Abstract
In arthropods, larger individuals tend to have more fat reserves, but data for many taxa are still missing. For the vinegar fly Drosophila melanogaster, only few studies have provided experimental data linking body size to fat content. This is rather surprising considering the widespread use of D. melanogaster as a model system in biology. Here, we hypothesized that fat content in D. melanogaster is positively correlated with body size. To test this, we manipulated the developmental environment of D. melanogaster by decreasing food availability. We then measured pupal size and quantified fat content of laboratory-reared D. melanogaster. We subsequently measured pupal size and fat content of several field-caught Drosophila species. Starvation, crowding, and reduced nutrient content led to smaller laboratory-reared pupae that contained less fat. Pupal size was indeed found to be positively correlated with fat content. The same correlation was found for field-caught Drosophila pupae belonging to different species. As fat reserves are often strongly linked to fitness in insects, further knowledge on the relationship between body size and fat content can provide important information for studies on insect ecology and physiology.
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Introduction
Body size is a key life history trait in insects1,2,3,4. Many experimental studies consequently use body size measurements as a proxy for fitness (e.g.5,6,7,8), because a larger size generally leads to a higher fecundity9,10 and longevity6,9. Insect body size depends on the environmental conditions experienced during development, such as temperature11, nutrition12,13 or population density14. As insects do not grow as adults, body size is determined entirely during the juvenile stages15. Final body size is thus reached after pupation or after the final molt for insects with complete or incomplete metamorphosis, respectively16,17,18,19. During the development of holometabolous insects, developing larvae need to reach several weight limits for metamorphosis to occur, i.e., the minimal viable weight and the critical weight20,21. The minimal viable weight corresponds to the weight at which nutrient reserves, such as fat, are sufficient to survive after metamorphosis, while the critical weight corresponds to a threshold after which metamorphosis can no longer be delayed, even if the larva is starved20. In Drosophila melanogaster, the minimal viable weight and the critical weight are occurring almost simultaneously during larval development20. Starvation before reaching the minimal weight impedes the initiation of metamorphosis, leading to death of the larva, while reduced food intake or starvation after reaching the critical weight leads to smaller adult individuals20,22. Reduced nutrition during development thus generally decreases size.
A range of traits are correlated to body size (i.e., allometry), including the relative dimensions of body parts, as well as physiological and behavioral traits23. In arthropods, one such key trait is the amount of fat reserves used for energy storage19,24,25,26,27,28,29,30. Lipids are essential macronutrients for nearly all living organisms, and most animals have the capacity to synthesize and store lipids when resources are abundant31. In insects, lipids are stored as lipid droplets in adipocytes of the fat body, mostly as triglycerides, i.e., fat3132. The fat body is an organ present in arthropods that consists of loose tissues mostly located in the abdomen. The fat body is involved in numerous metabolic functions and plays a key role in storage and release of lipids, with a function similar to the liver in vertebrates31. In insects, most lipids are accumulated during the larval stage and fat content generally peaks right before pupation33,34. Part of the fat reserves are then used for metamorphosis (~ 35% in D. melanogaster34; up to 50% in the fruit fly Ceratitis capitata35). Fat reserves available at the onset of the adult stage thus depend on the amount of fat accumulated during the larval stage and the amount consumed during metamorphosis33,34. For adult insects, fat reserves can have a large effect on fitness, as high fat reserves can positively affect key life history traits, such as longevity36 and fecundity24,25. Lipid reserves further play a key role for many other functions, such as stress resistance, survival during overwintering37, resistance to drought and starvation36,38,39, and increased immunity40.
Several studies have investigated the link between fat content and size in Drosophila melanogaster. For instance, Bryk et al.41 investigated the role of a protein (MAP4K3) on the regulation of body size in D. melanogaster, showing that flies where MAP4K3 was knocked down were smaller and had a lower fat content than controls. In another study, Gasser et al.42 created lines selected for high mortality and compared body size and fat content with lines selected for low mortality conditions. Only slight differences were observed, but generally the high-mortality selected flies were smaller and leaner than flies from the control line. Chippindale et al.43 also compared artificially selected lines, with unselected flies being smaller and containing less fat compared to individuals selected for starvation resistance. Kristensen et al.44 selected flies for 17 generation on a high protein diet and observed that these individuals had a greater body size and contained more fat than their counterparts selected on a standard diet. Juarez-Carreño et al.45 studied the role of candidate genes in body-fat sensing, and showed that larvae where the gene Sema1a (a gene regulating lipid transport and ribosome maturation) was knocked down were bigger and contained more fat than control larvae. These mutants were, however, unable to pupate and initiate metamorphosis. All these studies indeed report that smaller individuals have lower fat reserves (or conversely that bigger individuals have higher fat reserves), but for each of these studies treatments were compared to a control and variation in fat content was not directly correlated to size. Overall, relatively few studies have determined the relationship between body size and fat content in adult D. melanogaster, and immature developmental stages are rarely studied. This lack of knowledge is surprising given that D. melanogaster is a widely used model species in biology46,47,48,49, and a promising emerging model for studying lipid metabolism and obesity50,51,52,53,54,55.
To the best of our knowledge, no data yet exists on the relationship between size and fat content of non-mutant Drosophila prior to emergence. Here, we aimed to test whether fat reserves and size are positively correlated in Drosophila pupae. We further aimed to establish a non-invasive method to estimate pupal fat content (i.e., without destructive sampling) based on size. By manipulating developmental conditions, experienced by D. melanogaster larvae reared under laboratory conditions, in terms of nutrient content and availability, we produced a gradient of pupal sizes that were subsequently measured for total fat content. To be able to expand our findings to more ecologically relevant conditions, we further collected wild Drosophila species and tested for a correlation between pupal size and fat content. Our results show that there is a strong positive correlation between pupal size and fat content, both in laboratory-reared and field-caught Drosophila. Estimates of pupal size thus provide a good proxy for pupal fat content in several Drosophila species.
Methods
Insect maintenance and developmental conditions
Our Drosophila melanogaster (Diptera: Drosophilidae) stock originated from a culture that was set up in 1994 from field collections in Sainte-Foy-les-Lyon (France), kindly provided by Patricia Gibert (Claude Bernard University, Lyon, France) in 2016. Larvae were maintained in flasks with continuous access to food medium (60 ml/flask; composition: 20 g agar, 35 g yeast, 50 g sugar, 5 ml nipagin containing 100 g 4-methyl hydroxyl benzoate in 1 l 96% alcohol, and 5 ml propionic acid per liter water). After emergence, adults were maintained in cages (50 × 50 × 50 cm) with continuous access to the same food medium that was replaced every 3 to 4 days. Individuals were kept at a temperature of 23 °C, a relative humidity of 75%, and a photoperiod of L:D 16:8, unless stated otherwise.
To generate pupae that varied in size, we manipulated nutrient content or availability during fly development using three methods: starvation, crowding, and modification of the sugar content in the medium. To do so, flies were allowed to lay eggs during 24 h. Approximately 100 eggs were then collected using a fine paintbrush and distributed in vials (containing 10 ml food medium, a surplus quantity of food to avoid competition between larvae) that differed in nutrient content. To change the nutrient content of the food medium, the standard medium (described above; denoted as 1/1 = 1 part sugar/1 part yeast) was modified to contain either twice more (i.e., 2/1; n = 3 vials) or no sugar (i.e., 0/1; n = 1 vial). As a control, 4 vials of the standard 1/1 medium were also prepared. Nutrient availability was altered by allowing 100 larvae per vial to feed on the standard 1/1 medium for 2 or 3 days. Larvae were then starved by transferring them to a new vial containing a medium without sugar or yeast (starvation after 2d and 3d, respectively; n = 3 vials for both treatments). A third treatment was added where the number of individuals per vial was increased, leading to crowding, and hence a reduction in nutrient availability. To create crowding conditions, 300 eggs were counted under a stereomicroscope and transferred to a vial containing 1 ml of the standard 1/1 medium. This egg density (300 eggs ml−1) was chosen, because previous work showed that D. melanogaster experiences strong crowding under these conditions56. All vials were inspected twice a day for newly formed pupae to ensure that pupae were collected within one day after pupation. Pupae were collected individually, their development time recorded, and then frozen at − 20 °C until further processing. Only rarely vials could not be inspected within 24 h, which was taken into account in the statistical analyses (see below).
Collection of Drosophila pupae from the field
In addition to manipulating the size of laboratory-reared D. melanogaster, we aimed to investigate the correlation between size and fat content of wild Drosophila individuals. To do so, we collected individuals from the field using banana-bait traps. Each trap consisted of a 0.75 l plastic box, with an opening in the lid. The opening was covered by a fine net mesh of ~ 1 mm. Three traps were prepared, each containing half a banana, as well as a mixture of live baker’s yeast and apple cider vinegar. Each trap was then attached to a tree with the opening facing downward for 1 week in a backyard in Leuven (Belgium). Temperature and humidity were monitored with a thermometer-hygrometer Ibutton (Maxim Integrated) placed inside one of the traps (Supplementary Fig. 1a). For each trap, bananas containing eggs and larvae were placed in a flask containing the standard 1/1 food medium (three flasks in total). In addition to banana-bait traps, we also collected cherries infested by Drosophila suzukii from a cherry tree at the same location in Leuven. Cherries were distributed among 6 flasks containing the standard 1/1 food medium. All flasks were then kept in a cage and placed outside our facility (facing North and in continuous shadow). Temperature and humidity inside the cage were also monitored using an Ibutton device (Supplementary Fig. 1b). Pupae were subsequently collected as described above for laboratory-reared flies. Visual species identification based on pupae is very difficult for most drosophilid species, but the pupal shape of D. suzukii is easily recognizable57. Drosophila obtained from cherries were, therefore, identified as D. suzukii based on the shape of their respiratory tubes. Pupae obtained from banana-bait traps remained largely unidentified (referenced hereafter as “other species”). To have an estimation of the species present in each trap, adults were collected, killed and stored at − 20 °C, after which each species was identified58,59. Adult D. melanogaster, D. simulans, D. hydei and D. subobscura were present in the traps.
Pupal size measurement and neutral lipid quantification
To measure pupal size, a similar procedure as described in Ref.18 was followed. In short, each pupa was photographed individually using a camera linked to a stereomicroscope (Leica, SAPO). The total area of the pupal case was then measured using FiJi software (ImageJ v2.1.051;60). Quantification of the neutral lipid fraction (i.e., fat or triglycerides) was done using the protocol described in Ref.61. Briefly, pupae were dried in an oven at 60 °C for 3 days, after which dry weight was determined using a microbalance (Mettler Toledo, MT5). Each pupa was then placed into a glass tube with 4 ml of diethyl ether for 24 h. Pupae were then dried again at 60 °C for 3 days and pupal dry weight determined again, giving the neutral lipid-free dry weight. The absolute amount of fat (total neutral lipids in µg/pupae) was then obtained by subtracting the lipid-free dry weight from the lipid-containing dry weight. Ether extraction is an efficient method for fat quantification, as it extracts predominantly neutral lipids62, including triglycerides that represent lipids for energy storage31.
Statistical analyses
All analyses were performed with R version 4.0.263. For laboratory-reared pupae, time to pupation, pupal size, and fat content were analyzed using generalized linear mixed-effects models (GLMM) with poisson (pupation time) or gamma (size and lipid content) error distributions. For all GLMMs, the fixed effect diet was analyzed using analysis of deviance with the “Anova” function from the “car” package64. Differences between diet groups were then identified using estimated marginal means comparisons (EMMs) using the “emmeans” function65. Vial number and pupa collection time were included as random factors (for pupation time, pupal size, and lipid content), as was the extraction run (for fat content). For each GLMM, models were simplified by removing random factors step by step, and compared using an anova with the “model.sel” function from the “MuMIn” package66. When models did not differ significantly and the AIC was smaller (delta AIC ≥ 2), the simplified model was kept based on methods of model simplification presented in Ref.67. The final model for the time to pupation included vial number as a random factor and the final model for fat content included pupae collection time and extraction run as random factors. No random factor was included in the final model for size. Final models are presented in Table 1.
The correlation between pupal size and fat content was analyzed using linear mixed-effects models (LMM) for laboratory and field-collected pupae separately. For laboratory pupae, the fixed factor diet was used as a co-variable. Vial number, pupae collection time, and extraction run were included as random factors. The LMM was then simplified as described above for GLMMs, and only vial number was kept as a random factor in the final model (Table 1). The LMM was further simplified for interactions between fixed factors. For field pupae, species was used as a co-variable (D. suzukii or “other species”). Flask number, pupae collection time and extraction run were included as random factors. The LMM was simplified as described above, but all random effects and interactions were included in the final model (Table 1). For both LMMs, statistical significance of each variable was determined using analysis of deviance. R2 were then calculated for both LMMs using the “r.squaredGLMM” function from the “MuMIn” package. For mixed-effects models, R2 comes in two types: marginal and conditional. Marginal R2 represents the variance explained by the fixed effects of the model, while conditional R2 is interpreted as a variance explained by the entire model, including both fixed and random effects. Both the marginal and conditional R2 are reported.
Results
Developmental conditions affect pupation time, pupal size and lipid content
The first aim of this study was to produce pupae of different sizes and fat content. To do so, we manipulated the environmental conditions of developing D. melanogaster larvae by decreasing nutrient content or availability of the food medium. All developmental conditions allowed individuals to pupate (see Table 2 for the number of pupae formed per condition, and Supplementary Table 1 for details on the number of pupae formed in each replicate). Diet strongly affected pupal size (Fig. 1; GLMM, χ2 = 665.43, df = 5, p value < 0.001). The biggest pupae developed on the high-sugar (2/1; mean ± sd = 2.82 mm2 ± 0.49; Fig. 1b) and standard medium (control, 1/1; 2.31 mm2 ± 0.32; Fig. 1b), while the smallest pupae were found in the starvation treatments (starvation following 2 or 3 days feeding; 1.38 mm2 ± 0.26 and 1.33 mm2 ± 0.15, respectively; Fig. 1b). Crowding and the no-sugar medium (0/1) led to pupae of intermediate sizes (1.62 mm2 ± 0.29 and 1.95 ± 0.37 mm2, respectively; Fig. 1b).
Diet also had a major effect on pupal fat content (GLMM, χ2 = 412.92, df = 5, p.value < 0.001; Fig. 2), with pupae that developed on the nutrient-rich media (2/1 and 1/1; mean ± sd = 106.76 µg ± 26.21 and 92.81 µg ± 30.14, respectively; Fig. 2) showing the highest lipid content, while the leanest pupae (i.e., with the lowest lipid content) were obtained when larvae were starved (starvation after 2d and 3d; 31.73 µg ± 18.58 and 37.36 µg ± 11.31; Fig. 2) and under crowding conditions (41.12 µg ± 18.39; Fig. 2). Pupae developing on the no-sugar medium (0/1) showed an intermediate lipid content (55.34 µg ± 24.49; Fig. 2).
Dietary conditions (nutrient content and/or availability) during development also affected time to pupation, which ranged on average from 6.64 (± 0.49) days for larvae that developed on the sugar-rich (2/1) medium to 17.73 (± 2.68) days for starved larvae following 2 days of feeding (Table 2, GLMM, χ2 = 77.196, df = 5, p value < 0.001). Starvation after 2 days of feeding and the no-sugar medium increased development time compared to controls (standard medium, 1/1). Starvation after 3 days of feeding, crowding, and the sugar-rich medium (2/1) did not induce significant changes in development time compared to controls (1/1), but the sugar-rich medium (2/1) shortened development time compared to crowding and starvation after 3 days (Table 2).
Pupal size and fat content are correlated in laboratory-reared and field-caught Drosophila pupae
We used the variation in pupal size and fat content produced by the different environmental conditions of developing D. melanogaster larvae to test for a positive correlation between the two traits. Pupal size and fat content were strongly correlated for laboratory-reared individuals, with bigger pupae containing more fat (LMM, χ2 = 190.86, df = 1, p value < 0.001, marginal R2 = 0.57, conditional R2 = 0.85, N = 375, Fig. 3a). Diet did not affect the correlation between pupal size and fat content, meaning that the correlation coefficient was similar when different diets were compared (LMM, χ2 = 2.47, df = 5, p value = 0.78).
We further measured pupal size and fat content of several field-caught Drosophila species (D. suzukii pupae, n = 78; other species, n = 732; see Supplementary Table 1 for the number of pupae formed in each replicate). For field-collected pupae, we also found a strong correlation between size and fat content (LMM, χ2 = 688.08, df = 1, p.value < 0.001, marginal R2 = 0.47, conditional R2 = 0.78, N = 810, Fig. 3b). Of all species measured, D. suzukii pupae contained the most fat (LMM, χ2 = 27.43, df = 1, p value < 0.001; Fig. 3b). We further found an interaction effect between pupal size and species, as the slope of the regression was higher for D. suzukii pupae compared to the other species (LMM, χ2 = 4.13, df = 1, p value < 0.05, Fig. 3b).
Discussion
In this study, we manipulated nutrient content and nutrient availability of developing D. melanogaster larvae to produce phenotypes that differ in pupal size and fat content. Starvation, dietary restriction, and crowding generally increased development time, but reduced pupal size and fat content. These conditions are indeed known to decrease body size and fat reserves in adult D. melanogaster20,68,69,70, but also in other insects71,72,73,74. In this paper, the conditions that produced the smallest and leanest individuals were crowding and starvation. Crowding both decreases food availability (nutrients are consumed by conspecifics) and food quality (e.g., overconsumption of accumulated toxic waste produced by conspecifics in the food)75. For starvation, previous work showed that starvation of D. melanogaster larvae before 3 days of age (i.e., 70 h) can provoke major hormonal dysregulation that impedes reaching the minimal viable weight to initiate metamorphosis, thus leading to death20,22. In our study, indeed, only few individuals pupated under the harshest starvation condition (only 48 h of feeding after egg laying). Crowding and starvation are complex multifactorial stressors that can explain why these treatments led to the smallest pupae. The crowding conditions used in this study (300 eggs for 1 ml of food) represent a high density for developing individuals, but D. melanogaster larvae can develop in conditions of up to 1000 eggs for 1 ml of food. Survival, however, decreases dramatically under those conditions (with only 1.25% of individuals pupating56). It would be interesting to test how even higher densities than the one used in this study would influence size and fat content of D. melanogaster.
Our data further showed no significant differences in size, fat content and development time between controls (1/1) and individuals developing on the sugar-rich medium (2/1). This is in contrast to several studies on D. melanogaster that showed that sugar-rich diets increased the fat content of adults76,77 and larvae55, decreased body size70,76, and significantly increased time to pupation52,77. High sugar diets can further have deleterious effects on fitness, reducing fecundity76 or egg to pupa viability52. Absence of significant differences between our sugar-rich medium (2/1) and controls (1/1) can result from the concentration of sugar that was used for our sugar-rich medium (10%). Indeed, in the study of Klepsatel et al.76, major differences in body size and fecundity were observed for Drosophila that developed on a medium containing 25% sugar. Furthermore, when compared with individuals from the no-sugar medium (0/1), 2/1 pupae were bigger, contained more fat and developed faster, showing that variation in sugar quantity had a significant impact on these life history traits. An increase in fat reserves in relation to sugar quantity in the diet can be expected, because excess sugars from the diet are metabolized through glycolysis and the tricarboxylic acid cycle to produce acetyl coenzyme A (acetyl-CoA). Acetyl-CoA is then converted to triglycerides that are stored in lipid droplets within adipocytes in the fat body31,78. Overall, our findings, together with previous studies20,68,69,70, suggest that the nutritional conditions experienced by D. melanogaster individuals during the larval stage directly affect the size and fat content of pupae and adults that can have major consequences for fitness.
Pupal and adult size are intimately linked with fitness in insects2,79. In Aedes mosquitoes, for example, an increase in size of 25% leads to an increase in egg production of 100%25,80, while in D. melanogaster an increase in size of 43% can increase ovariole numbers by 290%81. In moths, larger individuals live longer, with a body size (wing length) increase of 25% leading to an increase of 79% percent in lifespan82. Similarly, in D. pseudoobscura a 24% increase in size can lead to a 32% increase in longevity9, while conversely decreasing sugar proportions in the diet decreases fat content of the flies by 26%, in turn decreasing median longevity by 122%36. Having large fat reserves can, however, also comes at a cost. D. melanogaster was found to show obesity-like phenotypes, with pathologies similar to those associated with obesity in humans, such as heart (dorsal vessel) failure, decreased endurance or metabolic dysregulation50,55. Despite these considerations, higher fitness in Drosophila is often associated with size and fat content (females with larger body size and higher fat reserves lay more eggs and live longer). This is clearly demonstrated by numerous studies where individuals artificially selected for a larger body size had higher fitness traits, including female fecundity10,83, longevity84 or male reproductive success10,84,85. The positive relationship between body size and fitness is not limited to insects, but has also been found in vertebrates, including reptiles86,87,88 and mammals89,90.
The main aim of this study was to use variation in size and fat content of Drosophila pupae (as a consequence of modifying nutritional conditions during developmental) to test the hypothesis that fat content is positively correlated with pupal size. We further collected wild pupae to observe if this correlation was also found in natural Drosophila populations. We observed that for field-caught Drosophila, fat content of 96% of D. suzukii pupae and 100% of pupae from other species were within the range of fat contents produced by our treatments (mainly induced by the nutrient-rich media, i.e., 2/1 and 1/1; see Fig. 3). Validating our hypothesis, pupal fat content was indeed positively correlated to pupal size both in laboratory-reared D. melanogaster and field-caught Drosophila species. Other works however, showed that some conditions can promote an opposite trend. Indeed, Kristensen et al.44 showed that flies developing on a protein enriched medium were larger than controls (measured as dry weight), but had fewer lipid reserves (in % of fly body mass). In our study, we increased the proportion of sugar in the diet, but not the proportion of protein. It would be interesting to test how a gradient of protein concentrations can affect the relationship between body size and total fat content in D. melanogaster.
In this study, we did not separate male and female pupae. Yet, we observed a strong relationship between fat content and size, with low variability for both laboratory and field pupae, as highlighted by the high conditional R2 (0.85 and 0.78, respectively). In arthropods, females tend to have higher fat reserves than males, but females also generally have a larger body size; hence the relationship between size and fat reserves is expected to be similar for males and females30. Pupal size, an easy and reliable measurement, is therefore a robust predictor of fat content for both laboratory-reared and field-caught Drosophila species. In insects, several body measurements, such as head width91, tibia length5, wing length92, thorax or abdomen length93, and pupal size17 are used as a proxy for body size, fat content, and fitness. As insect pupae are easy to handle in general, measuring pupal size represents a convenient and non-invasive method to estimate fat content and potentially fitness. By linking pupal size to fat content, our results can be of interest for future eco-evolutionary and physiological studies, because this method allows to estimate an individual’s energetic reserves with the option to use the same individual for further experimentation and life history measurements.
Drosophila larvae and pupae are hosts to numerous parasitic wasp (i.e., parasitoid) species94. During development, a parasitoid consumes fat from only a single host insect and fat stores are often not replenished during adult life95,96. Host resources available for developing parasitoids can thus in turn have major consequences for adult wasp life history traits97. Measuring pupal size of Drosophila, therefore, offers the possibility to estimate the amount of fat available for developing parasitoids98. Our data can also be of interest from an applied perspective, as the spotted wing drosophila (D. suzukii) is a major pest of red berries. Several studies are ongoing to develop biological control methods to fight this pest, such as the sterile or incompatible insect techniques99,98,101. The success of these techniques, which require mass release of sterile/incompatible males in infested areas, relies on the quality of released males102. In mass rearing facilities, measurements are made at several critical points to check insect quality. Our data shows that pupal size is a reliable estimate of fat content of Drosophila flies, including D. suzukii, and could, therefore, represent a good quality measurement of mass-produced individuals.
Data availability
All data used on this article are available on demand from the corresponding author.
References
Parker, J. & Johnston, L. A. The proximate determinants of insect size. J. Biol. 5, 15 (2006).
Honěk, A. Intraspecific variation in body size and fecundity in insects: A general relationship. Oikos 66, 483 (1993).
Kingsolver, J. G. & Huey, R. B. Size, temperature, and fitness: Three rules. Evol. Ecol. Res. 10, 251–268 (2008).
Beukeboom, L. W. Size matters in insects—An introduction. Entomol. Exp. Appl. 166, 2–3 (2018).
West, S. A., Flanagan, K. E. & Godfray, H. C. J. The relationship between parasitoid size and fitness in the field, a study of Achrysocharoides zwoelferi (Hymenoptera: Eulophidae). J. Anim. Ecol. 65, 631–639 (1996).
Sagarra, L. A., Vincent, C. & Stewart, R. K. Body size as an indicator of parasitoid quality in male and female Anagyrus kamali (Hymenoptera: Encyrtidae). Bull. Entomol. Res. 91, 363–367 (2001).
Ellers, J., Alphen, J. J. M. V. & Sevenster, J. G. A field study of size–fitness relationships in the parasitoid Asobara tabida. J. Anim. Ecol. 67, 318–324 (1998).
Armbruster, P. & Hutchinson, R. A. Pupal mass and wing length as indicators of fecundity in Aedes albopictus and Aedes geniculatus (Diptera: Culicidae). J. Med. Entomol. 39, 699–704 (2002).
Tantawy, A. O. & Vetukhiv, M. O. Effects of size on fecundity, longevity and viability in populations of Drosophila pseudoobscura. Am. Nat. 94, 395–403 (1960).
Lefranc, A. & Bundgaard, J. The influence of male and female body size on copulation duration and fecundity in Drosophila melanogaster. Hereditas 132, 243–247 (2004).
Atkinson, D. Temperature and organism size: A biological law for ectotherms? Adv. Ecol. Res. 25, 1–58 (1994).
Poças, G. M., Crosbie, A. E. & Mirth, C. K. When does diet matter? The roles of larval and adult nutrition in regulating adult size traits in Drosophila melanogaster. J. Insect Physiol. 139, 104051. https://doi.org/10.1016/j.jinsphys.2020.104051 (2020).
Tammaru, T. Determination of adult size in a folivorous moth: constraints at instar level? Ecol. Entomol. 23, 80–89 (1998).
Miller, R. S. & Thomas, J. L. The effects of larval crowding and body size on the longevity of adult Drosophila melanogaster. Ecology 39, 118–125 (1958).
Nijhout, H. F. The control of body size in insects. Dev. Biol. 261, 1–9 (2003).
Shingleton, A. W., Mirth, C. K. & Bates, P. W. Developmental model of static allometry in holometabolous insects. Proc. R. Soc. B 275, 1875–1885 (2008).
Koenraadt, C. J. M. Pupal dimensions as predictors of adult size in fitness studies of Aedes aegypti (Diptera: Culicidae). J. Med. Entomol. 45, 331–336 (2008).
Stillwell, R. C., Dworkin, I., Shingleton, A. W. & Frankino, W. A. Experimental manipulation of body size to estimate morphological scaling relationships in Drosophila. JoVE 56, 3162. https://doi.org/10.3791/3162 (2011).
Shin, S.-M., Akram, W. & Lee, J.-J. Effect of body size on energy reserves in Culex pipiens pallens females (Diptera: Culicidae). Entomol. Res. 42, 163–167 (2012).
Mirth, C. K. & Riddiford, L. M. Size assessment and growth control: How adult size is determined in insects. BioEssays 29, 344–355 (2007).
Chown, S. L. & Gaston, K. J. Body size variation in insects: A macroecological perspective. Biol. Rev. 85, 139–169 (2010).
Beadle, G. W., Tatum, E. L. & Clancy, C. W. Food level in relation to rate of development and eye pigmentation in Drosophila melanogaster. Biol. Bull. 75, 447–462 (1938).
Gayon, J. History of the concept of allometry1. Am. Zool. 40, 748–758 (2000).
Takken, W. et al. Larval nutrition differentially affects adult fitness and Plasmodium development in the malaria vectors Anopheles gambiae and Anopheles stephensi. Parasit. Vectors 6, 345 (2013).
Briegel, H. Metabolic relationship between female body size, reserves, and fecundity of Aedes aegypti. J. Insect Physiol. 36, 165–172 (1990).
Ellers, J. Fat and eggs: An alternative method to measure the trade-off between survival and reproduction in insect parasitoids. Neth. J. Zool. 3, 227–235 (1996).
González-Tokman, D. et al. Energy storage, body size and immune response of herbivore beetles at two different elevations in Costa Rica. Rev. Biol. Trop. 67, 608–620 (2019).
Timmermann, S. E. & Briegel, H. Larval growth and biosynthesis of reserves in mosquitoes. J. Insect Physiol. 45, 461–470 (1999).
Strohm, E. Factors affecting body size and fat content in a digger wasp. Oecologia 123, 184–191 (2000).
Lease, H. M. & Wolf, B. O. Lipid content of terrestrial arthropods in relation to body size, phylogeny, ontogeny and sex. Physiol. Entomol. 36, 29–38 (2011).
Arrese, E. L. & Soulages, J. L. Insect fat body: Energy, metabolism, and regulation. Annu. Rev. Entomol. 55, 207–225 (2010).
Kühnlein, R. P. Lipid droplet-based storage fat metabolism in Drosophila. J. Lipid Res. 53, 1430–1436 (2012).
Church, R. B. & Robertson, F. W. A biochemical study of the growth of Drosophila melanogaster. J. Exp. Zool. 162, 337–351 (1966).
Merkey, A. B., Wong, C. K., Hoshizaki, D. K. & Gibbs, A. G. Energetics of metamorphosis in Drosophila melanogaster. J. Insect Physiol. 57, 1437–1445 (2011).
Nestel, D., Tolmasky, D., Rabossi, A. & Quesada-Allué, L. A. Lipid, carbohydrates and protein patterns during metamorphosis of the Mediterranean fruit fly, Ceratitis capitata (Diptera: Tephritidae). Ann. Entomol. Soc. Am. 96, 237–244 (2003).
Lee, K. P. & Jang, T. Exploring the nutritional basis of starvation resistance in Drosophila melanogaster. Funct. Ecol. 28, 1144–1155 (2014).
Hahn, D. A. & Denlinger, D. L. Meeting the energetic demands of insect diapause: Nutrient storage and utilization. J. Insect Physiol. 53, 760–773 (2007).
Tejeda, M. T. et al. Effects of size, sex and teneral resources on the resistance to hydric stress in the tephritid fruit fly Anastrepha ludens. J. Insect Physiol. 70, 73–80 (2014).
Hoffmann, A. A., Hallas, R., Anderson, A. R. & Telonis-Scott, M. Evidence for a robust sex-specific trade-off between cold resistance and starvation resistance in Drosophila melanogaster. J. Evol. Biol. 18, 804–810 (2005).
Alaux, C., Ducloz, F., Crauser, D. & Le Conte, Y. Diet effects on honeybee immunocompetence. Biol. Lett. 6, 562–565 (2010).
Bryk, B., Hahn, K., Cohen, S. M. & Teleman, A. A. MAP4K3 regulates body size and metabolism in Drosophila. Dev. Biol. 344, 150–157 (2010).
Gasser, M., Kaiser, M., Berrigan, D. & Stearns, S. C. Life-history correlates of evolution under high and low adult mortality. Evolution 54, 1260–1272 (2000).
Chippindale, A. K., Chu, T. J. F. & Rose, M. R. Complex trade-offs and the evolution of starvation resistance in Drosophila melanogaster. Evolution 50, 753 (1996).
Kristensen, T. N., Overgaard, J., Loeschcke, V. & Mayntz, D. Dietary protein content affects evolution for body size, body fat and viability in Drosophila melanogaster. Biol. Lett. 7, 269–272 (2011).
Juarez-Carreño, S. et al. Body-fat sensor triggers ribosome maturation in the steroidogenic gland to initiate sexual maturation in Drosophila. Cell Rep. 37, 109830 (2021).
Markow, T. A. The secret lives of Drosophila flies. Elife 4, e06793 (2015).
Choma, M. A., Suter, M. J., Vakoc, B. J., Bouma, B. E. & Tearney, G. J. Physiological homology between Drosophila melanogaster and vertebrate cardiovascular systems. Dis. Model. Mech. 4, 411–420 (2011).
Morgan, T. H., Sturtevant, A. H., Muller, H. J. & Bridges, C. B. The Mechanism of Mendelian Heredity (H. Holt, 1923).
Dobzhansky, T. The influence of the quantity and quality of chromosomal material on the size of the cells in Drosophila melanogaster. Wilhelm Roux Arch. Entwickl Mech. Org. 115, 363–379 (1929).
Musselman, L. P. & Kühnlein, R. P. Drosophila as a model to study obesity and metabolic disease. J. Exp. Biol. 221, 163881 (2018).
DiAngelo, J. R. & Birnbaum, M. J. Regulation of fat cell mass by insulin in Drosophila melanogaster. Mol. Cell. Biol. 29, 6341–6352 (2009).
Rovenko, B. M. et al. High sucrose consumption promotes obesity whereas its low consumption induces oxidative stress in Drosophila melanogaster. J. Insect Physiol. 79, 42–54 (2015).
Hardy, C. M. et al. Obesity-associated cardiac dysfunction in starvation-selected Drosophila melanogaster. Am. J. Physiol.-Regul. Integr. Compar. Physiol. 309, R658–R667 (2015).
Hardy, C. M. et al. Genome-wide analysis of starvation-selected Drosophila melanogaster—A genetic model of obesity. Mol. Biol. Evol. 35, 50–65 (2018).
Musselman, L. P. et al. A high-sugar diet produces obesity and insulin resistance in wild-type Drosophila. Dis. Model. Mech. 4, 842–849 (2011).
Henry, Y., Renault, D. & Colinet, H. Hormesis-like effect of mild larval crowding on thermotolerance in Drosophila flies. J. Exp. Biol. 221, 169342 (2018).
Bulletin, E. P. P. O. Drosophila suzukii. EPPO Bull. 43, 417–424 (2013).
Bächli, G., Vilela, C. R., Escher, S. A. & Saura, A. The Drosophilidae (Diptera) of Fennoscandia and Denmark (Brill Academic Publishers, 2004).
Markow, T. A. & O’Grady, P. M. Drosophila: A Guide to Species Identification and Use (Elsevier, 2006).
Schindelin, J. et al. Fiji: An open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).
Visser, B. et al. Variation in lipid synthesis, but genetic homogeneity, among Leptopilina parasitic wasp populations. Ecol. Evol. 8, 7355–7364 (2018).
Williams, C. M., Thomas, R. H., MacMillan, H. A., Marshall, K. E. & Sinclair, B. J. Triacylglyceride measurement in small quantities of homogenised insect tissue: Comparisons and caveats. J. Insect Physiol. 57, 1602–1613 (2011).
R Core Team. R: A Language and Environment for Statistical Computing. (R Foundation for Statistical Computing, 2020).
Fox, J. & Weisberg, S. An R Companion to Applied Regression 2nd edn. (Sage, 2011).
Lenth, R., Singmann, H., Love, J., Buerkner, P. & Herve, M. Emmeans: Estimated marginal means, aka least-squares means. R Package Version 1, 3 (2018).
Burnham, K. P. & Anderson, D. R. A practical information-theoretic approach. In Model Selection and Multimodel Inference (ed. Burnham, K. P.) (Springer, 2002).
Crawley, M. J. The R Book (Wiley, 2007).
Borash, D. J. & Ho, G. T. Patterns of selection: Stress resistance and energy storage in density-dependent populations of Drosophila melanogaster. J. Insect Physiol. 47, 1349–1356 (2001).
Klepsatel, P., Procházka, E. & Gáliková, M. Crowding of Drosophila larvae affects lifespan and other life-history traits via reduced availability of dietary yeast. Exp. Gerontol. 110, 298–308 (2018).
Henry, Y., Overgaard, J. & Colinet, H. Dietary nutrient balance shapes phenotypic traits of Drosophila melanogaster in interaction with gut microbiota. Comp. Biochem. Physiol. A: Mol. Integr. Physiol. 241, 110626 (2020).
Ireland, S. & Turner, B. The effects of larval crowding and food type on the size and development of the blowfly, Calliphora vomitoria. Forensic Sci. Int. 159, 175–181 (2006).
Saunders, D. S. & Bee, A. Effects of larval crowding on size and fecundity of the blow fly, Calliphora vicina (Diptera: Calliphoridae). EJE 92, 615–622 (2013).
Ziegler, R. Changes in lipid and carbohydrate metabolism during starvation in adult Manduca sexta. J. Comp. Physiol. B 161, 125–131 (1991).
Ojeda-Avila, T., Arthur Woods, H. & Raguso, R. A. Effects of dietary variation on growth, composition, and maturation of Manduca sexta (Sphingidae: Lepidoptera). J. Insect Physiol. 49, 293–306 (2003).
Borash, D. J., Gibbs, A. G., Joshi, A. & Mueller, L. D. A genetic polymorphism maintained by natural selection in a temporally varying environment. Am. Nat. 151, 148. https://doi.org/10.1086/286108 (1998).
Klepsatel, P., Knoblochová, D., Girish, T. N., Dircksen, H. & Gáliková, M. The influence of developmental diet on reproduction and metabolism in Drosophila. BMC Evol. Biol. 20, 93 (2020).
Matzkin, L. M., Johnson, S., Paight, C., Bozinovic, G. & Markow, T. A. Dietary protein and sugar differentially affect development and metabolic pools in ecologically diverse Drosophila. J. Nutr. 141, 1127–1133 (2011).
Musselman, L. P. et al. Role of fat body lipogenesis in protection against the effects of caloric overload in Drosophila. J. Biol. Chem. 288, 8028–8042 (2013).
Reeve, M. W., Fowler, K. & Partridge, L. Increased body size confers greater fitness at lower experimental temperature in male Drosophila melanogaster. J. Evol. Biol. 13, 836–844 (2000).
Lounibos, L. P. et al. Does temperature affect the outcome of larval competition between Aedes aegypti and Aedes albopictus?. J. Vector Ecol. 27, 86–95 (2002).
Bergland, A. O., Genissel, A., Nuzhdin, S. V. & Tatar, M. Quantitative trait loci affecting phenotypic plasticity and the allometric relationship of ovariole number and thorax length in Drosophila melanogaster. Genetics 180, 567–582 (2008).
Holm, S. et al. A comparative perspective on longevity: The effect of body size dominates over ecology in moths. J. Evol. Biol. 29, 2422–2435 (2016).
Nunney, L. The response to selection for fast larval development in Drosophila melanogaster and its effect on adult weight: An example of a fitness trade-off. Evolution 50, 1193–1204 (1996).
Partridge, L. & Farquhar, M. Lifetime mating success of male fruitflies (Drosophila melanogaster) is related to their size. Anim. Behav. 31, 871–877 (1983).
Markow, T. A. & Ricker, J. P. Male size, developmental stability, and mating success in natural populations of three Drosophila species. Heredity 69, 122–127 (1992).
Wikelski, M. & Romero, L. M. Body size, performance and fitness in galapagos marine iguanas. Integr. Comp. Biol. 43, 376–386 (2003).
van Buskirk, J. & Crowder, L. B. Life-history variation in marine turtles. Copeia 1994, 66–81 (1994).
Broderick, A. C., Glen, F., Godley, B. J. & Hays, G. C. Variation in reproductive output of marine turtles. J. Exp. Mar. Biol. Ecol. 288, 95–109 (2003).
Wauters, L. A. et al. Effects of spatio-temporal variation in food supply on red squirrel Sciurus vulgaris body size and body mass and its consequences for some fitness components. Ecography 30, 51–65 (2007).
Lindström, J. Early development and fitness in birds and mammals. Trends Ecol. Evol. 14, 343–348 (1999).
Reim, C., Teuschl, Y. & Blanckenhorn, W. U. Size-dependent effects of temperature and food stress on energy reserves and starvation resistance in yellow dung flies. Evol. Ecol. Res. 8, 1215–1234 (2006).
Kölliker-Ott, U. M., Blows, M. W. & Hoffmann, A. A. Are wing size, wing shape and asymmetry related to field fitness of Trichogramma egg parasitoids? Oikos 100, 563–573 (2003).
Knapp, M. Relative importance of sex, pre-starvation body mass and structural body size in the determination of exceptional starvation resistance of Anchomenus dorsalis (Coleoptera: Carabidae). PLoS ONE 11, e0151459 (2016).
Lue, C.-H. et al. DROP: Molecular voucher database for identification of Drosophila parasitoids. Mol. Ecol. Resour. 21, 2437–2454 (2021).
Visser, B. et al. Loss of lipid synthesis as an evolutionary consequence of a parasitic lifestyle. Proc. Natl. Acad. Sci. 107, 8677–8682 (2010).
Visser B et al. Why do many parasitoids lack adult triglyceride accumulation, despite functioning fatty acid biosynthesis machinery? EcoEvoRxiv: https://doi.org/10.32942/osf.io/zpf4j
Arakawa, R., Miura, M. & Fujita, M. Effects of host species on the body size, fecundity, and longevity of Trissolcus mitsukurii (Hymenoptera: Scelionidae), a solitary egg parasitoid of stink bugs. Appl. Entomol. Zool. 39, 177–181 (2004).
Visser, B., Alborn, H.T., Rondeaux, S. et al. Phenotypic plasticity explains apparent reverse evolution of fat synthesis in parasitic wasps. Sci Rep 11, 7751 (2021). https://doi.org/10.1038/s41598-021-86736-8.
Krüger, A. P. et al. Effects of irradiation dose on sterility induction and quality parameters of Drosophila suzukii (Diptera: Drosophilidae). J. Econ. Entomol. 111, 741–746 (2018).
Nikolouli, K. et al. Sterile insect technique and Wolbachia symbiosis as potential tools for the control of the invasive species Drosophila suzukii. J. Pest Sci. 91, 1–15 (2017).
Nikolouli, K., Sassù, F., Mouton, L., Stauffer, C. & Bourtzis, K. Combining sterile and incompatible insect techniques for the population suppression of Drosophila suzukii. J. Pest Sci. 93, 647–661 (2020).
Calkins, C. O. & Parker, A. G. Sterile insect quality. In Sterile Insect Technique (eds Dyck, V. A. et al.) 269–296 (Springer, 2005).
Funding
This work was supported by the Fyssen foundation and the Fonds National de la Recherche Scientifique F.R.S.-FNRS (Grants “J.0190.21” and “T.0186.20”).
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T.E. and B.V. acquired the funding. T.E., C.M.N. and B.V. conceptualized the study. T.E. and V.L. performed the experiments. T.E. performed all statistical analysis and prepared all figures. T.E. and B.V. wrote the first version of the manuscript, which was validated by all authors.
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Enriquez, T., Lievens, V., Nieberding, C.M. et al. Pupal size as a proxy for fat content in laboratory-reared and field-collected Drosophila species. Sci Rep 12, 12855 (2022). https://doi.org/10.1038/s41598-022-15325-0
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DOI: https://doi.org/10.1038/s41598-022-15325-0
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