Behavioral Ecology Vol. 13 No. 5: 607-614
© 2002 International Society for Behavioral Ecology
Reduced reproductive effort in male field crickets infested with parasitoid fly larvae
Department of Biology, University of California, Riverside, CA 92521, USA
Address correspondence to G.R. Kolluru, who is now at the Department of Organismic Biology, Ecology and Evolution, University of California-Los Angeles, 621 Charles E. Young Drive South, Los Angeles, CA 90095-1606, USA. E-mail: gkolluru{at}obee.ucla.edu.
Received 27 July 1999; revised 23 May 2001; accepted 18 December 2001.
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ABSTRACT |
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Some populations of the field cricket Teleogryllus oceanicus are parasitized by the phonotactic fly Ormia ochracea. Flies locate crickets by their song and deposit larvae onto them. The larvae develop inside the cricket for 1 week before killing the host upon emergence. The reproductive compensation hypothesis predicts that parasitized crickets should increase their reproductive effort during the initial stages of infestation to offset the loss of fitness resulting from their shortened life span. An alternative hypothesis predicts that parasitized crickets will decrease reproduction, either because they are unable to reproduce or because selection acting on the parasitoid favors decreased host reproduction. In laboratory experiments, parasitized male crickets had reduced reproductive effort (spermatophore production, calling, mating activity, and mass allocated to reproductive tissue) compared to unparasitized males. Parasitized males fed ad libitum showed no evidence of allocating a greater proportion of their resources to reproduction. Parasitized and healthy males did not differ significantly in resting or maximal metabolic rates, although this may have been due to the substantial contribution of larval respiration to the metabolic rate of the host—parasitoid complex. These results are consistent with previous studies and suggest that T. oceanicus males parasitized by O. ochracea do not increase their reproductive effort. We discuss potential reasons that crickets do not increase reproductive effort in response to fly larvae and address difficulties in demonstrating altered life-history patterns in response to parasitism.
Key words: crickets, mating, phonotactic parasitoids, reproductive compensation, reproductive effort, Teleogryllus.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Parasitism can significantly influence host life-history patterns (Agnew et al., 2000
;
Forbes, 1993;
Minchella, 1985).
Traditionally, changes in patterns of reproduction in parasitized animals were
viewed as either byproducts of damage or as parasite adaptations to reduce
predation and to free host resources for parasite growth
(Fritz, 1982;
Horton and Moore, 1993;
Minchella, 1985). Indeed, some
parasites are castrators, completely destroying the host's reproductive
machinery (Kuris, 1974).
However, more recent studies have explored whether changes in host
reproduction represent host, rather than parasite, adaptations
(Forbes, 1993;
Poulin, 1995).
Minchella and Loverde
(1981) hypothesized that
parasitized animals should be selected to increase current reproductive effort
to offset future losses and demonstrated that snails exposed to trematode
parasites increased egg laying. Their "reproductive compensation
hypothesis" rests on the concept of residual reproductive value
(Fisher, 1958;
Williams, 1966). Because the
residual reproductive value of a parasitized individual is decreased, it would
be adaptive for that individual to shift more of its resources into the
current reproductive event. This hypothesis has since been supported in
several studies (reviewed by Agnew et al.,
2000). For example, crickets infected with bacteria,
Drosophila infested with mites, and amphipods harboring trematodes
all increase reproductive activity (Adamo,
1999; McCurdy et al.,
2000; Polak and Starmer,
1998).
Teleogryllus oceanicus is an Australian field cricket that has
been introduced into Hawaii, where it is parasitized by the New World tachinid
fly Ormia ochracea, also introduced into Hawaii
(Otte and Alexander, 1983;
Zuk et al., 1993). Gravid
female O. ochracea acoustically orient to T. oceanicus
calling song and deposit larvae on the cricket
(Cade, 1975). The larvae
develop inside the host for 1 week and kill the host upon emergence
(Adamo et al., 1995). Ormiine
flies significantly shorten host life span
(Lehmann and Heller, 1997),
and there is a sharp decrease in the frequency of older male crickets in
populations parasitized by O. ochracea compared with unparasitized
populations (Murray and Cade,
1995; Simmons and Zuk,
1994).
Parasitized crickets mount an encapsulation response in which O.
ochracea larvae are enclosed by layers of hemocytes
(Vinson, 1990). Contrary to
the reproductive compensation hypothesis, parasitized female Gryllus
crickets produced fewer eggs than control females
(Adamo, 1999;
Adamo et al., 1995). In the
most detailed study to date of the effects of O. ochracea on male
cricket reproduction, Adamo et al.
(1995) showed that latency to
courtship singing was unaltered in Gryllus species. However, these
authors did not examine other, potentially more direct measures of
reproductive effort. Other studies have demonstrated that infestation with
late-stage ormiine parasitoid larvae depresses male orthopteran calling
activity (Cade, 1984;
Zuk et al., 1995) and
spermatophylax weight (Lehmann and
Lehmann, 2000).
We tested the reproductive compensation hypothesis against the alternative
hypothesis that parasitization depresses host resource allocation to
reproduction (either as a byproduct of infestation or as an adaptation on the
part of the parasite; Adamo et al.,
1995; Agnew et al.,
2000; Cade and Wyatt,
1984). We examined the reproductive effort of parasitized and
unparasitized T. oceanicus males by experimentally infesting groups
of males and comparing their calling activity, spermatophore production, sperm
viability, and mating activity to that of unparasitized males. Changes in host
reproductive effort could be masked by parasitoid-induced changes in
reproductive activity or physiology (Adamo,
1999; Forbes,
1996; Perrin et al.,
1996). Accordingly, we also examined the effects of parasitoids on
the host's capacity for metabolic power output (which may impact reproductive
activities such as singing or courtship) and resting rates of energy
expenditure and on allocation of resources to storage versus reproductive
tissues.
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METHODS |
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Experimental infestation
We obtained parasitoid flies used in the calling activity and fecundity experiments from a laboratory colony maintained by R. R. Hoy at Cornell University. We trapped flies used in all other experiments on the grounds of the University of Hawaii, Hilo, using ceramic tiles coated with Tanglefoot insect trap coating and baited with tape-recorded, synthesized T. oceanicus song. Unless otherwise stated, we collected crickets to be parasitized by hand from the same location as the flies and anesthetized them with CO2 (100% for approximately 15 s) immediately before infestation. Larvae were dissected from the flies, and a dissecting pin was used to transfer three to five mobile larvae onto the membranous area around the front legs of each cricket (Cade, 1984
). This number of larvae is within the natural range found
parasitizing the wild Hilo population of T. oceanicus
(Zuk et al., 1995).
Unparasitized males were anesthetized and handled under a dissecting
microscope. At least one parasitoid larva emerged from each parasitized male, and unparasitized males did not harbor any parasitoid larvae. All infested crickets died within hours of larval emergence. The weights of pupae resulting from experimental infestations were heavier than those resulting from natural infestations, probably due to greater food intake by the laboratory-reared crickets (t test; n = 19 pupae from natural infestation, 20 from experimental infestation; t = 5.23; p < .0001).
Calling activity
We monitored calling activity using an electronic circuit connected to a
Macintosh IIsi computer (after Kidder and
Sakaluk, 1989). The device sampled each of eight microphones,
sensitive to the frequency of T. oceanicus calling song (4-5 kHz),
once per second, so that we were able to determine whether or not each male
was calling in 1-s increments throughout the night. The crickets and the
monitoring device were housed at the University of California, Riverside, in
an anechoic chamber maintained at 31 ± 3°C and 12:12 h light: dark
schedule.
Pilot tests using tape-recorded cricket song allowed for the adjustment of
microphone sensitivity. To ensure that the microphones accurately recorded
calling activity, we conducted a test of the monitoring device using four
laboratory-reared T. oceanicus males monitored for 6 nights (12 h per
night; after Bertram and Johnson,
1998). The eight microphones were grouped into pairs, and each
pair was threaded through a rubber stopper with a hole in it and hung 3 cm
from the top of each of four 0.5-1 plastic containers with air holes. Each
container housed one male cricket, a cardboard shelter, and ad libitum food
and water. We grouped data from each container's two microphones into
half-hour segments and compared them using correlation analysis. All of the 23
resulting correlations were significant after applying a Bonferroni correction
for multiple correlation tests (radjusted = 0.88 ±
0.05; p = 2.63 x 10-8 ± 1.87 x
10-8).
Calling activity experiments were conducted from February to May 1999. We obtained virgin male crickets from our laboratory colony as last-instar nymphs and housed them in a group container until they eclosed. The males were 1-7 weeks old at the start of the experiment, and males in the parasitized and control groups did not differ in age. Each male was individually isolated in a 0.5-1 plastic container with food, water, and a cardboard shelter. One microphone was placed in each container as described above, and calling activity was monitored for seven continuous nights (approximately 12 h per night). On the eighth day of the experiment, the males were anesthetized using CO2, handled under a dissecting microscope, and measured to the nearest 0.1 mm with digital calipers. Anesthesia and handling were performed to examine their effects on subsequent calling activity. Males were allowed to recover, and calling activity was monitored again for 2 nights. Males were then experimentally infested with parasitoid larvae as described above, and calling activity was monitored until the larvae emerged.
We repeated the experiment with a control group of males to examine the effects of repeated anesthesia and handling on calling activity. All experimental conditions including the ages of the males were similar to those in the parasitized male tests, except that these males were anesthetized and handled twice without being infested with parasitoid larvae. At the end of each experiment the males were individually housed until pupae emerged from all parasitized males. All males were then frozen, their pronotum width was measured using dial calipers, and they were dissected under a dissecting microscope to detect additional larvae. We compared the mean calling activity of males before and after parasitization (before and after second anesthetization for the control males) using Wilcoxon paired-sample tests.
Spermatophore production
We conducted spermatophore production experiments in Hilo, Hawaii, during
July and August 1997. We collected crickets as adults from the same location
as the flies a few days before each experiment. Although cricket ages were
unknown, all males were calling at the time of collection. The crickets were
maintained under the natural light: dark schedule (approximately 13:11 h) and
kept on an ad libitum diet of dry cat food and water. T. oceanicus
produce sperm in discrete spermatophores, which are small and simple, lacking
the nutrient-rich spermatophylax portion found in other orthopterans such as
katydids (Loher and Dambach,
1989). Because male crickets exhibit diel periodicity in
spermatophore production and do not usually transfer spermatophores during
daylight hours (Loher, 1989;
McFarlane, 1968), we collected
spermatophores from shortly before sunset (1700 h) until shortly after first
light (0500 h). Collection involved visual examination of the male and female
for protruding spermatophores, examination of the container floor for any
dislodged spermatophores, and gentle squeezing of the abdomen of each male to
extrude spermatophores that had not yet protruded
(Cade and Wyatt, 1984;
Zuk, 1987). We assumed that
spermatophores from parasitized and unparasitized males were equally likely to
be eaten.
We examined the effects of contact with females on spermatophore production in two separate experiments, each with a different set of crickets. In each case, we infested males between 1300 and 1500 h, and that evening was designated day 1 post-infestation. We collected spermatophores on days 4 and 6 post-infestation in the first experiment (n = 15 unparasitized, 12 parasitized males), and on days 1, 2, 3, and 5 post-infestation in the second experiment (n = 14 parasitized, 14 unparasitized males). Males in the first experiment were individually isolated for the few days between capture and the experiment, and males in the second experiment were housed in mixed-sex groups during those few days.
In both experiments, each male was placed in a 0.9-1 Styrofoam container with one unparasitized female within 1 h of infestation. On the indicated nights post-infestation, we checked each pair once every hour and removed any spermatophores detected with forceps. Females were rotated regularly in both experiments to minimize female effects on mating activity, so that in the first experiment each male was with three different females, and in the second experiment each male was with five different females.
Spermatophore data were normally distributed (analyses of residuals by
Shapiro-Wilk test; experiment 1: W = 0.9453; p = .1805;
experiment 2: W = 0.9473, p = .2431) and were analyzed using
repeated-measures ANOVA (SAS Institute,
1990).
Mating activity
We examined mating activity in the laboratory in Hilo, Hawaii, in June
1998. We collected crickets as adults from the same location as the flies a
few days before the start of the experiment and maintained them at 25 ±
1°C, 60% humidity and 12:12 h light:dark schedule throughout the
experiment. All males were calling at the time of capture and were therefore
assumed to be reproductively active. Each male (n = 19 parasitized,
16 unparasitized males) was individually housed in a 0.6-1 plastic container
with water, cat food, and a paper shelter. Just before the start of the first
day of observations one mature female was introduced into each male's
container. We rotated females regularly before the start of each day's
observations so that each male was with four different females by the end of
the experiment. We conducted behavioral observations every day between 2100
and 0030 h for 6 days. The parasite status of crickets was not known to the
observer. Each pair of crickets was examined and behavior recorded every 10
min during the observation period. If the male was courtship singing during an
observation period, then we monitored that couple either until mounting took
place or until the next scheduled observation occurred. Mounting was scored if
the female mounted the male and remained on him for more than 10 s. Failure
was scored if the male performed courtship singing for two or more
observations (
20 min) without being mounted. Males in this category were
usually not mounted for the remainder of the night.
Mating activity data were not normally distributed (mounting: W =
0.92, p = .01; failure: W = 0.87, p = .0005), and
there was no significant difference across nights (Kruskal-Wallis test;
mountings: 
2 = 4.50, p = .48; failures:
2 = 8.57, p = .20). Therefore, we performed
nonparametric tests on the totals for each male across the 6 nights of the
experiment.
Fertilization success
We examined the effects of parasitoid infestation on male fertilization
success by comparing the fertility and fecundity of females mated to
parasitized (days 2-4 post-infestation) and unparasitized males. The
experiment was conducted in an environmental chamber (27 ± 1°C, 70%
humidity, and 12:12 h light:dark schedule) from February to May 1999. We
obtained crickets from our laboratory colony as last-instar nymphs and housed
them in single-sex group containers until they eclosed. Males were either
parasitized or handled at 4-13 days of age, after which each male was housed
with two virgin, unparasitized females (with the exception of one
unparasitized male who was housed with only one female) for 2 nights in a
0.9-L plastic container with cat food, water, and shelter. Females were in
either the young (4-13 days old) or old (33-60 days old) age class and ranged
in size (pronotum width) from 5.11 to 5.99 mm. To minimize female age and size
effects on fecundity, we distributed females in a stratified, random fashion
between parasitized and unparasitized males, such that each group of males had
an approximately equal number of females of each age and size class. After 2
nights the females were removed into individual 0.9-l plastic containers with
food, water, shelter, and an egg-laying dish.
T. oceanicus females exhibit an egg flood, an increase in
oviposition immediately after mating
(Vaughan, 1995). Therefore, we
replaced egg-laying dishes after 2 days, and each female had two dishes,
representing the first 2 days of laying and the subsequent 13 days of laying.
Because T. oceanicus females lay unfertilized eggs
(Vaughan, 1995), we used
hatchlings and not eggs to assess male fertilization success. However, because
some clutches had no hatchlings, all unhatched eggs were also counted as an
indication of female fecundity. Although pigmented eyespots can usually be
used to determine if eggs contain embryos, the eggs from this experiment were
often dark due to age, and eyespots were therefore not used as indicators of
fertilization. At the end of the experiment we dissected all females to
determine whether they had eggs ready to be fertilized. We obtained
fertilization success data by taking the mean number of hatchlings and the
mean number of hatchlings plus eggs laid by the two females mated to each
male. The data were analyzed using Mann-Whitney U tests.
Metabolic rate
We measured resting metabolic rate (RMR) for 12 unparasitized and 11
parasitized, laboratory-reared male crickets descended from individuals
captured in Hawaii. Crickets were housed in 0.5-l plastic containers with
shelter and ad libitum food (Fluker's cricket food, cat chow) and water and
were housed and tested at 30 ± 1°C. Repeated RMR measurements were
taken on these crickets once per day on days 1-5 after infestation of the
parasitized crickets.
We used open-flow respirometry to measure rates of CO2
production (
CO2; ml/g/h).
Measurements were made using a LiCor 6251 CO2 analyzer capable of
resolving differences of 0.2-0.4 ppm of CO2 in air. Flow rates of
dry, CO2-free air (100-200 ml/min) were maintained at ±1% by
a Tylan mass flow controller. Excurrent air from the chamber was dried
(magnesium perchlorate) before entering the LiCor 6251. Outputs from both
instruments (as well as ambient temperature measured with thermocouples) were
recorded on Macintosh computers equipped with National Instruments A/D
converters and custom software for data acquisition and analysis (WartHog
Systems, written by M. A. Chappell and available at
www.warthog.ucr.edu).
We measured RMR in chambers constructed from 0.5-l plastic containers and
maintained at 30 ± 1°C in an environmental cabinet. RMRs were
determined as the mean minimal steady-state
CO2 during periods of at least 10
min when activity (indicated by abrupt changes in
CO2) was absent. We calculated
CO2 as:
 | (1) |
is flow rate corrected to standard
temperature and pressure (STP; 0°C and 101.3 kPa),
FICO2 is the initial fractional concentration of
CO2 (zero in these experiments), FECO2 is the
final fractional concentration of CO2, and RQ is the respiratory
quotient (the ratio of CO2 produced/O2 consumed). We
used an RQ of 0.85, which was previously measured in an independent group of
T. oceanicus (Chappell, unpublished data). The value of RQ used in
Equation 1 had little effect on calculated
CO2 because
FECO2 was very small (<.002).
We measured maximal metabolic rate (MMR) as
CO2 during intense, forced exercise
for 8 unparasitized and 11 parasitized wild-caught males from Hilo, Hawaii.
These crickets had been captured 3 days before testing and were kept in 0.5-l
plastic containers with shelter and ad libitum food (Fluker's cricket food,
cat chow) and water. They were housed and tested at 27 ± 1°C.
Repeated MMR measurements were taken on days 2, 4, and 6 after
infestation.
Because of the field location, we used a simpler closed system to measure
MMR. We placed single crickets inside a 140-ml syringe equipped with a
stopcock valve and flushed the syringe with dry, CO2-free air
(scrubbed with Dryerite and soda lime). The syringe was then sealed and shaken
in a uniform motion for exactly 5 min, forcing the cricket to exercise
vigorously. At the conclusion of exercise, the air in the syringe was injected
through a small tube of desiccant (magnesium perchlorate) into the
CO2 analyzer, and the maximum CO2 concentration
(FECO2) was recorded. Since FICO2 was
zero, we computed CO2 as:
 | (2) |
CO2, since FECO2
never exceeded 0.004).
Before analysis, we corrected all metabolic rates to 30.0°C using a
Q10 of 2.5 (Chappell, unpublished data). In both treatment groups,
metabolism varied linearly with mass over the small mass range of the
crickets. Accordingly, we corrected for mass effects by dividing metabolic
rates by mass and tested for differences between groups with repeated-measures
ANOVA (JMP; SAS Institute,
1995).
Validation of CO2 as a
metabolic index
We present data for CO2 rather
than rates of O2 consumption
(O2) because the sensitivity and
stability of the LiCor 6251 CO2 analyzer are approximately 100-fold
better than that of the best available O2 analyzer. However, using
CO2 to measure metabolism requires
careful consideration because the energy equivalence of
CO2 strongly depends on whether
lipid, protein, or carbohydrate is used to fuel respiration, and because
released CO2 may come from buffered storage in body fluids as well
as directly from respiration. Therefore, we conducted pilot studies during
which both O2 and
CO2 were measured during rest and
exercise in a low flow-rate, open system, which generated concentration
changes in both gases that were large enough for accurate measurements. We
used modified 0.5-l plastic containers (for RMR) and 30-ml syringes (for MMR)
as metabolic chambers, with flow rates of dry, CO2-free air of
100-120 ml/min. Excurrent gas was dried using magnesium perchlorate, passed
through the LiCor 6251, scrubbed of CO2 with Ascarite, redried, and
passed through an Applied Electrochemistry S-3A oxygen sensor. Changes in gas
content were recorded as described above. We calculated
O2 as:
 | (3) |
CO2 using Equation 1. We performed
validation tests on five crickets at rest and during forced exercise. The RQ
during rest was constant over time (mean ± SD = 0.78 ± 0.09) and
did not significantly differ between exercise and rest (0.75 and 0.78,
respectively; t = -0.26, p = .80), demonstrating that
CO2 is an accurate index of energy
expenditure in T. oceanicus.
Energy allocation
To determine whether parasitized crickets allocate energy resources
differently from unparasitized crickets, we dissected MMR crickets under
40x magnification and obtained dry weights of the wings, head, thoracic
exoskeleton, abdominal exoskeleton, legs, thoracic muscles, testes plus
accessory glands, and fat body by drying each part to a constant weight in a
55°C drying oven and weighing each to 0.01 mg using a Cahn 21
electrobalance. We removed all parasitoid larvae and associated breathing
tubes before weighing.
We also determined the dry weights of a different group of male crickets, each of which was exercised only once, on one of days 1, 2, 3, or 4 after infestation (using the methods described above) and euthanized on the day after exercise. These males were captured in Hilo, Hawaii, 3 days before the first day of testing (day 1: n = 2 parasitized and 5 unparasitized, day 2: n = 2 parasitized and 6 unparasitized, day 3: n = 3 parasitized and 3 unparasitized, day 4: n = 4 parasitized and 4 unparasitized).
We combined dry masses to obtain measures of reproductive tissues (testes plus accessory glands plus thoracic muscle) and storage tissue (fat body). We calculated the proportion of resources allocated to each tissue type as: proportion devoted to reproduction = (thoracic muscle mass + testis mass + accessory gland mass)/(head mass + thorax mass + abdomen mass + leg mass + wing mass + thoracic muscle mass + testis mass + accessory gland mass); proportion devoted to storage = fat body mass/(head mass + thorax mass + abdomen mass + leg mass + wing mass + thoracic muscle mass + testis mass + accessory gland mass).
Our measures of dry mass do not account for variation in per-gram energy
content of tissues. However, we assume that the relative fractions of lipid
and lipid-free mass were not significantly different between comparison
groups. Residuals of dry masses were normally distributed (group exercised
once: reproductive tissue proportion, W = 0.96, p = .39,
storage tissue proportion, W = 0.97, p = .65; group
exercised repeatedly: reproductive tissue proportion, W = 0.98,
p = .99, storage tissue proportion, W = 0.98, p =
.95), and differences between parasitized and unparasitized males were
evaluated using ANOVA (JMP; SAS Institute,
1995).
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RESULTS |
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Calling activity
Males in the infested group (n = 8) called significantly more before parasitoid infestation than after infestation (Wilcoxon paired-sample test; mean percentage of the night spent calling before = 50%, mean after = 1%, T = 0, p < .01; Figure 1). Although males in the control group (n = 6) exhibited a slight decrease in calling activity after the second anesthetization, this was not statistically significant (mean before = 37%, mean after = 23%, n = 6, T = 7, p > 0.5; Figure 1).
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Spermatophore production
Unparasitized males produced significantly more spermatophores per night
than parasitized males (repeated-measures ANOVA; experiment 1: unparasitized
mean ± SE = 3.3 ± 1.8, parasitized = 0.5 ± 1.2,
F1,24 = 29.78, p = .0001; experiment 2:
unparasitized = 1.51 ± 1.50, parasitized = 0.31 ± 0.70,
F1,19 = 7.22, p = .01;
Figure 2), although the two
groups did not differ in body size (pronotum width; experiment 1: F =
0.05, p = .82; experiment 2: F = 0.08; p = .78).
Males in experiment 1, who were individually isolated before the start of the
experiment, produced more spermatophores and peaked in spermatophore
production earlier in the night than males in experiment 2. There was no
significant variation in spermatophore production across nights (experiment 1:
F = 0.03, p = .86; experiment 2: F = 0.385,
p = .77), but there was a significant night x parasitization
status interaction in the first experiment (experiment 1: F = 5.21,
p = .03; experiment 2: F = 0.71, p = .56) because
unparasitized males increased spermatophore production from night 4 to night
6, whereas parasitized males decreased spermatophore production.
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Mating activity
Unparasitized males (n = 16) were mounted more and failed at
mounting solicitation attempts more than parasitized males (n = 19;
Kruskal-Wallis test; mountings: unparasitized mean ± SD = 4.00 ±
2.85, parasitized = 1.10 ± 1.52, Z = 3.41, p = .0007;
failures: unparasitized = 1.19 ± 0.75, parasitized = 0.37 ±
0.60, Z = 3.09, p = .002). However, the proportion of
attempts that resulted in a successful mounting did not differ between groups
(unparasitized mean ± SD = 0.71 ± 0.26, n = 16;
parasitized = 0.70 ± 0.39, n = 11, Z = 0.60,
p = .54), suggesting that the differences resulted from fewer
attempts by parasitized males rather than from rejection of parasitized males
by females. As previously described for T. oceanicus
(Burk, 1983), courtship song
always preceded mounting, and males never successfully forced a
copulation.
Fertilization success
Unparasitized and parasitized males did not differ in the number of
hatchlings they produced (Mann-Whitney U test: unparasitized mean
± SE = 26.42 ± 68.24, parasitized = 27.10 ± 52.73,
U = 16, p > .05). There was a great deal of variability
in hatchling production among females (mean ± SE = 28.10 ±
13.06), with several females producing no hatchlings. The low hatchling
production could not be explained by age group (U = 47, p
> .05) or body size (pronotum width; regression: r2 =
.024, F = 0.46, p = .51). Because the low hatching success
may have resulted from environmental conditions in the chamber, unhatched eggs
were also counted and a Mann-Whitney U test performed on hatchlings
plus eggs laid. The results were identical to the first test, so that there
was no difference between unparasitized and parasitized males in either the
fertility (hatchling production) or the fecundity (all eggs laid) of the
females they were mated with. Dissections revealed that 17 of the 21 females,
including most of those that had no hatchlings, retained at least 50 eggs in
their ovaries.
Metabolic rate
Parasitized and unparasitized crickets did not differ significantly in
either resting or maximal metabolic rates (repeated-measures ANOVA; resting:
F1,25 = 0.21, p = .65; maximal:
F1,32 = 0.04, p = .85; Figures
3 and
4). However, there was a
significant decrease in maximal metabolic rate across days for both treatment
groups (resting: F2,21 = 1.15, p = .33; maximal:
F2,17 = 9.80, p = .0005). There was no
significant parasite treatment x day interaction for resting or maximal
metabolic rates (p > .10 for both).
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Energy allocation
Parasitized males in the MMR group, who were exercised repeatedly and
weighed on day 6 after infestation (i.e., late in the course of parasitoid
development), had devoted a significantly smaller proportion of the total mass
to reproductive tissue (testes + accessory glands + thoracic muscle;
unparasitized mean ± SE = 0.068 ± 0.005, parasitized = 0.019
± 0.004; ANOVA: F1,15 = 56.47, p <
.0001) and storage tissue (fat body; unparasitized mean ± SE = 0.109
± 0.011, parasitized = 0.064 ± 0.009; ANOVA:
F1,15 = 10.18, p = .0061) than unparasitized
males. However, parasitized and unparasitized males in the other group, who
were exercised once and weighed on either day 1, 2, 3, or 4 after infestation,
did not differ in the proportion of reproductive or storage tissues (ANOVA:
reproductive tissue, F1,17 = 0.03, p = .87;
storage tissue, F1,17 = 0.39, p = .54;
Figure 5). There was a
significant decline in storage tissue with time (F3,17 =
4.13, p = .03) but no significant day x treatment group effect
(reproductive tissue: F3,17 = 0.61, p = .62;
storage tissue: F3,17 = 2.47, p = .10). There was
no evidence that the proportion of mass devoted to reproduction increased with
time in either treatment group (Figure
5).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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The reproductive compensation hypothesis predicts that parasitized animals should increase reproductive effort to offset losses resulting from their shortened expected life span (Minchella and Loverde, 1981
). Ormia ochracea larvae do not consume
host tissues during the first few days of infestation
(Adamo et al., 1995).
Parasitized crickets are therefore expected to increase reproductive effort
during this time to partially compensate for losses incurred later, when
larvae migrate to the abdomen and cause serious injuries to the host
(Adamo et al., 1995;
Forbes, 1993). Our
observations of parasitized T. oceanicus crickets contradict this
hypothesis. The reproductive effort (spermatophore production, calling, mating
activity, and proportion of mass allocated to reproductive tissue) of
parasitized males was significantly lower than that of unparasitized males.
Calling, spermatophore production, and mating were reduced even during the
initial stages of infestation. O. ochracea is therefore similar to
parasites that castrate their hosts by consuming reproductive tissues, by
interfering biochemically with the host, or by limiting host nutrients so that
investment into reproduction is impossible
(Hurd and Webb, 1997;
Kuris, 1974;
Reed and Beckage, 1997).
O. ochracea larvae consume thoracic muscles and fat body (but not
reproductive tissue; Adamo et al.,
1995). The decrease in fat body points to an energy trade-off as
the likely source of reduction in host reproductive effort. However, other
effects such as alteration of the hormonal composition of cricket hemolymph
(Adamo, 1994) may also be
responsible for reduced host reproduction.
Parasitoid-induced decreases in reproductive activity may obscure the
detection of an increase in host reproductive effort if only reproductive
activity is measured (Forbes,
1996; Perrin et al.,
1996). Therefore, we also examined metabolic rates and mass
allocation patterns of infested and healthy males. Parasitized males do not
spend less time feeding and do not consume less food than healthy males
(Adamo et al., 1995; Kolluru,
personal observation). When given ad libitum food and water, parasitized males
that were exercised repeatedly devoted a significantly smaller proportion of
total mass to both reproductive and storage tissues than did healthy males.
Contrary to the idea of reproductive compensation, these parasitized males did
not increase the proportion of mass allocated to reproductive tissues. Males
that were only exercised once, including those examined early during
infestation, did not differ from healthy males in energy allocation patterns.
These males also did not show any evidence of adaptive increases in energy
devoted to reproduction.
Our metabolic rate measurements provided an estimate of the minimal energy
expenditure (resting metabolic rate) and maximal capacity for aerobic
metabolic power output (maximal metabolic rate) of crickets in each treatment
group. To our knowledge, this is the first examination of the effects of a
dipteran parasitoid on host metabolism, although studies of hymenopteran
parasitoids have demonstrated decreased host metabolic rate
(Alleyne et al., 1997;
Rivers and Denlinger, 1994).
Surprisingly, we found no significant difference in the resting or maximal
metabolic rates of parasitized and unparasitized males, suggesting that there
is only a minimal metabolic cost of parasitism. However, parasitoid larvae may
account for as much as 30% of the mass of the host—parasitoid complex on
the day of emergence. Therefore, it is likely that parasitoid larvae
contributed a substantial fraction of the total resting metabolic rate of the
host—parasitoid complex (e.g.,
Alleyne et al., 1997). We
measured the CO2 production of newly emerged O. ochracea
larvae and found an average VCO2 of 1.07 ml/h/g, corresponding to a
potentially large fraction of the resting metabolism of the host-parasitoid
complex (Figure 4). This
implies that parasitized males have lower resting metabolic rates than our
data indicate (assuming the metabolic rate of newly-emerged larvae is similar
to that of preemergence larvae). Therefore, infested males may be unable to
increase reproductive effort because of a reduced metabolic capacity for
breaking down storage tissue for reallocation to reproduction. Our finding
that maximal aerobic power output during forced exercise was not affected by
parasitoids seems inconsistent with that hypothesis. However, those
measurements concerned very brief bouts of activity and may not reflect
parasitoid-induced constraints on metabolic performance over longer intervals
that are more relevant to reproductive output.
Our study aimed to determine the impact of parasitoid infestation on
optimal host life-history patterns. Some experimental studies addressing this
issue have shown that reproductive effort increases as predicted (e.g.,
Adamo, 1999). However, it is
difficult to interpret a lack of increase in reproduction in response to
infestation. We used multiple measures of reproductive and somatic energy
expenditure to determine the allocation patterns of parasitized T.
oceanicus males, all of which contradicted the prediction. However,
because reproductive effort is extremely difficult to assess directly, our
approach cannot exclude with certainty the possibility that increased
reproductive effort by parasitized males was masked by declines in
reproductive output relative to healthy males.
There are several possible explanations for why parasitized T.
oceanicus males may be unable to reallocate resources to reproduction
(see also Adamo, 1999). The
association between the cricket and fly may be too recent for an adaptive host
response to have evolved. Although T. oceanicus was introduced into
Hawaii at least 125 and possibly as many as 1000 years ago
(Kevan, 1990;
Otte and Alexander, 1983), it
is not known when O. ochracea reached Hawaii. However, many cricket
and fly generations have undoubtedly passed since the two came together in
Hawaii, and T. oceanicus song characteristics have had sufficient
time to evolve in response to the parasitoid
(Kolluru, 1999;
Rotenberry et al., 1996;
Zuk et al., 1993).
Alternatively, the encapsulation response may be too general to elicit an
increase in reproductive effort (Adamo,
1999; Vinson,
1990). This is supported by Adamo's
(1999) finding that bacterial
infection, which induces antimicrobial humoral immune responses, caused
reproductive compensation, but that neither O. ochracea larvae nor
Sephadex beads (both of which induce encapsulation) did so.
Our results demonstrate that parasitoid infestation constrains a male
cricket's ability to successfully reproduce and suggest that eavesdropping by
the fly should lead to adaptations by the cricket to avoid infestation
(Zuk and Kolluru, 1998). Both
amount of calling and spermatophore production rate are important for male
reproductive success (Kolluru,
1999; Sakaluk and Cade,
1980,
1983;
Zuk, 1987), and fly
infestation therefore represents a significant fitness cost to crickets even
before death. However, further studies are needed to establish the effects of
the parasitoid under more natural conditions. For example, although
wild-caught silent T. oceanicus males are more likely to harbor large
fly larvae than calling males (Zuk et al.,
1995), one field study showed no significant difference in calling
activity between parasitized and unparasitized males
(Kolluru, 1999). However, this
study did not control for cricket age or for intensity or stage of
infestation. Our results also differ somewhat from Cade's
(1984) study of calling by
parasitized crickets. He found a more gradual decrease in calling activity,
possibly because his crickets were less heavily infested, or because the
association between O. ochracea and T. oceanicus is more
recent than that with his Gryllus crickets. Similarly, although
reproductive tissue mass measurements and field data (Zuk, unpublished
observation) show pronounced degeneration of the seminiferous tubules of
parasitized crickets, our data suggest that parasitized males are able to
produce viable sperm (see also Adamo et
al., 1995). Therefore, parasitized males may experience limited
reproductive success if they are able to produce the occasional spermatophore
and successfully attract females. However, under field conditions, parasitized
males may be unable to successfully compete for access to females or may be
selected against by searching females.
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ACKNOWLEDGEMENTS |
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We are grateful to G. F. Grether for statistical advice, S. Brown, L. Nunney, and D. Price for the use of laboratory facilities in Hilo and Riverside, R. R. Hoy for providing parasitoid flies, B. Bergeron for building the electronic circuit, and N. M. Waser, D. F. Westneat, and three anonymous reviewers for their comments on earlier versions of the manuscript. We also thank K. Kettleson and the staff at the University of Hawaii and the Dolphin Bay Hotel staff for their assistance. This research was supported by the National Science Foundation (NSF) (DEB-9257749) and University of California Academic Senate grants to M.Z., NSF (IBN-9902255) grant to G.R.K. and M.Z., University of California at Riverside Intramural Funds to M.A.C., and Animal Behavior Society, Orthopterists' Society, Sigma Xi, and University of California Graduate Dean's Dissertation Research grants to G.R.K.
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