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 dissol...

<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 hou...

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.

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			<td height="42" style="height:42px;width:216px;">Welch&#39;s Original 100% Grape Juice</td>
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			<td height="21" style="height:21px;width:216px;">Sodium chloride</td>
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			<td height="21" style="height:21px;">Potassium phosphate monobasic</td>
			<td>Sigma</td>
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			<td height="21" style="height:21px;width:216px;">Sodium phosphate anhydrous</td>
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			<td height="21" style="height:21px;">Potassium chloride</td>
			<td>Fisher</td>
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			<td height="21" style="height:21px;width:216px;">35 x 10mm petri plate</td>
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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 co...

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 ...

The overall goal of this assay is to determine any changes in fat storage levels that may be caused by a change in genetics, diet, or treatment. These methods can help answer key questions in the field of metabolism such as genetic or neuronal contributions to obesity predisposition. The main advantage of this technique is that it's fast, economical, and very reproducible when compared to other methods currently used.Individuals new to this method or new to working with wandering stage larva may struggle with this protocol because the developmental timeline of different genotypes can be highly variable. To begin this procedure, add a small dollop of yeast paste to the middle of a grate plate. Smooth down any rough edges with a spatula.Next, transfer the flies of interest into an egg collection chamber, and place the grate plate with yeast paste on top facing inwards. Then tape the grate plate to the egg collection chamber to keep it from falling out. Make sure to write important genotype and date information on the bottom of the grate plate.Subsequently, upend the egg collection chamber to allow the flies to lay eggs on the grate plate. After that, cover the egg collection chamber with an open box and place the covered egg collection chamber in an incubator at 25 degrees celsius for four to six hours to allow the flies to lay eggs. When it is done, take the grate plate out of the egg collection chamber and transfer the flies back into their original bottle of food.Use a spatula to wipe off any excess yeast paste from the grate plate. Subsequently, store the grate plate in a larger Petri dish in a humidified incubator at 25 degrees celsius for 24 hours. In this procedure, score the top layer of the food by plunging it with a spatula several times to allow the larvae to burrow and feed.22 to 24 hours after egg collection use a small paintbrush to collect 50 first instar larvae of the same approximate size and place the larvae in the experimental food vial. It's very important that larvae of the same approximate size are selected and transferred to the experimental food. This will ensure tight coordination of larval development.Next, clean the paintbrush with a laboratory wipe and ensure that there are no remaining larvae on the paintbrush before continuing to another genotype. Afterward, place the vial of larvae in an incubator at 25 degrees celsius and allow them to develop. Remove the vial of larvae from the incubator once there are 20 to 40 larvae wandering up the sides of the vial.Record the date and time of the assay as well as the number of wandering larvae. Then prepare the experimental solution by adding 11.5 milliliters of PBS and nine milliliters of 20%sucrose to a 50 milliliter conical tube. Add 20%sucrose to the vial until it reaches one inch from the top of the vial.Use a spatula to gently stir the food top layer to free the larvae that are still burrowing in it. All the larvae will rise to the surface of the sucrose solution. Afterward, gently transfer the larvae into the initial sucrose solution and gently stir them.While transferring the larvae to the experimental solution, be careful not to squeeze or puncture the larvae. This may alter the results of the assay. Cap the conical tube and upend it several times to thoroughly mix the solution.Then uncap the conical tube and swirl it to create a gentle vortex. Allow two to five minutes for the larvae to settle, either floating to the top of the solution or sinking to the bottom. Record the number of floating larvae, and the specific sucrose concentration tested.Then add one milliliter of 20%sucrose to the experimental solution to increase its density. Subsequently, use a spatula to stir the larvae gently. Following this, cap the conical tube and upend it several times to thoroughly mix the solution.Subsequently, uncap the conical tube and swirl to create a gentle vortex. Record the number of floating larvae and the new concentration. Continue this process until at least 95%of the larvae are floating.Add 20%sucrose to the conical vial until it is one inch from the top. Next collect all the larvae in a dissection dish filled with PBS. Examine the larvae under a microscope for any small larvae, prepupae, pupae, or any other differences between genotypes to ensure that the procedures were done appropriately.Then record the total number of larvae. Afterward, transfer 10 larvae on a laboratory wipe for drying. Label a microcentrifuge tube and place the larvae in the tube.Subsequently, flash freeze the larvae in liquid nitrogen and store the tube at minus 20 degrees celsius. Shown here is an example of how the buoyancy-based assay can detect both higher and lower fat stored in the genetically manipulated larvae. This figure shows that excitation or silencing of a certain subset of neurons in the larval brain induces lower and higher stored fat respectively.The same genetic background control was used in both cases. Here is an example of how this phenotype obtained by the density assay is corroborated by neutral lipid extraction and quantification by GCMS. Once mastered, this technique can be done within 30 minutes if properly performed, and may be performed in tandem with other genotypes.While attempting this procedure, it's important to remember to synchronize the development of the experimental larvae as much as possible. Following this procedure, larvae can be snap frozen in liquid nitrogen to be used in other methods like gas chromatography mass spectrometry in order to obtain additional information like the specific classes of lipid types in the larvae. After its development, this technique paved the way for researchers in metabolism to explore the genetic and or neuronal contributions to obesity in Drosophila.After watching this video, you should have a good understanding of how to quickly screen genotypes for changes in levels of stored fats.

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8.5K vues.  University of Colorado Anschutz Medical Campus. The overall goal of this assay is to determine any changes in fat storage levels that may be caused by a change in genetics, diet, or treatment. These methods can help answer key questions in the field of metabolism such as genetic or neuronal contributions to obesity predisposition. The main advantage of this technique is that it's fast, economical, and very reproducible when compared to other methods currently used.Individuals new to this method or new to working with wandering stage...