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Impact of Edhazardia aedis (Microsporidia: Culicosporidae) on a Seminatural Population of Aedes aegypti (Diptera: Culicidae)
Biological Control 18, 39–48 (2000) doi:10.1006/bcon.1999.0805, available online at http://www.idealibrary.com on
Impact of Edhazardia aedis (Microsporidia: Culicosporidae) on a Seminatural Population of Aedes aegypti (Diptera: Culicidae) James J. Becnel1 and Margaret A. Johnson U.S. Department of Agriculture, Agricultural Research Service, Center for Medical, Agricultural, and Veterinary Entomology, Gainesville, Florida 32604 E-mail: [email protected] Received June 15, 1999, accepted November 28, 1999
aegypti has relied on source reduction and broad spectrum chemical larvicides and adulticides. Both methods can be effective but it is difficult to sustain longterm source reduction efforts. The development of resistance to insecticides by mosquitoes and the need for environmental conservation also indicate a necessity for alternative control strategies. Edhazardia aedis (Kudo) Becnel, Sprague, and Fukuda is one of the few pathogens that has been isolated from A. aegypti. This microsporidium was originally observed and described from Puerto Rico (Kudo, 1930) and reisolated in Thailand (Hembree, 1979). E. aedis has a complex life cycle involving both horizontal and vertical transmission, with four different spore types produced in the mosquito host (Becnel et al., 1989; Johnson et al., 1997). Unlike some other polymorphic mosquito-pathogenic microsporidia, there is not an obligate intermediate host involved in the life cycle (Becnel and Andreadis, 1999). A schematic representation of the basic life cycle of E. aedis is shown in Fig. 1. One sporulation sequence occurs in the adult female (infected orally as a larva) and results in the formation of binucleate (transovarial) spores. These spores are involved in vertical transmission of E. aedis to the subsequent generation via infected eggs. In infected progeny, there are two sporulation sequences in larval fat body. One sporulation sequence involves meiosis but this process aborts, rarely forming meiospores (Becnel et al., 1989). The other sporulation sequence involves nuclear dissociation resulting in larval death and the release of large numbers of uninucleate spores that are responsible for horizontal transmission when ingested by larvae. Within 72–96 h postinfection, a small binucleate spore (autoinfective spore) is formed in the gastric caecum as the result of a fourth sporulation sequence (Johnson et al., 1997). These spores germinate intracellularly and serve to spread the parasite within the host to larval oenocytes (autoinfection). In the adult mosquito, the binucleate (transovarial) spores are formed and infect
The effectiveness of the microsporidium Edhazardia aedis (Kudo) to control a seminatural population of Aedes aegypti (L.) was evaluated over a 2-year period. The tests were conducted in a large screened enclosure against an established population of A. aegypti provided caged rabbits as an ad lib. blood supply. In year 1, inoculative release of E. aedis resulted in dispersal of the microsporidium by infected A. aegypti females to all containers within the enclosed study site over a 20-week period. In the second year of the study, inundative release of E. aedis produced high larval and adult infections and successfully eliminated the population of A. aegypti within 11 weeks of introduction. In both years, a deviation from the typical life cycle that produced horizontally infectious spores was critical for persistence of E. aedis within containers. This study has demonstrated that E. aedis is superbly adapted to A. aegypti, having evolved a number of strategies that ensure long-term survival and make it a serious candidate for introduction as a classical biological control agent. Key Words: Aedes aegypti; Edhazardia aedis; biological control; microsporidia; Culicidae; mosquito.
INTRODUCTION
Dengue is the most important human viral disease transmitted by arthropod vectors, with an estimated 50–100 million cases of dengue and 250,000–500,000 cases of dengue hemorrhagic fever annually (RigauPe´rez et al., 1998). Aedes aegypti (L.) is the major vector of dengue in urban tropical and subtropical regions of the world (Gubler and Clark, 1995). Presently, there is no vaccine for dengue and the only effective method to prevent transmission is vector control. Control of A.
1 To whom correspondence should be addressed at USDA/ARS, CMAVE, P.O. Box 14565, Gainesville, Florida 32604. Fax: (904) 374-5966. E-mail: [email protected].
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FIG. 1. The life cycle of Edhazardis aedis in the larval and adult stages of the mosquito Aedes aegypti. Two important deviations are indicated; a vertical–vertical cycle and a horizontal–horizontal cycle.
the oocytes to complete the cycle. Thus, adults disseminate the pathogen, via vertical transmission, to new habitats where horizontal amplification produces additional infected adults (Becnel, 1990). There are two known deviations from this parental host–filial host alternation that may play important roles in maintenance of E. aedis under natural circumstances (Becnel et al., 1989). In these instances, the parasite completes its development through repeated cycles of vertical transmission or, alternatively, by repeated cycles of horizontal transmission. In the vertical–vertical cycle (deviation 1, Fig. 1), some infected progeny from infected adults survive to the adult stage. Binucleate spores are formed in female adults and the infection is transmitted vertically to progeny. In the horizontal–horizontal cycle (deviation 2, Fig. 1), parental generation larvae sometimes fail to pupate and the larval life is extended for up to 2 weeks. The entire developmental sequence occurs in these larvae to pro-
duce horizontally infectious uninucleate spores in fat body. This is the first ecological and epizootiological study conducted with E. aedis against a large contained population of A. aegypti. The optimism regarding the role of E. aedis as a component of a program to control A. aegypti focuses on a number of desirable traits determined in laboratory studies. E. aedis is host specific (Becnel and Johnson, 1993; Andreadis, 1994), safe for nontarget organisms (Becnel, 1992), and has detrimental effects on the reproductive capacity of A. aegypti (Becnel et al., 1995). In addition, the vertical and horizontal components of the life cycle of E. aedis are highly efficient, providing the means for the parasite to become established, persist, and spread in populations of A. aegypti (Hembree, 1982; Hembree and Ryan, 1982; Becnel et al., 1989; Becnel, 1990). This report describes a 2-year study to evaluate E. aedis against A. aegypti under seminatural conditions in
IMPACT OF Edhazardia aedis
Florida. The objectives of the first year were to determine spore survival in enclosure containers and the establishment, persistence, and dissemination of E. aedis in the population after inoculative release. The second year was designed to measure the impact of E. aedis on A. aegypti density after inundative release. MATERIALS AND METHODS
Spore survival/infectivity. A study was undertaken to determine the survival of the horizontally infectious uninucleate spores of E. aedis in enclosure containers held within the screened enclosure described below. Plastic buckets with 15 liters of well water plus 200 g of autoclaved leaves were allowed to steep for 3 days. The infusion was strained through a 230-mesh sieve (63 µm) and 1 liter was placed into each of 33 1.4-liter plastic containers. Water temperature was recorded hourly during the test period. Twenty-two of the containers were contaminated with three decapitated, E. aedis-infected A. aegypti larvae (1.4 ⫻ 106 spores/container) and covered with a screened lid. Three healthy A. aegypti larvae were added to each of the remaining 11 containers as controls. Each day, 2 exposed and 1 control container were randomly selected, into which 20 larvae (48 h old) were placed. Larvae were removed after 4 days, counted, and individually smeared and stained with a 10% Giemsastain solution to determine the percentage of infection. Probit analysis was used to estimate the infective time (IT50 and IT90 ) values for spore survival over the 10 days of the test. Inoculative release. Tests were conducted in an 8 ⫻ 6.5 ⫻ 3 m screened enclosure located in Gainesville, Florida, beginning May 1996. Four rows of golf cart tires (26 total) were positioned on racks 1 m off the ground. One-gallon plastic containers were placed into each tire, to which 10 g autoclaved leaves and 1 liter of well water were added. Seed germination paper strips (19 ⫻ 12.5 cm) lined each container for oviposition. Two rabbits in 1.2 ⫻ 1 ⫻ .5 m cages were located in opposite corners and served as an ad lib. blood source (University of Florida Animal Use Approval No. 9062). Approximately 5500 pupae of A. aegypti (eight generations in the laboratory) were allowed to emerge in the cage. After 4 weeks, larval populations were present in all containers. In June 1996, two containers were randomly selected from each row (eight total). One container from each row was selected to serve as covered controls fitted with screened lids to prevent escape and oviposition; two of these were contaminated with six decapitated A. aegypti larvae infected with E. aedis (a dose of 2.2 ⫻ 106 spores) and two with six decapitated healthy A. aegypti. The other selected container from each row (four total) was contaminated with six decapitated infected A. aegypti larvae and left uncovered to
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serve as the foci for production of infected adults. Pupae were removed from the covered containers daily, set up individually in vials for emergence, and held for 48 h. The sex was recorded and the infection status of each adult that survived 48 h was determined by microscopic examination for the presence of spores. All dead individuals were examined for infection. Sampling of open containers began 2 weeks postinoculation. Egg papers were collected biweekly from each container and cut in half. One half was returned to the container and flooded. Eggs on the other half were hatched in the laboratory, and the larvae were reared to the fourth instar and examined for vertical infection. Absolute counts of larvae and pupae from each container were determined biweekly starting on week 13. For each sample, patently infected larvae were examined with a dissecting microscope to determine whether infection was as a result of vertical transmission or horizontally induced (Becnel et al., 1989). Containers and all of their contents were returned to the enclosure immediately after examination. Water was added to containers biweekly to maintain the original level. Differences of the means between control and exposed groups were compared using Student’s t test. Inundative release. Tests were conducted in the same 8 ⫻ 6.5 ⫻ 3 m screened enclosure as those in year 1 (inoculative release). Containers from the previous year’s study (as described above) were left in place over the winter and flooded in April 1997 to initiate the A. aegypti population. Ovipositional data (half egg sheets) were collected beginning April 18 with a total of seven samples through July 3. As of April 30, A. aegypti larvae were present in all containers. The setup was the same as that for year 1 (inoculative release). Containers were treated on July 17, 1997, as follows: One container was randomly selected from each row to serve as covered controls (four total). Two of these were contaminated with 25 infected A. aegypti larval equivalents (3.2 ⫻ 107 spores) and two with 25 healthy A. aegypti larval equivalents. All open containers were contaminated with 25 infected A. aegypti larval equivalents and left uncovered. Pupae were removed from the covered containers daily, set up individually in vials for emergence, and held for 48 h. The infection status of each adult was determined by the presence of spores. Egg papers were collected weekly from the open containers and cut in half; one half was returned to the container and submerged, the other half hatched in the laboratory, reared to the fourth instar, and examined for vertical infection. Absolute counts of immatures in each container were determined weekly and the number of patent infections (either vertically or horizontally induced) recorded. Containers and all of their contents were returned to the enclosure immediately after examination. Differences of the means between control and exposed groups were compared using Student’s t test.
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FIG. 2.
The effect of enclosure conditions over time on the infectivity of uninucleate spores of E. aedis for A. aegypti. Values are means ⫾ SE.
RESULTS
Spore survival/infectivity. During the test period, water temperatures averaged 19.5 ⫾ 1.2°C in October 1995 (inoculative release) and 26.4 ⫾ 0.4°C in May 1996 (inundative release) (mean ⫾ SE). The two replicates of the spore survival test were not significantly different (t ⫽ 0.48, df ⫽ 10, P ⫽ 0.64) and therefore the data were combined. Simple linear regression (Fig. 2) revealed that time had a significant negative effect on spore longevity (df ⫽ 10, r2 ⫽ 0.858, y ⫽ 10.9 ⫹ 102.2). The IT90 value was 1.8 days and the IT50 value was 4.2 days (95% confidence limits). This indicates that spores introduced into enclosure containers can be expected to be highly infectious for the first 2 days but that spore survival falls rapidly and infectivity was reduced by 50% in less than 5 days. Inoculative release. The average daily emergence of adults from the closed containers was significantly greater for the unexposed group than the exposed group (t ⫽ 3.4; df ⫽ 24; P ⬍ 0.002). An average of 278.0 adults emerged from each of the two unexposed closed
containers with a 48-h survival rate of 99.3% (Table 1). In each of the two closed containers exposed to E. aedis, an average of 167.5 adults emerged, with a 48-h survival rate of 72.0%. Infection levels for adults with E. aedis in closed containers were slightly lower in males (72.6%) than in females (78.0%). The emergence pattern of healthy and infected female adults from the covered containers was cyclic over the 25-day period (Fig. 3). The number of uninfected females emerging from containers was significantly higher than that of infected females (t ⫽ 3.8; df ⫽ 18; P ⬍ 0.001). The first infected females emerged 4 days postexposure followed by four emergence peaks occurring at approximately 1-week intervals. E. aedis spread via vertical transmission from the original 4 inoculated open containers to all containers in the study site within 20 weeks postintroduction. Larvae from eggs collected from the enclosure and hatched in the laboratory confirmed oviposition by infected females 2 weeks after introduction in 4 of 22 open containers (18%) and again on weeks 4 and 6 but
TABLE 1 Comparative Emergence (Inoculative Release) of Healthy (Unexposed) and Edhazardia aedis-Infected Aedes aegypti Adults from Covered Containers Percentage of infection
Unexposed Exposed a
Mean ⫾ SE.
No. females emerged/container a
No. males emerged/container a
Percentage of adult survival (combined) a
Females a
Males a
98.5 ⫾ 4.1 71.5 ⫾ 5.3
179.5 ⫾ 5.8 96.0 ⫾ 7.3
99.3 ⫾ 0.34 72.0 ⫾ 0.10
0 78.0 ⫾ 9.6
0 72.6 ⫾ 3.6
IMPACT OF Edhazardia aedis
FIG. 3.
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Emergence patterns (inoculative release) of healthy and E. aedis-infected A. aegypti females from covered containers.
at very low levels (⬍0.6% vertical infections, Fig. 4). By week 6, a cumulative total of 11 of 22 containers (50%) were positive for infected eggs. Overall oviposition decreased to less than 50 larvae/half egg sheet on weeks 8 and 10 and infected progeny were not present in any of those samples. Infected progeny were again present on week 12 (Fig. 4), with 10 of 22 containers
positive and vertical infection levels had increased substantially to 8.0 ⫾ 5.1% of the larvae examined. Infection levels remained relatively high throughout the remainder of the test, with maximum rates on weeks 18 (12.2 ⫾ 8.6%) and 24 (18.6 ⫾ 14.6%) at a time when overall oviposition was declining (Fig. 4). By week 20, all containers were positive for infected eggs
FIG. 4. Ovipositional activity (inoculative release) of A. aegypti females based on larvae from enclosure-collected egg sheets hatched in the laboratory. Percentages of infection with vertically transmitted fat body infections (mean ⫾ SE) are indicated on the graph.
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at least once during the sampling period. Vertical infection levels from individual egg sheets ranged from 0.3 to 100.0% over the course of the study. Larval densities in containers remained relatively constant for weeks 12–18, attaining their highest level on week 16 at 231.5 larvae/container (Fig. 5). A steady reduction in larval densities was observed for the remainder of the test as Fall approached and temperatures decreased (Fig. 5). Examination of larvae from the open containers (weeks 12–26) revealed patently infected individuals on each sampling date (Fig. 6). These patently infected larvae were produced by two different pathways. One pathway was via vertical transmission from infected females to progeny. The other was the horizontal–horizontal cycle (deviation 2, Fig. 1), where horizontally infected larvae fail to pupate and eventually produce fat body infections with uninucleate spores. Larvae with horizontally induced fat body infections were present in containers throughout the sampling period at substantial levels peaking at 18.1% on week 18 (Fig. 6). Vertically infected larvae were also present in containers throughout the sampling period, with peak infections on weeks 14 (8.3%), 20 (7.9%), and 26 at 8.5% (Fig. 6). Inundative release. E. aedis did not become reestablished in the A. aegypti population between years 1 and 2 of this study. The first ovipositional data after the winter of 1996–1997 (April 18) produced 1 container of
21 that contained one patently infected larva (n ⫽ 1500) as a result of vertical transmission. Subsequent ovipositional data (six samples, April 30–July 3) did not produce larvae infected with E. aedis. The average daily emergence of adults from the closed containers was significantly greater for the unexposed group than for the exposed group (t ⫽ 3.7; df ⫽ 12; P ⬍ 0.003). Unexposed covered containers (combined) had an average of 85.5 adults emerge, of which 47.0 were female (Table 2). Exposed covered containers (combined) had an average of 10.5 adults emerge, of which 5.0 were female. This represents a reduction in adult production of approximately 78.1%. Seventy percent of the female adults and 32.1% of the male adults that emerged from the exposed containers were infected with E. aedis (Table 2). On July 16, there was an average of 100.7 ⫾ 10.7 larvae/egg sheet collected from the open containers (Fig. 7). This increased to 135.8 ⫾ 15.9 larvae/egg sheet on week 2 but declined dramatically thereafter, with all oviposition activity ceasing by week 11 (Fig. 7). Vertically infected larvae were first detected 2 weeks postexposure but at low levels (2 of 18 containers, 0.4 ⫾ 0.3% infection). Vertical infections were not found on weeks 3 and 4 but infection levels dramatically increased to 73.6 ⫾ 12.9% on week 5 and 34.6 ⫾ 10.1% on week 6 (Fig. 7). Eggs were not collected from any of the containers on weeks 7 and 8 and eggs were laid in only
FIG. 5. Biweekly A. aegypti larval densities (inoculative release) in open containers and the average water temperature for weeks 12–26 (means ⫾ SE).
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IMPACT OF Edhazardia aedis
FIG. 6. Biweekly percentages of E. aedis infection (inoculative release) in larval A. aegypti from open containers (means ⫾ SE). Patent infections are a result of vertical transmission from infected adults or the horizontal–horizontal cycle.
one container on week 9 that produced 24 larvae with 100% infection. Temperature (mean ⫾ SE) for the study period (July 16–October 9) was 24.5 ⫾ 0.26°C (range ⫽ 20.6– 31.1°C). Larval density on July 16 in open containers averaged 210.8 larvae/container (Fig. 8). Larval densities remained high for the following 3 weeks but declined dramatically thereafter, with no larvae present in any open containers on week 11 (Fig. 8). Patently infected larvae in the open containers were produced by two different pathways (horizontal– horizontal and vertical) as described above. The horizontally induced patent infections were the predominant type with substantial infection levels (11.1–61.1%) for the first 7 weeks of the test (Fig. 8). Vertically infected larvae did not appear in appreciable numbers until weeks 5, 7, and 8, with 7.3, 12.1, and 6.3% infection rates, respectively (Fig. 8). Combined infection rates (horizontal plus vertical) varied from a low of 24.5% on
week 2 to a high of 73.2% on week 7. There were very few larvae alive in containers after week 7. One container had six larvae on week 10, all of which were vertically infected. The adult and larval populations were completely eliminated by week 11 (Figs. 7 and 8). DISCUSSION
In this study, we examined two methods (inoculative and inundative release) for the introduction of E. aedis into a contained population of A. aegypti. Inoculative release exploits vertical transmission of E. aedis to spread the parasite to new, often cryptic, habitats. Inundative release severely impacts both larvae and adults and exploits the persistence of the parasite within a container via horizontal transmission, with two possible outcomes. One of these involves benign infections in larvae to produce infected adults that distribute the parasite to new habitats; the other is the
TABLE 2 Comparative Emergence (Inundative Release) of Healthy (Unexposed) and E. aedis-Infected A. aegypti Adults from Covered Containers Percentage of infection
Unexposed Exposed a
Mean ⫾ SE.
No. females emerged/container a
No. males emerged/container a
Percentage of adult survival (combined) a
Females a
Males a
47.0 ⫾ 7.2 5.0 ⫾ 0.0
38.5 ⫾ 4.3 5.5 ⫾ 0.9
90.8 ⫾ 4.4 44.4 ⫾ 2.0
0 70.0 ⫾ 10.0
0 32.1 ⫾ 17.9
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FIG. 7. Ovipositional activity (inundative release) of A. aegypti females based on larvae from enclosure collected egg sheets hatched in the laboratory. Percentages of infection with vertically transmitted fat body infections (mean ⫾ SE) are indicated on the graph.
continued production of uninucleate spores in infected larvae to maintain an infectious titer of spores within containers (deviation 2, Fig. 1). This is particularly important given the finding that spores released into the aquatic environment are relatively short-lived. Under ideal conditions of storage, purified uninucleate spores of E. aedis maintain infectivity for up to 30 days
at between 20 and 30°C (Undeen et al., 1993). However, exposure of the extracorporeal spores of E. aedis to the biotic and abiotic factors found in the aquatic environment reduced infectivity by 10% after 2 days and by 50% within 5 days. Inoculative release resulted in dispersal of E. aedis by infected A. aegypti females to 18% (4 of 22) of the
FIG. 8. Weekly A. aegypti larval densities (inundative release) and percentages of Edhazardia aedis infection in larval A. aegypti from open containers (means ⫾ SE). Patent infections are a result of vertical transmission from infected adults or the horizontal–horizontal cycle.
IMPACT OF Edhazardia aedis
containers in 2 weeks, 50% of the containers in 6 weeks, and 100% of containers in 20 weeks. Infected females deposited eggs with E. aedis during the first 2 weeks postexposure but infection levels were low and remained low through week 10. Infection levels increased dramatically on week 12 and initiated the first of four cycles of ovipositional activity by infected females. These cycles may reflect the emergence pattern of adults that is also cyclic as well as the previous finding that most infected adults would only survive through one gonotrophic cycle (Becnel et al., 1995). Vertical infection levels (based on ovipositional data) peaked on weeks 12 (8%), 18 (12%), and 24 (19%), which corresponded (allowing for a 2-week developmental time) to the maximum rates of vertically infected larvae observed in containers on weeks 14, 20, and 26. These are substantial infection levels that provided inoculum (spores) for horizontal transmission to individuals in containers, as demonstrated by the observed horizontal infection levels in larvae. However, overall horizontal levels are higher than indicated by the patent horizontal infection levels, which do not reflect the benign infections in larvae that produce infected adults. An interesting feature of the infected larvae within the containers was the presence of horizontally induced patent infections throughout the sampling period. This deviation (type 2) from the typical life cycle (Becnel et al., 1989) assured the continued presence of spores within containers that are infectious orally for mosquito larvae. This was an unexpected finding but is apparently important for the long-term persistence of the pathogen within a container without the need for continued input of vertically infected individuals. The survival strategy of E. aedis therefore involves original contamination of a container by vertical transmission to establish the infection followed by continued horizontal transmission to both maintain the infection in the container and produce infected adults to disseminate the pathogen to other containers. A. aegypti survived the winter between years 1 and 2 of this study, but E. aedis did not survive in sufficient numbers to become reestablished. The uninucleate spores of E. aedis are extremely cold sensitive, with infectivity for A. aegypti larvae completely lost after storage at 5°C for 24 h (Undeen et al., 1993). It is unlikely that uninucleate spores survived the winter temperatures at the test location, which included a number of hard freezes. Developmental stages of the microsporidium in eggs and larvae are less sensitive to cold (Undeen et al., 1993), and it is possible that E. aedis could have survived within either infected eggs or adults. Nevertheless, the detrimental effect of cold on E. aedis indicates that introduction against A. aegypti populations near the temperature limits of its range would require periodic augmentation. Inundative release of E. aedis produced high larval
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and adult infections and successfully eliminated a contained population of A. aegypti within 11 weeks of introduction. For the first 3 weeks of the study, the population slightly increased, probably due to the large numbers of healthy adults present at the time of introduction. This was followed by a dramatic decline in the larval and adult populations. The decline coincided with 74% vertical infection rates in progeny of female adults and 46% patent infection rates in larvae on week 5. With infection levels of this magnitude, the population was unable to recover and all activity ceased by week 11. As in the inoculative release, horizontal transmission played an important role in maintenance of the parasite within containers throughout the duration of the study. It is likely that the rapid elimination of A. aegypti in this test was due, in part, to the isolation of the population, preventing both immigration of healthy adults and emigration of infected adults. Nevertheless, the results demonstrate that the inundative introduction of E. aedis can severely impact both larval and adult populations. The intensity and duration of this impact can only be determined by careful evaluation of controlled field releases with E. aedis into unrestricted populations of A. aegypti. Andreadis (1989) was the first to successfully introduce a polymorphic microsporidium, Amblyospora connecticus Andreadis, into a larval field population of the salt marsh mosquito Aedes cantator Coquillett. This was accomplished via the release of the infected copepod host and resulted in infection rates ranging from 16 to 24%. Because E. aedis does not require an intermediate host for horizontal transmission, introduction is greatly simplified and long-term persistence does not depend on the presence of two hosts. For successful introduction as a classical biological control agent, E. aedis must be able to persist and spread within a population of A. aegypti and regulate mosquito abundance to levels that would prevent disease transmission. Based on this study, the most immediate impact using E. aedis would be realized via an inundative release to challenge a large portion of an A. aegypti population. This would not only effectively suppress the exposed population but also establish a presence in containers to suppress new input to the site as well as assure that adults produced would harbor the pathogen. This would serve to spread the pathogen to other sites and limit the longevity of adults (Becnel et al., 1995), thereby reducing the likelihood of more than one blood meal and also reducing the risk of disease transmission. In conclusion, we believe that based on the results of this and previous studies, E. aedis should be considered a serious candidate for introduction as a classical biological control agent against A. aegypti. E. aedis is a
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parasite superbly adapted to A. aegypti, having evolved a number of strategies to ensure long-term survival. Two of these strategies involve the highly efficient mechanisms of vertical and horizontal transmission. Vertical infection spreads the pathogen to new, often cryptic, habitats and horizontal transmission amplifies the infection by producing additional infected adults and larvae. Use of E. aedis for management of A. aegypti is proposed not as a sole method but rather as part of the natural complex of regulatory factors utilized for classical biological control. This approach recognizes that eradication of the target mosquito is an unrealistic expectation but with a combination of physical, cultural, chemical, and biological control methods, populations of A. aegypti can regulated. ACKNOWLEDGMENTS The authors thank Charles S. Apperson, Theodore G. Andreadis, and Wayne M. Brooks for critically reviewing the manuscript.
REFERENCES Andreadis, T. G. 1989. Infection of a field population of Aedes cantator with a polymorphic microsporidium, Amblyospora connecticus via release of the intermediate copepod host, Acanthocyclops vernalis. J. Am. Mosquito Control Assoc. 5, 81–85. Andreadis, T. G. 1994. Host range tests with Edhazardia aedis (Microsporida: Culicosporidae) against northern Nearctic mosquitoes. J. Invertebr. Pathol. 64, 46–51. Becnel, J. J. 1990. ‘‘Edhazardia aedis (Microsporida: Amblyosporidae) as a Biocontrol Agent of Aedes aegypti (Diptera: Culicidae).’’ Proc. Int. Colloq. Invertebr. Pathol. Microb. Control, Adelaide, Australia, Vol. 5, pp. 56–60. Becnel, J. J. 1992. Safety of Edhazardia aedis (Microspora, Amblyosporidae) for nontarget aquatic organisms. J. Am. Mosquito Control Assoc. 8, 256–260.
Becnel, J. J., and Andreadis, T. G. 1999. Microsporidia in insects. In ‘‘The Microsporidia and Microsporidiosis’’ (M. Wittner, Ed.), pp. 447–501. Am. Soc. for Microbiol., Washington, DC. Becnel, J. J., and Johnson, M. A. 1993. Mosquito host range and specificity of Edhazardia aedis (Microspora: Culicosporidae). J. Am. Mosquito Control Assoc. 9, 269–274. Becnel, J. J., Sprague, V., Fukuda, T., and Hazard, E. I. 1989. Development of Edhazardia aedis (Kudo, 1930) N. G., N. Comb. (Microsporida: Amblyosporidae) in the mosquito Aedes aegypti (L.) (Diptera: Culicidae). J. Protozool. 36, 119–130. Becnel, J. J., Garcia, J. J., and Johnson, M. A. 1995. Edhazardia aedis (Microspora: Culicosporidae) effects on the reproductive capacity of Aedes aegypti (Diptera: Culicidae). J. Med. Entomol. 32, 549–553. Gubler, D. J., and Clark, G. G. 1995. Dengue/dengue hemorrhagic fever: The emergence of a global health problem. Emerg. Infect. Dis. 1, 55–57. Hembree, S. C. 1979. Preliminary report of some mosquito pathogens from Thailand. Mosquito News 39, 575–582. Hembree, S. C. 1982. Dose-response studies of a new species of per os and vertically transmittable microsporidian pathogens of Aedes aegypti from Thailand. Mosquito News 42, 55–61. Hembree, S. C., and Ryan, J. R. 1982. Observations on the vertical transmission of a new microsporidian pathogen of Aedes aegypti from Thailand. Mosquito News 42, 49–54. Johnson, M. A., Becnel, J. J., and Undeen, A. H. 1997. A new sporulation sequence in Edhazardia aedis (Microsporidia: Culicosporidae), a parasite of the mosquito Aedes aegypti (Diptera: Culicidae). J. Invertebr. Pathol. 70, 69–75. Kudo, R. 1930. Studies on microsporidia parasitic in mosquitoes. VIII. On a microsporidian, Nosema aedis nov. spec., parasitic in a larva of Aedes aegypti of Puerto Rico. Arch. Protistenkd. 69, 23–28. Rigau-Pe´rez, J. G., Clark, G. G., Gubler, D. J., Reiter, P., Sanders, E. J., and Vorndam, A. V. 1998. Dengue and dengue haemorrhagic fever. Lancet 352, 971–977. Undeen, A. H., Johnson, M. A., and Becnel, J. J. 1993. The effects of temperature on the survival of Edhazardia aedis (Microspora, Amblyosporidae), a pathogen of Aedes aegypti. J. Invertebr. Pathol. 61, 303–307.
Impact of Edhazardia aedis (Microsporidia: Culicosporidae) on a Seminatural Population of Aedes aegypti (Diptera: Culicidae) James J. Becnel1 and Margaret A. Johnson U.S. Department of Agriculture, Agricultural Research Service, Center for Medical, Agricultural, and Veterinary Entomology, Gainesville, Florida 32604 E-mail: [email protected] Received June 15, 1999, accepted November 28, 1999
aegypti has relied on source reduction and broad spectrum chemical larvicides and adulticides. Both methods can be effective but it is difficult to sustain longterm source reduction efforts. The development of resistance to insecticides by mosquitoes and the need for environmental conservation also indicate a necessity for alternative control strategies. Edhazardia aedis (Kudo) Becnel, Sprague, and Fukuda is one of the few pathogens that has been isolated from A. aegypti. This microsporidium was originally observed and described from Puerto Rico (Kudo, 1930) and reisolated in Thailand (Hembree, 1979). E. aedis has a complex life cycle involving both horizontal and vertical transmission, with four different spore types produced in the mosquito host (Becnel et al., 1989; Johnson et al., 1997). Unlike some other polymorphic mosquito-pathogenic microsporidia, there is not an obligate intermediate host involved in the life cycle (Becnel and Andreadis, 1999). A schematic representation of the basic life cycle of E. aedis is shown in Fig. 1. One sporulation sequence occurs in the adult female (infected orally as a larva) and results in the formation of binucleate (transovarial) spores. These spores are involved in vertical transmission of E. aedis to the subsequent generation via infected eggs. In infected progeny, there are two sporulation sequences in larval fat body. One sporulation sequence involves meiosis but this process aborts, rarely forming meiospores (Becnel et al., 1989). The other sporulation sequence involves nuclear dissociation resulting in larval death and the release of large numbers of uninucleate spores that are responsible for horizontal transmission when ingested by larvae. Within 72–96 h postinfection, a small binucleate spore (autoinfective spore) is formed in the gastric caecum as the result of a fourth sporulation sequence (Johnson et al., 1997). These spores germinate intracellularly and serve to spread the parasite within the host to larval oenocytes (autoinfection). In the adult mosquito, the binucleate (transovarial) spores are formed and infect
The effectiveness of the microsporidium Edhazardia aedis (Kudo) to control a seminatural population of Aedes aegypti (L.) was evaluated over a 2-year period. The tests were conducted in a large screened enclosure against an established population of A. aegypti provided caged rabbits as an ad lib. blood supply. In year 1, inoculative release of E. aedis resulted in dispersal of the microsporidium by infected A. aegypti females to all containers within the enclosed study site over a 20-week period. In the second year of the study, inundative release of E. aedis produced high larval and adult infections and successfully eliminated the population of A. aegypti within 11 weeks of introduction. In both years, a deviation from the typical life cycle that produced horizontally infectious spores was critical for persistence of E. aedis within containers. This study has demonstrated that E. aedis is superbly adapted to A. aegypti, having evolved a number of strategies that ensure long-term survival and make it a serious candidate for introduction as a classical biological control agent. Key Words: Aedes aegypti; Edhazardia aedis; biological control; microsporidia; Culicidae; mosquito.
INTRODUCTION
Dengue is the most important human viral disease transmitted by arthropod vectors, with an estimated 50–100 million cases of dengue and 250,000–500,000 cases of dengue hemorrhagic fever annually (RigauPe´rez et al., 1998). Aedes aegypti (L.) is the major vector of dengue in urban tropical and subtropical regions of the world (Gubler and Clark, 1995). Presently, there is no vaccine for dengue and the only effective method to prevent transmission is vector control. Control of A.
1 To whom correspondence should be addressed at USDA/ARS, CMAVE, P.O. Box 14565, Gainesville, Florida 32604. Fax: (904) 374-5966. E-mail: [email protected].
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FIG. 1. The life cycle of Edhazardis aedis in the larval and adult stages of the mosquito Aedes aegypti. Two important deviations are indicated; a vertical–vertical cycle and a horizontal–horizontal cycle.
the oocytes to complete the cycle. Thus, adults disseminate the pathogen, via vertical transmission, to new habitats where horizontal amplification produces additional infected adults (Becnel, 1990). There are two known deviations from this parental host–filial host alternation that may play important roles in maintenance of E. aedis under natural circumstances (Becnel et al., 1989). In these instances, the parasite completes its development through repeated cycles of vertical transmission or, alternatively, by repeated cycles of horizontal transmission. In the vertical–vertical cycle (deviation 1, Fig. 1), some infected progeny from infected adults survive to the adult stage. Binucleate spores are formed in female adults and the infection is transmitted vertically to progeny. In the horizontal–horizontal cycle (deviation 2, Fig. 1), parental generation larvae sometimes fail to pupate and the larval life is extended for up to 2 weeks. The entire developmental sequence occurs in these larvae to pro-
duce horizontally infectious uninucleate spores in fat body. This is the first ecological and epizootiological study conducted with E. aedis against a large contained population of A. aegypti. The optimism regarding the role of E. aedis as a component of a program to control A. aegypti focuses on a number of desirable traits determined in laboratory studies. E. aedis is host specific (Becnel and Johnson, 1993; Andreadis, 1994), safe for nontarget organisms (Becnel, 1992), and has detrimental effects on the reproductive capacity of A. aegypti (Becnel et al., 1995). In addition, the vertical and horizontal components of the life cycle of E. aedis are highly efficient, providing the means for the parasite to become established, persist, and spread in populations of A. aegypti (Hembree, 1982; Hembree and Ryan, 1982; Becnel et al., 1989; Becnel, 1990). This report describes a 2-year study to evaluate E. aedis against A. aegypti under seminatural conditions in
IMPACT OF Edhazardia aedis
Florida. The objectives of the first year were to determine spore survival in enclosure containers and the establishment, persistence, and dissemination of E. aedis in the population after inoculative release. The second year was designed to measure the impact of E. aedis on A. aegypti density after inundative release. MATERIALS AND METHODS
Spore survival/infectivity. A study was undertaken to determine the survival of the horizontally infectious uninucleate spores of E. aedis in enclosure containers held within the screened enclosure described below. Plastic buckets with 15 liters of well water plus 200 g of autoclaved leaves were allowed to steep for 3 days. The infusion was strained through a 230-mesh sieve (63 µm) and 1 liter was placed into each of 33 1.4-liter plastic containers. Water temperature was recorded hourly during the test period. Twenty-two of the containers were contaminated with three decapitated, E. aedis-infected A. aegypti larvae (1.4 ⫻ 106 spores/container) and covered with a screened lid. Three healthy A. aegypti larvae were added to each of the remaining 11 containers as controls. Each day, 2 exposed and 1 control container were randomly selected, into which 20 larvae (48 h old) were placed. Larvae were removed after 4 days, counted, and individually smeared and stained with a 10% Giemsastain solution to determine the percentage of infection. Probit analysis was used to estimate the infective time (IT50 and IT90 ) values for spore survival over the 10 days of the test. Inoculative release. Tests were conducted in an 8 ⫻ 6.5 ⫻ 3 m screened enclosure located in Gainesville, Florida, beginning May 1996. Four rows of golf cart tires (26 total) were positioned on racks 1 m off the ground. One-gallon plastic containers were placed into each tire, to which 10 g autoclaved leaves and 1 liter of well water were added. Seed germination paper strips (19 ⫻ 12.5 cm) lined each container for oviposition. Two rabbits in 1.2 ⫻ 1 ⫻ .5 m cages were located in opposite corners and served as an ad lib. blood source (University of Florida Animal Use Approval No. 9062). Approximately 5500 pupae of A. aegypti (eight generations in the laboratory) were allowed to emerge in the cage. After 4 weeks, larval populations were present in all containers. In June 1996, two containers were randomly selected from each row (eight total). One container from each row was selected to serve as covered controls fitted with screened lids to prevent escape and oviposition; two of these were contaminated with six decapitated A. aegypti larvae infected with E. aedis (a dose of 2.2 ⫻ 106 spores) and two with six decapitated healthy A. aegypti. The other selected container from each row (four total) was contaminated with six decapitated infected A. aegypti larvae and left uncovered to
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serve as the foci for production of infected adults. Pupae were removed from the covered containers daily, set up individually in vials for emergence, and held for 48 h. The sex was recorded and the infection status of each adult that survived 48 h was determined by microscopic examination for the presence of spores. All dead individuals were examined for infection. Sampling of open containers began 2 weeks postinoculation. Egg papers were collected biweekly from each container and cut in half. One half was returned to the container and flooded. Eggs on the other half were hatched in the laboratory, and the larvae were reared to the fourth instar and examined for vertical infection. Absolute counts of larvae and pupae from each container were determined biweekly starting on week 13. For each sample, patently infected larvae were examined with a dissecting microscope to determine whether infection was as a result of vertical transmission or horizontally induced (Becnel et al., 1989). Containers and all of their contents were returned to the enclosure immediately after examination. Water was added to containers biweekly to maintain the original level. Differences of the means between control and exposed groups were compared using Student’s t test. Inundative release. Tests were conducted in the same 8 ⫻ 6.5 ⫻ 3 m screened enclosure as those in year 1 (inoculative release). Containers from the previous year’s study (as described above) were left in place over the winter and flooded in April 1997 to initiate the A. aegypti population. Ovipositional data (half egg sheets) were collected beginning April 18 with a total of seven samples through July 3. As of April 30, A. aegypti larvae were present in all containers. The setup was the same as that for year 1 (inoculative release). Containers were treated on July 17, 1997, as follows: One container was randomly selected from each row to serve as covered controls (four total). Two of these were contaminated with 25 infected A. aegypti larval equivalents (3.2 ⫻ 107 spores) and two with 25 healthy A. aegypti larval equivalents. All open containers were contaminated with 25 infected A. aegypti larval equivalents and left uncovered. Pupae were removed from the covered containers daily, set up individually in vials for emergence, and held for 48 h. The infection status of each adult was determined by the presence of spores. Egg papers were collected weekly from the open containers and cut in half; one half was returned to the container and submerged, the other half hatched in the laboratory, reared to the fourth instar, and examined for vertical infection. Absolute counts of immatures in each container were determined weekly and the number of patent infections (either vertically or horizontally induced) recorded. Containers and all of their contents were returned to the enclosure immediately after examination. Differences of the means between control and exposed groups were compared using Student’s t test.
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FIG. 2.
The effect of enclosure conditions over time on the infectivity of uninucleate spores of E. aedis for A. aegypti. Values are means ⫾ SE.
RESULTS
Spore survival/infectivity. During the test period, water temperatures averaged 19.5 ⫾ 1.2°C in October 1995 (inoculative release) and 26.4 ⫾ 0.4°C in May 1996 (inundative release) (mean ⫾ SE). The two replicates of the spore survival test were not significantly different (t ⫽ 0.48, df ⫽ 10, P ⫽ 0.64) and therefore the data were combined. Simple linear regression (Fig. 2) revealed that time had a significant negative effect on spore longevity (df ⫽ 10, r2 ⫽ 0.858, y ⫽ 10.9 ⫹ 102.2). The IT90 value was 1.8 days and the IT50 value was 4.2 days (95% confidence limits). This indicates that spores introduced into enclosure containers can be expected to be highly infectious for the first 2 days but that spore survival falls rapidly and infectivity was reduced by 50% in less than 5 days. Inoculative release. The average daily emergence of adults from the closed containers was significantly greater for the unexposed group than the exposed group (t ⫽ 3.4; df ⫽ 24; P ⬍ 0.002). An average of 278.0 adults emerged from each of the two unexposed closed
containers with a 48-h survival rate of 99.3% (Table 1). In each of the two closed containers exposed to E. aedis, an average of 167.5 adults emerged, with a 48-h survival rate of 72.0%. Infection levels for adults with E. aedis in closed containers were slightly lower in males (72.6%) than in females (78.0%). The emergence pattern of healthy and infected female adults from the covered containers was cyclic over the 25-day period (Fig. 3). The number of uninfected females emerging from containers was significantly higher than that of infected females (t ⫽ 3.8; df ⫽ 18; P ⬍ 0.001). The first infected females emerged 4 days postexposure followed by four emergence peaks occurring at approximately 1-week intervals. E. aedis spread via vertical transmission from the original 4 inoculated open containers to all containers in the study site within 20 weeks postintroduction. Larvae from eggs collected from the enclosure and hatched in the laboratory confirmed oviposition by infected females 2 weeks after introduction in 4 of 22 open containers (18%) and again on weeks 4 and 6 but
TABLE 1 Comparative Emergence (Inoculative Release) of Healthy (Unexposed) and Edhazardia aedis-Infected Aedes aegypti Adults from Covered Containers Percentage of infection
Unexposed Exposed a
Mean ⫾ SE.
No. females emerged/container a
No. males emerged/container a
Percentage of adult survival (combined) a
Females a
Males a
98.5 ⫾ 4.1 71.5 ⫾ 5.3
179.5 ⫾ 5.8 96.0 ⫾ 7.3
99.3 ⫾ 0.34 72.0 ⫾ 0.10
0 78.0 ⫾ 9.6
0 72.6 ⫾ 3.6
IMPACT OF Edhazardia aedis
FIG. 3.
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Emergence patterns (inoculative release) of healthy and E. aedis-infected A. aegypti females from covered containers.
at very low levels (⬍0.6% vertical infections, Fig. 4). By week 6, a cumulative total of 11 of 22 containers (50%) were positive for infected eggs. Overall oviposition decreased to less than 50 larvae/half egg sheet on weeks 8 and 10 and infected progeny were not present in any of those samples. Infected progeny were again present on week 12 (Fig. 4), with 10 of 22 containers
positive and vertical infection levels had increased substantially to 8.0 ⫾ 5.1% of the larvae examined. Infection levels remained relatively high throughout the remainder of the test, with maximum rates on weeks 18 (12.2 ⫾ 8.6%) and 24 (18.6 ⫾ 14.6%) at a time when overall oviposition was declining (Fig. 4). By week 20, all containers were positive for infected eggs
FIG. 4. Ovipositional activity (inoculative release) of A. aegypti females based on larvae from enclosure-collected egg sheets hatched in the laboratory. Percentages of infection with vertically transmitted fat body infections (mean ⫾ SE) are indicated on the graph.
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at least once during the sampling period. Vertical infection levels from individual egg sheets ranged from 0.3 to 100.0% over the course of the study. Larval densities in containers remained relatively constant for weeks 12–18, attaining their highest level on week 16 at 231.5 larvae/container (Fig. 5). A steady reduction in larval densities was observed for the remainder of the test as Fall approached and temperatures decreased (Fig. 5). Examination of larvae from the open containers (weeks 12–26) revealed patently infected individuals on each sampling date (Fig. 6). These patently infected larvae were produced by two different pathways. One pathway was via vertical transmission from infected females to progeny. The other was the horizontal–horizontal cycle (deviation 2, Fig. 1), where horizontally infected larvae fail to pupate and eventually produce fat body infections with uninucleate spores. Larvae with horizontally induced fat body infections were present in containers throughout the sampling period at substantial levels peaking at 18.1% on week 18 (Fig. 6). Vertically infected larvae were also present in containers throughout the sampling period, with peak infections on weeks 14 (8.3%), 20 (7.9%), and 26 at 8.5% (Fig. 6). Inundative release. E. aedis did not become reestablished in the A. aegypti population between years 1 and 2 of this study. The first ovipositional data after the winter of 1996–1997 (April 18) produced 1 container of
21 that contained one patently infected larva (n ⫽ 1500) as a result of vertical transmission. Subsequent ovipositional data (six samples, April 30–July 3) did not produce larvae infected with E. aedis. The average daily emergence of adults from the closed containers was significantly greater for the unexposed group than for the exposed group (t ⫽ 3.7; df ⫽ 12; P ⬍ 0.003). Unexposed covered containers (combined) had an average of 85.5 adults emerge, of which 47.0 were female (Table 2). Exposed covered containers (combined) had an average of 10.5 adults emerge, of which 5.0 were female. This represents a reduction in adult production of approximately 78.1%. Seventy percent of the female adults and 32.1% of the male adults that emerged from the exposed containers were infected with E. aedis (Table 2). On July 16, there was an average of 100.7 ⫾ 10.7 larvae/egg sheet collected from the open containers (Fig. 7). This increased to 135.8 ⫾ 15.9 larvae/egg sheet on week 2 but declined dramatically thereafter, with all oviposition activity ceasing by week 11 (Fig. 7). Vertically infected larvae were first detected 2 weeks postexposure but at low levels (2 of 18 containers, 0.4 ⫾ 0.3% infection). Vertical infections were not found on weeks 3 and 4 but infection levels dramatically increased to 73.6 ⫾ 12.9% on week 5 and 34.6 ⫾ 10.1% on week 6 (Fig. 7). Eggs were not collected from any of the containers on weeks 7 and 8 and eggs were laid in only
FIG. 5. Biweekly A. aegypti larval densities (inoculative release) in open containers and the average water temperature for weeks 12–26 (means ⫾ SE).
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IMPACT OF Edhazardia aedis
FIG. 6. Biweekly percentages of E. aedis infection (inoculative release) in larval A. aegypti from open containers (means ⫾ SE). Patent infections are a result of vertical transmission from infected adults or the horizontal–horizontal cycle.
one container on week 9 that produced 24 larvae with 100% infection. Temperature (mean ⫾ SE) for the study period (July 16–October 9) was 24.5 ⫾ 0.26°C (range ⫽ 20.6– 31.1°C). Larval density on July 16 in open containers averaged 210.8 larvae/container (Fig. 8). Larval densities remained high for the following 3 weeks but declined dramatically thereafter, with no larvae present in any open containers on week 11 (Fig. 8). Patently infected larvae in the open containers were produced by two different pathways (horizontal– horizontal and vertical) as described above. The horizontally induced patent infections were the predominant type with substantial infection levels (11.1–61.1%) for the first 7 weeks of the test (Fig. 8). Vertically infected larvae did not appear in appreciable numbers until weeks 5, 7, and 8, with 7.3, 12.1, and 6.3% infection rates, respectively (Fig. 8). Combined infection rates (horizontal plus vertical) varied from a low of 24.5% on
week 2 to a high of 73.2% on week 7. There were very few larvae alive in containers after week 7. One container had six larvae on week 10, all of which were vertically infected. The adult and larval populations were completely eliminated by week 11 (Figs. 7 and 8). DISCUSSION
In this study, we examined two methods (inoculative and inundative release) for the introduction of E. aedis into a contained population of A. aegypti. Inoculative release exploits vertical transmission of E. aedis to spread the parasite to new, often cryptic, habitats. Inundative release severely impacts both larvae and adults and exploits the persistence of the parasite within a container via horizontal transmission, with two possible outcomes. One of these involves benign infections in larvae to produce infected adults that distribute the parasite to new habitats; the other is the
TABLE 2 Comparative Emergence (Inundative Release) of Healthy (Unexposed) and E. aedis-Infected A. aegypti Adults from Covered Containers Percentage of infection
Unexposed Exposed a
Mean ⫾ SE.
No. females emerged/container a
No. males emerged/container a
Percentage of adult survival (combined) a
Females a
Males a
47.0 ⫾ 7.2 5.0 ⫾ 0.0
38.5 ⫾ 4.3 5.5 ⫾ 0.9
90.8 ⫾ 4.4 44.4 ⫾ 2.0
0 70.0 ⫾ 10.0
0 32.1 ⫾ 17.9
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FIG. 7. Ovipositional activity (inundative release) of A. aegypti females based on larvae from enclosure collected egg sheets hatched in the laboratory. Percentages of infection with vertically transmitted fat body infections (mean ⫾ SE) are indicated on the graph.
continued production of uninucleate spores in infected larvae to maintain an infectious titer of spores within containers (deviation 2, Fig. 1). This is particularly important given the finding that spores released into the aquatic environment are relatively short-lived. Under ideal conditions of storage, purified uninucleate spores of E. aedis maintain infectivity for up to 30 days
at between 20 and 30°C (Undeen et al., 1993). However, exposure of the extracorporeal spores of E. aedis to the biotic and abiotic factors found in the aquatic environment reduced infectivity by 10% after 2 days and by 50% within 5 days. Inoculative release resulted in dispersal of E. aedis by infected A. aegypti females to 18% (4 of 22) of the
FIG. 8. Weekly A. aegypti larval densities (inundative release) and percentages of Edhazardia aedis infection in larval A. aegypti from open containers (means ⫾ SE). Patent infections are a result of vertical transmission from infected adults or the horizontal–horizontal cycle.
IMPACT OF Edhazardia aedis
containers in 2 weeks, 50% of the containers in 6 weeks, and 100% of containers in 20 weeks. Infected females deposited eggs with E. aedis during the first 2 weeks postexposure but infection levels were low and remained low through week 10. Infection levels increased dramatically on week 12 and initiated the first of four cycles of ovipositional activity by infected females. These cycles may reflect the emergence pattern of adults that is also cyclic as well as the previous finding that most infected adults would only survive through one gonotrophic cycle (Becnel et al., 1995). Vertical infection levels (based on ovipositional data) peaked on weeks 12 (8%), 18 (12%), and 24 (19%), which corresponded (allowing for a 2-week developmental time) to the maximum rates of vertically infected larvae observed in containers on weeks 14, 20, and 26. These are substantial infection levels that provided inoculum (spores) for horizontal transmission to individuals in containers, as demonstrated by the observed horizontal infection levels in larvae. However, overall horizontal levels are higher than indicated by the patent horizontal infection levels, which do not reflect the benign infections in larvae that produce infected adults. An interesting feature of the infected larvae within the containers was the presence of horizontally induced patent infections throughout the sampling period. This deviation (type 2) from the typical life cycle (Becnel et al., 1989) assured the continued presence of spores within containers that are infectious orally for mosquito larvae. This was an unexpected finding but is apparently important for the long-term persistence of the pathogen within a container without the need for continued input of vertically infected individuals. The survival strategy of E. aedis therefore involves original contamination of a container by vertical transmission to establish the infection followed by continued horizontal transmission to both maintain the infection in the container and produce infected adults to disseminate the pathogen to other containers. A. aegypti survived the winter between years 1 and 2 of this study, but E. aedis did not survive in sufficient numbers to become reestablished. The uninucleate spores of E. aedis are extremely cold sensitive, with infectivity for A. aegypti larvae completely lost after storage at 5°C for 24 h (Undeen et al., 1993). It is unlikely that uninucleate spores survived the winter temperatures at the test location, which included a number of hard freezes. Developmental stages of the microsporidium in eggs and larvae are less sensitive to cold (Undeen et al., 1993), and it is possible that E. aedis could have survived within either infected eggs or adults. Nevertheless, the detrimental effect of cold on E. aedis indicates that introduction against A. aegypti populations near the temperature limits of its range would require periodic augmentation. Inundative release of E. aedis produced high larval
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and adult infections and successfully eliminated a contained population of A. aegypti within 11 weeks of introduction. For the first 3 weeks of the study, the population slightly increased, probably due to the large numbers of healthy adults present at the time of introduction. This was followed by a dramatic decline in the larval and adult populations. The decline coincided with 74% vertical infection rates in progeny of female adults and 46% patent infection rates in larvae on week 5. With infection levels of this magnitude, the population was unable to recover and all activity ceased by week 11. As in the inoculative release, horizontal transmission played an important role in maintenance of the parasite within containers throughout the duration of the study. It is likely that the rapid elimination of A. aegypti in this test was due, in part, to the isolation of the population, preventing both immigration of healthy adults and emigration of infected adults. Nevertheless, the results demonstrate that the inundative introduction of E. aedis can severely impact both larval and adult populations. The intensity and duration of this impact can only be determined by careful evaluation of controlled field releases with E. aedis into unrestricted populations of A. aegypti. Andreadis (1989) was the first to successfully introduce a polymorphic microsporidium, Amblyospora connecticus Andreadis, into a larval field population of the salt marsh mosquito Aedes cantator Coquillett. This was accomplished via the release of the infected copepod host and resulted in infection rates ranging from 16 to 24%. Because E. aedis does not require an intermediate host for horizontal transmission, introduction is greatly simplified and long-term persistence does not depend on the presence of two hosts. For successful introduction as a classical biological control agent, E. aedis must be able to persist and spread within a population of A. aegypti and regulate mosquito abundance to levels that would prevent disease transmission. Based on this study, the most immediate impact using E. aedis would be realized via an inundative release to challenge a large portion of an A. aegypti population. This would not only effectively suppress the exposed population but also establish a presence in containers to suppress new input to the site as well as assure that adults produced would harbor the pathogen. This would serve to spread the pathogen to other sites and limit the longevity of adults (Becnel et al., 1995), thereby reducing the likelihood of more than one blood meal and also reducing the risk of disease transmission. In conclusion, we believe that based on the results of this and previous studies, E. aedis should be considered a serious candidate for introduction as a classical biological control agent against A. aegypti. E. aedis is a
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parasite superbly adapted to A. aegypti, having evolved a number of strategies to ensure long-term survival. Two of these strategies involve the highly efficient mechanisms of vertical and horizontal transmission. Vertical infection spreads the pathogen to new, often cryptic, habitats and horizontal transmission amplifies the infection by producing additional infected adults and larvae. Use of E. aedis for management of A. aegypti is proposed not as a sole method but rather as part of the natural complex of regulatory factors utilized for classical biological control. This approach recognizes that eradication of the target mosquito is an unrealistic expectation but with a combination of physical, cultural, chemical, and biological control methods, populations of A. aegypti can regulated. ACKNOWLEDGMENTS The authors thank Charles S. Apperson, Theodore G. Andreadis, and Wayne M. Brooks for critically reviewing the manuscript.
REFERENCES Andreadis, T. G. 1989. Infection of a field population of Aedes cantator with a polymorphic microsporidium, Amblyospora connecticus via release of the intermediate copepod host, Acanthocyclops vernalis. J. Am. Mosquito Control Assoc. 5, 81–85. Andreadis, T. G. 1994. Host range tests with Edhazardia aedis (Microsporida: Culicosporidae) against northern Nearctic mosquitoes. J. Invertebr. Pathol. 64, 46–51. Becnel, J. J. 1990. ‘‘Edhazardia aedis (Microsporida: Amblyosporidae) as a Biocontrol Agent of Aedes aegypti (Diptera: Culicidae).’’ Proc. Int. Colloq. Invertebr. Pathol. Microb. Control, Adelaide, Australia, Vol. 5, pp. 56–60. Becnel, J. J. 1992. Safety of Edhazardia aedis (Microspora, Amblyosporidae) for nontarget aquatic organisms. J. Am. Mosquito Control Assoc. 8, 256–260.
Becnel, J. J., and Andreadis, T. G. 1999. Microsporidia in insects. In ‘‘The Microsporidia and Microsporidiosis’’ (M. Wittner, Ed.), pp. 447–501. Am. Soc. for Microbiol., Washington, DC. Becnel, J. J., and Johnson, M. A. 1993. Mosquito host range and specificity of Edhazardia aedis (Microspora: Culicosporidae). J. Am. Mosquito Control Assoc. 9, 269–274. Becnel, J. J., Sprague, V., Fukuda, T., and Hazard, E. I. 1989. Development of Edhazardia aedis (Kudo, 1930) N. G., N. Comb. (Microsporida: Amblyosporidae) in the mosquito Aedes aegypti (L.) (Diptera: Culicidae). J. Protozool. 36, 119–130. Becnel, J. J., Garcia, J. J., and Johnson, M. A. 1995. Edhazardia aedis (Microspora: Culicosporidae) effects on the reproductive capacity of Aedes aegypti (Diptera: Culicidae). J. Med. Entomol. 32, 549–553. Gubler, D. J., and Clark, G. G. 1995. Dengue/dengue hemorrhagic fever: The emergence of a global health problem. Emerg. Infect. Dis. 1, 55–57. Hembree, S. C. 1979. Preliminary report of some mosquito pathogens from Thailand. Mosquito News 39, 575–582. Hembree, S. C. 1982. Dose-response studies of a new species of per os and vertically transmittable microsporidian pathogens of Aedes aegypti from Thailand. Mosquito News 42, 55–61. Hembree, S. C., and Ryan, J. R. 1982. Observations on the vertical transmission of a new microsporidian pathogen of Aedes aegypti from Thailand. Mosquito News 42, 49–54. Johnson, M. A., Becnel, J. J., and Undeen, A. H. 1997. A new sporulation sequence in Edhazardia aedis (Microsporidia: Culicosporidae), a parasite of the mosquito Aedes aegypti (Diptera: Culicidae). J. Invertebr. Pathol. 70, 69–75. Kudo, R. 1930. Studies on microsporidia parasitic in mosquitoes. VIII. On a microsporidian, Nosema aedis nov. spec., parasitic in a larva of Aedes aegypti of Puerto Rico. Arch. Protistenkd. 69, 23–28. Rigau-Pe´rez, J. G., Clark, G. G., Gubler, D. J., Reiter, P., Sanders, E. J., and Vorndam, A. V. 1998. Dengue and dengue haemorrhagic fever. Lancet 352, 971–977. Undeen, A. H., Johnson, M. A., and Becnel, J. J. 1993. The effects of temperature on the survival of Edhazardia aedis (Microspora, Amblyosporidae), a pathogen of Aedes aegypti. J. Invertebr. Pathol. 61, 303–307.