Anopheles Quadrimaculatus - an overview (2023)

Anopheles quadrimaculatus complex: Anopheles quadrimaculatus (Say, 1824) and Anopheles smaragdinus (Reinert, 1997)

The Anopheles quadrimaculatus complex comprises five species that overlap in range, and only An. quadrimaculatus and An. smaragdinus have been recorded in Texas. All species are very similar and can only be distinguished by slight morphological differences. Anopheles quadrimaculatus is a medium-sized mosquito that can be identified by the four darker spots of dense scales on their wings where vein 1 is entirely dark. Their legs are entirely dark, and the femora and tibiae are tipped with pale scales. It has more setae on the scutal fossa, prealar area, and the interocular area than An. smaragdinus.

Bionomics. Eggs are deposited singly on the surface of water in patterns of vegetation chosen more frequently than others. Larvae occur in fresh water, slow moving streams, canals, ponds, and lakes containing emerging vegetation or floating debris, rice fields, and occasionally in temporary pools and permanent water swamps. Adults have multiple generations and overwinter in the hibernating adult stage. They are very active at dusk and at night in search of a blood meal and before dawn where they go into daytime resting places. They feed on humans, wild, and domestic animals. The species in this group have been infected successfully with P. vivax, P. falciparum, and P. malariae, and infection in wild-caught mosquitoes has been well established. Anopheles quadrimaculatus was regarded as the most important vector prior to the eradication of malaria in the United States (Carpenter and LaCasse, 1955) and a potential vector of other viruses (Reinert et al., 1997).

Mosquito Vectors and Epidemiology

Malaria has been viewed in the context of stable or unstable transmission, reflecting in part attributes of Anopheles species that affect their vectorial capacity. These include density, longevity, tendency to feed on humans, and duration of the extrinsic incubation period of the parasite in the vector. Stable malaria is most often associated with P. falciparum infection in highly endemic settings. It is characterized by low fluctuations in parasite incidence in human and vector populations, high prevalence, and high seroprevalence for antibodies. Epidemics are unlikely under these conditions, even though transmission continues at high rates. In such settings, vectors tend to be highly anthropophagic, exhibit greater longevity, and have relatively low, stable densities but still exhibit considerable seasonal variation. Unstable malaria tends to be associated with P. vivax infections in endemic settings of high fluctuation in disease incidence. Vectors tend to be zoophagic, have seasonally profound variation in population densities, have low or nondetectable field infection rates, and may have shorter longevity than do those in stable malaria settings. Epidemics can occur in conditions of unstable malaria if environmental changes favor increased vector–human contact, e.g., during civil strife, following water projects such as dams or irrigation schemes, or when a new vector is introduced into an area.

5.6 Establishment of Laboratory-Bred Anopheline Colonies

The requirement for mosquito-induced infections for malariotherapy was initially met with locally caught anophelines (James and Shute, 1926; Yorke and Macfie, 1924b). However, as the number of malariotherapy patients grew, a year-round supply of susceptible uninfected mosquitoes became indispensable. Thus were established the first insectaries for mass rearing of anophelines: Anopheles maculipennis (Shute, 1936), Anopheles quadrimaculatus (Boyd et al., 1935), and thereafter other species. The availability of mass-reared anophelines, to quote Boyd et al. (1935), ‘afford opportunities for research on entomological, parasitological, and epidemiological problems of malaria that have heretofore either been approached with difficulty or not at all’. Thus, a wealth of data accrued on the factors concerned with the transmission of malaria from humans to mosquitoes and from mosquitoes to humans (see below), and on anopheline biology, bionomics, and ecology. This illuminated the nature of malaria endemicity, and significantly benefited the design and efficiency of malaria control measures.

7.7 Light trap collections

Edhazardia aedis

Edhazardia aedis, a pathogen of Ae. aegypti, was isolated in Thailand (Hembree, 1979). This pathogen has a complex life cycle involving both horizontal and vertical transmission affecting two successive generations of the host (Hembree, 1982; Hembree and Ryan, 1982). Usually, one sporulation sequence occurs in the adult female (infected orally as a larva) and results in the formation of binucleate 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 but, owing to the abortion of one of the sequences, only one viable spore type is produced (Becnel et al., 1989). Larval death results in the release of uninucleate spores that are responsible for horizontal transmission when ingested by larvae. This developmental sequence leads to the formation of binucleate spores in the adult to complete the cycle.

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 horizontal transmission or, alternatively, by repeated cycles of vertical transmission.

Edhazardia aedis was transmitted to its natural host, Ae. aegypti, and to eight alternate hosts in the laboratory: Ae. albopictus, Ae. triseriatus, Ae. taeniorhynchus, Ae. atropalpus, Ae. vexans, Anopheles quadrimaculatus, Orthopodomyia signifera, and Toxorhynchites rutilus rutilus (Becnel and Johnson, 1993; Andreadis, 1994). Thirteen other mosquito species were not susceptible to E. aedis, including all species of Culex tested. In all susceptible hosts, the microsporidium underwent normal development but transovarial transmission was successful only in Ae. aegypti. Therefore, while a variety of mosquito species representing diverse genera is physiologically susceptible to E. aedis per os, the pathogen is specific for Ae. aegypti. Common non-target aquatic organisms were not susceptible to infection by E. aedis and, hence, there was no mortality due to E. aedis (Becnel, 1992b).

Edhazardia aedis causes larval death as a result of the highly efficient mechanism of vertical transmission, with approximately 95% of the progeny infected (Hembree, 1982; Hembree and Ryan, 1982; Becnel et al., 1995). Spores released from these infected progeny are infectious to all instars of Ae. aegypti and result in infected adults. The influence of the microsporidium E. aedis on the survival and reproduction of its mosquito host Ae. aegypti was studied in the laboratory (Becnel et al., 1995). Survival, fecundity, egg hatch, and percentage emergence for four gonotrophic cycles were compared for control and infected mosquitoes. Infected females oviposited 70% fewer eggs and percentage hatch was lower than for control females. Emergence in progeny of infected female Ae. aegypti was significantly less than for control mosquitoes in all gonotrophic cycles. The reproductive capacity (R_o) for control and infected adults was 168 and four, respectively, representing a 98% decrease. A semi-field study that involved the inoculative and inundative release of E. aedis produced encouraging results (Becnel and Johnson, 2000). Limitations to incorporating E. aedis into an integrated control program are the high costs involved in production in host mosquitoes and methods to store the fragile spore stage. The only possible field application of this microsporidian parasite would be as part of a classical biological control program to establish E. aedis in naïve host populations for long-term control (Becnel, 1990).

Ecological and epizootiological studies of E. aedis in natural populations of Ae. aegypti have not been completed. Optimism regarding the role of E. aedis as part of a program to control Ae. aegypti is based on a number of desirable traits determined in laboratory and semi-field studies. These studies have demonstrated that both the vertical and horizontal components of the life cycle of E. aedis are highly efficient in providing the means for the parasite to become established, persist, and spread in populations of Ae. aegypti. The profound effect on reproductive capacity of infected adults suggest the E. aedis can have a strong influence on host population dynamics (Becnel et al., 1995). Both inoculative and inundative release strategies were evaluated using E. aedis against a semi-natural population of Ae. aegypti (Becnel and Johnson, 2000). Inoculative release resulted in dispersal of E. aedis to all containers within the study site over a 20-week period. Inundative release eliminated the population of Ae. aegypti within 11 weeks of introduction. Good persistence is expected in release sites owing to life cycle flexibility with dissemination to other mosquito-inhabiting sites by means of vertical transmission. Survival during dry periods occurs within the mosquito eggs, where the pathogen can survive for the life of the egg (Becnel et al., 1989). In addition, an obligatory intermediate host is not required for horizontal transmission.

These desirable traits of E. aedis, including host specificity, support the belief that this pathogen can play an important role as a classical biocontrol agent in developing strategies to control Ae. aegypti. Ideally, E. aedis would be introduced via inoculative releases (rather than inundative releases), lessening the need for mass production. For this method to be successful, the pathogen must become permanently established or augmented seasonally to maintain acceptable levels of control.

5.1 Chitin synthase (CS)—genes

A large number of genes encoding the chitin polymerizing enzyme (CS) in yeast and filamentous fungi have been isolated and sequenced in the past three decades (Bullawa, 1993; Yarden and Yanofsky, 1991). Only in the last decade a number of CS genes from invertebrates notably nematodes (Fanelli et al., 2005; Foster et al., 2005; Harris et al., 2000; Veronico et al., 2001; Zhang et al., 2005) have been reported. Also recently, a CS gene from a marine bivalve mollusc (Atrina rigida) was cloned, sequenced and characterized (Weiss et al., 2006). The first cloned and sequenced CS cDNA (5757bp) from an insect was reported for the blowfly L. cuprina (Tellam et al., 2000). Since then, a number of other cloned and sequenced CS genes have been available from dipteran, lepidopteran and coleopteran species including the mosquitoes Aedes aegypti (Ibrahim et al., 2000; Kato et al., 2006) and Anopheles quadrimaculatus (Zhang and Zhu, 2006), the fruit fly Drosophila melanogaster (Gagou et al., 2002), the tobacco hornworm M. sexta (Hogenkamp et al., 2005), the fall armyworm Spodoptera frugiperda (Bolognesi et al., 2005), the beet armyworm Spodoptera exigua (Chen et al., 2007; Kumar et al., 2008), the diamondback moth Plutella xylostella (Ashfaq et al., 2007), and the red flour beetle Tribolium castaneum (Arakane et al., 2004). CS genes of the honey bee Apis mellifera, the mosquito A. gambiae and the fruit fly Drosophila pseudo-obscura were deduced from their respective genomic libraries. As more insect genomes are sequenced, additional CS genes are expected to become available in the future.

Multiple CS genes (grouped into two divisions and seven classes) encoding for a variety of CS isoenzymes with different biochemical properties and physiological functions were reported. Three and four CS isozymes were cloned in the yeasts Saccharomyces cerevisiae and Candida albicans, respectively (Lenardon et al., 2007; Lesage and Bussey, 2006), six genes in Botrytis cinerea (Choquer et al., 2004), and according to amino acid sequence similarities, up to 10 isozymes were identified in various filamentous fungi (Abramczyk et al., 2009). Similarly, multiple isozymes were described in the cloned (CS analogous) cellulose polymerizing enzyme genes (Pear et al., 1996). At least 10 isoforms of cellulose synthase isozymes were identified in Arabidopsis thaliana (Lindeboom et al., 2008), while 12 members of the gene family were detected in maize (Appenzeller et al., 2004).

In contrast to multiple fungal CS or cellulose synthase genes, basically only two tissue specific genes were discovered in insects (Gagou et al., 2002), derived perhaps from a gene duplication episode (Merzendorfer, 2006). It was found that the two genes are positioned adjacent to and on either side in the centromere of Drosophila third chromosome (Gagou et al., 2002). One gene (CS1) expressed in ectodermal cells encodes the integumental enzymes engaged in the formation of cuticles, while the other (CS2), which is relatively smaller in size, is restricted to midgut epithelial cells and is involved in the formation of PM chitinous matrices (Arakane et al., 2005; Hogenkamp et al., 2005; Zhu et al., 2002). A close analysis by Zimoch et al. (2005) revealed that although CS1 is expressed in M. sexta midgut tissue, its expression is due to the presence of tracheal cells developing in between midgut columnar cells. An inverse pattern of CS gene expression was demonstrated as CS1 mRNA levels are high during moulting and the wandering larval stage, while CS2 transcripts are elevated in feeding larvae during the inter-moult periods. Analyses using RNAi methodology, show that T. castaneum CS1 and CS2 are the contributors of cuticular and PM chitin, respectively (Arakane et al., 2005). When CS2 mRNA transcripts were knocked down in the mosquito A. aegypti, the PM was almost completely absent (Kato et al., 2006). The integumental gene (CS1) but not the gut gene (CS2) is expressed in two alternatively spliced forms referred as A and B. The spliced variants of M. sexta, which differ in amino acid sequence (a segment of 59 amino acid residues), are expressed differently during development and between tissues such as epidermis and ectodermal cells in tracheae (Arakane et al., 2004; Hogenkamp et al., 2005; Merzendorfer, 2006). It has been suggested that CS1B present in M. sexta midgut tracheae provide a potential glycosylation site that confer some unknown enzymatic function different from that of its CS1A counterpart (Hogenkamp et al., 2005). The two alternatively spliced transcripts of CS1 of the diamondback moth, P. xylostella, are expressed in all its developmental stages (Ashfaq et al., 2007). The CS1A variant is expressed during most of the larval and pre-pupal and pupal stages, while the CS1B form is expressed mainly during the pupal stage of T. castaneum (Arakane et al., 2004, 2005). S. exigua CS1 gene is highly expressed in early and late stages of each larval instar and high levels of transcripts throughout the pupal stage were detected (Chen et al., 2007). The CS2 gene from S. exigua gut was similar to the isolated one from M. sexta including intron/exons organization and absence of alternate exons (Kumar et al., 2008). The CS2 genes in M. sexta, T. castaneum and S. frugiperda are expressed in the midgut, with a peak of expression in the actively feeding stages (Arakane et al., 2005; Bolognesi et al., 2005; Hogenkamp et al., 2005; Zimoch and Merzendorfer, 2002). The transcripts' pattern in T. castaneum CS2 reveals increased level of expression in late larval and adult stages (Arakane et al., 2004). Increased levels of A. aegypti CS2 transcripts following blood meal were detected (Ibrahim et al., 2000; Kato et al., 2006). Blood feeding by the mosquito adults also induced transcriptional up-regulation of glutamine: fructose-6-phosphate amidotransferase, a key precursor in the pathway of the CS substrate formation (Kato et al., 2006).