at the bottom of the cage. Mean indicates the mean number of larvae per infected mosquito. a - three killed and five naturally dead, b - four killed and three naturally dead, c - five killed and three naturally dead, d - one killed and three naturally dead larvae.
Fig. 2. Morphological identification of the developmental stages of Dirofilaria immitis in dog and Anopheles stephensi. (A) Microfilariae of D. immitis in the dog blood. (B) Microfilariae of D. immitis in the abdomen of A.
stephensi.
incubation period. The vector efficiency index (VEI) was calculated as the average number of L3 developed in the mosquitoes after extrinsic development period, multiplied by 100 and divided by the mean number of ingested mf as described before (Kartman, 1954; Silaghi et al., 2017). The extrinsic development time of D. immitis to the L3 stage, based on the Dirofilaria development unit, was determined to be 10 d at the temperature of our insectary.
Bioinformatics analysis. Sequencing of the amplified gene of samples was performed at the Bioneer Coporation (Seoul, Korea). The obtained sequences were analysed and manually edited using BioEdit software (version 5.0.6; North Carolina State University). Sequence homology analyses were accomplished using the NCBI databases with BLAST search tool (http://www.ncbi.nlm.nih.gov/).
RESULTS
The morphological and molecular analyses demonstrated that only A. stephensi was present. The amplified sequence of A. stephensi was deposited in the GenBank with the accession number of KY863454. KCT and morphometric analysis indicated that the suspected dog sample was positive for D. immitis infection (Fig. 2A).
The molecular results confirmed the morphological identification of D. immitis larvae. The BLAST search identified our sequence as D. immitis and showed 99% homology with other sequences of D. immitis available in the NCBI database (e.g., AF217800). The amplified sequence
was deposited in the GenBank with the accession number of KY863453. After feeding A. stephensi with blood meal containing 1,500 mf ml-1, different percentages of the fed female mosquitoes were observed in two groups (64% and 85% in T and C, respectively). The average number of ingested mf per female was 9.6 (ranged from 1 to 18) in T group (n = 90, excluding the three mosquitoes sacrificed for the calculation of the mf intake). The mortality rates of A. stephensi in groups T and C at the end of study were 59% (n = 51) and 25% (n = 19), respectively (P < 0.05), being higher during the first 10 dpi, i.e., 44% in group T vs 13% in group C; P < 0.05 (Fig. 3).
Different developmental stages of D. immitis (Fig. 2B) in killed and naturally dead mosquitoes are shown in Table 1. The larvae were discriminated based on a morphological identification key as the sausage-like larvae (L1) as well as second (L2) and third (L3) stage larvae. Different parts of A. stephensi were screened for D. immitis larvae using molecular analyses, which confirmed the morphological identification of D. immitis larvae. The L1 began to be identified at 3 d post infection (dpi) and persisted until 13 dpi. L1 was found melanized and dead at 6 dpi but L3 was identified from 10 to 16 dpi. These infective larvae were observed emerging from the proboscis of four mosquitoes. The average number of infective larvae developed in the mosquitoes after extrinsic development period was 1.5. At the end of the study, 16 mosquitoes developed L3, and 49 mosquitoes survived at the end of incubation period, giving an estimated IR of 32.6% and VEI of 17%.
Fig. 3. The mortality rate of test and control groups of the Anopheles stephensi.
DISCUSSION
We found that A. stephensi mosquitoes isolated from Iran was susceptible to infection with D. immitis. To the best of our knowledge, this is the first study to evaluate the susceptibility of wild caught mosquitoes from Iran to D. immitis in the laboratory. The mortality rates of A. stephensi in T and C groups were 59% and 25%, respectively. The higher rate of mortality in the T group may be due to the development of larvae in the mosquitoes (Montarsi et al., 2015). The rate of mortality in our study is similar to the rates reported for C. quinquefasciatus (Say) and A. aegypti (Linnaeus) in the previous studies (Serrao et al., 2001; Carvalho et al., 2008). The melanization process only occurred in L1 but not in other stages of D. immitis larvae; this result is consistent with the results of other investigations (Nayar & Knight, 1999; Montarsi et al., 2015). The probable role of A. stephensi, as a competent vector for D. immitis, is also determined using our IR and VEI values (32.6% and 17%, respectively). A mosquito species is assumed to be a competent vector for infection when having the IR value of 10% and VEI of 9% (Kartman, 1953). In this regard, C. quinquefasciatus, which was fed with
blood containing 1,913 mf ml-1 and showing IR of 46% and VEI of 7.8%, was considered as 'refractory' (Carvalho et al., 2008). Recently, A. koreicus has been regarded as a competent vector of D. immitis that indicates high values of IR and VEI (68.2% and 25.2%, respectively) (Montarsi et al., 2015). Vectorial capacity is a measure not only of vector competence (the ability to become infected and transmit the pathogen) but also of vector density, host preference, feeding frequency, longevity, etc. (Grieve et al., 1983). Despite the probable susceptibility of A. stephensi, more investigations on host preference and host-seeking activity are needed to designate this mosquito as a competent vector for canine and human dirofilariasis.
In conclusion, A. stephensi, which is a widely distributed species in the Middle East, could be a competent vector of D. immitis in that region. The result obtained from the present study demonstrates that the A. stephensi represents great transmission potential for D. immitis and could be considered as a competent vectors for canine dirofilariasis or probably human dirofilariasis in the southern parts of Iran. Nevertheless, experimental and field evidences are necessary to confirm the role of A. stephensi in transmission of human and canine
dirofilariasis. As illustrated in Fig. 1, the distribution of heartworm disease and A. stephensi population overlaps only in the southern part of Iran. Therefore, more studies are required to determine the potential vectors of dirofilariasis in other areas of the country.
ACKNOWLEDGEMENTS
The present study was a part of a dissertation (R.S.) submitted in partial fulfillment of the requirements for a Ph.D. degree at the Shiraz University of Medical Sciences, Shiraz, Iran. This investigation received a financial support from Shiraz University of Medical Sciences, Shiraz, Iran and Tehran University of Medical Sciences, Tehran, Iran (project nos 7568 and 27251, respectively). We also thank M. Saffari and the Centre of Consultation and Research at Shiraz University of Medical Sciences for useful suggestions.
REFERENCES
Aranda, C., Panyella, O., Eritja, R. & Castella, J. 1998. Canine filariasis: importance and transmission in the Baix Llobregat area, Barcelona (Spain). Veterinary Parasitology 77: 267-275. Azari-Hamidian, S. 2007. Checklist of Iranian mosquitoes (Diptera: Culicidae). Journal of Vector Ecology 32: 235-242. Azari-Hamidian, S. & Harbach, R.E. 2009. Keys to the adult females and fourth-instar larvae of the mosquitoes of Iran (Diptera: Culicidae). Zootaxa 2078: 1-33.
Azari Hamidian, S., Yaghoobi Ershadi, M., Javadian, E., Abai, M., Mobedi, I., Linton, Y.M. & Harbach, R. 2009. Distribution and ecology of mosquitoes in a focus of dirofilariasis in northwestern Iran, with the first finding of filarial larvae in naturally infected local mosquitoes. Medical and Veterinary Entomology 23: 111-121. Bradley, T.J. & Nayar, J.K. 1985. Intracellular melanization of the larvae of Dirofilaria immitis in the Malpighian tubules of the mosquito, Aedes sollicitans. Journal of Invertebrate Pathology 45: 339-345.
Carvalho, G. A. D., Alves, L.C., Maia, R.T., Andrade, C.F.S.D., Ramos, R.A.D.N. & Faustino, M.A.D.G. 2008. Vector competence of Culex quinquefasciatus Say, 1823 exposed to different densities of microfilariae of Dirofilaria immitis (Leidy, 1856). Revista Brasileira de Entomologia 52: 658-662.
Chavshin, A.R., Oshaghi, M.A., Vatandoost, H., Hanafi-Bojd, A.A., Raeisi, A. & Nikpoor, F. 2014. Molecular characterization, biological forms and
sporozoite rate of Anopheles stephensi in southern
Iran. Asian Pacific Journal of Tropical Biomedicine 4: 47-51.
Cosgrove, J., Wood, R., Petric, D., Evans, D. & Abbott, R. 1994. A convenient mosquito membrane feeding system. Journal of the American Mosquito Control Association 10: 434-436.
Ferreira, C.A.C., De Pinho Mixäo, V., Novo, M.T.L.M., Calado, M.M.P., Goncalves, L.A.P., Belo, S.M.D. & De Almeida, A.P.G. 2015. First molecular identification of mosquito vectors of Dirofilaria immitis in continental Portugal. Parasites and Vectors 8: 139.
Grieve, R.B., Lok, J.B. & Glickman, L.T. 1983. Epidemiology of canine heartworm infection. Epidemiologic Reviews 5: 220-246.
Jafari, S., Gaur, S. & Khaksar, Z. 1996. Prevalence of Dirofilaria immitis in dogs of Fars province of Iran. Journal of Applied Animal Research 9: 27-31.
Kartman, L. 1953. Factors influencing infection of the mosquito with Dirofilaria immitis (Leidy, 1856). Experimental Parasitology 2: 27-78.
Kartman, L. 1954. Suggestions concerning an index experimental filarial infection in mosquitoes. American Journal of Tropical Medicine and Hygiene 3: 329-337.
Khedri, J., Radfar, M.H., Borji, H., Azizzadeh, M. & Akhtardanesh, B. 2014. Canine heartworm in southeastern of Iran with review of disease distribution. Iranian Journal of Parasitology 9: 560-567.
Khodabakhsh, M., Malmasi, A., Mohebali, M., Zarei, Z., Kia, E.B. & Azarm, A. 2016. Feline dirofilariasis due to Dirofilaria immitis in Meshkin Shahr District, northwestern Iran. Iranian Journal of Parasitology 11: 269-273.
Knott, J. 1939. A method for making microfilarial surveys on day blood. Transactions of the Royal Society of Tropical Medicine and Hygiene 33: 191-196.
Mirahmadi, H., Maleki, A., Hasanzadeh, R., Ahoo, M.B., Mobedi, I. & Rostami, A. 2017. Ocular dirofilariasis by Dirofilaria immitis in a child in Iran: a case report and review of the literature. Parasitology International 66: 978-981.
Montarsi, F., Ciocchetta, S., Devine, G., Ravagnan, S., Mutinelli, F., Frangipane Di Regalbono, A., Otranto, D. & Capelli, G. 2015. Development of Dirofilaria immitis within the mosquito Aedes (Finlaya) koreicus, a new invasive species for Europe. Parasites and Vectors 8: 177.
Nayar, J. & Knight, J. 1999. Aedes albopictus (Diptera: Culicidae): an experimental and natural host of Dirofilaria immitis (Filarioidea: Onchocercidae) in Florida, USA. Journal of Medical Entomology 36: 441-448.
Negahban, S., Daneshbod, Y., Atefi, S., Daneshbod, K., Sadjjadi, S.M., Hosseini, S.V., Bedayat, G.R. & Abidi, H. 2007. Dirofilaria repens diagnosed by the presence of microfilariae in fine needle aspirates: a case report. Acta Cytologica 51: 567-570.
Rishniw, M., Barr, S.C., Simpson, K.W., Frongillo, M.F., Franz, M. & Domínguez Alpizar, J.L. 2006. Discrimination between six species of canine microfilariae by a single polymerase chain reaction. Veterinary Parasitology 135: 303-314.
Sadjjadi, S.M., Mehrabani, D. & Oryan, A. 2004. Dirofilariasis of stray dogs in Shiraz, Iran. Journal of Veterinary Parasitology 18: 181-182.
Salahi-Moghaddam, A., Khoshdel, A., Dalaei, H., Pakdad, K., Nutifafa, G. & Sedaghat, M. 2017. Spatial changes in the distribution of malaria vectors during the past 5 decades in Iran. Acta Tropica 166: 45-53.
Serrao, M.L., Labarthe, N. & Lourenco-De-Oliveira, R. 2001. Vectorial competence of Aedes aegypti (Linnaeus 1762) Rio de Janeiro strain, to Dirofilaria immitis (Leidy 1856). Memórias do Instituto Oswaldo Cruz 96: 593-598.
Silaghi, C., Beck, R., Capelli, G., Montarsi, F. & Mathis, A. 2017. Development of Dirofilaria immitis and Dirofilaria repens in Aedes japonicus and Aedes geniculatus. Parasites and Vectors 10: 94.
Simon, F., Lopez-Belmonte, J., Marcos-Atxutegi, C., Morchon, R. & Martín-Pacho, J.R. 2005. What is happening outside North America regarding
human dirofilariasis? Veterinary Parasitology 133: 181-189.
Simon, F., Morchon, R., Gonzalez-Miguel, J., Marcos-Atxutegi, C. & Siles-Lucas, M. 2009. What is new about animal and human dirofilariosis? Trends in Parasitology 25: 404-409.
Simon, F., Siles-Lucas, M., Morchon, R., GonzalezMiguel, J., Mellado, I., Carreton, E. & Montoya-Alonso, J.A. 2012. Human and animal dirofilariasis: the emergence of a zoonotic mosaic. Clinical Microbiology Reviews 25: 507-544.
Subbarao, S.K., Vasantha, K., Adak, T., Sharma, V.P. & Curtis, C.F. 1987. Egg-float ridge number in Anopheles stephensi: ecological variation and genetic analysis. Medical and Veterinary Entomology 1: 265-271.
Taylor, A.E.R. 1960. The development of Dirofilaria immitis in the mosquito Aedes aegypti. Journal of Helminthology 34: 27-38.
Webber, W.A. & Hawking, F. 1955. Experimental maintenance of Dirofilaria repens and D. immitis in dogs. Experimental Parasitology 4: 143-164.
Zarei, Z., Kia, E.B., Heidari, Z., Mikaeili, F., Mohebali, M. & Sharifdini, M. 2016. Age and sex distribution of Dirofilaria immitis among dogs in Meshkin-Shahr, northwest Iran and molecular analysis of the isolates based on COX1 gene. Veterinary Research Forum 7: 325-330.
URL: http://www.ncbi.nlm.nih.gov/ (accessed: August 15, 2017).
R. Solgi, S.M. Sadjjadi, M. Mohebali, N.D. Djadid, A. Raz, S. Zakeri and Z. Zarei. Способность Anopheles stephensi (Diptera: Culicidae) переносить Dirofilaria immitis (Spirurida: Onchocercidae). Резюме. Нематоды Dirofilaria immitis обычны в южных провинциях Ирана, где складываются походящие условия для развития комаров Anopheles stephensi. Проведено исследование способности A. stephensi служить вектором-переносчиком для нематод D. immitis в лабораторных условиях. Кровь собак с добавлением антикоагулянтов, содержащие 1,500 микрофилярий в 1 мл была использвоана для заражения голодавших самок комаров. Всего 140 комаров было использовано для эксперимента (Т группа) и 94 особи комаров служили контролем (C группа). Комаров из Т группы вскрывали через определенные промежутки времени для морфологического и молекулярного выявления развивающихся стадий D. immitis. Результаты показали, что в среднем каждая самка комара заглатывала 9.6 микрофилярий. В конце эксперимента было установлено, что 49 самок пережили условия данного опыта (32.6%) и у 16 самок комаров развились инвазионные личинки 3-й стадии D. immitis. Таким образом, индекс эффективности переноса составил 17%. Сделан вывод о способности A. stephensi служить вектором для D. immitis.
