K.E.H. was supported by the Training Grant in Molecular Biology NIH-T32-GM08730. This work was supported by NIH, NIDDK Grant 5K01DK095932 and AHA Award 12BGIA11930014 to T.R.

1. Collect Eggs from Flies with Genotypes of Interest
NOTE: Ideal crosses consist of 150 virgin flies and at least 75 males. Stock collections should consist of at least 200 flies.
<ol>
	<li>Prepare grape plates.
	<ol>
		<li>Add 37.5 g agar to 1.5 L distilled water in a 4 L flask.</li>
		<li>Autoclave water/agar mix for 50 min at 121 &#186;C (250 &#186;F).</li>
		<li>While water/agar mix is autoclaving, add 50 g sucrose to 500 ml grape juice and stir with a stir bar on a heated stir plate until sucrose is dissolved.</li>
		<li>Turn off the heat and continue stirring to cool the grape juice/sucrose mixture to below 70 &#186;C. Test the internal temperature with an alcohol thermometer.</li>
		<li>Add 3 g Drosophila anti-fungal agent (e.g., Tegosept) to warm grape juice/sucrose mixture and stir to dissolve.</li>
		<li>When water/agar mixture is out of the autoclave, allow the flask to cool to the touch by swirling occasionally.</li>
		<li>Combine water/agar mixture and grape juice/sucrose mixture and stir until well mixed.</li>
		<li>Use a serological pipette to add 8 - 10 ml of the grape mixture to the lid of a small petri plate (35 x 10 mm style). Allow the mixture to crown higher than the height of the lid walls to form a convex shape.</li>
		<li>Allow grape plates to cool for 2 hr at room temperature and then store in an airtight container at 4 &#186;C.</li>
	</ol>
	</li>
	<li>Prepare yeast paste.
	<ol>
		<li>Add 6 ml phosphate buffer solution (PBS) to 4 g live active dried yeast in a 50 ml conical tube and mix with a spatula until smooth.</li>
		<li>Store yeast paste at 4 &#186;C.</li>
	</ol>
	</li>
	<li>Add a small dollop (approximately 1/8 tsp) of yeast paste to the middle of a grape plate. Smooth down any rough edges with a spatula.</li>
	<li>Place grape plates in an incubator until they have reached ambient temperature.</li>
	<li>Transfer flies of interest into an egg collection chamber and place grape plate with yeast paste facing inwards on top. Tape grape plate to the egg collection chamber to avoid letting the plate fall out. Make sure to write important genotype and date information on the bottom of the grape plate.</li>
	<li>Upend the egg collection chamber to allow the flies to lay eggs on the grape plate.</li>
	<li>Place a box or cover over the egg collection chambers and place them in an incubator at 25 &#186;C.</li>
	<li>Allow flies to lay eggs for 4 - 6 hr.</li>
	<li>End collection by taking the grape plate out of the egg collection chamber and transferring the flies back into their bottle of food. Use a spatula to wipe off any excess yeast paste from the grape plate. Store grape plate in a larger petri dish in a humidified incubator at 25 &#186;C for ~ 24 hr.</li>
</ol>
2. Transfer Larvae to Experimental Food
<ol>
	<li>Prepare experimental food. 
	NOTE: This food recipe is based on the Bloomington stock center recipe18. Important changes include increased amount of yeast to optimize the nutritional value of the food as assessed by timing of wandering stage (between 112 - 120 hr after egg collection at 25 &#186;C). While we have found this recipe ideal for the health and development of our experimental larvae, any food recipe is acceptable given calibration with positive and negative controls, examples of which are listed in step 3.10.
	<ol>
		<li>Measure 1,600 ml distilled water.</li>
		<li>Mix 134 g cornmeal, 10.6 g Drosophila Agar Type II, and some of the water (~ 500 ml) in a 1 L beaker. Blend for 2 min with a motorized hand mixer.</li>
		<li>Mix 70 g live active dry yeast, 18.4 g soy flour, 84.8 g malt extract, and some of the water (~ 500 ml) in another 1 L beaker. Blend for 2 min with a motorized hand mixer.</li>
		<li>Swirl each beaker and pour into a 4 L flask. Use remaining water to rinse out the beakers and add into the flask.</li>
		<li>Measure 140 ml light corn syrup and add to the flask, mix by swirling.</li>
		<li>Autoclave for 50 min at 121 &#186;C (250 &#186;F) in the liquid setting.</li>
		<li>After autoclaving, pour food into a 4 L beaker and allow to cool. Help the cooling process by blending with a motorized hand mixer.</li>
		<li>Once the flask has cooled enough to touch, add 8.8 ml propionic acid and 16.8 ml Drosophila anti-fungal agent/ethanol solution. Mix well. (Make Drosophila anti-fungal agent solution by adding 380 g Drosophila anti-fungal agent to 1 L 200 proof ethanol and stirring on a warm plate until dissolved, making sure the solution does not exceed 70 &#186;C.)</li>
		<li>Pour food into vials until the food is approximately 1 inch deep and allow to cool at RT for 2 - 3 hr. Plug vials and store in an incubator at 25 &#186;C. Use within a week.</li>
	</ol>
	</li>
	<li>Plunge a spatula several times into the top layer of the vial of food to score it. This will help the larvae to burrow and feed.</li>
	<li>22 - 24 hr after egg collection has ended, use a small paintbrush to collect 50 first instar (L1) larvae of the same approximate size. Place larvae in the experimental food. Clean the paintbrush on a laboratory wipe and ensure that there are no remaining larvae on the paintbrush before continuing to another genotype.</li>
	<li>Place vial of larvae in an incubator at 25 &#186;C and allow to develop.</li>
</ol>
3. Determine Fat Levels in Larvae
<ol>
	<li>Prepare solutions.
	<ol>
		<li>Prepare 1 L 10x PBS stock.
		<ol>
			<li>Add 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4, and 2.4 g KH2PO4 to 800 ml water in a 2 L beaker containing a stir bar. Stir with no heat until all salts are incorporated. Bring pH to 7.4. Increase volume to 1 L with water.</li>
		</ol>
		</li>
		<li>Prepare 2 L 1x PBS. Add 200 ml 10x PBS stock solution to 1,800 ml water.</li>
		<li>Use this stock of 1x PBS to make 20% w/v sucrose.</li>
	</ol>
	</li>
	<li>Remove the vial of larvae from the incubator once there are 20 - 40 larvae wandering up the sides of the vial (generally on the fifth day after egg collections). Record the date and time of the assay as well as the number of wandering larvae.</li>
	<li>Prepare experimental solution by adding 11.5 ml PBS and 9 ml 20% sucrose to a 50 ml conical tube.</li>
	<li>Add 20% sucrose to the vial until it is 0.5 - 1 inch from the top of the vial.</li>
	<li>Use a spatula to gently stir up the top layer of food to free larvae that are still burrowing in the food. All larvae will rise to the surface of the sucrose solution.</li>
	<li>Use forceps to gently transfer the larvae into the initial sucrose concentration from step 3.3.</li>
	<li>Use a spatula to gently stir the larvae in this solution.</li>
	<li>Cap the conical tube and upend several times to thoroughly mix the solution.</li>
	<li>Uncap the conical tube and swirl to create a gentle vortex.</li>
	<li>Allow the larvae to settle, either floating to the top of the solution or sinking to the bottom. Wait 2 - 5 min for larvae to settle. 
	NOTE: If there are still larvae that are not definitely either floating or sinking, pick a line to use as a cutoff point to label the larvae as either a floater or a sinker. Apply this same criterion to all genotypes. We use the angle at the bottom of the conical tube as our boundary. adp or sir217 mutant larvae may be used as a positive control and lsd219 mutants may be used as a negative control for calibration of the assay.</li>
	<li>Record the number of floating larvae and the specific concentration tested.</li>
	<li>Add 1 ml 20% sucrose to the experimental solution to increase its density.</li>
	<li>Repeat steps 3.7 to 3.10. Record the number of floating larvae and this new concentration.</li>
	<li>Continue steps 3.12 and 3.13 until at least 95% of the larvae are floating.</li>
	<li>Use forceps to collect all larvae into a dissection dish filled with PBS.</li>
	<li>Observe larvae under a microscope and note if there are any small larvae, pre-pupae, pupae, or any other differences between genotypes. 
	NOTE: Ideally, all larvae should appear homogeneous and at the same developmental stage. We have only observed consistent fat measurements under these conditions.</li>
	<li>Record the total number of larvae.</li>
	<li>Using forceps, transfer 10 larvae to a laboratory wipe to dry. Label a microcentrifuge tube and place 10 larvae in the tube.</li>
	<li>Flash freeze the larvae in liquid nitrogen. Store the tube at -20 &#186;C.</li>
</ol>

<ol>
	<li>Bier, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila,+the+golden+bug,+emerges+as+a+tool+for+human+genetics.">Drosophila, the golden bug, emerges as a tool for human genetics.</a> Nat Rev Genet. 6, 9-23 (2005).</li><li>Liu, Z., Huang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+metabolism+in+Drosophila+development+and+disease+Using+Drosophila+System+to+Study+Lipid+Metabolism+Lipids+Function+in+Drosophila+Early+Development.">Lipid metabolism in Drosophila development and disease Using Drosophila System to Study Lipid Metabolism Lipids Function in Drosophila Early Development.</a> Acta Biochim Biophys Sin. 45 (1), 44-50 (2013).</li><li>Ruden, D. M., Lu, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+conservation+of+metabolism+explains+howDrosophila+nutrigenomics+can+help+us+understand+human+nutrigenomics.">Evolutionary conservation of metabolism explains howDrosophila nutrigenomics can help us understand human nutrigenomics.</a> Genes Nutr. 1 (2), 75-84 (2006).</li><li>Wolf, M. J., Rockman, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila,+genetic+screens,+and+cardiac+function.">Drosophila, genetic screens, and cardiac function.</a> Circ. Res. 109, 794-806 (2011).</li><li>Trinh, I., Boulianne, G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+obesity+and+its+associated+disorders+in+Drosophila.">Modeling obesity and its associated disorders in Drosophila.</a> Physiology (Bethesda). 28 (2), 117-124 (2013).</li><li>Azeez, O. I., Meintjes, R., Chamunorwa, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fat+body,+fat+pad+and+adipose+tissues+in+invertebrates+and+vertebrates:+the+nexus.">Fat body, fat pad and adipose tissues in invertebrates and vertebrates: the nexus.</a> Lipids Health Dis. 13, 71 (2014).</li><li>Dourlen, P., Sujkowski, A., Wessells, R., Mollereau, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fatty+acid+transport+proteins+in+disease:+New+insights+from+invertebrate+models.">Fatty acid transport proteins in disease: New insights from invertebrate models.</a> Prog. Lipid Res. 60, 30-40 (2015).</li><li>Hildebrandt, A., Bickmeyer, I., K&#252;hnlein, R. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reliable+Drosophila+body+fat+quantification+by+a+coupled+colorimetric+assay.">Reliable Drosophila body fat quantification by a coupled colorimetric assay.</a> PLoS One. 6 (9), e23796 (2011).</li><li>Tennessen, J. M., Barry, W. E., Cox, J., Thummel, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+studying+metabolism+in+Drosophila.">Methods for studying metabolism in Drosophila.</a> Methods. 68, 105-115 (2014).</li><li>Al-Anzi, B., Zinn, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Colorimetric+measurement+of+triglycerides+cannot+provide+an+accurate+measure+of+stored+fat+content+in+Drosophila.">Colorimetric measurement of triglycerides cannot provide an accurate measure of stored fat content in Drosophila.</a> PLoS One. 5 (8), e12353 (2010).</li><li>Thanh, N. T. K., Stevenson, G., Obatoml, D., Bach, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+Lipids+in+Animal+Tissues+by+High-Performance+Thin-Layer+Chromatography+with+Densitometry.">Determination of Lipids in Animal Tissues by High-Performance Thin-Layer Chromatography with Densitometry.</a> J. Planar Chromatogr. 13, 375-381 (2000).</li><li>Borrull, A., Lopez-Martinez, G., Poblet, M., Cordero-Otero, R., Rozes, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+method+for+the+separation+and+quantification+of+neutral+lipid+species+using+GC-MS.">A simple method for the separation and quantification of neutral lipid species using GC-MS.</a> Eur. J. Lipid Sci. Technol. 117, 274-280 (2015).</li><li>Shui, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toward+one+step+analysis+of+cellular+lipidomes+using+liquid+chromatography+coupled+with+mass+spectrometry:+application+to+Saccharomyces+cerevisiae+and+Schizosaccharomyces+pombe+lipidomics.">Toward one step analysis of cellular lipidomes using liquid chromatography coupled with mass spectrometry: application to Saccharomyces cerevisiae and Schizosaccharomyces pombe lipidomics.</a> Mol. Biosyst. 6, 1008-1017 (2010).</li><li>Farvid, M. S., Ng, T. W. K., Chan, D. C., Barrett, P. H. R., Watts, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Association+of+adiponectin+and+resistin+with+adipose+tissue+compartments,+insulin+resistance+and+dyslipidaemia.">Association of adiponectin and resistin with adipose tissue compartments, insulin resistance and dyslipidaemia.</a> Diabetes. Obes. Metab. 7 (4), 406-413 (2005).</li><li>Urbanchek, M. G., Picken, E. B., Kalliainen, L. K., Kuzon, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Specific+force+deficit+in+skeletal+muscles+of+old+rats+is+partially+explained+by+the+existence+of+denervated+muscle+fibers.">Specific force deficit in skeletal muscles of old rats is partially explained by the existence of denervated muscle fibers.</a> J. Gerontol. A. Biol. Sci. Med. Sci. 56A (5), B191-B197 (2001).</li><li>Mosher, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coordination+between+Drosophila+Arc1+and+a+specific+population+of+brain+neurons+regulates+organismal+fat.">Coordination between Drosophila Arc1 and a specific population of brain neurons regulates organismal fat.</a> Dev. Biol. 405 (2), 280-290 (2015).</li><li>Reis, T., van Gilst, M. R., Hariharan, I. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+buoyancy-based+screen+of+drosophila+larvae+for+fat-+storage+mutants+reveals+a+role+for+Sir2+in+coupling+fat+Storage+to+Nutrient+Availability.">A buoyancy-based screen of drosophila larvae for fat- storage mutants reveals a role for Sir2 in coupling fat Storage to Nutrient Availability.</a> PLoS Genet. 6 (11), e1001206 (2010).</li><li>Teixeira, L., Rabouille, C., R&#248;rth, P., Ephrussi, A., Vanzo, N. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Perilipin/ADRP+homologue+Lsd2+regulates+lipid+metabolism.">Drosophila Perilipin/ADRP homologue Lsd2 regulates lipid metabolism.</a> Mech. Dev. 120 (9), 1071-1081 (2003).</li><li>Perez, C. L., Van Gilst, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+13C+Isotope+Labeling+Strategy+Reveals+the+Influence+of+Insulin+Signaling+on+Lipogenesis+in+C.+elegans.">A 13C Isotope Labeling Strategy Reveals the Influence of Insulin Signaling on Lipogenesis in C. elegans.</a> Cell Metab. 8, 266-274 (2008).</li></ol>

The authors declare that they have no competing financial interests.

There are a multitude of techniques that have been developed to measure lipid levels8-10. However, each of these methods comes with some major drawbacks that are addressed by the buoyancy-based assay outlined above. First, this assay is extremely quick. Testing a full concentration gradient takes no more than 30 - 60 min. This is a huge improvement on most of the techniques currently in use. For example, lipid quantitation by MS takes 7 - 9 hr to isolate the lipids and another several hours to analyze them. This is a strong deterrent when performing a screen on a large number of animals. By measuring a quick buoyancy phenotype, one can quickly focus on the genotypes that have the phenotype of interest. Second, the density assay requires only common reagents such as salts (for PBS) and sucrose. This makes it much easier and cheaper to screen a large number of larvae or quickly test an interesting genotype. Finally, this protocol is non-invasive and the larvae are still alive at the end. This allows further experimentation to be performed on the animals such as specific lipid quantitation or transcript or protein analysis. Additionally, recovered larvae may be allowed to continue growth to adulthood.
An additional benefit of this technique over others is increased sensitivity to small changes in fat levels in a population. By performing a whole concentration gradient on a population of 50 larvae, slight shifts in the population fat levels will be identified by a change in the intermediate concentrations, although the concentration at which the larvae begin floating and are totally floated may not be different than controls. This small shift in the population may not always be identified by other fat quantitation methods as they require a large group of larvae to analyze. The density assay on the other hand interrogates individuals within the population and can consequently identify these small changes or shifts in the overall population distribution.
As this technique relies on the density of the solution to produce reliable results, it is extremely important that the PBS and sucrose solutions are made consistently. We have found that different PBS recipes produce differences in solution density, leading to varying results. We use the recipe outlined above for consistent results. Furthermore, making the 20% sucrose from the same batch of 1x PBS is important as it results in the same background solution density. If the two solutions were made separately, slight variations in the PBS density could bring in additional density variable beyond the addition of sucrose to the experimental conical vial. Lastly, evaporation can be a problem, because over time the solutions will become more concentrated with the evaporation of water. It is therefore important to cap the bottles tightly and to remake the solution every week so that the experimental solutions remain consistent.
Among the steps outlined above, there are several critical steps that if altered, could result in inaccurate results by the buoyancy assay. First, it is important that the larvae transferred into the experimental food vial be the same age. Transferring larvae that vary in size will result in larvae at different developmental stages at the time the buoyancy assay is performed. This protocol has been developed to determine changes in fat levels in wandering L3 larvae and will not work accurately for early L3 or pre-pupae or later. Early third instar larvae have a tendency to float independently of fat levels while pre-pupae and pupae sink even at the highest concentrations used. For this reason, the age of the L1 larvae transferred is critical. Similarly, the point at which the vial is taken to perform the buoyancy assay is important. For the same developmental timing reasons stated above, the vials should be taken only when 20 - 40 larvae are wandering to ensure accurate density measurements. Additionally, the number of L1 transferred can affect results as well. The wide vials used in this protocol allow 50 larvae to eat without competition during development. A much larger number of larvae transferred than this will result in competition for food and may affect the amount of fat stored independent of genotype.
There may be genotypes with such extremely high fat levels that most or all of the larvae will float at the first concentration. In this case, the protocol may be modified by decreasing the concentration of the starting solution in order to obtain a full spectrum of the population density distribution. On the other hand, a specific genotype may be so lean that no larva floats in the initial sucrose. In this case, the initial steps may be omitted and a higher starting concentration used. This assay is very flexible and can be modified to better fit a wide range of fat levels. We advise the use of controls such as appropriate wild type animals (controlled for genetic background), positive controls (established fat mutants, such as adp or Sir217), and negative controls (published lean mutants such as lsd219). Appropriate use of this protocol will allow future screening of collections to determine new regulators of metabolism and obesity predisposition. Furthermore, this protocol provides an easy way for researchers in any field to test whether their genetic manipulation of interest causes a change in fat levels, without the need for complex protocols or expensive equipment. This protocol will help to elucidate the complex relationship between genetics and obesity.

Drosophila melanogaster has been used for over a century in the study of genetics and other basic biological questions. In the last few decades, it has become clear that Drosophila is a powerful tool in the study of many human diseases. As 70 - 80% of genes associated with human diseases have an identified fly ortholog1-4, Drosophila provides a simplified yet translatable system in which to study complex diseases. Metabolism in particular has benefited from such study. Not only are the genetics of metabolism well conserved between flies and humans, but the relevant organs and cell types are also very similar2,5. For example, the fat body of the fly stores both triacylglycerides (TAG) and glycogen, functions analogous to those performed in mammalian liver and white adipose tissue6. Using Drosophila as a model for human obesity has vastly improved our understanding of lipid metabolism and the genetics of obesity7. The larval stage of development is particularly useful for studying the segregation of nutrients to storage or utilization as it is dedicated to feeding and the storage of energy to be used during pupariation.
Currently, there are many different quantitative methods of determining fat storage levels in Drosophila. The most widely used method is the coupled colorimetric assay (CCA)8,9. CCA was developed for determining TAG levels in human serum and operates on the premise that glycerol liberated from the backbone of triglycerides will undergo several reactions, ultimately resulting in a redox-coupled reaction generating a colored product. Absorbance of specific wavelengths of light is then measured to determine the initial amount of glycerol present. However, glycerol can also be liberated from mono- and diacylglycerides in addition to TAG and therefore may not be an accurate measure of stored body fat9. Furthermore, eye pigment of crushed adult flies can interfere with some absorbance readings and complicate results9,10. Therefore, CCA must be accompanied and validated by thin layer chromatography (TLC), which allows for the separation of most lipid families that can be quantitated by densitometry10,11. However, some lipid classes like sterols cannot be analyzed and must be quantified a different way12. Mass spectrometry (MS) is an accurate way to quantitate all classes of major cellular lipids12,13. However, the lipid extraction procedures necessary to analyze by MS are both time consuming (most taking nearly a full day) and costly. Here we present an alternative method to quickly and cheaply determine organismal fat levels in the L3 larvae of Drosophila melanogaster.
The method presented below exploits the difference in density between fat tissue and lean tissue. Mammalian fat tissue has a density of approximately 0.9 g/ml14 while skeletal muscle as a density of 1.06 g/ml15. This difference means that animals with higher stores of fat will have lower density than animals with lower stores of fat, which will allow them to float better in a solution of fixed density. This property allows for extremely quick screening of a large number of animals while being both inexpensive and non-invasive. Buoyancy-based analysis has been used both to confirm the phenotypes of altering known regulators of fat levels as well as to identify new genetic and neurological regulators of obesity16,17.

<table><tbody><tr><td>Bacto Agar</td><td>VWR</td><td>214030</td><td></td></tr><tr><td>Welch&#039;s Original 100% Grape Juice</td><td></td><td></td><td></td></tr><tr><td>Sucrose</td><td>Fisher</td><td>S512</td><td></td></tr><tr><td>Tegosept</td><td>Genesee Scientific</td><td>20-259</td><td></td></tr><tr><td>Ethanol</td><td></td><td></td><td>200 proof</td></tr><tr><td>Baker&#039;s yeast</td><td>Fleichmann&#039;s</td><td></td><td></td></tr><tr><td>Yellow Corn Meal</td><td>Quaker</td><td></td><td>Enriched degerminated</td></tr><tr><td>Drosophila Agar Type II</td><td>Apex</td><td>66-104</td><td></td></tr><tr><td>Soy Flour</td><td>ADM Specialty Ingredients</td><td>062-100</td><td></td></tr><tr><td>Malt Extract</td><td>Breiss</td><td>5748</td><td></td></tr><tr><td>Light Corn Syrup</td><td>Karo</td><td></td><td></td></tr><tr><td>Propionic Acid</td><td>VWR</td><td>U330-09</td><td></td></tr><tr><td>Sodium chloride</td><td>Fisher</td><td>50147491</td><td></td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma</td><td>P5655-100G</td><td></td></tr><tr><td>Sodium phosphate anhydrous</td><td>Fisher</td><td>S3933</td><td></td></tr><tr><td>Potassium chloride</td><td>Fisher</td><td>P330-500</td><td></td></tr><tr><td>35 x 10 mm petri plate</td><td>Fisher</td><td>08-757-100A</td><td></td></tr><tr><td>50 ml Conical</td><td>Fisher</td><td>50-869-569</td><td></td></tr></tbody></table>

a buoyancy-based method of determining fat levels in drosophila

Drosophila melanogaster is a key experimental system in the study of fat regulation. Numerous techniques currently exist to measure levels of stored fat in Drosophila, but most are expensive and/or laborious and have clear limitations. Here, we present a method to quickly and cheaply determine organismal fat levels in L3 Drosophila larvae. The technique relies on the differences in density between fat and lean tissues and allows for rapid detection of fat and lean phenotypes. We have verified the accuracy of this method by comparison to body fat percentage as determined by neutral lipid extraction and gas chromatography coupled with mass spectrometry (GCMS). We furthermore outline detailed protocols for the collection and synchronization of larvae as well as relevant experimental recipes. The technique presented below overcomes the major shortcomings in the most widely used lipid quantitation methods and provides a powerful way to quickly and sensitively screen L3 larvae for fat regulation phenotypes while maintaining the integrity of the larvae. This assay has wide applications for the study of metabolism and fat regulation using Drosophila.

Figure 1 represents an example of how the buoyancy-based assay can detect both higher and lower fat stores in genetically manipulated larvae. Figure 1A shows that excitation or silencing of a certain subset of neurons (E347) in the larval brain induces lower and higher fat stores respectively. The same genetic background control was used in both cases. Figure 1B provides an example of how this phenotype obtained by the density assay is corroborated by neutral lipid extraction and quantification by GCMS.
<img alt="Figure 1" src="/files/ftp_upload/54744/54744fig1.jpg" /> 
Figure 1. Neuronal Function in Specific Regions of the Larval Brain is Necessary and Sufficient for Body Fat Regulation. Neuronal activity in the indicated E347-GAL4 line was silenced by overexpression of ShiDN, or activated by overexpression of NB, as indicated in the panels, and effects on body fat were assessed. (A) Percent of floating larvae at equilibrium in different density solutions. Fifty larvae were examined per biological replicate, n= 7 - 9 biological replicates per genotype. (B) Percent body fat (total neutral lipids including triacylglycerides and phospholipids divided by body weight) as measured by GCMS17,20 (n= 8). Blue lines or bars represent GAL4-only (no UAS) animals crossed to w1118 to control for effects of genetic background. P values represent results from two-tailed t tests. **** P &#60; 0.0001, *** P = 0.0001 to 0.001, ** P = 0.001 to 0.01, * P = 0.01 to 0.05. Error bars show SEM. This figure has been modified16. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54744/54744fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Buoyancy-based Method of Determining Fat Levels in Drosophila at JoVE.com

A Buoyancy-based Method of Determining Fat Levels in Drosophila

Here we present a method to measure organismal fat levels in the third instar (L3) larval stage of Drosophila melanogaster. This method exploits the comparatively low density of fat tissue to differentiate between larvae with altered fat stores. Buoyancy-based analysis is a valuable tool for rapid, reproducible, and economical screening.

A Buoyancy-based Method of Determining Fat Levels in Drosophila

Drosophila melanogaster is a key experimental system in the study of fat regulation. Numerous techniques currently exist to measure levels of stored fat ...

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Biology

8.6K Views.  University of Colorado Anschutz Medical Campus. Here we present a method to measure organismal fat levels in the third instar (L3) larval stage of Drosophila melanogaster. This method exploits the comparatively low density of fat tissue to differentiate between larvae with altered fat stores. Buoyancy-based analysis is a valuable tool for rapid, reproducible, and economical screening.

Video: A Buoyancy-based Method of Determining Fat Levels in Drosophila

Quantifying Abdominal Pigmentation in Drosophila melanogaster

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for understanding the development and evolution of morphological phenotypes. Abdominal pigmentation in Drosophila melanogaster has been particularly useful, allowing researchers to identify the loci that underlie inter- and intraspecific variations in morphology. Hitherto, however, D. melanogaster abdominal pigmentation has been largely assayed qualitatively, through scoring, rather than quantitatively, which limits the forms of statistical analysis that can be applied to pigmentation data. This work describes a new methodology that allows for the quantification of various aspects of the abdominal pigmentation pattern of adult D. melanogaster. The protocol includes specimen mounting, image capture, data extraction, and analysis. All the software used for image capture and analysis feature macros written for open-source image analysis. The advantage of this approach is the ability to precisely measure pigmentation traits using a methodology that is highly reproducible across different imaging systems. While the technique has been used to measure variation in the tergal pigmentation patterns of adult D. melanogaster, the methodology is flexible and broadly applicable to pigmentation patterns in myriad different organisms.

Quantifying Abdominal Pigmentation in Drosophila melanogaster

This work presents a method to quickly and precisely quantify the abdominal pigmentation of Drosophila melanogaster using digital image analysis. This method streamlines the procedures between phenotype acquisition and data analysis and includes specimen mounting, image acquisition, pixel value extraction, and trait measurement.

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for ...

Genetics

High Fat Diet Feeding and High Throughput Triacylglyceride Assay in Drosophila Melanogaster

Heart disease is the number one cause of human death worldwide. Numerous studies have shown strong connections between obesity and cardiac malfunction in humans, but more tools and research efforts are needed to better elucidate the mechanisms involved. For over a century, the genetically highly tractable model of Drosophila has been instrumental in the discovery of key genes and molecular pathways that proved to be highly conserved across species. Many biological processes and disease mechanisms are functionally conserved in the fly, such as development (e.g., body plan, heart), cancer, and neurodegenerative disease. Recently, the study of obesity and secondary pathologies, such as heart disease in model organisms, has played a highly critical role in the identification of key regulators involved in metabolic syndrome in humans.
Here, we propose to use this model organism as an efficient tool to induce obesity, i.e., excessive fat accumulation, and develop an efficient protocol to monitor fat content in the form of TAGs accumulation. In addition to the highly conserved, but less complex genome, the fly also has a short lifespan for rapid experimentation, combined with cost-effectiveness. This paper provides a detailed protocol for High Fat Diet (HFD) feeding in Drosophila to induce obesity and a high throughput triacylglyceride (TAG) assay for measuring the associated increase in fat content, with the aim to be highly reproducible and efficient for large-scale genetic or chemical screening. These protocols offer new opportunities to efficiently investigate regulatory mechanisms involved in obesity, as well as provide a standardized platform for drug discovery research for rapid testing of the effect of drug candidates on the development or prevention of obesity, diabetes and related metabolic diseases.

High Fat Diet Feeding and High Throughput Triacylglyceride Assay in Drosophila Melanogaster

This is a high fat diet feeding protocol to induce obesity in Drosophila, a model for understanding fundamental molecular mechanisms implicated in lipotoxicity. It also provides a high throughput triacylglyceride assay for measuring fat accumulation in Drosophila and potentially other (insect) models under various dietary, environmental, genetic or physiological conditions.

Heart disease is the number one cause of human death worldwide. Numerous studies have shown strong connections between obesity and cardiac malfunction in ...

Combining Quantitative Food-intake Assays and Forcibly Activating Neurons to Study Appetite in Drosophila

Food consumption is under the tight control of the brain, which integrates the physiological status, palatability, and nutritional contents of the food, and issues commands to start or stop feeding. Deciphering the processes underlying the decision-making of timely and moderate feeding carries major implications in our understanding of physiological and psychological disorders related to feeding control. Simple, quantitative, and robust methods are required to measure the food ingestion of animals after experimental manipulation, such as forcibly increasing the activities of certain target neurons. Here, we introduced dye-labeling-based feeding assays to facilitate the neurogenetic study of feeding control in adult fruit flies. We review available feeding assays, and then describe our methods step-by-step from setup to analysis, which combine thermogenetic and optogenetic manipulation of neurons controlling feeding motivation with dye-labeled food intake assay. We also discuss the advantages and limitations of our methods, compared with other feeding assays, to help readers choose an appropriate assay.

Combining Quantitative Food-intake Assays and Forcibly Activating Neurons to Study Appetite in Drosophila

Quantitative food-intake assays with dyed food provide a robust and high-throughput means to evaluate feeding motivation. Combining the food consumption assay with thermogenetic and optogenetic screens is a powerful approach to investigate the neural circuits underlying appetite in adult Drosophila melanogaster.

Food consumption is under the tight control of the brain, which integrates the physiological status, palatability, and nutritional contents of the food, ...

Behavior

The CApillary FEeder Assay Measures Food Intake in Drosophila melanogaster

For most animals, feeding is an essential behavior for securing survival, and it influences development, locomotion, health and reproduction. Ingestion of the right type and quantity of food therefore has a major influence on quality of life. Research on feeding behavior focuses on the underlying processes that ensure actual feeding and unravels the role of factors regulating internal energy homeostasis and the neuronal bases of decision-making. The model organism Drosophila melanogaster, with its great variety of genetically traceable tools for labeling and manipulating single neurons, allows mapping of neuronal networks and identification of molecular signaling cascades involved in the regulation of food intake. This report demonstrates the CApillary FEeder assay (CAFE) and shows how to measure food intake in a group of flies for time spans ranging from hours to days. This easy-to-use assay consists of glass capillaries filled with liquid food that flies can freely access and feed on. Food consumption in the assay is accurately determined using simple measurement tools. Herein we describe step-by-step the method from setup to successful execution of the CAFE assay, and provide practical examples to analyze the food intake of a group of flies under controlled conditions. The reader is guided through possible limitations of the assay, and advantages and disadvantages of the method compared to other feeding assays in D. melanogaster are evaluated.

The CApillary FEeder Assay Measures Food Intake in Drosophila melanogaster

The CApillary FEeder (CAFE) assay is a simple, budget-friendly, highly reliable method for investigating mechanisms underlying food intake. Used with the highly versatile genetic model organism Drosophila melanogaster, it provides a powerful means of gaining new insights into regulatory mechanisms of food intake.

For most animals, feeding is an essential behavior for securing survival, and it influences development, locomotion, health and reproduction. Ingestion of ...

Neuroscience

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

The nematode C. elegans has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans metabolic research.

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

We present robust biochemical and microscopic methods for studying Caenorhabditis elegans lipid stores. A rapid, simple, fixing-staining procedure for fluorescent lipid droplet imaging leverages the spectral properties of the lipophilic dye Nile red. We then present biochemical measurement of triglycerides and phospholipids using solid phase extraction and gas chromatography-mass spectrometry.

The nematode C. elegans has emerged as an important  model for the study of conserved genetic pathways regulating fat metabolism as  it relates to human ...

Quantification of Macronutrients Intake in a Thermogenetic Neuronal Screen using Drosophila Larvae

Foraging and feeding behaviors allow animals to access sources of energy and nutrients essential for their development, health, and fitness. Investigating the neuronal regulation of these behaviors is essential for the understanding of the physiological and molecular mechanisms underlying nutritional homeostasis. The use of genetically tractable animal models such as worms, flies, and fish greatly facilitates these types of studies. In the last decade, the fruit fly Drosophila melanogaster has been used as a powerful animal model by neurobiologists investigating the neuronal control of feeding and foraging behaviors. While undoubtedly valuable, most studies examine adult flies. Here, we describe a protocol that takes advantage of the simpler larval nervous system to investigate neuronal substrates controlling feeding behaviors when larvae are exposed to diets differing in their protein and carbohydrates content. Our methods are based on a quantitative colorimetric no-choice feeding assay, performed in the context of a neuronal thermogenetic-activation screen. As a read-out, the amount of food eaten by larvae over a 1 h interval was used when exposed to one of the three dye-labeled diets that differ in their protein to carbohydrates (P:C) ratios. The efficacy of this protocol is demonstrated in the context of a neurogenetic screen in larval Drosophila, by identifying candidate neuronal populations regulating the amount of food eaten in diets of different macronutrient quality. We were also able to classify and group the genotypes tested into phenotypic classes. Besides a brief review of the currently available methods in the literature, the advantages and limitations of these methods are discussed and, also, some suggestions are provided about how this protocol might be adapted to other specific experiments.

Quantification of Macronutrients Intake in a Thermogenetic Neuronal Screen using Drosophila Larvae

Described here is a protocol that enables the colorimetric quantification of the amount of food eaten within a defined interval of time by Drosophila melanogaster larvae exposed to diets of different macronutrient quality. These assays are conducted in the context of a neuronal thermogenetic screen.

Foraging and feeding behaviors allow animals to access sources of energy and nutrients essential for their development, health, and fitness. Investigating ...

An Improved Method for Accurate and Rapid Measurement of Flight Performance in Drosophila

Drosophila has proven to be a useful model system for analysis of behavior, including flight. The initial flight tester involved dropping flies into an oil-coated graduated cylinder; landing height provided a measure of flight performance by assessing how far flies will fall before producing enough thrust to make contact with the wall of the cylinder. Here we describe an updated version of the flight tester with four major improvements. First, we added a &#34;drop tube&#34; to ensure that all flies enter the flight cylinder at a similar velocity between trials, eliminating variability between users. Second, we replaced the oil coating with removable plastic sheets coated in Tangle-Trap, an adhesive designed to capture live insects. Third, we use a longer cylinder to enable more accurate discrimination of flight ability. Fourth we use a digital camera and imaging software to automate the scoring of flight performance. These improvements allow for the rapid, quantitative assessment of flight behavior, useful for large datasets and large-scale genetic screens.

An Improved Method for Accurate and Rapid Measurement of Flight Performance in Drosophila

Here we describe a method for rapid and accurate measurement of flight performance in Drosophila, enabling high-throughput screening.

Drosophila has proven to be a useful model system for analysis of behavior, including flight. The initial flight tester involved dropping flies into an ...

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and diseases, such as obesity and fatty liver. Usually, lipids are stored in lipid droplets, which are the multifunctional lipid storage organelles in cells. Lipid droplets vary in size and number in different tissues and under different conditions. It has been reported that lipid droplets are tightly controlled through regulation of its biogenesis and degradation. In Drosophila melanogaster, the oenocyte is an important tissue for lipid metabolism and has been recently identified as a human liver analogue regarding lipid mobilization in response to stress. However, the mechanisms underlying the regulation of lipid droplet metabolism in oenocytes remain elusive. To solve this problem, it is of utmost importance to develop a reliable and sensitive method to directly visualize lipid droplet dynamic changes in oenocytes during development and under stressful conditions. Taking advantage of the lipophilic BODIPY 493/503, a lipid droplet-specific fluorescent dye, described here is a detailed protocol for the dissection and subsequent lipid droplet staining in the oenocytes of Drosophila larvae in response to starvation. This allows for qualitative analysis of lipid droplet dynamics under various conditions by confocal microscopy. Furthermore, this rapid and highly reproducible method can also be used in genetic screens for indentifying novel genetic factors involving lipid droplet metabolism in oenocytes and other tissues.

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Presented here are detailed methods for the dissection and lipid droplet staining of oenocytes in Drosophila larvae using BODIPY 493/503, a lipid droplet-specific fluorescent dye.

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and ...

Analyzing Starvation-Induced Autophagy in the Drosophila melanogaster Larval Fat Body

Autophagy is a cellular self-digestion process. It delivers cargo to the lysosomes for degradation in response to various stresses, including starvation. The malfunction of autophagy is associated with aging and multiple human diseases. The autophagy machinery is highly conserved-from yeast to humans.&#160;The larval fat body of Drosophila melanogaster, an analog for vertebrate liver and adipose tissue, provides a unique model for monitoring autophagy in vivo. Autophagy can be easily induced by nutrient starvation in the larval fat body. Most autophagy-related genes are conserved in Drosophila. Many transgenic fly strains expressing tagged autophagy markers have been developed, which facilitates the monitoring of different steps in the autophagy process. The clonal analysis enables a close comparison of autophagy markers in cells with different genotypes in the same piece of tissue. The current protocol details procedures for (1) generating somatic clones in the larval fat body, (2) inducing autophagy via amino acid starvation, and (3) dissecting the larval fat body, aiming to create a model for analyzing differences in autophagy using an autophagosome marker (GFP-Atg8a) and clonal analysis.

Analyzing Starvation-Induced Autophagy in the Drosophila melanogaster Larval Fat Body

The present protocol describes the induction of autophagy in the Drosophila melanogaster larval fat body via nutrient depletion and analyzes changes in autophagy using transgenic fly strains.

Autophagy is a cellular self-digestion process. It delivers cargo to the lysosomes for degradation in response to various stresses, including starvation. ...

Gut Acidification Monitoring Assay: A Technique to Study the Acidification of the Intestinal Lumen in Drosophila using Bromophenol Blue Dye

Source: Abu, F., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/63141">Monitoring Gut Acidification in the Adult Drosophila Intestine.</a> J. Vis. Exp. (2021).
This video demonstrates an assay to monitor gut acidification in Drosophila flies. After the ingestion of food supplemented with a pH indicator dye, the acidification of the intestinal lumen results in a change in the color of the dye. Visual inspection of the digestive tract of the flies to examine the color change provides evidence of acidification.

Source: Abu, F., et al. Monitoring Gut Acidification in the Adult Drosophila Intestine. J. Vis. Exp. (2021).
This video demonstrates an assay to monitor ...

A Buoyancy-based Method of Determining Fat Levels in Drosophila

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Chapters in this video

Quantifying Abdominal Pigmentation in Drosophila melanogaster

High Fat Diet Feeding and High Throughput Triacylglyceride Assay in Drosophila Melanogaster

Combining Quantitative Food-intake Assays and Forcibly Activating Neurons to Study Appetite in Drosophila

The CApillary FEeder Assay Measures Food Intake in Drosophila melanogaster

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

Quantification of Macronutrients Intake in a Thermogenetic Neuronal Screen using Drosophila Larvae

An Improved Method for Accurate and Rapid Measurement of Flight Performance in Drosophila

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Analyzing Starvation-Induced Autophagy in the Drosophila melanogaster Larval Fat Body

Gut Acidification Monitoring Assay: A Technique to Study the Acidification of the Intestinal Lumen in Drosophila using Bromophenol Blue Dye