Open Access

Diapause and quiescence: dormancy mechanisms that contribute to the geographical expansion of mosquitoes and their evolutionary success

  • Diego Felipe Araujo Diniz1,
  • Cleide Maria Ribeiro de Albuquerque2,
  • Luciana Oliveira Oliva2,
  • Maria Alice Varjal de Melo-Santos1 and
  • Constância Flávia Junqueira Ayres1Email author
Parasites & Vectors201710:310

https://doi.org/10.1186/s13071-017-2235-0

Received: 26 January 2017

Accepted: 8 June 2017

Published: 26 June 2017

Abstract

Mosquitoes are insects belonging to the order Diptera and family Culicidae. They are distributed worldwide and include approximately 3500 species, of which about 300 have medical and veterinary importance. The evolutionary success of mosquitoes, in both tropical and temperate regions, is due to the various survival strategies these insects have developed throughout their life histories. Of the many adaptive mechanisms, diapause and quiescence, two different types of dormancy, likely contribute to the establishment, maintenance and spread of natural mosquito populations. This review seeks to objectively and coherently describe the terms diapause and quiescence, which can be confused in the literature because the phenotypic effects of these mechanisms are often similar.

Keywords

CulicidaeSeasonalityMetabolismAdaptationDispersionDisease transmission

Background

Mosquitoes are arthropods that can cause considerable nuisance and affect human health worldwide [1, 2]. They are among the most prolific and invasive species, contributing to the spread of endemic diseases [3, 4]. These organisms are present in most places on the planet, from the Arctic to the most remote desert oases, except Antarctica due to its extremely low temperatures. Thus, mosquitoes are widely diverse and can easily be found in a wide variety of habitats, including forested, rural and urban environments [2, 5].

These insects have been intensely studied since the end of the nineteenth century due to their ability to act as hosts for many pathogens, including helminths, protozoans and arboviruses, that cause disease in humans and other vertebrates [2, 6]. However, only 10% of the approximately 3500 mosquito species are medically relevant [1, 711].

Mosquitoes, especially from the genera Anopheles, Aedes and Culex, include vectors for three major groups of human pathogens: parasites from the genus Plasmodium, which cause malaria; filarial worms from the genera Wuchereria and Brugia; and many arboviruses, including the agents of dengue, yellow fever, chikungunya, zika and others [1214]. Estimates by the World Health Organization (WHO) indicate that diseases transmitted by mosquitoes are among the major causes of morbidity and mortality in developing countries [15], and high densities of mosquitoes severely challenge vector control programs [16]. The explosive growth of natural mosquito populations is strongly related to the survival and dispersion strategies that some species have acquired over the course of their evolutionary history [17].

Dormancy is a biological trait that may play an important role in the maintenance of natural populations and refers to a physiological phenomenon characterised by the interruption or reduction of metabolic activity in an organism. In mosquitoes, dormancy can occur at different stages of the life-cycle [18]. Diapause and quiescence represent different types of dormancy found in many species of mosquitoes. In this review, these terms are analysed for their conceptual principles and their respective delayed developmental effects; in addition, the mosquito species found to exhibit these phenomena will be noted.

Insect dormancy and its various types

Dormancy is a physiological phenomenon defined as a state of suspended development or suppressed metabolic activity in an organism [19]. Dormancy can occur in both plants and animals; in insects, it can manifest in the embryonic (pharate larvae), immature (larvae and pupae) and adult stages [18]. This phenomenon can be triggered by climactic signals, especially the photoperiod for temperate climate insects and relative humidity for tropical insects. This adaptation seeks to promote survival during and after unfavourable environmental conditions and is known in the literature as heterodynamic development [20]. In 1869, the term dormancy was first described as a period of inactivity caused by low temperatures by the French researcher Duclaux, who was studying silkworms (Bombyx mori) [20, 21]. According to a literature review by Danks [20] on the definitions and terminology of dormancy in insects, dormancy is divided into two major categories: diapause and quiescence. The terms diapause and quiescence have been reported to be synonymous in the literature [8, 2227], but these survival strategies arise from distinct signalling pathways even though the strategies have the same goal: to ensure survival during and after environmental stress.

Mosquitoes belong to one of the most well-adapted taxa in the insect group; they are present across most of the planet, they occupy diverse niches and are potential disease vectors [2]. Diapause and quiescence are well characterised in several stages of the mosquito life-cycle. In the embryonic phase, for example, both strategies have the same effect: the inhibition of larval hatching. Conversely, only diapause drives dormancy in the larval and adult stages of mosquitoes [28].

Diapause in mosquitoes

Diapause is a well-studied seasonal survival strategy and is influenced by several factors, such as the species-specific ecological interactions, biogeography, life history and physiology of many insects [29]. The etymology of the word “diapause” comes from the Greek diapausis (pause), derived from the verb diapauein, which means to stop or to decrease activity at a time of constant action [30]. Biologically, Tauber et al. [31] defined the diapause phenomenon as a dynamic state of low metabolic activity that is genetically determined and mediated by neurohormones that phenotypically affect individuals by decreasing morphogenesis, blocking reproduction and metamorphosis, and increasing tolerance to extreme environmental conditions

The first studies on diapause in mosquitoes coincided with early studies of seasonality, diapause and photoperiod in other insects [17]. Early reviews on the topic were performed by Lees [32], Danilevskii [33], Tauber et al. [31] and Danks [20]. Studies at the time were motivated by the mosquito’s hematophagous habit, which is linked to its ability to transmit the causative agents of several diseases such as malaria, filariasis, and many arbovirus infections (yellow fever, Western equine, St. Louis and Japanese encephalitis, and West Nile fever) [34].

Diapause is common in insects and other arthropods, especially in areas with harsh winters. Many aspects of diapause are critical for understanding the transmission cycle of vector-borne diseases, as this survival strategy contributes to the maintenance, establishment, growth and dispersion of natural vector populations after the end of an unfavourable season to their development [29]. The process of diapause seeks to reactivate development via external signals that control the genetic factors underlying the dormant phenotype. This can occur in several phases of the life-cycle, but most often only one developmental stage enters diapause [34].

What is the environmental signal that induces diapause in mosquitoes?

Species exhibiting the phenotypic plasticity to undergo diapause have the required information encoded in their genomes. The major stimuli inducing diapause in natural populations are changing photoperiod (short days and long nights) and gradual decreases in temperature [31, 3540]. Mosquito species that use photoperiod to signal diapause include Aedes albopictus, Aedes atropalpus, Aedes sollicitans, Aedes taeniorhynchus, Culex pipiens and Culex restuans [38, 39, 4144].

Preparation for diapause occurs in mosquitoes when pupae and/or adult females, which are thought to be the determining stages for this biological trait, are stimulated by exposure to the seasonal changes that typically occur during transitions between a favourable and unfavourable season [29, 39, 4547]. For Ae. albopictus, for instance, induced females develop their offspring for diapause, which in turn present low metabolism in each life-cycle stage during the winter [48, 49]. However, for Cx. pipiens, the induced pupae females express diapause when they become adults [50]. Therefore, this ecological adaptation is indispensable for coordinating the growth, development and reproduction of mosquito species found in temperate zones [29].

Ecophysiological phases of diapause

The phenomenon of diapause consists of three ecophysiological phases [51]. The first is the diapause preparation or pre-diapause phase, which corresponds to the sensitive stage in which the insect is exposed to one or more environmental signals (token-stimuli) that trigger and initiate the phenomenon in the offspring in the following season [19, 48]. In some species, this phase is favourable for the storage of energetic reserves that will be used for basal maintenance of the insect during dormancy and the reinitiation of development at the end of the process. In addition, morphophysiological, biochemical and behavioural changes can be observed in the individuals at this phase [19, 29, 51, 52]. This occurs because some mosquito species extend the developmental time of a specific life-cycle stage (delayed developmental effects) to increase their exposure to the stimulus, which is a favourable event for ensuring that the dormancy phenotype occurs in the offspring [29].

Culex pipiens females programmed for adult diapause have a longer larval phase, resulting in larger pupae and adults that have more lipids than their non-diapausing homologs [53]. The fat levels in females of this same species destined for diapause continue to increase significantly during the week following the emergence of the adults, reaching twice the level observed in non-diapausing females [54]. At the molecular level, this increase in energetic reserves is accompanied by an increased expression of genes associated with lipid reserve synthesis [55]. In Ae. albopictus, eggs in diapause are larger and contain more lipids than non-diapausing eggs, which is likely due to the increased expression of genes involved in lipid storage during pre-diapause [56].

Diapause programming (Fig. 1) involves the capture of photoperiod information by the central nervous system (CNS) of gravid females, followed by a cascade of biochemical events and culminating in the transfer of a molecular diapause regulator that promotes a dormancy state in embryos [29]. Thus, clock genes can reasonably be assumed to be involved in the regulation of circadian rhythms and, consequently, in the seasonal response based on the length of day and night [57, 58]. The main clock genes in mosquitoes that are involved in circadian rhythm regulation but are not necessarily related to diapause have been characterised in Ae. aegypti, Ae. albopictus, Anopheles gambiae, Cx. quinquefasciatus and Wyeomyia smithii [5965].
Fig. 1

Embryonic diapause induction in mosquitoes. 1 Exposure of pupae and/or adult females to short days, long nights and gradual temperature drop, and abiotic factors that promote the preparation of the embryonic diapause. 2 Expression of specific genes transferred from the female to the offspring allows diapause to be triggered and the embryos (pharate larvae) to become refractory to the hatching stimulus

Diapause specifically refers to the actual time when development is interrupted or significantly slowed, and the insect does not respond to environmental stimuli [29]. This is the second phase and can be divided into the following sub-phases: (i) the responsive phase-the beginning of the process when development is stopped at a specific life stage; (ii) the initiation stage-the phenomenon is maintained and controlled by endogenous and/or exogenous factors, and (iii) termination-the time when the individuals receive the signal to return to normal metabolic activity [19]. During diapause, various endogenous changes can be observed, but these depend on the species studied. In Ae. albopictus embryos, Wy. smithii larvae and Cx. pipiens adults, for example, lower lipid degradation and higher tolerance to desiccation and low temperatures are present [48, 6670].

At the molecular level, a few genetic components that mediate these adaptive physiological traits have been reported in previous studies. In Ae. albopictus, resistance to desiccation, promoted by diapause, results from an increase in the external surface area of the egg with increased hydrocarbon levels, and this is caused by an overexpression of a transcript involved in lipid storage. However, the mechanisms responsible for cold tolerance in this species have not been determined [71]. In Cx. pipiens, increased tolerance to desiccation during diapause is primarily due to an increase in the hydrocarbon layer on the cuticular body surface of adults and an increase in trehalose production, which contribute to both desiccation and cold tolerance [66]. In contrast, the molecular events that promote the effects of diapause in Wy. smithii have yet to be discovered [29].

The last phase, termed post-diapause, is characterised by the complete reactivation of metabolism and development in the insect [51]. Although photoperiod is widely used as an environmental stimulus for entering diapause, it is less often used to signal the end of diapause; however, some exceptions exist, such as, for example, in Wy. smithii, where another change in photoperiod causes diapause to end [29, 72]. In Ae. albopictus, the termination of diapause in the eggs may be signalled by changes in photoperiod and by increasing temperature [73]. Another interesting characteristic, in addition to post-diapause, is a phenomenon known as post-diapause quiescence (Fig. 2), which is also present in Ae. albopictus [49, 73]. This process is considered to be a phenotypically indistinguishable phase from diapause. The insect remains in a state of dormancy, its metabolic rate continues to be low, and many of the same genes associated with diapause continue to be expressed. Thus, diapause and quiescence possibly have many molecular components in common, although the components for initial programming are exclusive to diapause [49]. Physiologically, the only difference is that during quiescence, the insect remains fully capable of responding to environmental stimuli [29, 74, 75].
Fig. 2

Termination of the embryonic diapause in the mosquito (post-diapause). 1 Return to normal conditions (temperature and photoperiod) that signal the end of diapause. 2 Post-diapause embryo under favorable conditions responds to the stimuli of relative humidity increase and optimal temperature in the environment, resulting in larval hatching. 3 Post-diapause embryo under non-favorable abiotic conditions is sensitive to environmental stimuli but remains dormant in a quiescent state (post-diapause quiescence) until the temperature and relative humidity become ideal for larval hatching. The dynamics of quiescence are the same as post-diapause quiescence

Diapause in different mosquito species/life-cycle stages

Diapause can occur in different phases of the mosquito life-cycle, i.e. in the embryo (pharate larvae inside the egg), larval and adult stages. However, this type of dormancy tends to occur in a single stage of the life-cycle for a given species [19, 29, 34]. Furthermore, in some species, diapause can occur in more than one stage, more precisely, between the embryonic and larval stages [36, 62, 7678].

Embryonic diapause

This is the most common type of dormancy and occurs in the mosquito embryo. Using Ae. albopictus, a model organism for diapause, as a reference, the embryo is completely formed inside the egg chorion, but a metabolic depression of post-embryonic development occurs due to genetic programming; thus, the larva is unable to respond to any abiotic signals, that is, the larva is refractory to hatching stimuli [29, 45, 46, 79]. Embryos in diapause are more tolerant to desiccation and tend to have a higher total lipid content than normal embryos [57, 68, 80]. The overexpression of ecdysteroid transcripts, found by transcriptomic analysis of mature oocytes, likely regulates embryonic diapause in Ae. albopictus and other mosquitoes [48, 49]. The genera of mosquitoes with embryonic diapause are Aedes, Anopheles, Psorophora and Ochlerotatus [29, 34], and the major species reported in the literature for each genus are listed in Table 1.
Table 1

Embryonic diapause in different mosquito species

Species

References

Aedes albopictus

[46]

Aedes atropalpus

[41]

Aedes campestris

[150]

Aedes canadensis

[151]

Aedes caspius

[76]

Aedes dorsalis

[152]

Aedes fitchii

[153]

Aedes geniculatus

[77]

Aedes hendersoni

[78]

Aedes hexodontus

[154]

Aedes impiger

[129]

Aedes japonicus

[155]

Aedes mariae

[35]

Aedes nigripes

[156]

Aedes nigromaculis

[152]

Aedes sierrensis

[36]

Aedes sticticus

[157]

Aedes taeniorhynchus

[44]

Aedes triseriatus

[158]

Aedes vexans

[159]

Anopheles walkeri

[160]

Psorophora ferox

[151]

Ochlerotatus dorsalis

[152]

Ochlerotatus nigromaculis

[152]

Ochlerotatus hexodontus

[154]

Ochlerotatus flavescens

[34]

Ochlerotatus triseriatus

[75]

Ochlerotatus togoi

[75]

Diapause in larval stages

This physiological process is known in the literature as the syndrome of larval diapause, which is characterized in mosquitoes by the prolongation of the third- or fourth-instar. The induction of diapause in larvae is directly stimulated by a gradual decrease in environmental temperature, and the metabolic activity rapidly returns to normal in response to its normalisation in the wild, although changes in photoperiod also play a role in its induction [34]. The behaviour of the larvae is characterised by reduced locomotor and feeding activities, consequently promoting an increased accumulation of body reserves that, in turn, provide increased cold tolerance [34]. Under normal conditions, the progression of the development of the larval stages occurs biochemically through the periodic release of the steroid hormone ecdysone by the prothoracic gland, which culminates in the moults. When the larvae are in diapause, ecdysone release is lacking, and therefore, the larvae do not advance from one stage to the next [29]. Currently, no molecular studies have explained the hormonal basis for diapause in mosquitoes, but some studies have reported the absence of ecdysone as a major cause of larval diapause in other insects, which is likely similar to mosquitoes [81]. Mosquito species in which this type of dormancy has been observed are listed in Table 2.
Table 2

Larval diapause in different mosquito species

Species

References

Aedes caspius

[76]

Aedes geniculatus

[77]

Aedes hendersoni

[78]

Aedes sierrensis

[36]

Aedes togoi

[68]

Aedes triseriatus

[159]

Anopheles barberi

[161]

Anopheles plumbeus

[162]

Anopheles pulcherrimus

[85]

Armigeres subalbatus

[163]

Culiseta melanura

[164]

Orthopodomyia alba

[165]

Orthopodomyia puchripalpis

[166]

Orthopodomyia signifera

[167]

Toxorhynchites rutilus

[168]

Wyeomyia smithii

[169]

Diapause in adult females

Diapause in adult female mosquitoes involves a set of important characteristics, such as the interruption of gonadal development, reduced biting behaviour, negative phototaxis and changes in total metabolism, leading to the gradual accumulation of body fat. Mosquitoes can enter diapause in many habitats, such as caves, soil cavities, burrows, vegetable store-houses, empty sheds, basements, and catacombs [34]. In adult females, a type of dormancy occurs, known as reproductive diapause, where sexual immaturity is prolonged because the ovarian follicles do not differentiate completely and hence, delay the blood feeding activity [8, 34, 74].

The majority of studies on diapause in adult mosquitoes has been performed on the species Cx. pipiens, which is considered a model organism [29]. Under normal conditions, after the emergence of the winged form, juvenile hormone (JH) is synthesised and released, promoting ovarian follicle growth within 3 days, and shortly after, the female is ready for its blood meal, which will contribute to oocyte maturation. In contrast, females programmed for diapause do not release JH immediately after emergence, and the follicles remain atrophied. The females also have a reduced aggressiveness [82]. Experiments on diapausing Cx. pipiens females treated with JH have exhibited ovarian growth stimulation, confirming the importance of inhibiting this hormone to initiate diapause in adult mosquitoes [54, 83]. It is important to highlight that in this species, males do not undergo diapause, thus, they inseminate females and then die, as they cannot overwinter [31, 33, 84]. The Anopheles and Culex species reported as exhibiting adult diapause are listed in Table 3 [38, 40, 75, 8594].
Table 3

Adult diapause in different mosquito species

Species

References

Anopheles atroparvus

[85]

Anopheles earlei

[86]

Anopheles freeborni

[87]

Anopheles superpictus

[85]

Anopheles gambiae

[88]

Anopheles hyracanus

[87]

Anopheles maculipennis

[85]

Anopheles messeae

[85]

Anopheles punctipennis

[89]

Anopheles sacharovi

[75]

Culex bitaeniorhynchus

[75]

Culex apicalis

[75]

Culex modestus

[75]

Culex pipiens

[39]

Culex restuans

[39]

Culex tarsalis

[90]

Culex tritaeniorhynchus

[91]

Culiseta alaskaensis

[92]

Culiseta impatiens

[93]

Culiseta inornata

[94]

The molecular biology of diapause in mosquitoes

Most studies on the genetic basis of diapause in mosquitoes have focused on two species, Ae. albopictus and Cx. pipiens, which are considered model organisms for this approach. Early studies were performed in the fly Drosophila melanogaster; however, although this species is a reference for basic genetic studies, it did not yield good results in the gene expression studies, as the insect showed highly variable responses and high variance between individuals [9597].

Diapause in Cx. pipiens, according to breeding experiments, is polygenetically regulated and involves genes on all three chromosomes [98, 99]. A more detailed study on the species using suppressive subtractive hybridization to determine the expression profile of diapause genes revealed that a set of 40 genes were differentially expressed. Most of these genes were implicated in the expression of structural components and responses to the environmental stress [100]. One of the upregulated genes was a stress tolerance gene expressing a heat-shock protein (HSP70), which functions as a chaperone to inhibit abnormal protein folding in harsh environmental conditions, including desiccation and cold [101]. In addition, metabolic genes are overexpressed in Cx. pipiens during diapause, including the mitochondrial malate dehydrogenase (mmd), methylmalonate-semialdehyde dehydrogenase and cytochrome oxidase (cox) genes. These genes may be involved in the specific metabolic events associated with diapause and have been implicated in increased cold tolerance. The expression of certain cytoskeletal genes was also upregulated by preparation for diapause. The actin gene, for example, is overexpressed during the diapause preparation stage, likely due to increased flying activity before dormancy begins, and the expression levels of this gene decrease gradually during diapause and are low at diapause termination. Downregulated genes included the ribosomal genes S3A, rpS6 and rpS24, which are involved in gene regulation (translation initiation) and inhibit or reduce the expression of several other metabolic genes [102].

Most information on changes in gene expression associated with diapause in mosquitoes is based on recent high-throughput sequencing studies (such as RNA-seq) examining the transcriptome of Ae. albopictus at different stages [29, 48, 49, 102]. Early studies on the differential expression of transcripts were performed in the ovary cells of this mosquito (oocytes), and later, the molecular mechanisms during embryogenesis were investigated.

A study by Poelchau et al. [48], who sequenced the oocyte transcriptome of diapausing Ae. albopictus females, and another more recent similar study from the same group, Poelchau et al. [49], who used diapausing embryos from Ae. albopictus, revealed the overexpression of genes involved in various biological processes. The following are included among these genes: the gene ing1, which encodes for the inhibitor of growth protein and is involved in the interruption of the cell division cycle [52, 103]; the gene rack1, a putative receptor for activated protein kinase C, which may bind to several signaling molecules, including transcription factors related to ecdysone (20-hydroxyecdysone), and is probably associated with the preparation for diapause [104, 105]; the gene pepck (phosphoenolpyruvate carboxykinase), whose product participates in the glycogen pathway to move from aerobic to anaerobic metabolism in diapausing mosquitoes [106, 107]; and the gene GPCR (G protein-coupled receptor), which is involved in increased resistance to environmental stress [108].

Quiescence in mosquitoes

Quiescence is a type of irregular dormancy (non-seasonal) characterised by slowed metabolism and directly resulting from unfavourable environmental conditions, including low humidity and high temperatures [22, 74, 109, 110]. This adaptive trait is often confused with diapause, especially when referring to embryonic dormancy, but quiescence is a less complex biological trait that does not depend on endogenous control for its initiation. Stimuli that trigger quiescence are referred to as acyclic environmental changes [19]. Quiescence also differs from diapause because it is neither a previously programmed event, nor is it hormonally controlled; once the stimulus that induces the process ceases, physiological activity is restored [29, 34, 73]. Because quiescence is controlled exogenously, it is possible that rapid gene activation and macromolecule synthesis or degradation are not required for entry into the quiescent state [109].

In mosquitoes, as in other organisms, the term quiescence is applied to various biological events. Most commonly studied in the egg, quiescence in mosquitoes can be stimulated in different stages or structures, enabling the insect to attain favourable conditions for survival. In the mosquito Cx. quinquefasciatus, for example, mature spermatozoids are maintained in quiescence in the male reproductive tract and are activated in response to specific chemical signals [111]. In this species, motility is stimulated by substances from the accessory glands in males and is possibly controlled by protein phosphorylation and Ca2+ levels [111]. In addition, in females, degenerative dilations may develop in the ovary, which contains granular material during winter, and the presence of these expansions in the ovaries is thought to be indicative of quiescence [112, 113].

In the family Culicidae, quiescence, unlike diapause, has been primarily observed in the egg, reflected in the resistance to desiccation that allows the embryo to survive in dry conditions. The process begins when the embryo (pharate larvae) receives an external stimulus, such as a rapid drop in humidity or change in temperature, which signals unfavourable environmental conditions and impedes larval hatching [19, 34, 74]. In this case, the developmental arrest is temporary and immediately reversible, as contact with water induces rapid hatching; that is, the quiescent embryo is not refractory to hatching stimuli as is found in diapausing embryos [18, 19, 34, 49, 114]. As shown in Table 4, the genera reported exhibiting quiescence are Aedes, Anopheles and Culex [23, 49, 68, 80, 115124]. The species Ae. aegypti is prominent among mosquitoes due to its strategy of prolonged viability by embryonic quiescence, significantly contributing to the constant expansion of populations in the wild [74, 75]. However, several studies have erroneously reported this trait as diapause [8, 2224, 26].
Table 4

Embryonic quiescence in different mosquito species

Species

References

Aedes aegypti

[68]

[23]

[116]

[170]

[118]

[119]

[120]

[121]

[122]

[115]

[123]

Aedes albopictus

[68]

[117]

[71]

[48]

Aedes flavopictus

[68]

Aedes galloisi

[68]

Aedes riversi

[68]

Anopheles aquasalis

[121]

[123]

Anopheles gambiae

[124]

Culex quinquefasciatus

[123]

[123]

Egg quiescence or embryonic desiccation resistance

Egg quiescence is commonly referred to as “embryonic desiccation resistance” (EDR) and depends on several factors that range from differences in eggshell composition and structure to physiological changes, resulting in reduced metabolism in the larvae contained within the egg [22, 116, 121, 125]. However, because the ability to resist desiccation is a property of the egg and not of the embryo and because desiccation can occur at other stages of development, the term “egg resistance to desiccation (ERD)” has been suggested as more appropriate for referring to this phenomenon [123].

The three layers that form the eggshell, the exochorion, endochorion and serosal cuticle, are particularly important for ERD [116, 123]. The first two layers are produced in the ovary, by females, and are, therefore, present at laying [74, 123, 126]. The serosal cuticle (the innermost layer), in turn, is an extracellular matrix produced by the extraembryonic serosa during early embryogenesis. In Ae. aegypti, secretion of the serosal cuticle occurs between 11 and 13 h after oviposition and approximately 8 h post-fertilization in An. gambiae [115, 124].

This cuticle likely secretes a chitin-containing material under the chorion, the external layer of the egg, making it impermeable and protecting the embryo from desiccation [116, 123]. Changes in the amounts of the eggshell components are associated with water loss regulation and are fundamental for determining the intensity of egg dehydration. Aedes albopictus females exposed to short day length in temperate regions produce eggs in photoperiodic diapause, unlike populations in tropical regions, which enter quiescence. One of the characteristics of the egg that permits this adaptation is the high quantity of fatty acyl-CoA elongase in the tissue of mature oocytes responsible for producing hydrocarbons in the eggshell [71]. These hydrocarbons regulate water loss in insect eggs, and the abundance of this enzyme varies in the eggs of Ae. albopictus exposed to long and short days in temperate populations but is maintained at relatively constant levels in tropical populations [80, 123]. In addition to several hydrocarbons in the eggshell, the amount of chitin is another factor involved in ERD in mosquitoes, such as Cx. quinquefasciatus, An. aquasalis and Ae. aegypti. Eggshells with higher amounts of chitin are more resistant to desiccation [123].

Quiescence patterns in container-inhabiting mosquitoes

ERD has been more commonly studied in container-inhabiting mosquitoes, including Ae. aegypti and Ae. albopictus. In urban areas, females often lay their eggs in containers with clean water, especially disposable containers, tires, plant pots and water storage containers [127, 128]. Because the eggs are laid near the water surface, this developmental phase is very susceptible to dehydration, particularly during the first few hours of development [129].

First-instar larvae that remain inside quiescent eggs have been referred to as pharate first-instar quiescence [34, 74]. Normal development finishes approximately 3 days after oviposition and larval survival depends on maternal reserves [119]. Throughout the quiescent period, the larval developmental period is significantly prolonged, and lipid reserves are reduced, incurring fitness costs for larval viability, compromising the reproductive performance of the adult [34, 74].

Minimally studied, quiescence in mosquito eggs does not appear to have a uniform pattern, exhibiting variability between species or even among populations of the same species [69, 123, 130]. Under similar low-moisture conditions, the pharate first instars of Cx. quinquefasciatus, An. aquasalis and Ae. aegypti can survive for a few hours, 1 day or several months, respectively [123]. These differences may be due to traits inherent to the eggs of each species, such as size, the structure of the exochorion and endochorion, differences in metabolite quantity and formation of the serosal cuticle [68, 121, 131, 132].

Brazilian colonies of Ae. aegypti maintained at a temperature of 28 ± 1 °C, a relative humidity of 80 ± 5% and a photoperiod of 12 h had a viability period of up to 492 days, with high hatching rates between three and 121 days [23]. A similar pattern with high larval hatching rates (80%) was reported by Diniz et al. [115] in quiescent Ae. aegypti eggs that had been stored for up to 150 days. The authors compared eggs from laboratory and wild populations with different susceptibilities to the insecticide temephos, which were then maintained for up to 180 days at 26 °C, with a photoperiod of 12 h and at 50–60% humidity. The high viability of quiescent eggs from temephos-resistant females suggests a high contribution to the maintenance of resistant individuals in the wild. Similarly, in Australia, quiescent Ae. aegypti eggs remained viable for more than a year with a hatching rate of approximately 2–15%, allowing its dispersion to new locations [133]. Species inhabiting forests have been shown to be less resistant to changes in humidity. Aedes riversi, Ae. galloisi and Ae. flavopietus eggs have different survivability rates in very humid conditions but were less resistant than Ae. aegypti and Ae. albopictus under low humidity. Intraspecific differences in ERD were also observed among these species, as Ae. riversi and Ae. flavopietus strains from subtropical regions had lower viability than strains from temperate regions [68].

The molecular biology of quiescence in mosquitoes

Although much is known about the metabolic mechanisms and molecular biology of diapause, very little is known about these aspects during quiescence in mosquitoes. A study by Poelchau et al. [49] that compared the transcriptomes of quiescent and diapausing Ae. albopictus eggs found that the genetic expression profile between these samples converged over time; that is, the transcription profile in eggs during late diapause (40 days) is similar to that in quiescent eggs [49]. An important aspect of this study is that expression levels of genes related to lipid metabolism were always higher in eggs in diapause, demonstrating the likely importance of this reserve for maintaining embryonic diapause and explaining why eggs in diapause have more lipid reserves than quiescent eggs [49].

Currently, the metabolic pathways or hormones associated with quiescence are unknown, and only the chitin synthase (CHS) gene has been described as being related to this phenomenon in mosquitoes. This gene promotes the synthesis of chitin, which is then secreted into the extracellular space of the egg, with direct implications for the formation of the serosal cuticle and consequently the resistance to desiccation [116, 124, 134]. Despite being primarily cited for An. gambiae (AgCHS), this gene is highly conserved in two other species of mosquitoes, An. quadrimaculatus and Ae. aegypti. The gene has two variants, but only the allele AgCHS1 is involved in embryogenesis. In Ae. aegypti, for example, the expression of the gene peaks between nine and 12 h after oviposition, coinciding with the acquisition of resistance to desiccation through the complete covering of the embryo by the chitinized serosal cuticle [116, 124].

Eco-epidemiological importance of quiescence

In Europe, a considerable increase in invasive mosquito propagation has been observed since the end of the 1990s, with the species Ae. albopictus, Ae. aegypti, Ae. japonicus, Ae. atropalpus and Ae. koreicus already established on the continent [131]. In addition to increased population densities, the distribution of Ae. albopictus has continued to increase, and several other species of Aedes are being reported in new countries each year [135]. For example, recently, a research group from Brock University reported the detection of Ae. aegypti for the first time in Canada [136]. In addition, Lima et al. [137] reported a permanent Ae. aegypti local population in the Capitol Hill neighbourhood in Washington DC that can overwinter. This is contrary to the previous hypothesis that different introductions of Ae. aegypti every year maintain that local population. All these species are well adapted to the urban environment, exploiting a variety of container habitats that proliferate near human settlements, and both quiescence and diapause may be contributing to the maintenance of these populations. In addition to the annoyance of their bites, these mosquitoes are potential vectors for agents that cause tropical diseases, including Zika, dengue, chikungunya and yellow fever [138]. Quiescence in Ae. aegypti may also allow the survival of infected embryos, favouring virus survival and its maintenance in nature [8, 122, 139]. For example, DENV-1 was detected and isolated in 8.33% of Ae. aegypti eggs in Florida, suggesting that maintenance of dengue outbreaks in 2009 and 2010 in Key West may have been facilitated by vertical transmission [140]. Transovarian transmission of DENV in the field was also detected in larvae and adults originating from larvae collected in domestic containers in Rajasthan, India. Approximately 1.09% of the reservoirs contained larvae with the virus, detected by the indirect fluorescence antibody test and reverse transcriptase polymerase chain reaction. In this case, dormant eggs may have contributed to prolonging dengue epidemics [141]. Furthermore, Zika virus, a flavivirus that has recently caused large outbreaks in several countries and has been linked to microcephaly cases and other neurological complications, has also been reported as being transferred via transovarian transmission by Ae. aegypti and Ae. albopictus [142144, 145].

Although the implications of quiescence on the ecology of mosquito vectors and public health are well established, its effects on physiology, behaviour and life history are less understood. Maternal reserves accumulated in eggs directly influence the period of dormancy in the first-instar larvae contained within the eggs [8, 120]. Thus, quiescent eggs pose an important problem for vector control because these eggs can directly contribute to the maintenance of mosquito populations in treated areas. Aedes aegypti eggs from a single laying, at the same age, and maintained under the same environmental conditions had different hatching rates during the same period of quiescence. Aedes aegypti embryos employ a hedge betting mechanism not all eggs hatch at the first stimulus; some need a second wetting stimulus to hatch [146]. This will ensure that in the event of sudden unfavourable conditions, such as cold temperatures or a dry spell, following the oviposition of the egg batch, the entire batch is not lost [34]. Another explanation could be that not all larvae hatch simultaneously because of competition for space and resources as noted by Livdahl et al. [147] for Ae. triseriatus. Furthermore, Ebrahimi et al. [148] showed that the eggs of An. gambiae embryos are not stimulated to hatch when the water surface is agitated, demonstrating that environmental factors could indicate the best time for hatching. Sota & Mogi [68] suggested that intraspecific variation in the survival time of eggs is an inherited trait dependent on environmental pressures. Variations in the length of quiescence of eggs and variable hatching rates may be mechanisms that Ae. aegypti employs to produce continuous, although fluctuating, populations of adults in the wild at various stages, depending on the existence of favourable or unfavourable environmental conditions [23].

Therefore, quiescence provides a high adaptive potential to Ae. aegypti and Ae. albopictus populations, increasing the viability of their eggs and the chances of surviving in nature [148, 149]. This trait has contributed to the geographical expansion of these two species at a global level, an issue that is closely related to the spread of diseases [122].

Conclusions

As presented in this review, dormancy, especially diapause and quiescence, has a significant impact on the life history of mosquitoes, as well as of many other arthropods. Dormancy is part of the life history of many mosquito species, providing a mechanism to overcome unfavorable seasons in tropical and temperate zones. This trait may have independently evolved several times in the family Culicidae, as the phenomenon occurs at various developmental stages in different species. These adaptive strategies provide, on an evolutionary scale, mechanisms for species survival, as offspring continue to be produced, even when exposed to the various types of stress found in a habitat, and this, in turn, contributes to the territorial expansion of natural populations, consequently increasing their invasive potential. Diapause and quiescence are not the same biological phenomenon but have been treated as synonymous in previous studies. In addition, these different types of dormancy likely aid the propagation of the transmission cycles of diseases caused by different types of arboviruses, as these etiological agents can be transferred via the transovarian route. Both of these biological phenomena could play important roles in the ecology and evolution of many insect species, such as, for example, the mosquito Ae. albopictus, which has both phenotypes. Thus, the phenotypic plasticity generated by these intrinsic characteristics results in the reproductive success and survival of mosquitoes in the face of adverse environmental conditions and the different control measures practised by humans. These mechanisms are also fundamental for adapting to more frequent changes in climate. These phenomena are possibly still developing and need to be more thoroughly studied, as the information generated from associated research may be applied to innovative control strategies.

Abbreviations

AgCHS: 

Chitin synthase of Anopheles gambiae

CHS: 

Chitin synthase

cox

Cytochrome oxidase

EDR: 

Embryonic desiccation resistance

GPCR: 

G protein-coupled receptor

ing1

Inhibitor of growth protein

JH: 

Juvenile hormone

MMD: 

Mitochondrial malate dehydrogenase

pepck

Phosphoenolpyruvate carboxykinase

rack1

Receptor for activated protein kinase C

Declarations

Acknowledgements

Not applicable.

Funding

Not applicable.

Availability of data and materials

Not applicable.

Authors’ contributions

DFAD, MAVMS and CFJA designed this study; DFAD conducted the review and wrote the manuscript; CMRA and LOO participated in writing the quiescence topic; and CFJA and MAVMS reviewed the manuscript. All authors read and approved the final manuscript.

Competing interests

The authors declare that they have no competing interests.

Consent for publication

Not applicable.

Ethics approval and consent to participate

Not applicable.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

(1)
Entomology Department, Aggeu Magalhães Institute, Oswaldo Cruz Foundation
(2)
Zoology Department, Federal University of Pernambuco

References

  1. Rueda LM. Global diversity of mosquitoes (Insecta: Diptera: Culicidae) in freshwater. Hydrobiologia. 2008;595:477–87.View ArticleGoogle Scholar
  2. Lozovel AL. Culicidae (Mosquitos). In: Marcondes CB, editor. Entomologia médica e veterinária. Brasil: Atheneu; 2011. p. 107–74.Google Scholar
  3. Gubler DJ. Resurgent vector-borne diseases as a global health problem. Emerg Infect Dis. 1998;4:442–50.PubMedPubMed CentralView ArticleGoogle Scholar
  4. Lounibos L. Invasions by insect vectors of human disease. Annu Rev Entomol. 2002;47:233–66.PubMedView ArticleGoogle Scholar
  5. Service M. Mosquito (Diptera:Culicidae) dispersal the long and short of it. Entomol Soc Am. 1997;34:579–88.Google Scholar
  6. Kraemer MUG, Sinka ME, Duda KA, Mylne AQN, Shearer FM, Barker CM, et al. The global distribution of the arbovirus vectors Aedes aegypti and Aedes albopictus. elife. 2015;4:1–18.View ArticleGoogle Scholar
  7. Almeida PG. Mosquitoes (Diptera, Culicidae) and their medical importance for Portugal: challenges for the 21st century. Acta Medica Port. 2011;24:961–74.Google Scholar
  8. Consoli RAGB, Lourenço de Oliveira R. Principais mosquitos de importância sanitária no Brasil. Editora FIOCRUZ; 1994.Google Scholar
  9. Goddard J. Physician ‘s guide 5th edition the public health image library. Emerg Infect Dis. 2007;13:2007.View ArticleGoogle Scholar
  10. Harwood F, James MT. Entomology in human and animal health. New York: Macmillan Publ. Co. Inc.; 1979.Google Scholar
  11. National Center for Infectious Diseases Centers for Disease. 2007. Available from: https://www.cdc.gov/dengue/entomologyecology/index.html. Accessed 21 Sept 2016.
  12. Wilke ABB, Gomes A de C, Natal D, Marrelli MT. Control of vector populations using genetically modifi ed mosquitoes. Rev Lit Arts Am. 2009;43:869–74.Google Scholar
  13. Donalisio MR, Freitas ARR. Chikungunya no Brasil: um desafio emergente. Rev Bras Epidemiol. 2015;18:283–5.PubMedView ArticleGoogle Scholar
  14. Diallo D, Sall AA, Diagne CT, Faye O, Faye O, Ba Y, et al. Zika virus emergence in mosquitoes in southeastern Senegal, 2011. PLoS One. 2014;9:4–11.View ArticleGoogle Scholar
  15. World Health Organisation. The European health report 2005: public health action for healthier children and populations. World Heal Organ; 2005. Available from: http://www.euro.who.int/__data/assets/pdf_file/0020/98300/E87399.pdf. Accessed 1 Dec 2016.
  16. Poopathi S, Tyagi B. The challenge of mosquito control strategies: from primordial to molecular approaches. Biotech Mol Bio Rev. 2006;1:51–65.Google Scholar
  17. Danks HV. The range of insect dormancy responses. Eur J Entomol. 2002;99:127–42.View ArticleGoogle Scholar
  18. Gordon G, Headrick DH. A dictionary of entomology. Oxford: CABI Publ series; 2001.Google Scholar
  19. Martins R, Lewinsohn T, Barbeiros M. Ecologia e comportamento dos insetos. Oecologia Bras. 2000;8:149–92.View ArticleGoogle Scholar
  20. Danks HV. Insect dormancy: an ecological perspective (Biological Survey of Canada Monograph Series, No 1 ). Broadway: Entomological Society of Canada; 1987.Google Scholar
  21. Duclaux E. De 1 “influence de froid d”hiver sur Iedevelopment de 1′ embryon du ver à sole et surI’éclosion de la graine. Oompt Eend Acad Sci. 1869;69:1021–2.Google Scholar
  22. Christophers R. Aedes aegypti, the yellow fever mosquito. 1st ed. London: Cambridge University Press; 1960.Google Scholar
  23. Silva HH, Silva IG. Influência do Período de Quiescência dos Ovos Sobre o Ciclo de Vida de Aedes aegypti (Linnaeus, 1762) (Diptera, Culicidae) em Condições de Laboratório. Rev Soc Bras Med Trop. 1999;32:349–55.PubMedView ArticleGoogle Scholar
  24. Soares-Pinheiro VC, Dasso-Pinheirob W, Trindade-Bezerrac JM, Tadeib WP. Eggs viability of Aedes aegypti Linnaeus (Diptera, Culicidae ) under different environmental and storage conditions in Manaus, Amazonas, Brazil. Braz J Biol. 2016;doi: dx.doi.org/10.1590/1519-6984.19815.Google Scholar
  25. Luz C, Tai MHH, Santos AH, Silva HHG. Impact of moisture on survival of Aedes aegypti eggs and ovicidal activity of Metarhizium anisopliae under laboratory conditions. Mem Inst Oswaldo Cruz. 2008;103:214–5.PubMedView ArticleGoogle Scholar
  26. Choi MY, Estep A, Sanscrainte N, Becnel J, Vander Meer RK. Identification and expression of PBAN/diapause hormone and GPCRs from Aedes aegypti. Mol Cell Endocrinol Elsevier Ireland Ltd. 2013;375:113–20.View ArticleGoogle Scholar
  27. Silva HHG Da, Silva lonizete G da, Lira KDS, lonizete Garcia da Silva. Metodologia de criação, manutenção de adultos e estocagem de ovos de Aedes aegypti (Linnaeus,1762) em laboratório. Rev Patol Trop 1998;27:53–63.Google Scholar
  28. Clements AN, Kerkut GA. The physiology of mosquitoes: international series of monographs on pure and applied biology. Oxford: Pergamon Press. 1963;17:410.Google Scholar
  29. Denlinger DL, Armbruster PA. Mosquito diapause. Annu Rev Entomol. 2014;59:73–93.PubMedView ArticleGoogle Scholar
  30. Mirriam G, Mirriam C. Collegiate dictionary. 20th ed. Encyclopaedia Britannica, Inc: Massachusetts; 2000.Google Scholar
  31. Tauber MJ, Tauber CA, Masaki S. Seasonal adaptations of insects. Oxford: Oxford University Press; 1986.Google Scholar
  32. Lees AD. The Physiology of diapause in arthropods. 1955;4:150.Google Scholar
  33. Danilevskii AS. Photoperiodism and seasonal development of insects. Minnesota: Leningr Univ Press; 1965.Google Scholar
  34. Vinogradova EB. Diapause in aquatic insects, with emphasis on mosquitoes. Monogr Biol. 2007;84:83–113.Google Scholar
  35. Coluzzi M, Di Deco M, Gironi A. The influence of photoperiod on the selection of oviposition sites in Aedes mariae (Diptera: Culicidae). Parassitologia. 1975;17:121–30.PubMedGoogle Scholar
  36. Jordan RG, Bradshaw WE. Geographic variation in the photoperiodic response of the western tree-hole mosquito, Aedes sierrensis. Ann Entomol Soc Am. 1978;71:787.View ArticleGoogle Scholar
  37. Hanson SM, Craig GB. Cold acclimation, diapause, and geographic origin affect cold hardiness in eggs of Aedes albopictus (Diptera: Culicidae). J Med Entomol. 1994;31:192–201.PubMedView ArticleGoogle Scholar
  38. Eldridge BF, Johnson MD, Bailey CL. Comparative studies of two North American mosquito species, Culex restuans and Culex salinarius: response to temperature and photoperiod in the laboratory. Mosq News. 1976;36:506–13.Google Scholar
  39. Eldridge B. The effect of temperature and photoperiod on blood ­ feeding and ovarian development in mosquitoes of the Culex pipiens Complex. Am J Trop Med Hyg. 1968;17:1968.View ArticleGoogle Scholar
  40. Spielman A, Wong J. Environmental control of ovarian diapause in Culex pipiens. Ann Entomol Soc Amer. 1973;66:905–7.View ArticleGoogle Scholar
  41. Anderson JF. Influence of photoperiod and temperature on the induction of diapause in Aedes atropalpus (Diptera, Culicidae). Entomol Exp Appl. 1968;11:321–30.View ArticleGoogle Scholar
  42. Pumpuni CB, Knepler JCG. Influence of temperature and larval nutrition on the diapause inducing photoperiod of Aedes albopictus. J Am Mosq Control Assoc. 1992;8:223–7.PubMedGoogle Scholar
  43. Anderson JF. Induction and termination of embryonic diapause in the Salt Marsh mosquito, Aedes sollicitans (Diptera: Culicidae). Bulletin of the Connecticut Agricultural Experiment Station. 1970. Available: http://www.ct.gov/caes/lib/caes/documents/publications/bulletins/b711.pdf. Accessed 16 Jun 2017.
  44. Parker BM. Effects of photoperiod on the induction of embryonic diapause in Aedes taeniorhynchus (Diptera: Culicidae). J Med Entomol. 1985;22:392–7.PubMedView ArticleGoogle Scholar
  45. Wang R-L. Observations on the influence of photoperiod on egg diapause in Aedes albopictus (Skuse). Acta Enomol Sin. 1966; 15:75-77.Google Scholar
  46. Mori A, Oda T, Wada Y. Studies on the egg diapause and overwintering of Aedes albopictus in Nagasaki. Trop Med. 1981;23:79–90.Google Scholar
  47. Shinkawa Y, Takeda SI, Tomioka K, Matsumoto A, Oda T, Chiba Y. Variability in circadian activity patterns within the Culex-Pipiens complex (Diptera, Culicidae). J Med Entomol. 1994;31:49–56.PubMedView ArticleGoogle Scholar
  48. Poelchau MF, Reynolds JA, Denlinger DL, Elsik CG, Armbruster PA. A de novo transcriptome of the Asian tiger mosquito, Aedes albopictus, to identify candidate transcripts for diapause preparation. BMC Genomics. 2011;12:619.PubMedPubMed CentralView ArticleGoogle Scholar
  49. Poelchau M, Reynolds J, Elsik C, Denlinger D, Armbruster P. RNA-Seq reveals early distinctions and late convergence of gene expression between diapause and quiescence in the Asian tiger mosquito, Aedes albopictus. J Exp Biol. 2013;216:4082–90.PubMedPubMed CentralView ArticleGoogle Scholar
  50. Faraji A, Gaugler R. Experimental host preference of diapause and non-diapause induced Culex pipiens pipiens (Diptera: Culicidae). Parasit Vectors. 2015;8:389.PubMedPubMed CentralView ArticleGoogle Scholar
  51. Koštál V. Eco-physiological phases of insect diapause. J Insect Physiol. 2006;52:113–27.PubMedView ArticleGoogle Scholar
  52. Denlinger DL. Regulation of diapause. Annu Rev Entomol. 2002;47:93–122.PubMedView ArticleGoogle Scholar
  53. Zhang Q, Denlinger D. Expression analysis of the prothoracicotropic hormone gene in the northern house mosquito, Culex pipiens, in association with diapause and blood feeding. Insect Mol Biol. 2011;20:201–13.PubMedView ArticleGoogle Scholar
  54. Sim C, Denlinger DL. Insulin signaling and FOXO regulate the overwintering diapause of the mosquito Culex pipiens. Proc Natl Acad Sci U S A. 2008;105:6777–81.PubMedPubMed CentralView ArticleGoogle Scholar
  55. Robich RM, Denlinger DL. Diapause in the mosquito Culex pipiens evokes a metabolic switch from blood feeding to sugar gluttony. Proc Natl Acad Sci U S A. 2005;102:15912–7.PubMedPubMed CentralView ArticleGoogle Scholar
  56. Reynolds JA, Poelchau MF, Rahman Z, Armbruster PA, Denlinger DL. Transcript profiling reveals mechanisms for lipid conservation during diapause in the mosquito, Aedes albopictus. J Insect Physiol. 2013;58:966–73.View ArticleGoogle Scholar
  57. Goto S, Shiga S, Numata H. Photoperiodism in insects: perception of light and the role of clock genes. Oxford: Oxford University Press; 2010. p. 258–86.Google Scholar
  58. Koštál V. Insect photoperiodic calendar and circadian clock: independence, cooperation, or unity? J Insect Physiol. 2011;57(5):538–56.PubMedView ArticleGoogle Scholar
  59. Gentile C, Rivas GBS, Meireles-filho ACA, Peixoto AA. Circadian expression of clock genes in two mosquito disease vectors: cry2 is different. J Biol Rhythm. 2009;24:444–51.View ArticleGoogle Scholar
  60. Ptitsyn AA, Reyes-solis G, Saavedra-rodriguez K, Betz J, Suchman EL, Carlson JO, et al. Rhythms and synchronization patterns in gene expression in the Aedes aegypti mosquito. BMC Genomics. 2011;12:1–16.View ArticleGoogle Scholar
  61. Summa K, Urbanski JM, Zhao X, Poelchau MF, Armbruster P. Cloning and sequence analysis of the circadian clock genes period and timeless in Aedes albopictus (Diptera: Culicidae). J Med Entomol. 2012;49:777–82.PubMedView ArticleGoogle Scholar
  62. Mathias D, Jacky L, Bradshaw WE, Holzapfel CM. Geographic and developmental variation in expression of the circadian rhythm gene, timeless, in the pitcher-plant mosquito, Wyeomyia smithii. J Insect Physiol. 2005;51:661–7.PubMedView ArticleGoogle Scholar
  63. Rund SSC, Hou TY, Ward SM, Collins FH, Duffield GE. Genome-wide profiling of diel and circadian gene expression in the malaria vector Anopheles gambiae. Proc Natl Acad Sci U S A. 2011;108:E421–30.PubMedPubMed CentralView ArticleGoogle Scholar
  64. Leming MT, Rund SSC, Behura SK, Duffield GE, O’Tousa JE. A database of circadian and diel rhythmic gene expression in the yellow fever mosquito Aedes aegypti. BMC Genomics. 2014;15:1128.PubMedPubMed CentralView ArticleGoogle Scholar
  65. Meuti ME, Stone M, Ikeno T, Denlinger DL. Functional circadian clock genes are essential for the overwintering diapause of the northern house mosquito, Culex pipiens. J Exp Biol. 2015;218:412–22.PubMedPubMed CentralView ArticleGoogle Scholar
  66. Benoit JB, Denlinger DL. Suppression of water loss during adult diapause in the northern house mosquito, Culex pipiens. J Exp Biol. 2007;210:217–26.PubMedView ArticleGoogle Scholar
  67. Hanson SM, Craig GB. Aedes albopictus (Diptera: Culicidae) eggs: field survivorship during northern indiana winters. J Med Entomol. 1995;32:599–604.PubMedView ArticleGoogle Scholar
  68. Sota T, Mogi M. Survival time and resistance to desiccation of diapause and non-diapause eggs of temperate Aedes (Stegomyia) mosquitoes. Entomol Exp Appl. 1992;63:155–61.View ArticleGoogle Scholar
  69. Evans KW, Brust RA. Induction and termination of diapause in Wyeomyia smithii (Diptera: Culicidae), and larval survival studies at low and subzero temperatures. Can Entomol. 1972;104:1937–50.View ArticleGoogle Scholar
  70. Rinehart JP, Li A, Yocum GD, Robich RM, Hayward SAL, Denlinger DL. Up-regulation of heat shock proteins is essential for cold survival during insect diapause. Proc Natl Acad Sci U S A. 2007;104:11130–7.PubMedPubMed CentralView ArticleGoogle Scholar
  71. Gray EM, Bradley TJ. Physiology of desiccation resistance in Anopheles gambiae and Anopheles arabiensis. Am J Trop Med Hyg. 2005;73:553–9.PubMedGoogle Scholar
  72. Lounibos LP, Bradshaw WE. A second diapause in Wyeomyia smithii: seasonal incidence and maintenance by photoperiod. Can J Zool. 1975;53:215–21.PubMedView ArticleGoogle Scholar
  73. Lacour G, Chanaud L, Ambert GL, Hance T. Seasonal synchronization of diapause phases in Aedes albopictus (Diptera : Culicidae ). PLoS One. 2015;10(12):1–16.View ArticleGoogle Scholar
  74. Clements A. The biology of mosquitoes: development, nutrition and reproduction. Florida: Chapman Hall; 1992.Google Scholar
  75. Vinogradova E. [Diapause of blood - sucking mosquitoes and its control.] Leningrad. Russia: Nauka Academic Publishers; 1969. (In Russian)Google Scholar
  76. Abdel-Rahman A, Adham F. The effect of photoperiod on diapause induction in Aedes caspius Pallus. J Egypt Soc Parasitol. 1983;13:343–7.PubMedGoogle Scholar
  77. Sims S, Munstermann L. Egg and larval diapause in two populations of Aedes geniculatus (Diptera: Culicidae). J Med Entomol. 1983;3:263–71.View ArticleGoogle Scholar
  78. Gallaway W. Larval diapause of Aedes hendersoni and Aedes trisereatus from southern Manitoba. J Am Mosq Contr Assoc. 1985;1:92–3.Google Scholar
  79. Poelchau MF, Huang X, Goff A, Reynolds J, Armbruster P. An experimental and bioinformatics protocol for RNA-seq analyses of photoperiodic diapause in the Asian Tiger mosquito, Aedes albopictus. J Vis Exp. 2014:1–13.Google Scholar
  80. Urbanski J, Aruda A, Armbruster P. A transcriptional element of the diapause program in the Asian tiger mosquito, Aedes albopictus, identified by suppressive subtractive hybridization. J Insect Physiol. 2010;56:1147–54.PubMedView ArticleGoogle Scholar
  81. Denlinger D, Yocum G, Rinehart J. Hormonal control of diapause. insect endocrinol. San Diego: Gilbert, ed. LI 2012;430–463.Google Scholar
  82. Readio J, Chen MH, Meola R. Juvenile hormone biosynthesis in diapausing and nondiapausing Culex pipiens (Diptera: Culicidae). J Med Entomol. 1999;36:355–60.PubMedView ArticleGoogle Scholar
  83. Spielman A. Effect of synthetic juvenile hormone on ovarian diapause of Culex pipiens mosquitoes. J Med Entomol. 1974;11:223–5.PubMedView ArticleGoogle Scholar
  84. Pener MP. Environmental cues, endocrine factors, and reproductive diapause in male insects. Chronobiol Int. 1992;9:102–13.PubMedView ArticleGoogle Scholar
  85. Vinogradova E. [An experimental investigation of ecological factors inducing imaginal diapause in blood-sucking mosquitoes (Diptera: Culicidae).] Entomol Obozr. 1960;39:327–340. (In Russian).Google Scholar
  86. Hudson J. Overwintering sites and ovarian development of some mosquitoes in central Alberta. Canada Mosq News. 1978;38:570–379.Google Scholar
  87. Depner K, Harwood R. Photoperiodic responses of two latitudinally diverse groups of Anopheles freeborni (Diptera: Culicidae). Ann Entomol Soc Amer. 1966;59:7–11.View ArticleGoogle Scholar
  88. Lehmann T, Dao A, Yaro A, Adamou A, Kassogue T, Al. E. Aestivation of the African malaria mosquito, Anopheles gambiae in the Sahel. Am J Trop Med Hyg. 2010;83:601–606.Google Scholar
  89. Washino R, Bailey S. Overwintering of Anopheles punctipennis (Diptera:Culicidae) in California. J Med Entomol. 1970;7:95–8.PubMedView ArticleGoogle Scholar
  90. Harwood R, Halfhill E. The effect of photoperiod on fat body and ovarian development of Culex tarsalis (Diptera: Culicidae). Ann Entomol Soc Amer. 1964;57:596–600.View ArticleGoogle Scholar
  91. Kawai S. Studies on the follicular development and feeding activity of the females of Culex tritaeniorhynchus with special reference to those in autumn. Trop Med. 1969;11(3):145–69.Google Scholar
  92. Frohne W. Biology of an Alaskan mosquito, Culiseta alaskaensis (Ludl.). Ann Entomol Soc Amer. 1954;47:9–24.View ArticleGoogle Scholar
  93. Frohne W. Natual history of Culiseta impatiens (Wlk.), (Diptera, Culicidae) in Alaska. Trans Amer Micr Sci. 1953;72:103–18.View ArticleGoogle Scholar
  94. Hudson J. Induction of diapause in female mosquitoes, Culiseta inornata by a decrease in day length. J Insect Physiol. 1977;23:1377–82.View ArticleGoogle Scholar
  95. Emerson K, Uyemura A, Mcdaniel K, Schmidt P, Bradshaw W, Holzapfel C. Environmental control of ovarian dormancy in natural populations of Drosophila melanogaster. J Comp Physiol. 2009;195:825–329.View ArticleGoogle Scholar
  96. Saunders D. Insect clocks. Amsterdam: Elsevier; 2002.Google Scholar
  97. Schmidt P, Paaby A, Heschel M. Genetic variance for diapause expression and associated life histories in Drosophila melanogaster. Evolution (NY). 2005;59:2616–2625.Google Scholar
  98. Wilton DP, Smith GC. Ovarian diapause in three geographic strains of Culex pipiens (Diptera: Culicidae). J Med Entomol. 1985;22:524–8.PubMedView ArticleGoogle Scholar
  99. Mori A, Romero-Severson J, Severson D. Genetic basis for reproductive diapause is correlated with life history traits within the Culex pipiens Complex. Insect Mol Biol. 2007;16:515–24.PubMedGoogle Scholar
  100. Robich R, Rinehart J, Kitchen L, Denlinger D. Diapause­specific gene expression in the northern house mosquito, Culex pipiens L., identified by suppressive subtractive. J Insect Physiol. 2007;53:235–45.PubMedView ArticleGoogle Scholar
  101. Feder J, Rossi J, Solomon J, Solomon N, Lindquist S. The consequences of expressing hsp70 in Drosophila cells at normal temperatures. Genes Dev. 1992;6:1402–13.PubMedView ArticleGoogle Scholar
  102. Poelchau M, Reynolds J, Elsik C, Denlinger D, Armbruster P. Deep sequencing reveals complex mechanisms of diapause preparation in the invasive mosquito, Aedes albopictus. Proc Biol Sci. 2013;280(1759):20130143.PubMedPubMed CentralView ArticleGoogle Scholar
  103. Lunardi A, Di Minin G, Provero P, Dal Ferro M, Carotti M, Del Sal G, et al. A genome-scale protein interaction profile of Drosophila p53 uncovers additional nodes of the human p53 network. Proc Natl Acad Sci U S A. 2010;107:6322–7.PubMedPubMed CentralView ArticleGoogle Scholar
  104. Kadrmas JL, Smith MA, Pronovost SM, Beckerle MC. Characterization of RACK1 function in Drosophila development. Dev Dyn. 2007;236:2207–15.PubMedView ArticleGoogle Scholar
  105. Quan GX, Krell PJ, Arif BM, Feng Q. Receptor of activated C kinase 1 (RACK1) is necessary for the 20-hydroxyecdysone-induced expression of the transcription factor CHR3 in the spruce budworm Choristoneura fumiferana. Insect Mol Biol. 2006;15:79–87.PubMedView ArticleGoogle Scholar
  106. Ragland GJ, Denlinger DL, Hahn DA. Mechanisms of suspended animation are revealed by transcript profiling of diapause in the flesh fly. Proc Natl Acad Sci U S A. 2010;107:14909–14.PubMedPubMed CentralView ArticleGoogle Scholar
  107. Hahn DA, Denlinger DL. Energetics of insect diapause. Annu Rev Entomol. 2011;56:103–21.PubMedView ArticleGoogle Scholar
  108. Lin Y, Seroude L, Benzer S. Extended life ­ span and stress resistance in the Drosophila mutant. Science. 1998;282(5390):943–6.PubMedView ArticleGoogle Scholar
  109. Hand SC, Podrabsky JE. Bioenergetics of diapause and quiescence in aquatic animals. Thermochim Acta. 2000;349:31–42.View ArticleGoogle Scholar
  110. Juliano S, O’Meara G, Morrill J, Cutwa M. Desiccation and thermal tolerance of eggs and the coexistence of competing mosquitoes. Oecologia. 2002;130:458–69.Google Scholar
  111. Thaler CD, Miyata H, Haimo LT, Cardullo RA. Waveform generation is controlled by phosphorylation and swimming direction is controlled by Ca2+ in sperm from the mosquito Culex quinquefasciatus. Biol Reprod. 2013;89:135.PubMedView ArticleGoogle Scholar
  112. Nelms BM, Macedo PA, Kothera L, Savage HM, Reisen WK. Overwintering biology of Culex (Diptera: Culicidae) mosquitoes in the Sacramento Valley of California. J Med Entomol. 2013;50:773–90.PubMedPubMed CentralView ArticleGoogle Scholar
  113. Reisen WK, Meyer RP, Milby MM. Overwintering studies on Culex tarsalis (Diptera: Culicidae) in Kern County, California: temporal changes in abundance and reproductive status with comparative observations on C. quinquefasciatus (Diptera: Culicidae). Ann Entomol Soc Am. 1986;79:677–85.View ArticleGoogle Scholar
  114. Almeida E. Fecundidade, fertilidade e quiescência dos ovos de Aedes aegypti Linnaeus, 1762 (Diptera: Culicidae) em resposta a variações de temperatura e umidade. UFPE; 2005. Available from: http://repositorio.ufpe.br/handle/123456789/808. Accessed 21 Feb 2017.
  115. Diniz DFA, Melo-Santos MAV de, Santos EM de M, Beserra EB, Helvecio E, Carvalho-Leandro D, et al. Fitness cost in field and laboratory Aedes aegypti populations associated with resistance to the insecticide temephos. Parasit Vectors 2015;8:662.Google Scholar
  116. Rezende GL, Martins AJ, Gentile C, Farnesi LC, Pelajo-Machado M, Peixoto AA, et al. Embryonic desiccation resistance in Aedes aegypti: presumptive role of the chitinized serosal cuticle. BMC Dev Biol. 2008;8:82.PubMedPubMed CentralView ArticleGoogle Scholar
  117. Juliano SA, O’Meara GF, Morrill JR, Cutwa MM. Desiccation and thermal tolerance of eggs and the coexistence of competing mosquitoes. Oecologia. 2002;130:458–69.PubMedPubMed CentralView ArticleGoogle Scholar
  118. McMeniman CJ, O’Neill SL. A virulent wolbachia infection decreases the viability of the dengue vector Aedes aegypti during periods of embryonic quiescence. PLoS Negl Trop Dis. 2010;4Google Scholar
  119. Perez MH, Noriega FG. Aedes aegypti Pharate 1st instar quiescence affects larval fitness and metal tolerance. J Insect Physiol. 2012;58:824–9.PubMedPubMed CentralView ArticleGoogle Scholar
  120. Perez MH, Noriega FG. Aedes aegypti Pharate 1st instar quiescence: a case for anticipatory reproductive plasticity. J Insect Physiol. 2013;59:318–24.PubMedPubMed CentralView ArticleGoogle Scholar
  121. Vargas HCM, Farnesi LC, Martins AJ, Valle D, Rezende GL. Serosal cuticle formation and distinct degrees of desiccation resistance in embryos of the mosquito vectors Aedes aegypti, Anopheles aquasalis and Culex quinquefasciatus. J Insect Physiol. 2014;62:54–60.PubMedView ArticleGoogle Scholar
  122. Yang HM. Assessing the influence of quiescence eggs on the Ddynamics of Mmosquito Aedes aegypti. Appl Math. 2014;5:2696–711.View ArticleGoogle Scholar
  123. Farnesi LC, Menna-Barreto RFS, Martins AJ, Valle D, Rezende GL. Physical features and chitin content of eggs from the mosquito vectors Aedes aegypti, Anopheles aquasalis and Culex quinquefasciatus: connection with distinct levels of resistance to desiccation. J Insect Physiol Elsevier Ltd. 2015;83:43–52.View ArticleGoogle Scholar
  124. Goltsev Y, Rezende GL, Vranizan K, Lanzaro G, Valle D, Levine M. Developmental and evolutionary basis for drought tolerance of the Anopheles gambiae embryo. Dev Biol. 2009;330:462–70.PubMedPubMed CentralView ArticleGoogle Scholar
  125. Urbanski JM, Benoit JB, Michaud MR, Denlinger DL, Armbruster P. The molecular physiology of increased egg desiccation resistance during diapause in the invasive mosquito, Aedes albopictus. Proc R Soc B Biol Sci. 2010;277:2683–92.View ArticleGoogle Scholar
  126. Chapman RF, Reginald F, Simpson SJ, Douglas AE. The insects: structure and function. 5th ed. Cambridge: Cambridge University Press; 2013.Google Scholar
  127. Teixeira MDG, Barreto ML, Guerra Z. Epidemiologia e medidas de prevenção do Dengue. Inf Epidemiol SUS. 1999;8:5–33.Google Scholar
  128. Pinheiro VCS, Tadei WP. Frequency, diversity, and productivity study on the Aedes aegypti most preferred containers in the city of Manaus, Amazonas, Brazil. Rev Inst Med Trop Sao Paulo. 2002;44:245–50.PubMedView ArticleGoogle Scholar
  129. Chadee DD, Corbet PS, Talbot H. Proportions of eggs laid by Aedes aegypti on different substrates within an ovitrap in Trinidad, West Indies. Med Vet Entomol. 1995;9:66–70.PubMedView ArticleGoogle Scholar
  130. Trpis M. Seasonal changes in the larval populations of Aedes aegypti in two biotopes in Dar es salaam, Tanzania. Bull World Health Organ. 1972;47:245–55.PubMedPubMed CentralGoogle Scholar
  131. Schaffner F, Kaufmann C, Pflüger V, Mathis A. Rapid protein profiling facilitates surveillance of invasive mosquito species. Parasit Vectors. 2014;7:142.PubMedPubMed CentralView ArticleGoogle Scholar
  132. Li JS, Li J. Major chorion proteins and their crosslinking during chorion hardening in Aedes aegypti mosquitoes. Insect Biochem Mol Biol. 2006;36:954–64.PubMedPubMed CentralView ArticleGoogle Scholar
  133. Faull KJ, Williams CR. Intraspecific variation in desiccation survival time of Aedes aegypti (L.) mosquito eggs of Australian origin. J Vector Ecol. 2015;40:292–300.PubMedView ArticleGoogle Scholar
  134. Merzendorfer H. Insect chitin synthases: a review. J Comp Physiol B. 2006;176:1–15.PubMedView ArticleGoogle Scholar
  135. Medlock JM, Hansford KM, Schaffner F, Versteirt V, Hendrickx G, Zeller H, et al. A review of the invasive mosquitoes in Europe: ecology, public health risks, and control options. Vector-Borne Zoonotic Dis. 2012;12:435–47.PubMedPubMed CentralView ArticleGoogle Scholar
  136. Bajer E. Brock team finds first Aedes aegypti mosquitoes in Canada. The BrockNews. 2016. https://brocku.ca/brock-news/2016/10/brock-team-finds-first-aedes-aegypti-mosquitoes-in-canada/. Accessed 13 Jan 2017.
  137. Lima A, Lovin DD, Hickner PV, Severson DW. Evidence for an overwintering population of Aedes aegypti in Capitol Hill neighborhood, Washington, DC. Am J Trop Med Hyg. 2016;94:231–5.PubMedPubMed CentralView ArticleGoogle Scholar
  138. Schaffner F, Medlock JM, Bortel WV. Public health significance of invasive mosquitoes in Europe invasive mosquito species in Europe. Clin Microbiol Infect. 2013;19:685–92.PubMedView ArticleGoogle Scholar
  139. Rosen L. Sexual transmission of dengue viruses by Aedes albopictus. Am J Trop Med Hyg. 1987;37:398–402.PubMedView ArticleGoogle Scholar
  140. Buckner EA, Alto BW, Lounibos LP. Vertical transmission of key West dengue-1 virus by Aedes aegypti and Aedes albopictus (Diptera: Culicidae) mosquitoes from Florida. J Med Entomol. 2013;50:1291–7.PubMedPubMed CentralView ArticleGoogle Scholar
  141. Angel A, Angel B, Joshi V. Rare occurrence of natural transovarial transmission of dengue virus and elimination of infected foci as a possible intervention method. Acta Trop. 2016;155:20–4.PubMedView ArticleGoogle Scholar
  142. Thangamani S, Huang J, Hart CE, Guzman H, Tesh RB. Vertical transmission of Zika virus in Aedes aegypti mosquitoes. Am J Trop Med Hyg. 2016;95:1169–73.PubMedPubMed CentralView ArticleGoogle Scholar
  143. Di Luca M, Severini F, Toma L, Boccolini D, Romi R, Remoli ME, et al. Experimental studies of susceptibility of Italian Aedes albopictus to Zika virus. Eur Secur. 2016;21Google Scholar
  144. Ciota AT, Bialosuknia SM, Ehrbar DJ, Kramer LD. Vertical transmission of Zika virus by Aedes aegypti and Aedes albopictus mosquitoes. Emerg Infect Dis. 2017;23:28277199.Google Scholar
  145. Smartt, CT, Stenn, TMS, Chen, TY, Teixeira, MG, Erivaldo P.et al. Evidence of Zika virus RNA fragments in Aedes albopictus (Diptera: Culicidae) field collected eggs from Camaçari, Bahia, Brazil. J Med Ent. 2017. (In press).Google Scholar
  146. Livdahl TP, Koenekoop RK, Futterweit SG. The complex hatching response of Aedes eggs to larval density. Ecol Entomol. 1984;9:437–42.View ArticleGoogle Scholar
  147. Ebrahimi B, Shakibi S, Foster WA. Delayed egg hatching of Anopheles gambiae (Diptera: Culicidae) pending water agitation. J Med Entomol. 2014;51:580–90.PubMedPubMed CentralView ArticleGoogle Scholar
  148. Benedict MQ, Levine RS, Hawley WA, Lounibos LP. Spread of the tiger: Gglobal risk of invasion by the mosquito Aedes albopictus. Vector-Borne Zoonotic Dis. 2007;7:76–85.PubMedPubMed CentralView ArticleGoogle Scholar
  149. Powell JR, Tabachnick WJ. History of domestication and spread of Aedes aegypti-a review. Mem Inst Oswaldo Cruz. 2013;108:11–7.PubMedPubMed CentralView ArticleGoogle Scholar
  150. Tauthong P, Brust R. The effect of photoperiod on diapause induction, and temperature on diapause termination in embryos of Aedes campestris Dyar and Knab (Diptera:Culicidae). Can J Zool. 1977;55:129–34.PubMedView ArticleGoogle Scholar
  151. Pinger R, Eldrige B. The effect of photoperiod on diapause induction in Aedes canadensis and Psorophora ferox (Diptera: Culicidae). Ann Entomol Soc Amer. 1977;70:437–41.View ArticleGoogle Scholar
  152. Telford A. The pasture Aedes of central and northern California. Seas Hist Ann Entomol Soc Amer. 1958;51:360–5.View ArticleGoogle Scholar
  153. Horsfall W, Lum P, Henderson L. Eggs of floodwater mosquitoes (Diptera: Culicidae) V. Effect of oxygen on hatching of intact eggs. Ann Entomol Soc Amer. 1958;51:209–13.View ArticleGoogle Scholar
  154. Beckel W. Investigations of permeability, diapause, and hatching in eggs of the mosquito Aedes hexodontus Dyar. Can J Zool. 1958;36:541–55.View ArticleGoogle Scholar
  155. Andreadis T, Anderson J, Munstermann L, Wolfe R, Florin D. Discovery, distribution and abundance of the newly introduced mosquito Ochlerotatus japonicus (Diptera: Culicidae) in Connecticut, USA. J Med Entomol. 2001;38:774–9.PubMedView ArticleGoogle Scholar
  156. Corbet P, Danks H. Egg-laying habits of mosquitoes in the high arctic. Mosq News. 1975;35:8–14.Google Scholar
  157. Parker B. Photoperiod-induced diapause in a North Carolina strain of Aedes sollicitans; photosensitivity of fully formed and developing embryos. J Am Mosq Contr Assoc. 1988;4:57–63.Google Scholar
  158. Kappus K, Venard C. The effects of photoperiod and temperature on the induction of diapuase in Aedes triseriatus say. J Insect Physiol. 1967;13:1007–19.View ArticleGoogle Scholar
  159. Wilson G, Horsfall W. Eggs of floodwater mosquitoes XII. Installment hatching of Aedes vexans (Diptera: Culicidae). Ann Entomol Soc Amer. 1970;63:1644–7.View ArticleGoogle Scholar
  160. Hurlbut H. Further notes on the overwintering of the eggs of Anopheles walkeri with a description of the eggs. J Parasitol. 1938;24:521–6.View ArticleGoogle Scholar
  161. Baker F. The effect of photoperiodism on resting tree-hole mosquito larvae. Can Entomol. 1935;67:149–53.View ArticleGoogle Scholar
  162. Bradshaw W, Holzapfel C. Fitness and habitat segregation of British tree-hole mosquito communities. Ecol Entomol. 1991;16:133–44.View ArticleGoogle Scholar
  163. Zhang X, Huang K, Wang L, Peng X, Fu X, Liu H. A study on the ecology of Armigeres subalbatus in Dawa area of the Mengshan mountain in Shandong Province. Chin J Parasit Dis. 1992;10:48–51.Google Scholar
  164. Wallis RC. Sex difference in pupation of larval Culiseta melanura. J Am Mosq Control Assoc. 1988;4:192–4.PubMedGoogle Scholar
  165. Baker F. A new species of Orthopodomyia, O. alba sp. n. (Diptera, Culicidae). Proc Entomol Soc Wash. 1936;38:1–7.Google Scholar
  166. Tate P. The larval instars of Orthopodomyia pulchripes. Parasitology. 1932;24:111–20.View ArticleGoogle Scholar
  167. Bradshaw W. Photoperiodism in Orthopodomyia signifera. Can J Zool. 1973;51:355–7.PubMedView ArticleGoogle Scholar
  168. Bradshaw W, Lounibos L. Evolution of dormancy and its photoperiodic control in pitcher - plant mosquitoes. Evolution. 1977;31:546–67.PubMedView ArticleGoogle Scholar
  169. Bradshaw WE, Lounibos LP. Photoperiodic control of development in the pitcher-plant mosquito, Wyeomyia smithii. Can J Zool. 1972;50:713–9.View ArticleGoogle Scholar
  170. Alto BW, Juliano SA. Precipitation and temperature effects on populations of Aedes albopictus (Diptera: Culicidae): implications for range expansion. J Med Entomol. 2001;38:646–56.PubMedPubMed CentralView ArticleGoogle Scholar

Copyright

© The Author(s). 2017

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.

Advertisement