Plant senescence cues entry into diapause in the gall fly Eurosta solidaginis: resulting metabolic depression is critical for water conservation
Department of Zoology, Miami University, Oxford, OH 45056, USA
* Author for correspondence at present address: Department of Biological Sciences, University of Nevada Las Vegas, 4505 Maryland Parkway, Las Vegas, Nevada 89154, USA (e-mail: jason.williams{at}ccmail.nevada.edu)
Accepted 20 September 2005
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Summary |
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Key words: desiccation resistance, diapause, Eurosta solidaginis, cuticular permeability, respiratory transpiration
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Introduction |
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To survive the desiccating conditions of winter, galling insects such as
cynipid wasps (Williams et al.,
2002) and the goldenrod gall fly Eurosta solidaginis
(Ramløv and Lee, 2000
),
have extremely low rates of water loss. These soft-bodied, immature insects
have rates of water loss that are similar to heavily sclerotized, adult desert
beetles (see Edney, 1977
and
references therein; in Hadley,
1994a
). To attain such high levels of desiccation resistance,
overwintering goldenrod gall fly larvae seasonally reduce their rates of water
loss from late summer to mid-winter. In particular, E. solidaginis
reduce their rate of water loss more than sixfold, within a two-week period in
early autumn (Williams et al.,
2004
). However, the mechanisms by which E. solidaginis
reduce their rate of water loss or the cues triggering the enhanced
desiccation resistance during this period are unknown.
The goldenrod gall fly (Diptera: Tephritidae) ranges throughout much of
central and eastern North America, from Texas to southern Canada. In late
spring or early summer, females oviposit in terminal buds (future stems) of
goldenrod plants (Solidago spp.)
(Uhler, 1951). Larvae feed and
grow within the moist gall tissue throughout the summer and early autumn. In
southwest Ohio goldenrod plants and gall tissues senesce and rapidly dry in
late September and early October (Irwin et
al., 2001
; Williams et al.,
2004
). In addition to reducing their rate of water loss, larvae
undergo other physiological changes as their gall tissue senesces. For
instance, larvae produce the cryoprotectant glycerol in response to plant
senescence (Baust and Lee,
1982
; Rojas et al.,
1986
), suggesting this low molecular mass polyol may be important
in preventing dehydration. Larvae also enter diapause during this period
(Irwin et al., 2001
). Diapause
is defined as a genetically determined state of low metabolic activity,
suppressed development and heightened resistance to environmental extremes,
which generally begins before, and lasts longer than, the adverse conditions
(Tauber et al., 1986
;
Danks, 1987
).
Overwintering and dormant insects lose water to the environment primarily
through cuticular and respiratory transpiration
(Edney, 1977;
Hadley, 1994a
;
Danks, 2000
). During the fall
and winter, larvae of E. solidaginis reduce their rates of cuticular
water loss by producing epicuticular lipids, which increase by 40-fold from
summer to mid-winter (Nelson and Lee,
2004
), and possibly by producing cryoprotectants
(Williams et al., 2004
). These
larvae may also reduce their rates of loss by lowering respiratory
transpiration, as their metabolism decreases when entering diapause. However,
reductions in metabolic rate for inactive insects frequently have a relatively
minor effect on desiccation resistance (for reviews, see
Chown, 2002
;
Chown and Nicolson, 2004
).
Regardless, larvae of E. solidaginis probably reduce both avenues of
water loss as they increase their desiccation resistance in early autumn as
gall tissue senesces, but the relative contributions of decreased cuticular
and respiratory transpiration are unknown.
The purpose of this study was to determine whether plant senescence and
diapause induction were closely associated with the seasonal reduction in
rates of larval water loss that occurs in early October as documented
previously (Williams et al.,
2004). Specifically, we measured the cuticular and estimated
respiratory contributions to total organismal water loss as gall tissue
naturally senesced. We also determined whether mild desiccation stress and
premature plant senescence would trigger enhanced larval desiccation
resistance. We measured total rates of water loss, rates of cuticular water
loss, carbon dioxide production (as an estimate of metabolic rate and diapause
induction), body water content, hemolymph osmolality and glycerol content of
larvae collected from the field just prior to, and after, natural plant
senescence. To determine if the field-collected larvae were taken from
desiccating and senescent plant tissue, we measured the water content and
water activity of gall tissues.
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Materials and methods |
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Gall measurements
Hydric conditions of the gall tissue were assessed by measuring the total
water content and water vapor potential of the gall tissue immediately
surrounding the larvae. Gall water content was determined by weighing 10 galls
that had contained larvae to ±0.1 mg using a Mettler Toledo AG245
balance (Columbus, OH, USA), before and after drying in an oven at 65°C
until they reached a constant mass. Water vapor potential of the gall tissue
was determined by the psychrometric vapor pressure depression technique
described by Hølmstrup and Westh
(1994). Immediately after
opening an occupied gall, 1020 mg of gall tissue surrounding the larval
chamber was transferred to a Wescor C-52 sample chamber (Logan, Utah, USA) and
allowed to equilibrate for 30 min. Water potential was then determined with a
Wescor HR 33T Dewpoint Microvoltmeter (Logan, Utah, USA) operated in the
dewpoint mode. Measurements were taken on 10 randomly selected galls collected
on 1 October. Galls in the stem cutting treatment and those collected on 20
October were too dry to accurately measure water activity.
Measurements of desiccation resistance
Resistance to desiccation was determined by measuring rate of water loss,
in µg mm2 h1; body water content, as a
ratio of wet mass to dry mass; and rate of cuticular water loss. To determine
rate of organismal water loss, which includes both respiratory and cuticular
components, larvae (N=10) were weighed to ±0.01 mg to obtain a
fresh mass. Larvae were then re-weighed after being desiccated
non-convectively over Drierite (W. A. Hammond Drierite Co., Ohio, USA) at 4%
relative humidity (RH) and 20°C until they lost 510% of their fresh
mass. Cuticular surface area was estimated from initial wet mass using an
equation determined by Williams et al.
(2004):
y=0.912x+4.204, r2=0.804, where
y = surface area in mm2 and x = mass in mg. Body
water content was determined by placing the desiccated larvae in an oven at
65°C until a constant dry mass was obtained.
Rates of cuticular water loss (µg mm2 h1) were measured to determine the relative contributions of respiratory and cuticular components of overall organismal water loss. Cuticular water loss was assessed by weighing larvae (N=10) before and after exposure to 4% RH at 20°C as described above, however, prior to testing, the spiracles of each larva were topically blocked with a small amount of Thomas Scientific Lubriseal stopcock grease (Swedesboro, New Jersey, USA), to eliminate respiratory water loss. Stopcock grease was carefully applied with a 10 ml syringe to ensure that only the spiracles were covered with the stopcock grease. In addition, larvae were examined after the experiment to ensure that the grease did not spread to other areas of the cuticle. This procedure is extremely effective at blocking the four larval spiracles of E. solidaginis, as CO2 emission is completely eliminated after application of only a small amount of stopcock grease to the spiracles (data not shown).
Hemolymph osmolality and cryoprotectant concentration
Hemolymph osmolality (N=10) was determined by drawing 7-10 µl
of hemolymph into a capillary tube through a small incision in the larval
cuticle. The hemolymph was then analyzed in a Wescor Vapro 5520 Hemolymph
Osmometer (Logan, Utah, USA). To measure glycerol concentration, larvae
(N=7) were frozen at 80°C until whole body measurements
were performed by enzymatic assay (Sigma Chemical Co., St Louis, Missouri,
USA, no. 337) as described by Hølmstrup et al.
(1999).
Measurement of metabolism
To determine larval diapause status, we assessed metabolic rate by
measuring CO2 emission. Larvae (N=7 per treatment) were
weighed and individually placed into 14 ml3 glass respirometry
chamber kept within a temperature-controlled bath held at 20°C. Carbon
dioxide was measured using a flow through (50 ml min1)
respirometer (TR-3 model, Sable Systems, Las Vegas, Nevada, USA). Room air was
used for respirometry measures after it was drawn through a column of ascarite
(Thomas Scientific, Swedesboro, New Jersey, USA) to remove CO2 and
drierite to reduce water content of the air. Larvae were equilibrated to the
chamber for 1 h prior to analysis. Metabolic rate data were converted into
units of µl CO2 emitted per mg fresh mass per hour using DATACAN
software (Sable Systems). We did not measure larval activity during metabolic
rate determinations because CO2 production was relatively stable
after the 1 h acclimation period, and larvae, regardless of diapause status,
rarely moved, unless prodded, after being removed from the gall.
Effect of mild and moderate desiccation stress on larval desiccation resistance
A second group of larvae were collected between September 28 and October 3,
prior to the dramatic seasonal decrease in rates of water loss, and were used
to determine the effect of mild and moderate desiccation stress on desiccation
resistance. Immediately after collection, larvae were removed from their galls
and placed in desiccators over either double distilled water to produce a RH
of 100%, a saturated solution of sodium sulfate producing a RH of 95% or a
saturated solution of sodium chloride producing a RH of 75%. Each desiccator
was placed in an incubator maintained at 20°C. After 3, 6 or 10 days
exposure to these conditions, larvae were removed and assessed for water loss
rate, body water content, cuticular permeability, hemolymph osmolality,
glycerol concentration and metabolic rate using the methods described
above.
Statistical analysis
After data were determined to be normally distributed with homogeneous
variances, a one-way analysis of variance followed by a Bonferroni multiple
comparisons (Statview 5.0, SAS Institute Inc., Cary, North Carolina, USA) was
used to determine differences in means between larvae collected on 1 and 20
Oct., the stem cutting treatment, and all relative humidity treatments. To
increase statistical power when examining the larval treatments exposed to
various relatively humidities, a one-way ANOVA followed by a Bonferroni
multiple comparisons was used to identify differences in RH treatments on a
given day. A significance level of =0.05 was used for all tests.
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Results |
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Measures of desiccation resistance
Although the larvae were exposed to desiccating conditions after gall
senescence, they were able to maintain their hydration levels, as the body
water content did not differ among the 1 October, stem cutting and 20 October
groups (Fig. 1). In addition,
body water content did not differ in the laboratory RH treated larvae and 1
October controls (Fig. 1).
Values ranged between 1.62±0.03 and 1.37±0.05 mg water mg dry
mass1 (5862% of fresh mass).
|
Rates of larval water loss were dramatically (P<0.05) lower for larvae in the stem cutting group and those collected on 20 October (averaging 0.7± 0.2 µg mm2 h1) compared to larvae collected on 1 October (6.0±1.8 µg mm2 h1; Fig. 2A). After just 3 days of exposure to 75% RH, larvae had rates of water loss (1.3±0.1 µg mm2 h1) that were statistically similar to those in the stem cutting and 20 October groups (Fig. 2A). Furthermore, humidity may influence E. solidaginis to increase desiccation resistance as 6-days exposure to 100% RH and 95% RH were required before rates of larval water loss were significantly lower that those collected and analyzed on 1 October (Fig. 2A). In addition, larvae exposed to 75% RH for 10 days had significantly lower rates of water loss (0.8±0.1 µg mm2 h1; P<0.05) than those in the 100% RH, 10 day treatment (2.1±0.5 µg mm2 h1).
|
Measures of metabolism
Metabolic rate was significantly lower (P<0.05) for larvae
analyzed on 20 October (post-plant senescence) and in the stem cutting group
(premature plant senescence), averaging 0.05±0.01 µl CO2
g1 h1 compared with larvae from the 1
October group (pre-senescent goldenrod; 0.31±0.06 µl CO2
g1 h1;
Fig. 2C). Interestingly,
metabolic rate lowered rapidly once the larvae were removed from their gall;
all larvae in the 100%, 95% and 75% RH treatments had similar metabolic rates
(0.09±0.05 µl CO2 g1
h1) to those in the stem cutting and 20 October group just 3
days after being removed from the gall. No larvae, in any experimental group,
demonstrated discontinuous gas exchange.
Hemolymph osmolality and cryoprotectant production
Hemolymph osmolality increased by an average of 105 mOsm
kg1 in the stem cutting and 20 October treatments, compared
with the 1 October collection, which averaged 534±14 mOsm
kg1 (Fig.
3A). Larvae exposed to 100%, 95% and 75% RH for 6 and 10 days
hadsignificantly higher hemolymph osmolalities, averaging 673±27 mOsm
kg1, than those in the 1 October collection
(P<0.05). Mildly desiccating conditions did not appear to
influence hemolymph osmolality as solute concentrations did not differ between
the 100% RH and 95 or 75% RH treatments during the 3, 6 or 10 days of exposure
(Fig. 3A).
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Discussion |
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Enhanced cuticular resistance plays a minor role in rapid increase in desiccation resistance
It is generally thought that increased desiccation resistance in arthropods
is primarily due to a reduction in cuticular water loss
(Hadley, 1994a;
Hadley, 1994b
). However,
reduced cuticular transpiration contributed only a small component to lowering
the rates of total water loss in this study
(Fig. 4). Three groups of
larvae significantly reduced both their rates of total water loss and rates of
cuticular water loss compared to the 1 October control (20 October field
group, 75% RH day 3, and 75% RH day 10 groups;
Fig. 2A,B). Larvae in these
groups averaged a decrease in their rate of total water loss of 5.1 µg
mm2 h1, but reduced their rate of
cuticular transpiration by only 0.8 µg mm2
h1 (Fig. 4).
Thus, reductions in rate of cuticular water loss only accounted for a small
portion (15%) of the overall decrease in rate of water loss in these groups.
Consequently, because the decrease in the rate of water loss for E.
solidaginis larvae at this time in their life cycle is only due to
changes in cuticular and respiratory transpiration, the majority (
85%) of
the overall reduction in rate of water loss was the result of a reduction in
respiratory transpiration (Fig.
4).
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Mechanisms of reduced cuticular water loss
Reductions in cuticular water loss were probably due to increases in
cuticular lipids. The integument of an insect is the primary barrier to
cuticular water loss (Hadley,
1994a; Gibbs,
1998
) and many insects at risk of dehydration enhance their
desiccation resistance by increasing epicuticular lipids when entering a
dormant stage or diapause (s.f. Manduca sexta,
Bell et al., 1975
;
Sarcophaga crassipalpis, Yoder et
al., 1992
; Mamestra configurata,
Hegdekar, 1979
). Eurosta
solidaginis larva also increases the amount of its epicuticular lipids by
40-fold over several months, from late summer to mid-winter
(Nelson and Lee, 2004
). Thus,
even though the abrupt decrease in rates of water loss for E.
solidaginis occurred within days, increased hydrocarbons probably reduced
rates of cuticular water loss of the larvae exposed to desiccating
conditions.
Less clear is the effect that cryoprotectant accumulation may have had on
reducing cuticular water loss. Cryoprotectants are so-called because they
enhance insect cold tolerance. Recently, several authors have suggested that
cryoprotectants may be beneficial for water conservation through colligative
action (Ring and Danks, 1994;
Block, 1996
;
Bayley and Hølmstrup,
1999
) or by binding water at the cuticular basement membrane (see
discussion in Williams et al.,
2004
). Synthesis of the cryoprotectant glycerol accounted for most
(
60%) of the overall increase in hemolymph osmolality in both field and
laboratory groups (Fig. 3A,B).
The observed increases in larval glycerol concentration confirmed earlier
reports that E. solidaginis produces this cryoprotectant in response
to gall tissue senescence (Baust and Lee,
1982
; Rojas et al.,
1986
), suggesting that glycerol may have a protective role when
entering a dormant state or desiccating environment. However, it is unclear if
glycerol production lowered rates of cuticular water loss. For instance,
larvae in the 75% RH treatments had lower rates of cuticular water loss than
larvae in the 100% RH treatments on days 3 and 10 of the study
(Fig. 2B) and larvae in the 75%
RH treatments were also the only laboratory groups to have an increased
glycerol concentration when compared to the 1 October control group
(Fig. 3B). By contrast, larvae
in the 95% RH treatment did not follow this trend. These larvae had lower
rates of cuticular water loss than the 100% RH treatment
(Fig. 2B) but did not have
increased glycerol levels compared with control values
(Fig. 3B). Thus, it appears
unlikely that glycerol production was a primary factor in reducing rates of
cuticular water loss during this study.
Reductions in cuticular water loss were triggered by environmental moisture, or more specifically, the presence of a water potential deficit between the larvae's hemolymph and the environment. A water potential deficit of only 1700 kPa (100 kPa=1 bar) between the hemolymph of the collembolan Folsomia candida and its environment induce a marked increase in its drought tolerance (Sjurnsen et al., 2001). On 1 October, larvae were in a potentially hydrating environment as their gall tissues were quite moist and had a higher water activity than their hemolymph (plant tissue averaged 1280 kPa; Table 1; larval hemolymph averaged 1300 kPa as calculated from osmolality measures). Larvae placed in the 100% RH treatment continued to experience a potentially hydrating environment and did not reduce their rate of cuticular water loss (Fig. 2B). In contrast, larvae placed in the 95% and 76% RH treatments were subjected to desiccating conditions with an average water potential deficit between their hemolymph and environmental water vapor of 5820 and 39 210 kPa, respectively. In addition, larvae in the stem cutting and 20 October groups also had similarly reduced rates of cuticular water loss after being subjected to water potential deficits in their galls (Fig. 2B; Table 1). Consequently, mild desiccation stress cued larvae to reduce their rate of cuticular water loss and did so in as little as 3 days.
Plant senescence triggers entry into larval diapause
Low metabolic rates for various field- and laboratory-treated larvae
indicate that they had entered diapause. Larvae from southwest Ohio reduce
their metabolic rate by more than 75% as they enter diapause and maintain a
metabolic rate of 0.1 µl CO2 g1
h1 at 20°C throughout diapause
(Irwin et al., 2001
). Carbon
dioxide production of larvae taken from post-senescent galls or larvae exposed
to various RH treatments decreased by an average of 75% (ranging between 61%
and 84%) compared to the 1 October control group, and averaged 0.07 µl
CO2 g1 h1. Taken together, the
reduction and resulting level of CO2 production indicate that these
larvae were in diapause. The induction of diapause occurred rapidly for larvae
placed in the RH treatments as they entered the dormant state within 3 days of
being removed from the gall.
Plant senescence triggered E. solidaginis larvae to enter
diapause. A variety of cues induce insects to enter diapause, including food
availability or quality, moisture, oxygen and pH. However, the most common
cues for temperate insects are related to temperature and/or photoperiod
(Tauber et al., 1986;
Danks, 1987
). Irwin et al.
(2001
) suggested that a
combination of low temperature, such as experiencing an initial frost,
photoperiod, or host plant senescence induced larvae to enter diapause. Yet in
our study, photoperiod and temperature did not appear to influence diapause
induction. For example, all larvae in the various RH treatments entered
diapause even though they were held at a relatively high constant temperature
(20°C) after collection (Fig.
2C). In addition, field groups experienced the same temperatures
and photoperiods prior to testing; however, larvae in the stem cutting group
had entered diapause by 1 October, whereas larvae from the control group had
not (Fig. 2C). However, the
fact that larvae in the stem cutting group, taken from dried gall tissue,
entered diapause by 1 October suggests that plant senescence induced these
larvae to enter the dormant state. As gall tissues senesce the environment
that the larva inhabits changes in two distinct ways. The nutritive gall layer
on which the larvae feed deteriorates, eliminating their only food source
(Uhler, 1951
). In addition,
the larvae are subjected to a desiccating environment for the first time
(Williams et al., 2004
).
Moisture is an important cue for many insects to enter diapause (see
references in Tauber et al.,
1998
), yet moisture, or more specifically a desiccating
environment, does not appear to influence diapause induction for E.
solidaginis. All larvae removed from their galls and placed in various
relative humidities entered diapause regardless of whether they were in a
non-desiccating environment (100% RH treatment) or a desiccating environment
(95% and 75% RH treatments; Fig.
2C). However, larvae in all the RH treatments were removed from
their food and water source, suggesting that food and water availability or
quality is the primary cue that triggers E. solidaginis to enter
diapause.
Importance of diapause in the water balance of dormant insects
Larvae of E. solidaginis probably reduce their respiratory
transpiration by lowering their metabolic rate as they enter diapause. It is
well established that elevated metabolic rate, due to activities such as
flight, is directly related to increased respiratory water loss and,
consequently, total water loss (Nicolson
and Louw, 1982; Lehmann,
2001
). However, for most inactive insects, respiratory water loss
constitutes a minor portion of their total water loss, 20% or less
(Chown, 2002
). Therefore, most
studies have found that reductions in basal metabolic rate have little effect
on maintaining water balance for these insects
(Quinlan and Hadley, 1993
;
Hadley, 1994b
; Djawden and
Bradley, 1997; Rourke, 2000
).
In spite of that, others contend that reductions in basal metabolic rate can
be important in limiting insect water loss
(Zachariassen, 1996
;
Davis et al., 2000
;
Addo-Bediako et al., 2001
).
Reductions in metabolism may have a greater impact on water conservation for
xeric-adapted insects, in which respiratory transpiration constitutes the
majority of their total water loss, such as the beetle Phrynocolus
petrosus, which loses 69% of its water through respiration
(Zachariassen, 1991
). For the
non-diapausing 1 October control larvae, respiratory water loss was 4.9 µg
mm2 h1 (estimated by subtracting the rate
of cuticular water loss, Fig.
2B, from the rate of total water loss,
Fig. 2A), or 80% of the total
water loss. However, transpiratory water loss was dramatically reduced, to
only 0.6 µg mm2 h1, for larvae in
diapause (Fig. 4). These larvae
also reduced their metabolic rate by 4.2-fold over the same testing periods.
Therefore, a substantially lowered metabolism due to diapause would reduce the
need for gas exchange and would allow the larvae to reduce the rates of
respiratory and total water loss by regulating their spiracular openings
(Gibbs et al., 2003
).
Temperate insects benefit from diapause in a variety of ways. Insects in
the state of diapause are often more resistant to adverse conditions. Diapause
also synchronizes spring emergence and prevents premature development, which
would be fatal, during unseasonably warm periods in late winter
(Tauber et al., 1986;
Danks, 1987
). The reduced
metabolic rate of diapausing insects also conserves stored energy needed for
spring development and reproduction (Danks,
1987
; Irwin and Lee, 2003; Williams et al., 2003). However, few
authors consider desiccation resistance as an important function of diapause
for temperate and polar insects. Our data suggest that a lowered metabolic
rate, due to diapause induction, is extremely important in conserving body
water. For instance, if the non-diapausing 1 October control larvae maintained
the respiratory transpiration rate of 4.9 µg mm2
h1 (Fig. 2A
minus Fig. 2B), we estimate it
would only take them 26 days to loose 10% of their body water to the
desiccating conditions used in this experiment through respiration
transpiration. In contrast, diapausing larvae had an average transpiration
rate of 0.6 µg mm2 h1 and it would take
these larvae 214 days to loose 10% of their body water through respiratory
transpiration. Since many temperate insects remain in diapause for 6 months or
more, a dramatically reduced rate of respiratory water loss, through a lowered
metabolic rate, would have a profound impact on their overwintering water
balance.
In summary, the large seasonal reduction in rate of water loss for E. solidaginis larvae is cued by the senescing of their gall tissue and is primarily due to reduced respiratory transpiration as the larvae enter diapause.
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Acknowledgments |
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References |
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