Nematoda. ISSN 2358-436X. Copyright © 2014. This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/3.0), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

1http://dx.doi.org/10.4322/nematoda.09015 Nematoda, 2015;2: e092015

Potential of entomopathogenic nematodes as biocontrol agents of immature stages of Aedes aegyptiDenise de Oliveira Cardosoa, Vicente Martins Gomesa, Claudia Dolinskia and Ricardo Moreira Souzaa*

a Grupo de Pesquisa em Nematologia, Laboratório de Entomologia e Fitopatologia (LEF), Centro de Ciências e Tecnologias Agropecuárias (CCTA), Universidade Estadual do Norte Fluminense Darcy Ribeiro (UENF), Campos dos Goytacazes (RJ) Brazil

*ricmsouza@censanet.com.br

HIGHLIGHTS• Heterorhabditisindica LPP35, H. indica LPP1 and H. baujardi LPP31 were virulent to Aedesaegypti L3 and L4.• Steinernemacarpocapsae All was not virulent to A.aegypti L3 and L4.• There was a positive correlation between larvae mortality rate and inoculum density.• A.aegypti L3 and L4 supported H. indica LPP35 progeny production.• H.indica LPP35, found in a bromeliad tank, seems to be the strain best fitted to control A.aegypti.

ABSTRACT: Heterorhabditisindica LPP35 caused more than 85% mortality of third (L3) and fourth (L4) instars of Aedesaegypti. However, the same nematode strain did not cause mortality of L1 and L2 instars. When compared to the virulence of different species/isolates of Heterorhabditis and Steinernema against the L3 and L4, there was greater mortality among larvae of A.aegypti when species of Heterorhabditis were used. Results from H.indica LPP35 stood out, as this strain caused 95 and 94% mortality in Assays 1 and 2, respectively. In the different dose‑response assay, the increase in concentration of H.indica LPP35 infective juveniles (IJs) against L3 and L4 presented a positive correlation with the mortality of the mosquito larvae. Doses of 10, 20 and 40 IJs/larva presented mortality below 50%, whereas doses of 80, 100 IJs/larva presented mortality above 75%, reaching 100% mortality at the dose of 160 IJs/larva of A.aegypti. H.indica LPP35 completed its biological cycle in larvae of A.aegypti and, on average, about 200 IJs emerged from the insect cadavers. In the laboratory assays, H.indica LPP35 presented potential to be used as an agent for biological control of A.aegypti L3 and L4 larvae.

Keywords: Heterorhabditis, Steinernema, mosquito, dengue fever, chykungunya fever, Heterorhabditisbaujardi LPP7, H.baujardi LPP31, H.indica LPP35, H.indica LPP1.

Cite as Cardoso DO, Gomes VM, Dolinski C, Souza RM. Potential of entomopathogenic nematodes as biocontrol agents of immature stages of Aedesaegypti. Nematoda.2015;2:e092015. http://dx.doi.org/10.4322/nematoda.09015

Received July 14, 2015 Accepted Aug. 06, 2015

INTRODUCTIONMosquitoes (Diptera: Culicidae) are of great importance in health, as they are vectors of parasitic and

viral diseases in humans and other animals. Aedes aegypti L. stands out as the vector for the viruses that cause dengue and chikungunya fevers, which have afflicted hundreds of thousands of people per year in more than 100 tropical and subtropical countries[1], as well as being a vector of yellow fever[2].

Various biological control agents, such as viruses, bacteria, fungi and nematodes have been tested in the control of urban pests as an alternative to using conventional insecticides with their attendant problems[3, 4, 5]. In recent decades, the potential of entomopathogenic nematodes (EPNs) has been harnessed for the biological control of insect pests[6]. In some countries, commercial products based on EPNs have already been incorporated into integrated pest management programs for some agricultural crops[7]. Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema

ORIGINAL ARTICLE

2http://dx.doi.org/10.4322/nematoda.09015

Cardoso et al.

Nematoda, 2015;2: e092015

Entomopathogenic nematodes for the control of Aedes aegypti larvae

(Rhabditida) are obligate parasites of insects. These nematodes have a symbiotic relationship with bacteria in the genera Photorhabdus and Xenorhabdus, respectively. Infective juveniles (IJs), the only stage of the nematodes found in the soil, enter hosts through natural openings such as the mouth, anus or spiracles, but heterorhabditid IJs can also enter through the cuticle. After penetrating into the host’s hemocoel, the nematodes release their symbiotic bacteria, which usually kill the host within 24 to 48 h. The bacteria are also responsible for antibiotic production and for providing nutrition for the nematodes. The nematodes feed, develop, mate, and often complete 2-3 generations within the host cadaver. When resources within the cadaver are depleted, a new generation of IJs is produced and leaves the cadaver to search for new hosts[6].

Welch[8] was the first to test the use of EPNs in controlling Aedes spp., including A. aegypti, in the laboratory and in the field. The application of Steinernema carpocapsae (Weiser) Wouts, Mracek, Gerdin & Bedding reduced the larval population and the emergence of adults. Several authors have examined the fundamental aspects of the relationship between EPNs and the larval stages of mosquitoes and black flies. Dadd[9] noticed that the larval stages L1 and L2 of Culex pipiens L. have mouth dimensions that prevent them from ingesting S. carpocapsae IJs, and few are ingested by L3. The L4 ingest hundreds of IJs, but only a small proportion of these are not damaged by the mouthparts and pharynx of the insect. According to Molloy et al.[10] and Gaugler & Molloy[11], some larvae of black flies (Simullium spp.) survive the ingestion of S. carpocapsae IJs and can defecate them still alive. On the other hand, many larvae of mosquitoes die even if the ingested IJs do not reach the hemocoel, presumably because they cause physical damage to the larval gut. According to Poinar & Kaul[12], the hemocoel invasion of C. pipiens L3 and L4 by H. bacteriophora Poinar IJs causes septicemia by the symbiotic bacteria P. luminescens (Thomas & Poinar) Boemare, Akhurst & Mourant or by Pseudomonas aeruginosa (Schröter) Migula, which normally colonizes the larval gut, but only enters the hemocoel after the peritrophic membrane has been perforated by the IJs.

Despite susceptibility to the symbiotic nematode-bacteria complex, larvae of Aedes spp., including A. aegypti, encapsulated part of the IJs of S. carpocapsae, S. feltiae (Filipjev) Wouts, Mracek, Gerdin & Bedding, and Heterorhabditis heliothidis Khan, Brooks & Hirschmann, causing their death. However, most of the larvae die too[13, 14]. The encapsulation and melanization of IJs by larvae of mosquitoes have been studied at histological and ultrastructural levels[15, 16, 17]. It is worth mentioning that some P. luminescens and Xenorhabdus nematophila (Poinar & Thomas) Thomas & Poinar strain have been shown to present oral toxicity to larvae of A. aegypti on their own[18, 19].

Other authors have assessed the virulence of EPNs to mosquito larvae, under laboratory conditions[12, 13, 14, 20, 21, 22, 23, 24]. Nonetheless, the discrepancy in methodological procedures and results limits generalizations. For instance, mortality rates varied from zero in assays involving H. bacteriophora, S. feltiae or S. rarum (Doucet) Mamiya against C. pipiens[20] up to nearly 100% in H. bacteriophora against the same C. pipiens[12]. The calculated LD50, useful to predict aspects related to application technology, has been found to vary widely, from 63-80 IJs/larva in H. bacteriophora-C. pipiens assays up to 1,263 IJs/larva for S. feltiae-A. aegypti[12, 14]. Some authors have reported reproduction of EPNs in their host[22, 23, 24], although no numbers were provided. Collectively, these reports suggest that regional experiments should be performed before one attempts to deploy a biocontrol program.

In the last three decades rampant epidemics of dengue fever have occurred in Brazil and other tropical and subtropical countries, despite efforts by health officials to treat larval sites with the application of insecticides, bacteria and transgenic male mosquitoes[25, 26]. Clearly, more options are needed. Our objective was to test the virulence of different EPNs against A. aegypti larvae, as well as determining the dose-response and progeny of the most virulent strain. We hypothesized that LPP35, a strain isolated from the phytotelma of the bromeliad Nidularium procerum Lindm, would be the most virulent. Insects, particularly dipterans, are the most abundant metazoans associated with bromeliad phytotelmata in the Atlantic Forest biome[27]. Therefore, it is plausible that an EPN strain adapted to this biome would be particularly effective in parasitizing mosquito larvae.

MATERIAL AND METHODS

Cultivating EPNs and obtaining eggs and larvae of Aedes aegyptiThe EPNs used in the experiments (Table 1) were multiplied in larvae of Galleria mellonella

L. (Lepidoptera: Pyralidae), according to a procedure described by Woodring & Kaya[28]. The IJs that emerged from the larvae were maintained in an aqueous suspension in cell culture flasks at a temperature of 16 °C in the dark, for use in the experiments.

3http://dx.doi.org/10.4322/nematoda.09015

Cardoso et al.

Nematoda, 2015;2: e092015

Entomopathogenic nematodes for the control of Aedes aegypti larvae

Eggs of A. aegypti were obtained from oviposition traps installed in residences in the city of Campos dos Goytacazes, Rio de Janeiro, Brazil. The traps were made in black plastic pots with 1,500 mL capacity, with six pieces of Eucatex wood (3 × 12 cm) attached to the edge with elastic bands, maintaining the rough side of the wood turned toward the inner part of the pot. Each trap received 250 mL of tap water. The piece of wood containing the eggs was put to dry in the shade for 48 h. After this period they were acclimatized in plastic Gerbox boxes and stored at room temperature. For egg hatching, the pieces of wood were immersed in deep trays containing about 1 L of tap water to which about 0.5 g of mouse feed had been added. The development from L1 through L4 took approximately 48 h at room temperature (25 ± 3 °C). To conclude the experiments, the larvae were classified at stages L1 to L4 in accordance with their mean lengths of 2.5, 3.9, 5 and 7.3 mm, respectively.

Virulence of Heterorhabditis indica LPP35 to different larval stages of Aedes aegyptiFor each larval stage (L1 to L4), plastic cups (5 × 3.5 cm or 50 mL) containing 10 mL of tap water

received 10 larvae and a suspension of 1 mL containing 1,000 IJs. Ten repetitions (cups) were used for each larval stage. The control followed the same procedure, except for the absence of nematodes (different larval stages). The experiment was conducted in a completely randomized design. The pots were covered and acclimatized in a growth chamber at 25 ± 2 °C, in the dark. The evaluations were done daily until pupae appeared (at about 7 days), with the number of live and dead larvae being counted. Dead larvae were those presenting a change in color, from white to brown, and the absence of movement. The dead larvae were observed under a stereoscopic microscope to check the presence of IJs inside them. The entire experiment was repeated after the first, with new nematode and insect batches, Assays 1 and 2.

Virulence of different EPNs to larvae of Aedes aegyptiAs stages L3 and L4 were the only ones susceptible to H. indica LPP35 (see Results and Discussion),

these stages were used in the following experiments. For each EPN (Table 1) 10 plastic cups were used containing 10 mL of water, 1,000 IJs and 10 specimens of a mixture of L3 and L4 (L3/L4) of A. aegypti larvae. The pots were closed and stored in a growth chamber at 25 ± 2 °C, 80% RH and photoperiod of 12 h. The control used the same procedures, except for the absence of nematodes. The experiment was conducted in a completely randomized design. The evaluations were carried out as described in the previous experiment. The entire experiment was repeated after the first, with new nematode and insect batches, Assays 1 and 2.

Dose-response relationship in the virulence of Heterorhabditis indica LPP35 to L3/L4 of Aedes aegypti

To plastic cups of 50 mL, containing 10 mL of tap water and one L3/L4, 1 mL of aqueous suspension was added, containing the doses (treatments) 10, 20, 40, 80, 100 or 160 IJs. The control treatment used the same, except for the absence of nematodes. Ten repetitions (cups) were used and the experiment was conducted in a completely randomized design. The cups were closed and kept in a growth chamber at a temperature of 25 ± 2 °C, in the dark. The evaluations were carried out as described in the previous experiment. The entire experiment was repeated after the first, with new nematode and insect batches, Assays 1 and 2.

Table 1. Entomopathogenic nematode strains used in the assays against Aedes aegypti larvae.

Species, strain Origin (state, country)

Steinernema carpocapsae ALL North Carolina, USA

Heterorhabditis indica LPP35 Rio de Janeiro, Brazil

Heterorhabditis baujardi LPP31 Alagoas, Brazil

Heterorhabditis indica LPP1 Rondônia, Brazil

Heterorhabditis baujardi LPP7 Rondônia, Brazil

4http://dx.doi.org/10.4322/nematoda.09015

Cardoso et al.

Nematoda, 2015;2: e092015

Entomopathogenic nematodes for the control of Aedes aegypti larvae

Heterorhabditis indica LPP35 progeny in Aedes aegypti L3/L4The larvae were infected in plastic cups that received 50 mL of water and 1 mL of an aqueous

suspension calibrated for 100 IJs/larva. Thirty L3/L4 that presented symptoms of infection and presence of hermaphrodites inside them were individually transferred to Petri dishes with diameter of 6 cm, which were kept for seven days in a growth chamber at 25 ± 2 °C, in the dark. After this period, the IJs emerging from each cadaver were collected and stored separately in test tubes. To count IJs, three aliquots of 0.1 mL were observed in a Peters chamber under a stereoscopic microscope. The progeny/larva was expressed by the mean, standard deviation, coefficient of variation and confidence interval (at 5%), also noting the minimum and maximum values obtained. The experiment was repeated once (Assays 1 and 2).

Statistical analysisFor the statistical analysis of all experiments, the original data were tested for homogeneity of

variances (Cochran and Bartlett tests) and normality of errors (Lilliefors test) at 5% probability, using the System of Statistical Analyses (SAEG)[29]. As the data met the presuppositions, ANOVA was applied. The means for treatments were compared by Tukey test (P< 0.05). For the dose-response experiment, the regression equations were obtained with the program Excel (2007) with alpha 0.05.

RESULTS AND DISCUSSIONAedes aegypti L1 and L2 instars were not affected by EPNs, only L3 and L4. Heterorhabditis indica

Poinar, Karunakar & David LPP35, H. baujardi Phan, Subbotin, Nguyen & Moens LPP31, and H. indica LPP1 were virulent to A. aegypti L3 and L4. There was a positive correlation between larva mortality rate and inoculum density. A. aegypti L3 and L4 supported progeny production of H. indica LPP35, the strain found in the bromeliad, which was also the most virulent strain tested.

H. indica LPP35 caused the death of more than 81% of the A. aegypti L3 and L4, but it was not virulent to L1 or L2 (Figure 1). Data were combined and analyzed (F= 47.1; df= 3, 79; P≤0.05). In the cups, mosquito

Figure 1. Mortality rate of larval stages of Aedes aegypti (L1, L2, L3, L4) following exposure to infective juveniles of Heterorhabditis indica LPP35. Values are means of twenty replicates for each stage, each with 10 larvae. The experiment was conducted twice (Assays 1 and 2) and the data combined for analysis. There were no death in any larval stage used in the control treatment. Means followed by the same letter do not differ according to Tukey test, at 5%.

5http://dx.doi.org/10.4322/nematoda.09015

Cardoso et al.

Nematoda, 2015;2: e092015

Entomopathogenic nematodes for the control of Aedes aegypti larvae

Figure 2. Fourth instar larvae of A. aegypti, healthy (top), and infected by Heterorhabditis indica LPP35 with typical light brown coloration and a hermaphrodite coming out of the cephalic region (bottom).

larvae and nematodes were at the bottom and we could visualize L3/L4 ingesting IJs. Moreover, only L3 and L4 presented the brown coloration typical of the infection caused by heterorhabditids with nematodes inside of them (Figure 2). These results corroborate the studies carried out by Dadd[9] and Poinar & Leutenegger[17], who noticed that IJs are ingested by L3 and L4 mosquitoes but not L1 or L2, due to the reduced dimensions of their mouth parts.

H. indica LPP35, H. baujardi LPP 31 and H. indica LPP1 were virulent to L3/L4 of A. aegypti, in both assays with mortality ranging from 80 to 95% in the first assay and 82 to 96% in Assay 2, both differing from the control (F= 157.95; df= 5, 119; P≤ 0.05 and F= 141.84; df= 5, 119; P≤ 0.05, respectively) (Figure 3). Results for H. baujardi LPP7 were inconclusive, because the mortality rate reached nearly 40% in the first assay, but it did not differ from the control in the second assay. Previous authors reported generally high virulence of Heterorhabditis spp. towards mosquito larvae, with the exceptions of H. bacteriophora vs. C. pipiens[20] and H. indica vs. C. gelidus Theobald[21].

In the present work, S. carpocapsae All was not virulent to L3/L4 of A. aegypti. Nonetheless, Welch[8] tested another isolate of S. carpocapsae – DD136 – and it was virulent to mosquito larvae. However, we believe this strain is, in reality, from S. feltiae as mentioned in other studies[30, 31]. Differences among isolates of the same EPN species in virulence towards a host may stem from differences in the tritrophic symbiotic interaction of nematode-bacteria-insect. Nonetheless, methodological differences (e.g. inoculation procedures) and errors (e.g. low viability of IJs) cannot be ruled out. S. feltiae may be marginally virulent to mosquito larvae[14, 20], while for S. rarum there are reports of no virulence towards C. pipiens[20] and a 75% mortality rate of C. apicinus Philippi, larvae[22].

In this study, we used on purpose a small quantity of water that concentrated the mosquito larvae and IJs, therefore maximizing their encounter and likelihood of infection. This is because we wanted to know the EPNs’ performance under the best conditions. In preliminary field experiments, we varied the water volume to find out whether it would affect the infection, and it did indeed have an effect[32].

Several authors[33, 34] have highlighted the importance of experiments that supply information about the EPN-host interaction. Dose-response curves indicate the potential of an EPN in relation to a pest. In the current work, the increase in the dose of IJs of H. indica LPP35/larva was positively correlated with the mortality rate of L3/L4 of A. aegypti (Figure 4). This trend was also reported for S. feltiae and H. heliotidis against A. aegypti[14] and for S. carpocapsae and H. bacteriophora against C. pipiens[12, 19].

As regards the progeny of H. indica LPP35 obtained in L3/L4 of A. aegypti, in the first assay, a mean of 197 IJs/larva was obtained (S2= 83.3; CV%= 22.3; confidence interval = 31), with minimum and maximum values of 80 and 377, respectively. In the second assay, the values were a mean of 245 IJs/larva (S2= 79.1; CV%= 23.9; confidence interval = 31.7), with minimum and maximum values

6http://dx.doi.org/10.4322/nematoda.09015

Cardoso et al.

Nematoda, 2015;2: e092015

Entomopathogenic nematodes for the control of Aedes aegypti larvae

Figure 4. Mortality rate of a mixture of L3 and L4 of A. aegypti following exposure to different concentrations of infective juveniles of Heterorhabditis indica LPP35/larva. Values are means of ten replicates for each concentration, each with one larva.

Figure 3. Mortality rate of a mixture of L3 and L4 of A. aegypti following exposure to infective juveniles of Steinernema carpocapsae All, Heterorhabditis baujardi LPP7, H. baujardi LPP31, H. indica LPP1, and H. indica LPP35. Values are means of ten replicates for each isolate, each with 10 larvae. The experiment was conducted twice (Assays 1 and 2). Means followed by the same letter do not differ according to the Tukey test, at 5%.

7http://dx.doi.org/10.4322/nematoda.09015

Cardoso et al.

Nematoda, 2015;2: e092015

Entomopathogenic nematodes for the control of Aedes aegypti larvae

of 82 and 355, respectively. This low progeny is probably due to the small number of whole IJs that managed to penetrate the hemocoel of the larvae of A. aegypti[9] and to the low mass (= few nutritional resources) of the host, which determines a smaller progeny of EPNs[35]. However, these results suggest that the EPNs recycling can take place in natural conditions. We also noticed that the size of IJs was the same as that of those produced in G. mellonella or A. aegypti larvae. However, we still need to investigate further to establish if their virulence is similar.

Collectively, these results suggest the potential of H. indica LPP1, H. baujardi LPP31 and particularly of H. indica LPP35 to reduce the density of L3 and L4 of A. aegypti in aquatic urban environments. Nevertheless, the near 100% mortality rate observed for LPP35 under laboratory conditions may not occur under field conditions. Of particular concern are water temperature peaks in small oviposition sites, which may reach unsuitable levels for IJs during summer daytime. Preliminary field experiments, however, have confirmed the potential of LPP35[32], and this is encouraging for new on-going trials.

CONCLUSIONSHeterorhabditis baujardi LPP31, H. indica LPP35 and H. indica LPP1 were highly virulent to L3 and L4

of A. aegypti under laboratory conditions, while S. carpocapsae All was not virulent at all. Results for H. baujardi LPP7 were inconsistent. A positive correlation was observed between the mortality rate of L3 and L4 and the inoculum density of H. indica LPP35 IJs/larva. A. aegypti L3 and L4 supported the production of a small progeny of H. indica LPP35, a strain obtained from a bromeliad tank. This strain seems very promising for A. aegypti larval control.

ACKNOWLEDGEMENTSThe authors are thankful to Rachel Campos-Herrera (University of Neuchâtel, Neuchâtel, Switzerland)

and David Shapiro-Ilan (USDA, Byron, USA) for providing reprints of some early publications on biocontrol of mosquitoes.

REFERENCES1. Centers for Disease Control and Prevention - CDC. 2015. [on line]. Accessed on June 8, 2015. Available from: http://

www.cdc.gov/dengue/

2. Mahesh Kumar P, Kovendan K, Murugan K. 2013. Integration of botanical and bacterial insecticide against Aedes aegypti and Anopheles stephensi. Parasitology Research, 112: 761-771. http://dx.doi.org/10.1007/s00436-012-3196-z. PMid:23242266.

3. Cameron M, Lorenz L. 2013. Biological and environmental control of disease vectors. Commonwealth Agricultural Bureau, Wallingford, 224 pp.

4. Scholte E-J, Knols BGJ, Samson RA, Takken W. 2004. Entomopathogenic fungi for mosquito control: a review. Journal of Insect Science, 4: 19. http://dx.doi.org/10.1093/jis/4.1.19. PMid:15861235.

5. Regis L, Silva-Filha MH, Nielsen-LeRoux C, Charles J-F. 2001. Bacteriological larvicides of dipteran disease vectors. Trends in Parasitology, 17: 377-380. http://dx.doi.org/10.1016/S1471-4922(01)01953-5. PMid:11685898.

6. Kaya HK, Gaugler R. 1993. Entomopathogenic nematodes. Annual Review of Entomology, 38: 181-206. http://dx.doi.org/10.1146/annurev.en.38.010193.001145.

7. Dolinski C, Choo HY, Duncan LW. 2012. Grower acceptance of entomopathogenic nematodes: case studies on three continents. Journal of Nematology, 44: 226-235. PMid:23482423.

8. Welch HE. 1961. Nematodes as agents for insect control. Proceedings of the Entomological Society of Ontario, 92: 11-19.

9. Dadd RH. 1971. Size limitations on the infectibility of mosquito larvae by nematodes during filter-feeding. Journal of Invertebrate Pathology, 18: 246-251. http://dx.doi.org/10.1016/0022-2011(71)90152-2. PMid:5092842.

10. Molloy D, Gaugler R, Jamnback H. 1980. The pathogenicity of Neoaplectana carpocapsae to blackfly larvae. Journal of Invertebrate Pathology, 36: 302-306. http://dx.doi.org/10.1016/0022-2011(80)90039-7.

11. Gaugler R, Molloy D. 1981. Instar susceptibility of Simulium vittatum (Diptera: Simuliidae) to the entomogenous nematode Neoaplectana carpocapsae. Journal of Nematology, 13: 1-5. PMid:19300712.

12. Poinar GO Jr, Kaul HN. 1982. Parasitism of the mosquito Culex pipiens by the nematode Heterorhabditis bacteriophora. Journal of Invertebrate Pathology, 39: 382-387. http://dx.doi.org/10.1016/0022-2011(82)90063-5.

13. Welch HE, Bronskill JF. 1962. Parasitism of mosquito larvae by the nematode, DD136 (Nematoda: Neoaplectanidae). Canadian Journal of Zoology, 40: 1263-1268. http://dx.doi.org/10.1139/z62-102.

14. Molta NB, Hominick WM. 1989. Dose- and time-response assessments of Heterorhabditis heliothidis and Steirnernema feltiae (Nem.:Rhabitida) against Aedes aegypti larvae. Entomophaga, 34: 485-493. http://dx.doi.org/10.1007/BF02374386.

8http://dx.doi.org/10.4322/nematoda.09015

Cardoso et al.

Nematoda, 2015;2: e092015

Entomopathogenic nematodes for the control of Aedes aegypti larvae

15. Bronskill JF. 1962. Encapsulation of rhabditoid nematodes in mosquitoes. Canadian Journal of Zoology, 40: 1269-

1275. http://dx.doi.org/10.1139/z62-103.

16. Andreadis TG, Hall DW. 1976. Neoaplectana carpocapsae: encapsulation in Aedes aegypti and changes in host

hemocytes and hemolymph proteins. Experimental Parasitology, 39: 252-261. http://dx.doi.org/10.1016/0014-

4894(76)90125-9. PMid:816666.

17. Poinar GO Jr, Leutenegger R. 1971. Ultrastructural investigation of the melanization process in Culex pipiens

(Culicidae) in response to a nematode. Journal of Ultrastructure Research, 36: 149-158. http://dx.doi.org/10.1016/

S0022-5320(71)80094-1. PMid:5568354.

18. da Silva OS, Prado GR, Silva JL, Silva CE, da Costa M, Heermann R. 2013. Oral toxicity of Photorhabdus luminescens

and Xenorhabdus nematophila (Enterobacteriaceae) against Aedes aegypti (Diptera: Culicidae). Parasitology Research,

112: 2891-2896. http://dx.doi.org/10.1007/s00436-013-3460-x. PMid:23728731.

19. Ahn J-Y, Lee J-Y, Yang E-J, Lee Y-J, Koo K-B, Song K-S, Lee K-Y. 2013. Mosquitocidal activity of anthraquinones isolated

from symbiotic bacteria Photorhabdus of entomopathogenic nematode. Journal of Asia-Pacific Entomology, 16:

317-320. http://dx.doi.org/10.1016/j.aspen.2013.04.005.

20. de Doucet MM, Bertolotti MA, Giayetto AL, Miranda MB. 1999. Host range, specificity, and virulence of Steinernema

feltiae, Steinernema rarum, and Heterorhabditis bacteriophora (Steinernematidae and Heterorhabditidae) from

Argentina. Journal of Invertebrate Pathology, 73: 237-242. http://dx.doi.org/10.1006/jipa.1998.4831. PMid:10222175.

21. Pandii W, Maharmart S, Boonchuen S, Silapanuntakul S, Somsook V. 2008. Efficacy of entomopathogenic nematodes

(Nematoda:Rhabditida) against Culex gelidus (Diptera: Culicidae) larvae. Journal of Vector Borne Diseases, 5: 24-35.

22. Cagnolo SR, Almirón WR. 2010. Capacity of the terrestrial entomopathogenic nematode Steinernema rarum

(Rhabditida: Steinernematidae) to parasite Culex apicinus larvae (Diptera: Culicidae). Revista de la Sociedad

Entomológica Argentina, 69: 141-145.

23. Peschiutta ML, Cagnolo SR, Almirón WR. 2014. Susceptibility of larvae of Aedes aegypti (Linnaeus) (Diptera: Culicidae)

to entomopathogenic nematode Heterorhabditis bacteriophora (Poinar) (Rhabditida: Heterorhabditidae). Revista

de la Sociedad Entomológica Argentina, 73: 99-108.

24. Zohdy NM, Shamseldean MM, Abd-El-Samie EM, Hamama HM. 2013. Efficacy of the steinernematid and

heterorhabditid nematodes for controlling the mosquito, Culex quinquefasciatus Say (Diptera: Culicidae). Journal

of Mosquito Research, 3: 33-44.

25. Fundação Oswaldo Cruz - FIOCRUZ. 2015. [on line]. Accessed on June 27, 2015. Available from: http://www.fiocruz.

br/rededengue/cgi/cgilua.exe/sys/start.htm?tpl=home

26. Brasil. Ministério da Saúde. 2015. [on line]. Accessed on June 27, 2015. Available from: http://portalsaude.saude.

gov.br/

27. Greeney HF. The insects of plant-held waters: a review and bibliography. Journal of Tropical Ecology. 2001;17:

241-260. http://dx.doi.org/10.1017/S026646740100116X.

28. Woodring L, Kaya HK. 1988. Steinernematid and heterorhabditid nematodes: a handbook of techniques. Series

Bull., 331. Arkansas Agricultural Experiment Station, Fayetteville, Arkansas, 30 pp.

29. Universidade Federal de Viçosa - UFV. 1997. Sistema para Análise Estatísticas: SAEG. Versão 9.1. Universidade

Federal de Viçosa, Viçosa.

30. Dunphy GB, Webster JM. 1985. Influence of Steinernema feltiae (Filipjev) Wouts, Mracek, Gerdin and Bedding DD136

strain on the humoral and haemocytic responses of Galleria mellonella (L.) larvae to selected bacteria. Parasitology,

91: 369-380. http://dx.doi.org/10.1017/S0031182000057437.

31. Ishibashi N, Kondo E. 1986. Steinernema feltiae (DD-136) and S. glaseri: Persistence in soil and bark compost and

their influence on native nematodes. Journal of Nematology, 18: 310-316. PMid:19294183.

32. Cardoso DO. 2014. Potencial e caracterização do isolado de nematoide entomopatogênico LPP35 como agente no

controle de formas imaturas de Aedes aegypti [dissertation]. Universidade Estadual do Norte Fluminense Darcy

Ribeiro, Campos dos Goytacazes, Brazil, 65 pp.

33. Bedding RA, Molyneux AS, Akhurst RJ. 1983. Heterorhabditis spp., Neoaplectana spp., and Steinernema kraussei:

interspecific and intraspecific differences in infectivity for insects. Experimental Parasitology, 55: 249-257. http://

dx.doi.org/10.1016/0014-4894(83)90019-X. PMid:6832283.

34. Molyneux AS, Bedding RA, Akhurst RJ. 1983. Susceptibility of larvae of the sheep blowfly Lucilia cuprina to various

Heterorhabditis spp., Neoaplectana spp., and in undescribed steinernematid (Nematoda). Journal of Invertebrate

Pathology, 42: 1-7. http://dx.doi.org/10.1016/0022-2011(83)90196-9. PMid:6886466.

35. Molina JP, Moino A Jr, Cavalcanti RS. 2004. Produção in vivo de nematodes entomopatogênicos em diferentes insetos

hospedeiros. Arquivos do Instituto Biologico, 71: 347-354.