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First published online October 5, 2007
Journal of Experimental Biology 210, 3547-3558 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.006924

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Natural variation in food acquisition mediated via a Drosophila cGMP-dependent protein kinase

Karla R. Kaun¹, Craig A. L. Riedl¹, Munmun Chakaborty-Chatterjee¹, Amsale T. Belay¹, Scott J. Douglas¹, Allen G. Gibbs² and Marla B. Sokolowski^1,*

¹ Department of Biology, University of Toronto, 3359 Mississauga Road, Mississauga, Ontario, L5L 1C6, Canada
² School of Life Sciences, University of Nevada, 4505 Maryland Parkway, Las Vegas, NV 89154-4004, USA

^* Author for correspondence (e-mail: marla.sokolowski{at}utoronto.ca)

Accepted 2 July 2007

	Summary

TOP Summary Introduction Materials and methods Results and Discussion References

 
In natural environments where food abundance and quality can change drastically over time, animals must continuously alter their food acquisition strategies. Although genetic variation contributes to this plasticity, the specific genes involved and their interactions with the environment are poorly understood. Here we report that natural variation in the Drosophila gene, foraging (for), which encodes a cGMP-dependent protein kinase (PKG), affects larval food acquisition in an environmentally dependent fashion. When food is plentiful, the wild-type rover (for^R) allele confers lower food intake and higher glucose absorption than both the wild-type sitter (for^s) allele and the mutant for^s2 allele. When food is scarce, for^R, for^s and for^s2 larvae increase food intake to a common maximal level, but for^R larvae retain their increased absorption efficiency. Changes in for expression can induce corrective behavioral modifications in response to food deprivation. When reared in environments with low food levels, for^R larvae have higher survivorship and faster development than for^s and for^s2 larvae. Together, these results show that natural variation in for has far reaching implications affecting a suite of phenotypes involved in the regulation of food acquisition.

Key words: Drosophila, feeding, foraging, cGMP-dependent, protein kinase

	Introduction

TOP Summary Introduction Materials and methods Results and Discussion References

 
Animals must feed to survive, but foraging often involves a delicate balance of benefits and risks. The benefits of nutrient acquisition are weighted against elevated exposure to threats from predators, parasites, exhaustion or toxins, for example. Many risk-mitigation adaptations, which help to increase the foraging benefit-to-risk ratio, have evolved among animals. These include behavioral strategies, such as foraging at times when predators are less active or abundant, or when food is less well defended. Other adaptations include altered metabolic or physiological traits. Some animals, for example, may gorge themselves to rapidly increase fat supplies when food is plentiful and then deplete these reserves in times of famine. Alternatively, other animals, such as reptiles, adjust their metabolism or gut absorption levels to maintain body mass in response to food scarcity (Stubbs and Tolkamp, 2006; Wang et al., 2006).

Within a species, individuals may exhibit polymorphic and/or plastic responses to fluctuating food availability. Naturally varying genes involved in plastic responses to changes in food availability are difficult to identify, often due to the subtlety of their influence. Nevertheless, within-species genetic polymorphisms affecting foraging behaviors have been identified and characterized in the fruit fly, Drosophila melanogaster (Osborne et al., 1997), and the nematode, Caenorhabditis elegans (de Bono and Bargmann, 1998). Here we establish a novel role for the D. melanogaster gene, foraging (for), which influences foraging locomotory behaviors, in the mediation of food acquisition strategies and plastic responses to food availability.

The fly gene, for, encodes a cGMP-dependent protein kinase (PKG), and natural allelic variation in for affects rover- or sitter-type foraging locomotion (de Belle et al., 1989). Larvae with the wild-type rover (for^R) allele travel further when feeding and move more between food patches than those with only sitter (for^s) alleles (Osborne et al., 1997; de Belle et al., 1989; Sokolowski et al., 1983; Sokolowski, 1980). Well-fed rovers have higher PKG enzyme activities compared to sitters (Osborne et al., 1997; de Belle et al., 1989), and transgenic expression of for in sitters is sufficient to induce rover-like elevations of PKG activity and foraging locomotion (Osborne et al., 1997).

We speculated that the increased foraging locomotion of rovers may be energetically costly (Berrigan and Lighton, 1993), and thus, we hypothesized that the rover variants would require more energy and thereby differ in food acquisition compared to the sitter variants. Food acquisition was assessed by measuring rates of both food intake and nutrient absorption. Intriguingly, rover allelic variants have lower food intake than sitters despite having similar sizes and metabolic and developmental rates. This apparent paradox is clarified by our further observation that rovers may counterbalance their reduced intake with increased rates of nutrient absorption.

Despite different foraging styles, rovers and sitters occur at stable frequencies in natural populations (70% rovers: 30% sitters) (Sokolowski et al., 1997). However, selection experiments have demonstrated a selective advantage of the rover allele (for^R) compared to the sitter allele (for^s) when rover and sitter larvae are reared together for many generations in crowded conditions (Sokolowski et al., 1997). We postulated that this rover advantage may stem in part from an increased ability to tolerate or adjust to the more depleted medium found in crowded culture conditions. Thus, we hypothesized (1) that rovers will have enhanced survival compared to sitters in nutrient-restricted conditions, and (2) that this advantage may arise from differences in nutrient uptake. Indeed, our results indicate that, when reared in environments with relatively low nutrient levels, rover larvae have higher survivorship and faster development than sitter larvae. Moreover, changes in for expression can induce corrective behavioral modifications in response to food. That is, when food is scarce both rover and sitter larvae express similarly reduced PKG activity and food intake is elevated to a common maximal level. Rover larvae, however, retain their relatively elevated nutrient absorption efficiency, perhaps contributing to their survival advantage under such conditions.

The mechanisms by which changes in gene function and expression are naturally selected to confer adaptive behavioral responses to varying environments is a central question in behavioral genetics. This work uses the D. melanogaster rover/sitter polymorphism to investigate this question by addressing the role of for in mediating plasticity in food acquisition strategies.

	Materials and methods

TOP Summary Introduction Materials and methods Results and Discussion References

 
Strains
The rover and sitter strains used in these experiments are homozygous for the for^R or for^s alleles on chromosome-2 and share co-isogenic third chromosomes from the for^R strain (de Belle and Sokolowski, 1987). The X-chromosomes shared by these strains were mostly from the for^R background. for^s2 is a sitter mutant generated on the rover genetic background and does not differ from for^R aside from the induced mutation within the for gene (Osborne et al., 1997; de Belle et al., 1989). In order to increase the expression of for, a DNA fragment encoding the complete forT2 amino acid sequence (Kalderon and Rubin, 1989) was subcloned into the transformation vector, pUAST, and transformed into w¹ embryos using standard techniques (Spradling and Rubin, 1982). The sitter second chromosome (containing the for locus) was substituted into the transgenic strain, creating w¹;for^s;UASforT2. These were then crossed to w¹;for^s;hsGAL4, resulting in an increase in expression of forT2 in an otherwise for^s genetic background. We relied on leaky expression of the hs promoter (e.g. Bainton et al., 2005); experiments were done at 23°C and no heat shock was applied.

Flies (Drosophila melanogaster) were maintained in 170 ml plastic culture bottles with 40 ml of standard culture medium at 25±1°C and a 12 h:12 h L:D photocycle. Standard culture medium contained 50 g Baker's yeast, 100 g sucrose, 16 g agar, 0.1 g KPO₄, 8 g sodium potassium tartrate, 0.5 g NaCl, 0.5 g MgCl₂ and 0.5 g Fe₂(SO₄)₃ per litre of tapwater. Larvae were reared from egg-hatch to mid-third instar (96±2 h post-hatch) at densities of 100 larvae per 35 ml of medium in 100 mmx15 mm Petri dishes.

Food intake
Method 1: image analysis
Whole body measurement. Larvae were removed from food plates, washed in distilled water and groups of 10 were placed into circular wells (86 mm in diameter and 0.5 mm deep) previously filled with yeast paste (2:1 water:yeast) mixed with 0.08% Brilliant Blue R dye (Sigma, Mississauga, ON, Canada). The wells were then covered with 9 cm Petri plate lids. Larvae remained on this dyed yeast paste for varying amounts of time depending on the experiment. They were then boiled for 10 s, aligned on a microscope slide, placed under a dissecting microscope (Zeiss, Toronto, ON, Canada) and imaged using Northern Eclipse software (Empix Imaging, Mississauga, ON, Canada). Food intake was measured as the number of pixels (square pixels were used for quantification) in the image colored by the dye relative to the total number of pixels in the whole larval body, taken as a percentage. Image J software was used (ImageJ V 1.28, 2002 and ImageJ V 1.32j, 2004) for the digital quantification. Thirty larvae/strain/condition combinations were assayed. Initially food intake was measured after feeding larvae for 10, 15 and 20 min. Further experiments were done using the 15 min feeding period, which was representative of strain differences. All larvae were staged to mid-third instar prior to testing food intake (Demerec, 1994). When larvae were reared in high-quality food conditions this was 96±2 h post-hatch. When larvae were reared in 25% food this was 144–168 h post-hatch, and when larvae were reared in 15% food this was 168–192 h post-hatch, depending on humidity in the rearing incubator.

Fructose–agarose and glucose–agarose intake. Fructose– agarose and glucose–agarose food intake experiments were performed as above with the exception of food substrate. For fructose–agarose food intake experiments, 100 mmx15 mm Petri plates containing 1% agarose, 2 mol l^–1 fructose and 0.5% Carmine dye (Sigma) that were prepared 4 h prior to the test. For glucose–agarose food intake experiments, 45 mmx10 mm plates were made 24 h prior to testing with 2.3% agarose, 10% glucose and 0.5% Carmine dye. Small cuts were made in glucose–agarose plates to aid in feeding on a solid substrate. Larvae were left on sugar–agarose food substrate for 15 min before food intake was measured.

Gut measurement. Larvae were reared as above and fed dyed yeast paste for 15 min. Guts were dissected in phosphate-buffered saline (PBS) and gently elongated on a Sylgard^TM base for imaging. Gut food content was quantified by calculating the amount of dye-colored pixels as a percentage of the total number of pixels in the gut. Twenty larval guts/strain/condition combinations were measured.

Method 2: spectrophotometric analysis
Methods for spectrophotometric analysis of food intake were modified from Edgecomb et al. (Edgecomb et al., 1994). Briefly, groups of 50 larvae were reared, as above, and fed yeast paste with 0.16% Erioglaucine dye (aka FD&C Blue No. 1, Sigma) for 15 min. After feeding, each group of larvae was washed 3x in distilled water, placed in 1.5 ml tubes and immediately frozen in liquid nitrogen. Larvae were then homogenized in 250 µl distilled water, centrifuged at 13 g for 10 min, and 225 µl of supernatant was transferred to a new 1.5 ml tube containing 50 µl 100% ethanol. Tubes were vortexed for 30 s and re-centrifuged for 10 min. 250 µl of supernatant was placed in a new tube, which was centrifuged at 13 g for 5 min. Then, 200 µl supernatant was placed in a 96-well crystal plate and the OD⁶³³ read (SpectraMax Plus 384, Molecular Devices, Sunnyvale, CA, USA). Nine groups of 50 larvae per strain were analyzed.

Mouthhook contractions
Mouthhook contractions were measured for 15 min per larva on a glucose–agarose substrate described above. Contractions were measured when larvae were scraping the glucose–agarose substrate in and/or along a crack in the substrate. Individual larvae were immediately boiled following mouthook measurements and food intake was measured.

Body size and mass measurements
Length and width measurements were obtained from the digital images using ImageJ software. To establish dry mass, groups of ten larvae were placed in 1.5 ml tubes and dried for 48 h in a desiccator, after which masses were determined for 30 groups. All larvae were staged to mid-third instar prior to measurement.

Food quality manipulation
Larvae were reared in food qualities defined as 100%, 75%, 50%, 25% or 15% until mid-third instar (Demerec, 1994) for food intake tests and size measurement. Food quality was manipulated by decreasing the proportion of both yeast and sucrose in the standard culture medium (see recipe above) while keeping all other constituents constant.

Survivorship and developmental time
Twenty newly hatched larvae were placed in a 9.5 mmx25 mm diameter glass vial with 10 ml of food medium (100% 75%, 50%, 25% or 15% quality). Animals were grown at 25°C on a 12 h:12 h L:D cycle with lights on at 08:00 h. Eclosed adult flies in each vial were counted and removed once every 24 h for 26 days.

Gut contractions
Third-instar larvae were removed from food plates, washed gently and stuck to a clear microscope slide using double-sided stick tape, ventral side facing up. The number of gut contractions in the anterior midgut, acidic region, and posterior midgut were measured for 2 min. This number was divided by two to yield the number of gut contractions per minute.

The anterior midgut was defined as the region of the gut immediately following the proventriculus. The acidic midgut was defined as the narrow region of midgut following the wide foregut region and is characterized by a low pH marked by a color change from blue to yellow after ingestion of Bromophenol Blue. The posterior midgut was defined as the muscular area of the midgut immediately following the acidic portion and preceding the hindgut.

Excretion rate
Third-instar larvae were removed from food plates, washed gently, then placed on a 100 mmx15 mm Petri dish with 5 ml yeast paste with 0.08% Fast Green FCF dye for 30 min. Larvae that had food in their gut were selected. Larvae were then placed on an agarose substrate for 3 h. The number of excretion spots were counted every 15 min and then added for a total number after 3 h. Total concentration of excretion was measured by soaking feces–agarose in 1 ml distilled water for 12 h then measuring concentration of dye at 625 nm in a spectrophotometer.

PKG assays
PKG enzyme assays were performed on 96 h post-hatch whole larval homogenates. Ten whole larvae were homogenized in 25 mmol l^–1 Tris (pH 7.4), 1 mmol l^–1 EDTA, 2 mmol l^–1 EGTA, 5 mmol l^–1 ß-mercaptoethanol, 0.05% Triton X-100 and protease inhibitor cocktail (Roche Diagnostics, Laval, QC, Canada) and microcentrifuged for 5 min. The supernatant was removed and total protein levels were quantified. Supernatants containing equal amounts of total protein were analysed for cGMP-dependent protein kinase (PKG) activity. The reaction mixture contained (at final concentration): 40 mmol l^–1 Tris-HCl (pH 7.4), 20 mmol l^–1 magnesium acetate, 0.2 mmol l^–1 [g³²P]ATP (500–1000 c.p.m. pmol^–1) (Amersham Pharmacia Biotech, Baie D'Urfe, QC, Canada), 113 mg ml^–1 heptapeptide (RKRSRAE), 3 mmol l^–1 cGMP (Promega, Burlington, ON, Canada) and a highly specific inhibitor of cAMP-dependent protein kinase (5-24, Calbiochem, San Diego, CA, USA). The reaction mixtures were incubated at 30°C for 10 min, followed by termination of the reaction by spotting 70 µl of the reaction mix onto Whatman P-81 filters, which were then soaked with 75 mmol l^–1 H₃PO₄ for 5 min and washed three times with 75 mmol l^–1 H₃PO₄ to remove any unreacted [³²P]ATP. Filters were rinsed with 100% ethanol and air dried before quantification. For quantification of PKG activity, counts were taken in a Wallac 1409 Liquid Scintillation Counter (Perkin Elmer, Woodbridge, ON, Canada) using universal scintillation cocktail (ICN). Specific activity of PKG was expressed as pmol of ³²P incorporated into the substrate min^–1 mg^–1 protein.

Larval respiration rate
Respiration rates were measured by indirect calorimetry (Gibbs et al., 2003). Groups of ten mid-third-instar larvae were placed in 5 ml glass–aluminum respirometry chambers containing a strip of medium, approximately 1 cmx2 cmx 0.2 cm. Larvae exhibited apparently normal behavior, eating and crawling in and around the food. The chambers were placed in a Sable Systems (Las Vegas, Nevada, USA) TR-2 respirometer, and CO₂-free air was pumped through the chamber at 100 ml min^–1. Placement of the larvae in the respirometer was staggered, so that they had been in place approximately 60 min before data were collected. Rates of CO₂ release were measured over a 15 min period with a Li-Cor LI-6262 infrared CO₂ sensor. Data acquisition and analysis were done using Datacan V software (Sable Systems, Las Vegas, NV, USA). Control readings from chambers containing medium alone indicated no CO₂ release from microbial contaminants.

Glucose and leucine absorption
Larvae were removed from food plates, washed in distilled water and placed in groups of 70 in 9 cmx1 mm circular wells. Wells were filled with dead yeast paste (2:1 water:yeast, autoclaved 20 min) mixed with 0.08% Brilliant Blue R dye (Sigma) and 2 µCi ml^{–1 14}C-6-glucose (specific activity 58 µCi mmol l^–1, Amersham Biosciences) or 2 µCi ml^–1 L-[U-¹⁴C]leucine (specific activity 318 µCi mmol l^–1, Amersham Biosciences). After 15 min of feeding, larvae were removed and washed gently with a constant stream of 50 ml distilled water. Groups of ten larvae were then placed into 1.5 ml tubes, frozen in liquid nitrogen and stored at –80°C. For absorption experiments [protocol modified from Riha and Luckinbill (Riha and Luckinbill, 1996)] larvae were purged of the radio-labeled yeast paste prior to collection by placing them on unlabeled, undyed, heat-killed yeast paste for 3 h, the time it took for larvae to have no visible dye remaining in their guts (data not shown). We found no differences in the rate of passage of food through the rover or sitter guts (data not shown). Larvae were then carefully removed from the yeast paste, washed, placed in groups of ten into 1.5 ml tubes, and frozen in liquid nitrogen, as above.

Twenty-four hours later, larvae were removed from –80°C, placed in scintillation vials in groups of ten and solubilized at 70°C for 10 min in 200 µl Solvable (PerkinElmer Life Sciences, Woodbridge, ON, Canada) with 100 µl perchloric acid (Sigma). Next, 100 µl H₂O₂ was added and samples were vortexed for 30 s. We then added 10 ml scintillation fluid, and samples were vortexed again for 30 s and shaken for 2 h, then left at room temperature for 24 h. Samples were then vortexed again, left for 2 h at room temperature, and the amount of ¹⁴C in each vial calculated using counts observed over 60 s per sample in a scintillation counter (Wallac 1409 Liquid Scintillation Counter). Sample sizes were ten larvae/vial with six vials/strain.

View larger version (39K): [in this window] [in a new window]   Fig. 1. for affects food intake. (A) Mid-third-instar rover (forR), sitter (fors) and sitter mutant (fors2) larvae with ingested dye visible through the ventral cuticle. (B) Well-fed rovers (forR) had significantly lower food intake than well-fed sitters after 10 min (ANOVA, F(2,84)=12.03, P<0.0001), 15 min (F(2,85)=11.46, P<0.0001) and 20 min (F(2,85)=7.09, P<0.001) on dyed yeast paste. A two-way ANOVA on strain (forR, fors, fors2) and time (10, 15, 20 min) showed a significant effect of strain (F(2,254)=12.12, P<0.001), no effect of time (F(2,254)=0.13, P=0.9) and no strain-by-time interaction (F(2,254)=0.23, P=0.9). (C) Rovers had significantly less blue area than sitters at early third (F(2,89)=4.24, P=0.02) and mid-third-instar (F(2,87)=10.52, P<0.0001) stages. (D) Spectrophotometric quantification of homogenates from larvae fed a Erioglaucine (FD and C Blue No. 1) dyed yeast paste for 15 min showed that forR larvae ingest significantly less dye than fors or fors2 larvae (F(2,26)=7.88, P<0.01).

Western blot
Western blots were carried out as described (Belay et al., 2007) with the following changes. Protein was extracted from fly heads of 3–7 day-old flies reared at 23°C. 20 µg of protein extract was electrophoretically separated on SDS–10% polyacrylamide gels. To control for equal protein loading and transfer, blots were probed with a 1:5000 diluted monoclonal anti-ß-actin antibody (Sigma).

Statistical analysis
The y-axis in all figures represents mean ± s.e.m. unless otherwise stated. SAS or JMP/IN 5.1 was used for all statistical analyses (SAS Institute Inc., Cary, NC, USA). Two-way and one-way analyses of variances (ANOVA) were performed followed by Student–Neuman–Keuls pairwise post-hoc comparisons using P<0.05 as significant. Non-parametric Kruskal–Wallis tests were performed followed by Wilcoxon two-group comparisons using P<0.05 as significant when sample size was smaller than six.

	Results and Discussion

TOP Summary Introduction Materials and methods Results and Discussion References

 
for affects food acquisition
We first asked whether variation in for elicits differences in food intake and absorption in well-fed rovers and sitters. To determine if the effects we saw were specific to for, we also measured food intake and absorption in the sitter mutant (for^s2), which was generated on a rover genetic background (Pereira and Sokolowski, 1993).

To measure food intake, well-fed third instar larvae were placed in food, a yeast paste colored with blue dye, and the amount of dye ingested was quantified. The amount of dye in the gut seen through the cuticle of each larva relative to the larval body area was used to estimate food intake (Fig. 1A). We quantified food intake in the natural rover (for^R) and sitter (for^s) allelic variants as well as the sitter mutant (for^s2) to establish the connection between for-PKG and food intake. Well-fed rovers (for^R) had significantly lower food intake than well-fed sitters after 10, 15 and 20 min on dyed yeast paste (Fig. 1B,C).

These data were confirmed using several assays including spectrophotometric quantification of dye (Fig. 1D), and visual quantification of dye in dissected guts (Fig. 2A). Quantification of the rate at which the gut filled with food calculated between 5 and 30 min of feeding on a yeast paste substrate demonstrated that sitter (for^s and for^s2) larvae fill their gut at a higher rate than rover (for^R) larvae (Fig. 2B). After approximately 30 min of feeding, the midguts of all larvae were filled with blue yeast paste, thus limiting further quantification of food intake. for^R larvae also showed decreased food intake compared to for^s and for^s2 larvae on glucose–agarose and fructose–agarose substrates, suggesting that the decreased food intake of for^R larvae is a general response to all food types (Fig. 2C).

View larger version (22K): [in this window] [in a new window]   Fig. 2. for affects feeding rate and food acquisition. (A) Gut dissections from larvae fed blue yeast paste for 15 min also indicated that forR larvae ingested significantly less food than fors or fors2 larvae (F(2,26)=5.32, P<0.05). (B) The rate that the gut filled with food was measured by feeding larvae on a yeast paste with Brilliant Blue dye, extracting the gut, and comparing the length of the gut with food to the length of the entire gut. Gut size did not differ significantly between strains (F(2,67)=0.86, P=0.43). Sitter (fors and fors2) larvae filled their gut more quickly than rover (forR) larvae (10–20 min feeding, two-way-ANOVA F(8,36)=70.53 P=0.0008; effect of strain, F(2,36)=5.82, P=0.007; effect of time, F(2,36)=5.52, P=0.008; one-way ANOVAs: 5 min, P=0.82; 10 min, P=0.10; 15 min, P=0.05; 20 min, P=0.04). This difference stabilized after about 25–30 min of feeding when the midguts of all larvae become saturated with the blue yeast paste, resulting in no measurable difference in food intake (one-way ANOVAs: 25 min, P=0.85; 30 min, P=0.68). (C) Rovers showed significantly less food intake on fructose–agarose (F(2,87)=12.20, P<0.0001; forR vs fors, P<0.0001; forR vs fors2, P<0.0002; fors vs fors2, P=0.92) and glucose–agarose (F(2,87)=3.98, P=0.02; forR vs fors, P=0.0078; forR vs fors2, P=0.048; fors vs fors2, P=0.47) when food intake was measured using Carmine dye. (D) Drosophila larvae use their mouthhooks for both food ingestion and locomotion. We found no significant correlation between mouthhook movements and amount of dye ingested when larvae fed on a glucose–agarose substrate (R2=0.18, P=0.11).

Traditionally, mouthhook movements have been used to quantify food intake in larvae (Sewell et al., 1975; Wu et al., 2005). However, our observations of larval behavior suggested that larvae use their mouthhooks for both food ingestion and locomotion. We found no significant correlation between mouthhook movements and amount of dye ingested when larvae fed on a glucose–agarose substrate (Fig. 2D). In summary, our results clearly show that for affects the amount of food consumed in well-fed larvae, where variants of for associated with higher PKG activity confer lower food intake than variants of for associated with lower PKG activity.

A paradox in energy consumption and energy use
Rovers were originally distinguished from sitters by their increased foraging locomotion (Sokolowski, 1980). We confirmed that after 15 min exposure to food, rovers travel significantly farther on yeast than do sitters (P<0.0001) (Fig. 3A). This difference is expressed in the presence of a nutritive (yeast) but not a non-nutritive (agar) substrate (Fig. 3A). Thus, it was unclear how this increased locomotion could be sustained by lower food intake especially since, when food is plentiful, rover and sitter larvae develop at similar rates, reach similar sizes (length, width, mass) and have respiration rates that do not differ (Table 1, Fig. 4A).

View larger version (33K): [in this window] [in a new window]   Fig. 3. for affects glucose absorption third-instar larvae. (A) No differences in path length between rover (forR), sitter (fors) and sitter mutant (fors2) larvae were found during a 15 min test on a non-nutritive substrate (agar; F(2,56)=0.34, P=0.71). However, as expected, rovers showed significantly longer foraging trails than sitter and sitter mutants during a 15 min test on a yeast substrate (F(2,53)=35.77, P<0.0001). (B) After 15 min of feeding on a yeast–water paste containing 14C-labeled glucose, rover larvae ingested significantly less 14C than sitters (F(2,17)=52.66, P<0.001). When subsequently exposed to unlabeled medium for 3 h, rovers retained significantly more 14C label, indicating a higher level of glucose absorption compared to sitters (F(2,17)=7.36, P<0.006). (C) After 15 min of feeding on a yeast–water paste containing 14C-labeled L-U-leucine, well-fed rover larvae ingested significantly less 14C than sitters (F(2,15)=11.03, P=0.001; forR vs fors, P=0.0003; forR vs fors2, P=0.03; fors vs fors2, P=0.04). When subsequently exposed to unlabeled medium for 3 h, rovers did not differ significantly from sitters in amount of 14C label (F(2,15)=1.92, P=0.18). (D) Rovers (forR) and sitters (fors and fors2) do not differ significantly in the number of contractions of the anterior midgut (F(2,27)=0.17, P=0.85), acidic region (F(2,27)=1.83, P=0.18) or posterior midgut (F(2,27)=1.37, P=0.27). (E) No significant differences were found in the amount excreted by forR, fors or fors2 larvae (F(2,57)=0.049, P=0.95). In addition, no significant interaction between the number of fecal spots and strain was found with a logistic regression analysis on excreted food concentration (F(2,5)=2.40, P=0.100).

View larger version (63K): [in this window] [in a new window]   Fig. 4. Food level affects survivorship to eclosion, development rate and food intake in rovers and sitters. (A) Larvae were reared from egg-hatch on media containing 100%, 75%, 50%, 25% or 15% of the yeast and sucrose content of standard medium (100%; see Materials and methods). Values are means ± s.e.m. Between-strain survivorship analysis revealed that differences were not significant at 100% (F(2,21)=0.91, P=0.4), 75% (F(2,20)=1.70, P=0.2), 50% (F(2,21)=1.17, P=0.3) or 25% (F(2,21)=0.73, P=0.5) food levels, but at 15% rovers differed significantly from fors (F(1,14)=6.00, P<0.03) and fors2 (F(1,14)=13.29, P<0.003). Between-strain comparisons of developmental delay revealed that rovers differed from both fors and fors2 at 15% (F(2,21)=4.33, P<0.03) and rovers differed from fors2 at 100% (F(2,21)=4.42, P<0.03), 50% (F(2,21)=7.06, P<0.005) and 25% (F(2,21)=3.80, P<0.04) food levels but not at 75% (F(2,20)=2.58, P=0.1). (B) Rovers (forR) ingested less yeast paste after 15 min feeding than sitters (fors and fors2) when raised on high levels of food (100%, 75%, 50%), but not when reared on low levels of food (25%, 15%) [two-way ANOVA for strain (F(2,435)=5.13, P<0.006) and strain-by-food level (F(8,435)=2.28, P<0.02); one-way ANOVA on 100% (F(2,87)=6.86, P<0.002), 75% (F(2,87)=14.33, P<0.0001), 50% (F(2,87)=6.34, P<0.003), 25% (F(2,87)=0.14, P=0.9), 15% (F(2,87)=0.11, P=0.9)]. (C) Rovers and sitters differ in food intake measured after 5 min feeding on yeast paste when reared at 100%, food quality but not when food-deprived (reared on 25% and 15% food) [ANOVA, at 100%, F(2,87)=5.53, P<0.0006, 25% F(2,87)=0.69, P=0.50), 15% F(2,87)=0.33, P=0.7]. (D) All larvae reared under low food levels were staged to mid-third instar prior to measuring food intake. Rover and sitter larvae reared in dilute (15% and 25%) food were smaller in size at mid-third instar compared to larvae reared in normal (100%) food. However, no within-strain differences were found at any food level [two-way ANOVA, F(8,260)=56.50, P<0.0001; strain, F(2,260)=0.19, P=0.82; food quality, F(2,260)=73.25, P<0.0001, with significantly smaller larvae at 25% (P<0.0001) and 15% (P<0.0001) compared to 100%; strainxfood quality F(2,4)=0.23, P=0.92]. (E) At 15% food, rovers, sitters and sitter mutants ingested similar amounts of 14C-labeled media in15 min (F(2,15)=0.67, P=0.3), but rovers absorbed more than sitter and sitter mutants (F(2,17)=45.51, P<0.0001). forR larvae had a twofold increase in absorption compared to fors and fors2 larvae at 15% food levels (F(2,12)=41.96, P<0.0001; forR vs fors, P<0.0001; forR vs fors2, P<0.0001; fors vs fors2, P=0.12). (F) After 15 min of feeding on a yeast–water paste containing 14C-labeled L-U-leucine, rover larvae reared on 15% food did not differ significantly from sitters in amount of 14C ingested (F(2,15)=0.37, P=0.70). When subsequently exposed to unlabeled medium for 3 h, rovers absorbed significantly more 14C label compared to sitter mutants (F(2,15)=4.55, P=0.03; forR vs fors, P=0.15; forR vs fors2, P=0.009; fors vs fors2, P=0.53).

Intriguingly, despite rovers showing increased locomotion on food, the similarity in metabolic rate between the natural variants suggests similar energy output. Notably, there was no significant behavioral difference observed in mouthhook extensions (Fig. 2D), which could be used for both locomotion and feeding. This suggests that active feeding in a smaller area by sitter larvae equalizes the total activity and therefore metabolic demand between the two natural variants. Thus for D. melanogaster larvae, reaching out and grabbing a mouthful of food seems to require as much energy as moving to a new location.

This was our first indication that even though rovers ingested less food than sitters, they do not suffer a fitness deficit relative to sitters. Thus, we hypothesized that rovers may absorb their food more efficiently than sitters.

for enhances glucose absorption
We hypothesized that rovers may offset their reduced food intake with enhanced glucose absorption. We tested this using ¹⁴C-6-glucose-labeled food to compare ingestion and absorption (Riha and Luckinbill, 1996; Carvalho et al., 2005) (see Materials and methods). We measured the total amount of ¹⁴C ingested and the total amount of ¹⁴C absorbed.

As described above, for^R larvae have lower food intake than sitter larvae, as measured by total ¹⁴C uptake levels with ¹⁴C-6-glucose (Fig. 3B). Notably, chasing the radio-labeled medium with unlabeled food resulted in significantly higher ¹⁴C counts in for^R animals relative to for^s (Fig. 3B). Thus, for^R larvae absorbed approximately 50% of the ¹⁴C-6-glucose ingested whereas for^s and for^s2 larvae absorbed approximately 15% of the ¹⁴C-6-glucose ingested. Rovers retain more of the radioactive label after clearing their guts with unlabeled medium, indicating higher glucose absorption in rovers than in sitter and sitter mutant larvae.

To determine if rovers and sitters differed in amino acid absorption we repeated the above experiment with L-[U-¹⁴C]leucine (Fig. 3C). Leucine was chosen because it is one of the most abundant amino acids in yeast, a major food source of D. melanogaster larvae (Yamada and Sgarbiera, 2005). No absorption differences were found with L-[U-¹⁴C]leucine in well-fed larvae (Fig. 3C). As for glucose, rovers had lower intake of radiolabelled leucine than sitters. This suggested that the for-mediated differences in absorption in well-fed larvae may be specific to carbohydrates. It is not known how rovers absorb more glucose in their guts than sitters. Measures of excretion concentration, gross gut morphology and gut contraction rate did not show obvious differences between for^R, for^s or for^s2 (Fig. 3D,E). Together, these results provide evidence that the rover/sitter natural polymorphism is a food-related phenomenon involving food intake and absorption.

for affects survivorship and development time under low-nutrient conditions
Our results demonstrate that for affects the nutrient acquisition strategies of well-fed larvae, where rovers have decreased food intake and higher glucose absorption compared to sitters. We hypothesized that these differences in energy acquisition could affect larval responses to changes in food availability. Because the opportunity for larval food deprivation in nature is high (Atkinson, 1979) we investigated whether larvae exhibited plastic responses to food deprivation and if so whether this plasticity was mediated by for.

In order to determine appropriate food deprivation conditions we reared larvae from egg-hatch to pupation on media containing 100%, 75%, 50%, 25% or 15% of the yeast and sucrose content of standard medium (see Materials and methods). We hypothesized that the differences in food acquisition between variants of for affect larval development and survivorship. Larval development and survivorship of for^R, for^s and for^s2 did not differ at 100%, 75% or 50%, suggesting that these food levels were not limiting. However, all strains were developmentally delayed when reared on 25% and 15%, and survivorship to eclosion also tended to be reduced (Fig. 4A). As a result, we used 25% and 15% as our food deprived conditions. Within-strain comparisons showed that all strains developed more slowly at 25% and 15%, and had significantly lower survival to eclosion at 15%. The magnitude of these changes was greatest in for^s and the for^s2 sitter mutant larvae. Between-strain survivorship analysis revealed that differences were not significant at 100%, 75%, 50% or 25% food levels, but at 15% rovers differed significantly from for^s and for^s2. Between-strain comparisons of developmental delay revealed that rovers differed from both for^s and for^s2 at 15% and rovers differed from for^s2 at 100%, 50% and 25% food levels, but not at 75%. These results suggest variation in for affects larval survivorship under low food conditions.

for affects plasticity of food intake in food-depleted environments
When larvae are reared to third instar under food deprivation conditions and then placed on a dyed yeast paste, for^R, for^s and for^s2 larvae exhibit plasticity in food intake, and this plasticity is mediated by for (see below). The food intake of all strains was significantly elevated when larvae were reared to mid-third instar on media containing 25% or 15% food compared to those reared on 100% food (Fig. 4B; two-way ANOVA for food level, F_(4,435)=18.94). Rovers (for^R) ingested less yeast paste than sitters when (for^s and for^s2) when raised on high levels of food (100%, 75%, 50%), but not when reared on low levels of food (25%, 15%) whether they were feeding for 15 min (Fig. 4B) or 5 min (Fig. 4C). When for^R, for^s and for^s2 larvae reared in food-deprived conditions (25% and 15%) reached mid-third instar, their sizes did not significantly differ from each other, despite being developmentally delayed and smaller than larvae reared in abundant food (50%, 75% and 100%) (Fig. 4D); for all food intake measures, larvae were staged to mid-third instar (see Materials and methods).

Glucose absorption differences persisted when larvae were reared under food-deprivation conditions (25% or 15%) demonstrating the persistence of increased glucose absorption by rovers compared to sitters and sitter mutants (Fig. 4E). As expected, in 15% food, for^R, for^s and for^s2 larvae ingested similar amounts of ¹⁴C-labeled media in 15 min. However, for^R still absorbed more than for^s and for^s2 larvae. Since both rovers and sitters ingest food at the same rate when reared under poor food conditions, these results also suggest that rovers are able to absorb more nutrients per unit food ingested, and differences in absorption are not due to the amount of food in the gut.

Intriguingly, the absorption efficiency, defined as the percentage of ¹⁴C-absorbed/¹⁴C-ingested remained at about 50% for for^R larvae and 17% for for^s and for^s2 larvae, regardless of the quality or dilution of food in which the larvae are reared. Although it looks as if the amount of ¹⁴C absorbed increased from 100% to 15% food in for^R larvae, their total food ingestion also increases, resulting in an unchanged absorption efficiency.

We also found a nearly significant difference in absorption of leucine between rovers and sitters when reared under 15% food (Fig. 4F), suggesting that larvae may alter their leucine absorption in response to food deprivation but further experiments are required to assess this hypothesis. Overall, both rovers and sitters ingest food at the same rate after being reared under food deprivation conditions; however, rovers absorb more glucose than sitters. This may contribute to their enhanced fitness under 25% and 15% food deprivation conditions.

From the data above, we conclude that food ingestion rates of D. melanogaster larvae are sensitive to the nutritional state of the individual. Larvae do not eat at maximal rates when well fed but may approach ceiling ingestion rates when placed on yeast paste after having been raised in the 25% and 15% food conditions provided here. Furthermore, we find that rover food intake is not fixed at lower levels than sitters; rather, the differential varies depending on the degree of food deprivation. Since maximal rates of food intake do not appear to differ between the variants, the greater plasticity of rovers can be explained by their lower intake rates under well-fed conditions, which may be balanced by their higher glucose absorption under both well-fed and food-deprived conditions. These data may help explain why long-term selection at high larval density favors rovers (Sokolowski et al., 1997).

Variation in for mediates food acquisition strategies
As the larval cells in which FOR are expressed are not yet known, we chose to manipulate for by ubiquitously increasing expression of for in sitters. To this end, we employed the transgenic sitter strain carrying a T2 transgene (UASforT2), which was expressed ubiquitously in a sitter genetic background using the leaky expression of a heat-shock promoter (hsGAL4). Such expression of forT2 in a sitter genetic background has been previously shown to result in rover-like foraging locomotion (Osborne et al., 1997) and increases the amount of protein produced by forT2 (Fig. 5A).

View larger version (25K): [in this window] [in a new window]   Fig. 5. Variation in for mediates nutrient acquisition strategies. (A) fors;hs-GAL4/UASforT2 flies reared at 23°C show an increase in T2 protein level (lane 1) compared to control flies reared under the same conditions (lanes 2 and 3). The actin bands show that equal amounts of protein were loaded in each lane. (B) Expression of a forT2 transgene in sitters resulted in a rover-like decrease in food intake (ANOVA: at 5 min, F(1,57)=3.03, P=0.09; 10 min, F(1,58)=7.60, P=0.008; 15 min, F(1,58)=11.92, P=0.001; 20 min, F(1,57)=5.46, P=0.02). (C) Ubiquitous expression of forT2 in sitters using a leaky hs-GAL4 transgene significantly decreased food intake (F(2,89)=19.56, P<0.0001) and (D) increased glucose absorption rate (F(2,17)=47.51, P<0.0001).

View larger version (36K): [in this window] [in a new window]   Fig. 6. Variation in for affects response to food quality. (A) Expression of forT2 was sufficient to increase developmental rate and survivorship in food-deprived sitter larvae raised on low nutrient media (25% or 15%) [development time: at 25% (F(2,27)=17.78, P<0.0001) and at 100% (F(2,26)=0.95, P=0.4); survivorship: at 15% (F(2,29)=4.07, P<0.03) and at 100% (F(2,28)=0.66, P=0.5)]. (B) PKG enzyme activity is inversely related to food intake. Rovers (forR) had significantly higher PKG activity than sitters (fors and fors2) when reared on high food levels, but this difference diminished when larvae were food-deprived [two-way ANOVA: for strain (F(2,63)=57.36, P<0.0001), food level (F(2,63)=95.26, P<0.0001), strain-by-food level (F(4,63)=11.67, P<0.0001); ANOVA: for 100% (F(2,21)=87.9, P<0.0001), 25% (F(2,21)=14.39, P<0.0001), 15% (F(2,21)=0.70, P=0.5) food level]. (C) Sitter larvae expressing a forT2 transgene expressed a rover-like pattern of food [two-way ANOVA: for strain (F(2,261)=3.78, P=0.02), food level (F(2,261)=2.85, P=0.06), strain-by-food level (F(4,261)=1.60, P=0.2); one-way ANOVA by strain: for 100% (F(2,87)=6.87, P<0.002), 25% (F(2,87)=0.32, P=0.7), 15% (F(2,87)=0.42, P=0.7) food level].

Interestingly, unlike food intake, the increased rate of glucose absorption in rovers did not vary in a food-dependent manner. Regardless of food level, higher levels of glucose absorption were found in rovers and sitters expressing the forT2 transgene compared with sitters and the sitter mutant. Elevated glucose absorption in rovers might partially account for their increased survivorship compared to sitters at 25% and 15%, despite similar PKG enzyme activity levels.

Our data do not address whether for acts throughout development or in an immediate nature to mediate food acquisition and plasticity in food intake. Thus, several possibilities could explain why glucose absorption does not vary with food level in a similar way as food intake and PKG activity. For example, food deprivation may not downregulate for activity in cell types important for glucose absorption, or the effects of PKG on glucose absorption may be fixed early in development. Future studies on specific expression of for and targeting manipulation of for in the cells in which for is expressed may elucidate both the developmental timescale and cellular mechanism through which for acts to mediate these responses.

Conclusions
The implications of this study are twofold. Firstly we demonstrate that natural variation in PKG affects food acquisition, and secondly we show for's role in plastic compensatory food intake responses to food deprivation. The rover/sitter system provides an enticing model for further investigations into the effects of `thrifty' genes that regulate efficiency of nutrient homeostasis (Zimmet and Thomas, 2003; Shmulewitz et al., 2006). Alleles of such genes may promote individual fitness under restrictive food regimes (Neel, 1962) just as for-PKG regulates normal individual differences in food intake. The role of for in food-specific behaviors is conserved in several species including honeybees, nematodes and harvester ants (Ben-Shahar et al., 2002; Fujiwara et al., 2002; Ingram et al., 2005). Future studies will determine whether similar gene-by-environment interactions prevail in the for orthologs identified in other species.

Intriguingly, a phylogeny constructed from available for-PKG protein sequences suggests that despite broad taxonomic breadth (nematodes to humans), there may be a widespread conserved association between PKG and food-related behaviors (Fitzpatrick and Sokolowski, 2004). If the role of for in the plasticity of food intake and regulation of food absorption is conserved in other animals including humans, then it may play an important role in regulation of energy homeostasis essential in maintaining a healthy body mass. The human homolog of for, cGK1, plays an important role in intestinal muscle function and thus may affect passage of intestinal content and ability to absorb food (Hofmann, 2005). Changes in PKG are also associated with disorders such as obesity and diabetes. For example, high expression of cGK1 has been associated with obesity, and a reduction of PKG has been associated with diabetes and high glucose concentrations (Wang et al., 2002; Wang et al., 2003; Wang et al., 2004; Engeli et al., 2004; Su et al., 2003; Zanetti et al., 2005; Chang et al., 2004). Although a polymorphism in cGKI has not yet been found between obese and healthy mass populations (Zakharkin et al., 2005), the evidence from D. melanogaster larvae suggests that cGKI may help sustain a healthy mass by maintaining a balance between energy input and output. Future research will help resolve whether a natural polymorphism in for also contributes to individual differences in energy balance in humans and across diverse taxa.

Little is known of the genetic or molecular basis for phenotypic plasticity, as genetic variability has intentionally been removed or reduced during the establishment of model organisms used in genetic analyses (DeWitt and Scheiner, 2004). Our findings provide a rare example of a naturally varying gene that affects plasticity providing a unique opportunity to investigate both the genetic and functional aspects of plasticity.

	Acknowledgments

 
We would like to thank J. Levine and the members of the Sokolowski lab for their support and comments on the manuscript. We would also like to thank two anonymous reviewers for their very helpful comments in revising this manuscript. Funding was provided by NIDDK grants 5R01DK070141-03 to M.B.S. and NSF grant IOB-0514402 to A.G.G. The authors declare no competing financial interests.

	References

TOP Summary Introduction Materials and methods Results and Discussion References

 

Atkinson, W. D. (1979). A field investigation of larval competition in domestic Drosophila. J. Anim. Ecol. 48,91 -102.[CrossRef]

Bainton, R. J., Tsai, L. T.-Y., Schwabe, T., DeSalvo, M., Gaul, U. and Heberlein, U. (2005). moody encodes two GPCRs that regulate cocaine behaviors and blood-brain barrier permeability in Drosophila. Cell 123,145 -156.[CrossRef][Medline]

Belay, A. T., Scheiner, R., So, A. K. C., Douglas, S. J., Chakaborty-Chatterjee, M., Levine, J. D. and Sokolowski, M. B. (2007). The foraging gene of Drosophila melanogaster: spatial expression analysis and sucrose responsiveness. J. Comp. Neurol. 504,570 -582.[CrossRef][Medline]

Ben-Shahar, Y., Robichon, A., Sokolowski, M. B. and Robinson, G. E. (2002). Influence of gene action across different time scales on behavior. Science 296,741 -744.[Abstract/Free Full Text]

Berrigan, D. and Lighton, J. R. B. (1993). Bioenergetic and kinematic consequences of limblessness in larval Diptera.J. Exp. Biol. 179,245 -259.[Abstract]

Carvalho, G. B., Kapahi, P. and Benzer, S. (2005). Compensatory ingestion upon dietary restriction in Drosophila melanogaster. Nat. Methods 2, 813-815.[CrossRef][Medline]

Chang, S., Hypolite, J. A., Velez, M., Changolkar, A., Wien, A. J., Chacko, S. and DiSanto, M. E. (2004). Downregulation of cGMP-dependent protein kinase-1 activity in the corpus cavernosum smooth muscle of diabetic rabbits. Am. J. Physiol. 287,R950 -R960.

de Belle, J. S. and Sokolowski, M. B. (1987). Heredity of rover-sitter alternative foraging strategies of Drosophila melanogaster larvae. Heredity 59, 73-84.

de Belle, J. S., Hilliker, A. J. and Sokolowski, M. B. (1989). Genetic localization of foraging (for): a major gene for larval behavior in Drosophila melanogaster.Genetics 123,157 -163.[Abstract/Free Full Text]

de Bono, M. and Bargmann, C. I. (1998). Natural variation in a neuropeptide Y receoptor homolog modifies social behavior and food response in C. elegans. Cell 94,679 -689.[CrossRef][Medline]

Demerec, M. (1994). Biology of Drosophila. Woodbury, NY: CSHL Press.

DeWitt, T. J. and Scheiner, S. M. (2004). Phenotypic Plasticity: Functional and Conceptual Approaches. Oxford: Oxford University Press.

Edgecomb, R. S., Harth, C. E. and Schneiderman, A. M. (1994). Regulation of feeding behavior in adult Drosophila melanogaster varies with feeding regime and nutritional state. J. Exp. Biol. 97,215 -235.

Engeli, S., Janke, J., Gorzelniak, K., Bohnke, J., Ghose, N., Lindschau, C., Luft, F. C. and Sharma, A. M. (2004). Regulation of the nitric oxide system in human adipose tissue. J. Lipid. Res. 45,1640 -1648.[Abstract/Free Full Text]

Fitzpatrick, M. J. and Sokolowski, M. B. (2004). In search of food: exploring the evolutionary link between cGMP-dependent protein kinase (PKG) and behaviour. Integr. Comp. Biol. 44,28 -36.[Abstract/Free Full Text]

Fujiwara, M., Sengupta, P. and McIntire, S. L. (2002). Regulation of body size and behavioral state of C. elegans by sensory perception and the EGL-4 cGMP-dependent protein kinase. Neuron 36,1091 -1102.[CrossRef][Medline]

Gibbs, A. G., Fukuzato, F. and Matzkin, L. M. (2003). Evolution of water conservation mechanisms in Drosophila. J. Exp. Biol. 206,1183 -1192.[Abstract/Free Full Text]

Hofmann, F. (2005). The biology of cyclic GMP-dependent protein kinases. J. Biol. Chem. 280, 1-4.[Free Full Text]

Ingram, K. K., Oefner, P. and Gordon, D. M. (2005). Task-specific expression of the foraging gene in harvester ants. Mol. Ecol. 14,813 -818.[CrossRef][Medline]

Kalderon, D. and Rubin, G. M. (1989). cGMP-dependent protein kinase genes in Drosophila. J. Biol. Chem. 264,10738 -10748.[Abstract/Free Full Text]

Neel, J. V. (1962). Diabetes mellitus: a `thrifty' genotype rendered detrimental by `progress'? Am. J. Human Genet. 14,353 -362.[Medline]

Osborne, K. A., Robichon, A., Burgess, E., Butland, S., Shaw, R. A., Coulthard, A., Pereira, H. S., Greenspan, R. J. and Sokolowski, M. B. (1997). Natural behavior polymorphism due to a cGMP-dependent protein kinase of Drosophila. Science 277,834 -836.[Abstract/Free Full Text]

Pereira, H. S. and Sokolowski, M. B. (1993). Mutations in the larval foraging gene affect adult locomotory behavior after feeding in Drosophila melanogaster. Proc. Natl. Acad. Sci. USA 90,5044 -5046.[Abstract/Free Full Text]

Riha, V. F. and Luckinbill, L. S. (1996). Selection for longevity favors stringent metabolic control in Drosophila melanogaster. J. Gerontol. A Biol. Sci. Med. Sci. 51,B284 -B294.[Abstract]

Sewell, D., Burnet, B. and Connolly, K. (1975). Genetic analysis of larval feeding behavior in Drosophila melanogaster.Genet. Res. 24,163 -173.

Shmulewitz, D., Heath, S. C., Blundell, M. L., Han, Z., Sharma, R., Salit, J., Auerback, S. B., Signorini, S., Breslow, J. L., Stoggel, M. et al. (2006). Linkage analysis of quantitative traits for obesity, diabetes, hypertension, and dyslipidemia on the island of Kosrae, Federated States of Micronesia. Proc. Natl. Acad. Sci. USA 103,3502 -3509.[Abstract/Free Full Text]

Sokolowski, M. B. (1980). Foraging strategies of Drosophila melanogaster: a chromosomal analysis. Behav. Genet. 10,291 -302.[CrossRef][Medline]

Sokolowski, M. B., Hansell, R. I. C. and Rotin, D. (1983). Drosophila larval foraging behavior. II. Selection in the sibling species, D. melanogaster and D. simulans. Behav. Genet. 13,169 -177.[CrossRef][Medline]

Sokolowski, M. B., Pereira, H. S. and Hughes, K. (1997). Evolution of foraging behavior in Drosophila by density-dependent selection. Proc. Natl. Acad. Sci. USA 94,7373 -7377.[Abstract/Free Full Text]

Spradling, A. C. and Rubin, G. M. (1982). Transposition of cloned P elements into Drosophila germ line chromosomes. Science 218,341 -347.[Abstract/Free Full Text]

Stubbs, R. J. and Tolkamp, B. J. (2006). Control of energy balance in relation to energy intake and energy expenditure in animals and man: an ecological perspective. Br. J. Nutr. 95,657 -676.[CrossRef][Medline]

Su, J., Zhang, S., Tse, J., Scholz, P. M. and Weiss, H. R. (2003). Alterations in nitric oxide-cGMP pathway in ventricular myocytes from obese leptin-deficient mice. Am. J. Physiol. 285,H2111 -H2117.

Wang, S., Shiva, S., Poczatek, M., Darkey-Usmar, V. and Murphy-Ullrich, J. E. (2002). Nitric-oxide and cGMP-dependent protein kinase regulation of glucose-mediated thrombospondin 1-dependent transforming growth factor-beta activation in mesangial cells. J. Biol. Chem. 277,9880 -9888.[Abstract/Free Full Text]

Wang, S., Wu, X., Lincoln, T. M. and Murphy-Ullrich, J. E. (2003). Expressing of constitutively active cGMP-dependent protein kinase prevents glucose stimulation of thrombospondin 1 expression and TGF-ß activity. Diabetes 52,2144 -2150.[Abstract/Free Full Text]

Wang, S., Skorczewski, J., Jeng, X., Mei, L. and Murphy-Ullrich, J. E. (2004). Glucose up-regulated thrombospondin 1 gene transcription and transforming growth factor-ß activity through antagonism of cGMP-dependent protein kinase repression via upstream stimulatory factor 2. J. Biol. Chem. 279,34311 -34322.[Abstract/Free Full Text]

Wang, T., Hung, C. C. Y. and Randall, D. J. (2006). The comparative physiology of food deprivation: from feast to famine. Annu. Rev. Physiol. 68,223 -251.[CrossRef][Medline]

Wu, Q., Zhang, Y., Xu, J. and Shen, P. (2005). Regulation of hunger-driven behaviors by neural ribosomal S6 kinase in Drosophila. Proc. Natl. Acad. Sci. USA 102,13289 -13294.[Abstract/Free Full Text]

Yamada, E. A. and Sgarbiera, V. C. (2005). Yeast (Saccaromyces cerevisiae) protein concentrate: preparation, chemical composition, and nutritional and functional properties. J. Agric. Food Chem. 53,3931 -3936.[CrossRef][Medline]

Zakharkin, S. O., Belay, A. T., Fernandez, J. R., De Luca, V., Kennedy, J. L., Sokolowski, M. B. and Allison, D. B. (2005). Lack of association between polymorphism of the human cyclic GMP-dependent protein kinase gene and obesity. Int. J. Obes. Lond. 29,872 -874.[CrossRef][Medline]

Zanetti, M., Barazzoni, R., Stebel, M., Roder, M., Roder, E., Biolo, G., Baralle, F. E., Cattin, L. and Guarnieri, G. (2005). Dysregulation of the endothelial nitric oxide synthase-soluble guanylate cyclase pathway is normalized by insulin in the aorta of the diabetic rat. Atherosclerosis 181, 69-73.[CrossRef][Medline]

Zimmet, P. and Thomas, C. R. (2003). Genotype, obesity and cardiovascular disease – has technical and social advancement outstripped evolution? J. Intern. Med. 254,114 -125.[CrossRef][Medline]

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