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First published online February 12, 2007
Journal of Experimental Biology 210, 854-864 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.02715
Larval feeding duration affects ecdysteroid levels and nutritional reserves regulating pupal commitment in the yellow fever mosquito Aedes aegypti (Diptera: Culicidae)
Department of Entomology and Center for Tropical and Emerging Global Diseases, University of Georgia, Athens, GA 30602, USA
* Author for correspondence at present address: University of Richmond, Department of Biology, Gottwald Science Center, Richmond, VA 23173, USA (e-mail: atelang{at}richmond.edu)
Accepted 11 January 2007
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Summary |
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Key words: larval nutrition, larval-pupal molt, metamorphosis, development, hormone
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Introduction |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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Data on the endocrine and non-endocrine mechanisms underlying insect
metamorphosis have been collected mainly from lepidopteran species, especially
the tobacco hornworm Manduca sexta and the silkmoth Bombyx
mori, and these data have come to form a central dogma of insect
endocrinology (Nijhout, 1994
):
larvae grow to a certain mass before molting to the next larval stage, and a
critical mass must be attained prior to committing to a metamorphic molt. Both
the larval and metamorphic molts are mediated by the release of a neuropeptide
from the larval brain, now considered to be prothoracicotropic hormone (PTTH),
which then stimulates the prothoracic glands (PG) to synthesize and release
ecdysteroids that cause the larva to stop feeding and commit to pupation.
However, the metamorphic molt only occurs when ecdysteroids are produced in
the absence of juvenile hormone (JH) during a critical period in the last
larval stage. If JH is present during the critical period, the insect molts
into another larval stage and retains larval characteristics. In the absence
of JH, the larva commits to its first metamorphic molt into its pupal
stage.
Using model insects such as M. sexta and Drosophila
melanogaster, progress has lead to an understand of the neural events
involved in the release of PTTH and the biochemical events of PTTH signal
transduction (Rybczynski,
2005). Likewise, the entire ecdysteroid biosynthetic pathway is
close to being characterized in larval insects
(Gilbert, 2004;
Gilbert et al., 2002).
Unfortunately, little more is known about what other than the `critical mass'
triggers the metamorphic molt. Nijhout stated that the `critical size is
merely a symptom of the underlying physiology and as such provides no
information on what factors are of importance to the animal in question, nor
is the critical size straightforwardly related to the size at which the molt
will ultimately occur' (Nijhout,
1981). So, what is the underlying physiological mechanism(s) and
what are the factors an animal uses to make molting decisions? Physical mass
may be a general parameter for the larva, but some correlate of mass, like
nutrient reserves, is surely relevant.
Past and recent studies implicate larval feeding and nutrient intake as
factors to cue metamorphic readiness. For example, nutrient intake and
accumulation of mass is important for two phases during the last larval instar
of M. sexta. In the first phase, nutritional input and growth allow a
larva to reach a critical mass for metamorphic commitment
(Nijhout, 2003). In the second
phase, after reaching its critical mass, additional larval growth occurs only
if food is available. If nutrient intake is reduced and the larva grows only a
little after meeting its critical mass, it will metamorphose at a smaller size
and subsequently form a smaller sized adult. This second larval growth phase
has major consequences for the individual during its adult stage because a
larger adult body size, especially for females, is often correlated with
greater reproductive success (Bronson,
1985; Iyengar and Eisner,
1999; Tu and Tatar,
2003). Additional evidence comes from studies in which M.
sexta were starved during the last larval stage, resulting in either
death or a delay in metamorphosis, depending on when in the last instar
starvation was initiated (Nijhout and
Williams, 1974b). Feeding the larvae after a period of starvation
usually result in a supernumery larval molt
(Bhaskaran and Jones, 1980;
Cymborowski et al., 1982;
Jones et al., 1980). This
delay in metamorphosis affords larvae an additional period of time to feed,
grow and meet a critical mass to commit to pupation. Recently, pupal
commitment by last instar M. sexta was shown to depend on sugar
intake, whereas the proliferation and growth of imaginal discs, cells destined
to be adult-specific structures, required both dietary sugar and protein
(MacWhinnie et al., 2005).
Juvenile animals ingest food not only for energy production and maintenance of
biochemical processes, but also to accumulate larval nutrient reserves.
Nutrient reserves acquired during the juvenile stage play a large role for the
adult toward its reproductive success
(Boggs, 1997;
Briegel, 1990a;
Briegel, 1990b;
O'Brien et al., 2000;
Rivero et al., 2001;
Telang et al., 2006;
Telang and Wells, 2004).
Threshold levels of larval nutrient stores are likely candidates that inform
the larva it is ready for metamorphosis. Recent studies using M.
sexta and D. melanogaster have further elaborated how larval
nutrition and hormones affect growth and metamorphosis in these model insects
(Colombani et al., 2005;
Truman et al., 2006).
Given the medical importance of mosquitoes, it is surprising how little is
known about the nutritional and hormonal regulation of their post-embryonic
development. What we do know indicates that endocrine regulation of molting
and metamorphosis in mosquitoes may differ from the model developed for
Lepidoptera and applied to higher dipterans. In Lepidoptera, the PG secretes
ecdysteroids, and in higher dipterans, such as the fleshfly
Neobellieria (formerly Sarcophaga) bullata and
D. melanogaster, the larval ring gland has the PG as one of its
components and secretes ecdysteroids
(Bollenbacher et al., 1976;
Dai and Gilbert, 1991;
Parvy et al., 2005;
Redfern, 1983). Contrary to
this classical view, the tissue identified as the PG complex in fourth (last)
instar Aedes aegypti does not release ecdysteroids but, instead,
tissues in the thorax and abdomen synthesize and release ecdysteroids in
vitro (Jenkins et al.,
1992). Although a few current studies have begun examining the
endocrine regulation of larval mosquito metamorphosis
(Lan and Grier, 2004;
Margam et al., 2006;
Nishiura et al., 2003), the
role of larval nutrition in regulating metamorphosis has not been
addressed.
In the present study, we have examined the development and metamorphic readiness of larval Aedes aegypti in response to larval nutrition. We manipulated the length of time that fourth instars had access to food and measured its effect on larval mass, nutritional reserves and profiles of ecdysteroid production and hemolymph titers. Our data suggest that the timing of ecdysteroid release is not necessarily critical to initiate the larval–pupal molt, but both the ecdysteroid titer and the nutritional status of the fourth instar are crucial factors in initiating this first metamorphic molt in mosquito larvae. These findings will allow us to eventually construct a conceptual model specific for the nutritional and hormonal regulation of mosquito post-embryonic development.
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Materials and methods |
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Larval staging
Under our standard colony rearing conditions, a majority of A.
aegypti larvae are late third instars 4 days post-hatching. For
experiments, in the late morning of day 4, late third instars were chosen
based on the presence of darkly tanned head capsules. Selected larvae were
then housed individually in 24-well cell culture plates with each well
containing 1.6 ml of distilled water. Third instar larvae were not given food
during this period. The next morning, newly molted fourth instars were
transferred individually to new wells containing fresh 1.6 ml distilled
water.
Measurement of growth
To monitor increase in mass when optimally fed, individual fourth instars
were fed ad libitum on a 2% (w/v) solution of A. aegypti
diet that was easily dispensed to individual larvae. In previous trials, this
feeding regimen supported the growth and metamorphosis of experimental larvae
at the same degree and schedule as that of larvae mass reared for the colony.
Wet mass of fourth instars was measured at different times during this stadium
(newly molted, 12, 24, 36 and 48 h) and in pupae (12 h after
larval–pupal ecdysis). Groups of three larvae were removed from
their feeding wells, rinsed in fresh water and blotted dry prior to weighing.
Each larval group was dried on a heat block at 90°C to determine dry mass.
Both fresh and dry masses were obtained using a microbalance (Mettler MT5,
Columbus, OH, USA. Three replicates were collected using different cohorts
(N=18).
Hemolymph ecdysteroid titer, in vitro ecdysteroid secretion and the ecdysteroid radioimmunoassay
To verify ecdysteroid release from both the thorax and abdomen
(Jenkins et al., 1992),
triplicate preparations of thorax and abdomen and hemolymph samples were
collected at different times during their fourth stadium (newly molted, 6, 12,
24, 30, 36, 48 and 52 h) and in pupae (12 h after larval–pupal ecdysis).
Dissections and tissue incubations were conducted in a buffered medium (139
mmol l–1 NaCl, 4.05 mmol l–1 KCl, 1.85 mmol
l–1 CaCl2, 12.5 mmol l–1 Hepes,
2.5 mmol l–1 trehalose, 0.3 mmol l–1
MgCl2 and 0.9 mmol l–1 NaHCO3; pH 6.5,
adjusted with NaOH) (Riehle and Brown,
1999) containing phenylthiourea (0.5 mg ml–1
buffered medium), added to minimize melanization. Hemolymph was collected from
the same set of larvae prior to body region separation for the in
vitro bioassay. To collect hemolymph, groups of four larvae were
decapitated and hemolymph was allowed to flow into 75 µl of buffered medium
with phenylthiourea, assisted by gentle pressure applied to each carcass.
After 5 min incubation, 50 µl of the hemolymph solution were collected and
assayed for ecdysteroids using a radioimmunoassay (RIA). Hemolymph ecdysteroid
titers are reported on a larva equivalent basis.
For isolation of body regions, the terminal abdominal segments (including
segment 8, the anal lobe, anal papillae and the respiratory siphon) were
discarded. Thoraces and abdomina were separated from each other to facilitate
removal of the alimentary canal from each section. Removal of PGs from
thoraces was not confirmed since it has already been determined that PGs are
not the source of ecdysteroids (Jenkins et
al., 1992). To measure the amounts of ecdysteroids released,
groups of four thoraces and four abdomina were incubated together in 60 µl
buffered medium with phenylthiourea in a 0.6 ml polypropylene tube lid at
27°C for 6 h. After incubation, 25 µl of medium was collected and
analyzed for ecdysteroid content using RIA. It was previously determined that
the amount of ecdysteroids secreted from body region incubations into 50 µl
of medium was beyond the linear range of the standard curve (see below), so
these samples were diluted by half to stay within the quantitation range
(10–250 pg). Ecdysteroid secretion by tissues is reported on a single
thorax and abdomen set basis.
The RIA uses an ecdysteroid antiserum (AS 4919, a gift from P. Poncheron,
Université P. et M. Curie, Paris, France) that recognizes ecdysone and
20-hydroxyecdysone equally (Porcheron et
al., 1989). Full details of the RIA were previously described
(Sieglaff et al., 2005).
Values for each tissue sample type are reported as `mean pg ecdysteroid',
because the secreted ecdysteroid species are unknown. For this experiment,
triplicates of four thorax plus abdomen preparations and four body hemolymph
collections were analyzed for all the time points. Values reported are means
of triplicate tissue preparations for larval age groups taken from one cohort
(N=27).
Critical feeding period and scoring of metamorphosis
Experimental fourth instars were allowed to feed on a 2% (w/v) solution of
A. aegypti standard diet for the following time periods only: 12, 18,
24, 30, 36, 42 or 48 h. After the allocated time period, larvae were
transferred into new wells containing fresh water only and scored for pupation
and adult eclosion. A control group consisted of larvae with access to food
for the entire fourth stadium. For both experimental and control animals, we
recorded duration of fourth instar development time (N=261), pupation
(N=335), and adult eclosion in response to food access period
(N=335). We did not determine the sex of larvae but recorded gender
of each individual at adult eclosion only. Lastly, wing length was used to
assess body size of females successfully emerging from each treatment and was
measured from the point of attachment to the wing tip, not including fringe,
under a dissecting microscope using an ocular micrometer (N=116).
Three replicates were set up using different cohorts.
For larvae that did not pupate, the duration of time they remained alive during their fasting period was monitored (N=49). To examine whether larvae retain their capacity to molt after a period of fasting, newly molted fourth instars were allowed to feed for 12 h, fasted for 7 days, and then given additional opportunities to feed for defined periods of time. Larvae were then scored for pupation and adult eclosion (N=140). Lastly, wing length was used to assess body size of females successfully emerging from each supplementary feeding treatment (N=37). Two replicates were examined using different cohorts.
Quantification of nutritional reserves
Groups of larvae, either newly molted fourth instars or larvae from the
three feeding regimens, were immediately frozen for analysis of nutritional
reserves. Dry mass was obtained by drying each larval group on a heat block at
90°C until no further water loss was detected. Whole-body homogenates of
four larvae per treatment were made to extract storage lipid, glycogen and
proteins using a published procedure (Van
Handel, 1965) modified for A. aegypti
(Zhou G. et al., 2004), except
that the isolation of amino acids and sugars was omitted. Fractions of lipid,
glycogen and protein were frozen until they could be quantified using
colormetric-based assays. The amount of storage lipid, triacylglycerol, was
determined by a modified vanillin reagent assay
(Van Handel, 1985b). Total
amount of glycogen was determined using a modified anthrone-based assay
(Van Handel, 1985a). Protein
was quantified using the BCA protein assay reagent kit (Pierce, Rockford, IL,
USA). Complete details of our assay procedures were previously described
(Telang and Wells, 2004). All
nutrients are reported on a per larva basis and three replicates of all
feeding regimens were collected (N=12).
Data analyses
Increase in wet and dry mass of larvae during their fourth stadium was
analyzed using analysis of variance (ANOVA). Both in vitro
ecdysteroid release by thoraces and abdomina and hemolymph titer during the
fourth larval stage and in pupae were analyzed by ANOVA. Likelihood of
pupation and successful male or female eclosion in response to periods of food
availability were examined using contingency analysis and the
2 statistic. Wing length was analyzed using ANOVA, with
periods of food availability as the explanatory variable. For the feeding
regimen experiment, amount of lipid, glycogen and protein per individual larva
was analyzed using analysis of covariance (ANCOVA), using larval dry mass as a
covariate. In vitro ecdysteroid release by thoraces and abdomina and
hemolymph titer for larval feeding regimen were analyzed by ANOVA. When
necessary, differences between means were further analyzed using the
Tukey–Kramer HSD test. All data were statistically analyzed using JMP IN
(version SAS Institute Inc.). Least square means (± s.e.m.) were
obtained from statistical models and used in all graphical illustrations.
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An in vitro bioassay was used to determine the capacity for
ecdysteroid production by thoraces and abdomina dissected from larvae at
different times during their fourth stadium and into their pupal stage. In
addition, hemolymph was collected from the same set of larvae prior to tissue
dissection for the in vitro bioassay. In an earlier study
(Jenkins et al., 1992),
ecdysteroid production by these tissues and hemolymph titers were measured
only for the first 36 h of the fourth instar, even though the duration of the
stadium was reported to be 36–52 h. In the present study we clarified
the ecdysteroid profile of the entire last larval and early pupal stage of
A. aegypti. Levels of ecdysteroid production by thorax and abdomen
preparations differed over the course of the fourth larval stage, including
the pupal stage (one-way ANOVA, P<0.0001)
(Fig. 2). Thoraces and abdomina
of newly molted fourth instars produced a detectable level of ecdysteroids
(Fig. 2). During the fourth
instar, this capacity increased over the period of 12–30 h to peak at
about 90 pg before falling to a basal level at 48 h (Tukey–Kramer HSD,
P
0.05). Ecdysteroid release by thoraces and abdomina again
increased in 12 h old pupae (Tukey–Kramer HSD, P0.05)
(Fig. 2). Hemolymph ecdysteroid
titer also differed over the course of the fourth larval stage (one-way ANOVA,
P=0.009), with high levels in 30 h old larvae and higher still in 12
h old pupae (Fig. 2). Hemolymph
ecdysteroid levels correlated well with that of tissue production
(r2=0.52), and both showed a similar rise and fall pattern
(Fig. 2).
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Critical feeding period for pupation and adult eclosion
For these experiments, we manipulated nutrient input by giving fourth
instars access to food for defined time periods, followed by a period of
fasting. Ingestion was not measured in this experiment, but larvae most likely
maintained high ingestion rates when food was present. Earlier studies
indicate that an A. aegypti larva fills its gut after just 1 h of
feeding on nutritive substances (Rashed
and Mulla, 1989; Sneller and
Dadd, 1977) and continues to ingest and move food through its gut
as long as additional nutritive substances are present
(Merritt et al., 1992). The
larval diet for colony rearing and for these experiments includes brewers
yeast, and yeast has been found to ensure high ingestion rates by larval
A. aegypti and other species (Aly,
1985; Rashed and Mulla,
1989). Under our rearing conditions, the probability of pupation
was influenced by feeding period (P<0.0001, 2
test), greater than 50% of fourth instars successfully pupated when they had
at least 24 h to feed (Fig.
3A). For larval groups that had access to food for 24 h or longer,
we found no evidence that feeding period affected the development time fourth
instars took to pupate (P=0.210). On average, fourth instars spent
2.6 days until pupation. Under our rearing conditions, mortality was minimal
for these groups, and only 14 of 288 total (<5%) died as either larvae or
pupae when given access to food for 24 h or longer. Most fourth instars were
developmentally arrested if given less than 24 h to feed
(Fig. 3A). There was no
statistical difference in mean survival time between developmentally arrested
larvae that were allowed to feed for 12 or 18 h (P=0.833). Both
groups of larvae survived on average for 14 days prior to death.
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We also examined whether developmentally arrested larvae retained the capacity for metamorphosis. Another group of newly molted fourth instars were allowed to feed for only 12 h and fasted for 7 days. When fed ad libitum after fasting, 100% of larvae successfully pupated and eclosed as adults (data not shown). However, when given only 12 h to feed after fasting, only 70% of fourth instars successfully pupated, and a greater proportion of the eclosed individuals were males (Fig. 3B). A 50/50 ratio of male and female emergence is observed when developmentally arrested larvae are given supplementary periods of feeding of 18 h or longer (Fig. 3B). Female body size, as assessed by wing length, increased with a longer supplementary period of feeding (data not shown).
The influence of larval age and nutritional condition toward metamorphosis
This experiment was carried out to distinguish cues for metamorphosis
related to a larva's nutritional state versus its age. For this
experiment, newly molted fourth instars were set up in specific feeding period
regimens, and tissues were collected from these larvae after 36 h. Since
thoraces and abdomina produced peak levels of ecdysteroids around 30–36
h (Fig. 2) and all individuals
pupated and eclosed as adults if fed for at least 30 h
(Fig. 3A), we focused our
feeding regimen experiments on 36 h old fourth instars. In one feeding
regimen, newly molted fourth instars were deprived of food for the entire 36 h
period. In a second regimen, fourth instars were given access to food for 12 h
but fasted until they were 36 h old. In the third regimen, fourth instars were
given access to food for the entire 36 h period. Specific correlates of larval
development, such as ecdysteroid levels, body mass and nutrient reserves, were
measured for each larval group.
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Fourth instar larvae that were given access to food for 36 h had a
significantly higher body mass compared to newly molted larvae and larval
groups given only 12 h to feed or not (Tukey–Kramer HSD,
P0.05) (Fig. 6A).
Levels of nutrient reserves in fourth instars, corrected for larval dry mass,
differed among the feeding treatment groups (ANCOVA, P<0.0001)
(Fig. 6B–D). Compared to
newly molted fourth instars, levels of lipid, glycogen and protein decreased
significantly in fourth instars that were deprived of food for 36 h
(Fig. 6B–D), although no
such decrease in dry mass was measured
(Fig. 6A). Fourth instars 36 h
old that had access to food for 12 h had significantly greater lipid levels
compared to newly molted fourth instars
(Fig. 6B) but similar levels of
glycogen and protein (Fig.
6C,D). Overall, fourth instars that were 36 h old and had access
to food for that length of time were the heaviest and contained significantly
higher levels of all nutrient reserves.
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Discussion |
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). Females of mosquito species that must ingest blood
meals for all egg cycles are considered anautogenous, whereas, females of
autogenous species can produce at least their first egg batch without a blood
meal. When larval mosquitoes were fed high or low amounts of food, both
autogenous and anautogenous females emerged with high or low levels of
nutritional reserves, respectively (Telang
et al., 2006; Telang and
Wells, 2004). In those studies, larvae restricted to a poor diet
emerged with low reserve levels, but they committed to pupation nonetheless.
In our current study, we wanted to explore the importance of larval nutrition
in more detail and specifically toward its importance for metamorphic
readiness. To begin examining the readiness of mosquito larvae to pupate, we
measured basic parameters of larval development such as critical mass,
nutritional reserves, and ecdysteroid production with hemolymph titers during
the fourth instar and early pupal stage of A. aegypti.
In mosquitoes, the nutritional environment experienced by a female when a
larva dictates her adult body size
(Briegel, 1990a;
Briegel, 1990b;
Telang et al., 2006;
Telang and Wells, 2004), as
for many insects (Calvo and Molina,
2005; Nijhout,
2003; Slansky and Scriber,
1985; Stern,
2003). An important physiological determinant of adult body size
is a minimum or critical mass that must be gained during the larval stage to
trigger the commitment to metamorphose. For the model insect M.
sexta, the critical mass has been defined as `the minimal weight in
which further feeding and growth are not required for a normal time course to
metamorphosis and pupation'
(Davidowitz et al., 2003;
Nijhout and Williams, 1974b).
This same definition was used to determine the critical mass for pupariation
in last instar D. melanogaster
(Mirth et al., 2005;
Zhou X. et al., 2004). For the
yellow fever mosquito Aedes aegypti, the critical mass has been
defined as the mass at which 50% of larvae pupate when they are starved
subsequent to being weighed (Chambers and
Klowden, 1990). Using both these definitions, we estimated the
critical mass to be 2.7–3.2 mg wet mass for larvae reared at 27°C
(Fig. 1). This critical mass
was achieved by 24 h old fourth instar larvae. This is also the minimum length
of time these larvae needed to access food so that at least 50% of them
pupated and emerged as adults (Fig.
3A), and the age at which ecdysteroid production and titer begins
to rise (Fig. 2). Our
measurement of critical mass is greater than previously reported for A.
aegypti strain Segemaganga (Chambers
and Klowden, 1990). In that study, the critical mass for A.
aegypti varied from 1.9–2.6 mg when larvae were reared at 22°C
to 1.6–1.9 mg when larvae were reared at 32°C
(Chambers and Klowden, 1990).
Differences in larval critical mass reported in both studies most likely
reflect genetic differences among the various strains of A. aegypti
currently under long-term colonization.
According to the model developed for M. sexta, when larvae reach
the critical mass, the corpora allata (CA), glands responsible for JH
synthesis and secretion, are turned off and JH levels begin to decrease
(Nijhout and Williams, 1974a),
with the elimination of JH facilitated by the catabolic enzyme JH esterase
(Browder et al., 2001). The
absence of JH for a metamorphic molt is important, and it is thought to render
the larval brain competent to secrete PTTH
(Truman, 1972;
Truman and Riddiford, 1974).
PTTH stimulates secretion of ecdysteroids by the PG that then causes larvae to
stop feeding and commit to pupation
(Nijhout, 1994). Therefore,
the size that a larva attains at its metamorphic molt determines the size of
its adult body. In our current study, fourth instar mosquito larvae reach
their critical mass 24 h post-ecdysis (Fig.
1), and we measured a single peak of ecdysteroid secretion by
tissues and hemolymph titers 30–36 h post-eclosion
(Fig. 2). For mosquitoes, it is
not known if a brain factor with PTTH-like activity is responsible for
stimulating ecdysteroid production by larval tissues, thereby initiating the
metamorphic molt. Whether JH plays a regulatory role in mosquito metamorphosis
is also not known. However, larvae treated with the JH mimic, methoprene,
exhibited delayed pupation, suggesting that a critical `JH sensitive' period
exists in mosquito larval development (Lan
and Grier, 2004), as has been shown in some some lepidopteran
studies (Nijhout, 1994).
Among past studies on A. aegypti metamorphosis, two main areas of
discrepancy exist (Chambers and Klowden,
1994; Fournet et al.,
1995; Jenkins et al.,
1992; Lan and Grier,
2004; Margam et al.,
2006). First, the length of time fourth instars take to pupate
differs greatly among the studies (51–72 h). This development time is
affected by several factors such as temperature, food quantity and rearing
densities. Unfortunately, the studies have employed different larval rearing
protocols or have not adequately described conditions to facilitate
experimental duplication. To assist future experiments, it is important to
report relevant larval rearing parameters such as temperature, rearing
densities, container dimensions and feeding regimen, as herein. Second, there
is disagreement about the number of ecdysteroid peaks that occur during the
last instar. In some studies, two or three peaks in ecdysteroid titer are
reported for the last instar (Fournet et
al., 1995; Lan and Grier,
2004; Margam et al.,
2006; Westbrook and Russo,
1985). Different ecdysteroid antisera and immunoassays (RIA
versus enzyme immunoassay) are used in these studies, and more
notably, whole body extracts taken at different times were assayed. The amount
of ecdysteroids in such extracts is greater than that in hemolymph taken at
similar time points (Jenkins et al.,
1992) (see also this study), and from personal experience, there
is a high degree of non-specific binding of ecdysteroid antisera to such
extracts that can only be resolved by the assay of serial dilutions of each
extract to identify its appropriate dilution for accurate quantification in
the linear range of the immunoassay. This important information is not to be
found in studies using whole body extracts of mosquito larvae. These studies
do discuss a `commitment peak' that occurs 14–28 h
(Lan and Grier, 2004) or 24 h
(Margam et al., 2006) post
ecdysis to the last instar larvae, which in this last instance is not a
significant increase in comparison to the larger peak occurring 30–36 h
post ecdysis. Among studies, there is a consensus that an ecdysteroid peak
occurs 24–36 h later in fourth instar development
(Fournet et al., 1995;
Lan and Grier, 2004;
Margam et al., 2006;
Westbrook and Russo, 1985) and
a larger one occurs during the early part of the pupal stage
(Fournet et al., 1995;
Margam et al., 2006;
Whisenton et al., 1989).
Similarly in a previous study (Jenkins et
al., 1992) and the present one, a significant increase in
ecdysteroid secretion by tissues in thoraces and abdomina of 30–36 h old
fourth instar larvae corresponded to an ecdysteroid increase in hemolymph
titer, which we believe is the pre-molt increase that triggers
larval–pupal apolysis. Ecdysteroid secretion by tissues and hemolymph
titer decline in 48–52 h old fourth instars, which are nearing pupation,
and greatly increase in pupae 12 h after the larval–pupal molt.
A more thorough examination of ecdysteroid titers at lower temperatures
(thereby prolonging larval development) or at shorter time intervals in last
instar larvae, may reveal an earlier peak between 6 and 9 h post last instar
ecdysis that would initiate the rise in gene expression for a variety of
ecdysteroid-regulated genes, such as the isoforms of the ecdysteroid receptor
and ultraspiracle recently reported
(Margam et al., 2006). It
should be noted, however, that a recent and exhaustive study of ecdysteroid
titer and related biosynthetic enzyme levels in whole bodies of last instar
Drosophila melanogaster failed to substantiate such an early
commitment peak (Warren et al.,
2006). This study did find surprisingly low whole body ecdysteroid
peaks at 20 and 28 h post ecdysis to the last instar that were correlated with
an increase in biosynthetic enzyme levels in this same study, along with a
sevenfold increase in ecdysteroid titer during the early phase of pupation.
For now, it appears that the timing of major ecdysteroid peaks during the last
instar and early pupae required for metamorphosis by these two dipteran
species is similar, but the tissue sources of the ecdysteroids are different,
as may be the regulation of ecdysteroid secretion.
For metamorphosis to occur, nutrient intake is necessary to facilitate
growth during the last instar (Nijhout,
2003). When fourth instars were given longer periods of time to
feed, a greater proportion of them successfully pupated and eclosed as adults.
However, most fourth instars allowed to feed for only 12 h and then starved
were developmentally arrested (Fig.
3A). Similar results were reported using a different experimental
design (Nishiura et al.,
2007). Larvae can live for up to 2 weeks when starved and still
retain the capacity to pupate if re-fed
(Fig. 3B). Our study is not the
first to observe the ability of A. aegypti to withstand starvation
for this period of time (Rasnitsyn and
Yasyukevich, 1989;
Wigglesworth, 1942). The
ability of larval A. aegypti to tolerate starvation is believed to
depend on stored energy, primarily lipids
(Gilpin and McClelland, 1979;
Wigglesworth, 1942).
Starvation tolerance over a long period of time is thought to be common among
container-inhabiting species in general
(Barrera, 1996;
Barrera and Medialdea, 1996),
and this period may be an important factor to be considered for control
measures and the study of their vectorial capacity.
When fed optimally during their fourth instar, the increase of ecdysteroids
to cue metamorphic commitment is seen in 30–36 h old fourth instar
larvae. When newly molted fourth instars were starved at ecdysis and deprived
of food for 36 h, they show no increase in ecdysteroid secretion or body mass
compared to newly molted fourth instars (Figs
5,
6A). These starved fourth
instars maintained similar body mass but had lower levels of lipid, glycogen
and protein compared to newly molted fourth instars
(Fig. 6A–D). When fourth
instars face starvation conditions, we expect them to catabolize storage
nutrients. Given that fourth instars remain alive for 14 days without food, it
is not surprising that we see no decrease in mass after just a 36 h fasting
period. Another study observed a major decrease in A. aegypti pupal
mass only when they were starved for at least 4 days as fourth instars
(Rasnitsyn and Yasyukevich,
1989). Lastly, our biochemical analyses do not account for sugars,
amino acids or storage excretion, but the dry mass of starved fourth instars
may include these materials. It is thought that the larval fat body can
function in storage excretion, and large deposits of uric acid have been
observed in fat body cells of starved A. aegypti larvae, presumably
as a product of deamination of amino acids or proteins
(Wigglesworth, 1942;
Wigglesworth, 1987). If
accumulation of uric acid occurred in starved fourth instars in our study,
this would undoubtedly contribute toward their mass maintenance. Overall,
fourth instar A. aegypti allowed to feed for the entire 36 h period
accumulated the highest levels of all three storage nutrients
(Fig. 6B–D). The fat
content of insect larvae of many species increases from early to late instars
(Slansky and Scriber, 1985).
Mosquito larvae may be no exception, but this awaits further study.
Our data suggest that metamorphic capacity is dependent on a larva's nutritional condition, not just the chronological age at which ecdysteroid increase occurs. Tissues in 36 h old fourth instars that had fed for 12 h secreted higher levels of ecdysteroids compared to newly molted larvae or larvae deprived of food for 36 h (Fig. 5). The level of ecdysteroids measured in the 12 h fed/36 h old fourth instars (65 pg per thorax and abdomen set per 6 h) falls at the lower end of the 60–100 pg range, which appears to commit metamorphosis in 30–36 h old fourth instars fed ad libitum (Fig. 2). However, that level of ecdysteroid secretion is clearly not sufficient to initiate pupation since we know fourth instars are developmentally arrested if given only a 12 h feeding period (Fig. 3A). Only in 36 h old fourth instars that had access to food for 36 h do we observe significant accumulation of nutrients and the necessary ecdysteroid increase, indicating that both conditions must be met for larvae to initiate metamorphosis (Fig. 5, Fig. 6A–D).
Strong positive correlations have been found between body mass and caloric
reserves of lipid and carbohydrates
(Chambers and Klowden, 1990),
and it was concluded that carbohydrates accumulated by larvae strongly
influenced readiness to pupate. According to our data, it is during the 12 h
and 36 h time spent feeding that the lipid, glycogen and protein thresholds
are met to trigger the metamorphic molt
(Fig. 6B–D). Nutrients
accumulated by larvae have to serve both body-building and energy needs. Thus,
the threshold for metamorphic readiness must encompass all three nutrient
reserves. To the best of our knowledge, data concerning nutritional reserves
accumulated by larvae during development have not been collected for the model
insects M. sexta or D. melanogaster.
Thus, the timing of ecdysteroid release may not be critical to initiate the
larval–pupal molt for mosquito larvae, but both the ecdysteroid titer
and the nutritional condition of fourth instars are crucial factors in
initiating the first metamorphic molt. De novo synthesis of
ecdysteroids by larval abdomen has been documented in the housefly Musca
domestica (Studinger and Willig,
1975), the mealworm beetle Tenebrio molitor
(Romer et al., 1974) and in
A. aegypti (Jenkins et al.,
1992). Currently it is not known what cells common to both the
thorax and abdomen of A. aegypti larvae are ecdysteroidogenic.
Ecdysteroid release from epidermal cells and oenocytes, which are associated
with the fat body in some insects
(Chapman, 1998), are dispersed
throughout the thorax and abdomen and have been implicated in these past
studies. Future studies of larval mosquitoes will discover how the fat body
and associated tissues convey information about nutrient levels to the nervous
system so that PTTH, JH and ecdysteroid production by non-prothoracic gland
cells are coordinated to induce metamorphosis.
Mosquitoes such as A. aegypti and Anopheles gambiae are
responsible for vectoring pathogens that cause serious human diseases, such as
dengue fever and malaria. A complimentary aspect of effective disease
intervention is suppression of the vector. Effective mosquito population
control is best targeted at the larval and pupal stages because these stages
are confined by their aquatic habitats relative to the highly mobile aerial
adult stage. We focus our work on the yellow fever mosquito A.
aegypti in the hope that studies regarding its larval development will
contribute to knowledge regarding its biology and control. Traditional vector
control methods have employed chemical pesticides. However, concerns over
non-target effects and target insect resistance have led to a shift in control
towards more biorational approaches, such as the bacterium Bacillus
thuringiensis israelensis (Bti), and insect growth regulators (IGRs),
such as JH mimics (methoprene) and ecdysone agonists. The normal larval
physiology of metamorphosis is disrupted by IGRs
(Beckage et al., 2004;
Nishiura et al., 2003;
Palli et al., 2005), but the
mechanisms by which they do so are not well understood. Full exploitation of
current and novel IGRs would be aided by an understanding of the regulation of
postembryonic development of mosquitoes.
|
Acknowledgments |
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References |
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