This work was made possible by the NIH grant: R01GM109028 (S.P.C.)

1. Nile Red (NR) Staining of Lipids
<ol>
	<li>Preparation of 5 mg/mL NR stock solution
	<ol>
		<li>In a 500 mL bottle, add 100 mg of NR powder to 200 mL of 100% acetone.</li>
		<li>Cover the bottle with aluminum foil to avoid any exposure to light.</li>
		<li>Before use, stir the solution for 2 h in the dark.</li>
		<li>For long term use, store the NR stock solution in a tightly sealed bottle without any light exposure. Scale NR stock solution according to research needs. Ensure that the same stock solution is used across NR-staining experiments to obtain consistent staining and imaging results.</li>
	</ol>
	</li>
	<li>Preparation of NR working solution
	<ol>
		<li>For every 1 mL of 40% isopropanol (v/v), add 6 &#181;L of NR stock solution.</li>
		<li>Prepare 600 &#181;L of NR working solution for each sample. 
		NOTE: Make fresh NR working solution right before staining. Depending on NR stock solution needs, use either a 15 mL or 50 mL graduated conical tube.</li>
	</ol>
	</li>
	<li>Preparation of worms for NR lipid staining
	<ol>
		<li>Grow worms to early L4 stage at 20 &#176;C on nematode growth medium (NGM) seeded with late-log OP50 E. coli.</li>
		<li>Wash the worms off the plate with 1 mL of 1x phosphate-buffered saline + 0.01% Triton X-100 (PBST) solution and put the worm suspension in a 1.5 mL microfuge tube.</li>
		<li>Centrifuge the worms at 560 x g for 1 min. Remove the supernatant and repeat this step until E. coli is cleared from suspension.</li>
		<li>Add 100 &#181;L of 40% isopropanol to the worm pellet and incubate it at room temperature for 3 min.</li>
		<li>Centrifuge the worms at 560 x g for 1 min and remove the supernatant without disrupting the worm pellet. 
		NOTE: Use higher volumes of PBST to wash the worms off the plates if using larger plates or staining more worms per plate. Do not wash the worms in PBST more than 15 min before the sample fixation. Perform additional washes if the supernatant is not cleared of bacteria. Carry out the incubation in 40% isopropanol using a nutator or rocker. Always monitor the tubes to make sure full sample agitation is occurring.</li>
	</ol>
	</li>
	<li>Lipid staining with NR
	<ol>
		<li>In the dark, add 600 &#181;L of NR working solution to each sample. Invert the tubes three times and fully mix the worms in NR solution.</li>
		<li>Rotate the sample in the dark at room temperature for 2 h.</li>
		<li>Following the incubation, centrifuge the worms at 560 x g for 1 min and remove the supernatant.</li>
		<li>Add 600 &#181;L of PBST and incubate the samples in the dark for 30 min to remove excess NR stain.</li>
		<li>Centrifuge the samples at 560 x g for 1 min and remove all but approximately 50 &#181;L of supernatant. 
		NOTE: NR incubation does not require agitation. Settling of worms is common during this step.</li>
	</ol>
	</li>
	<li>Preparation of slides for microscope imaging
	<ol>
		<li>Resuspend the worm pellet in remaining supernatant.</li>
		<li>Place 5 &#181;L of worm suspension on a microscope slide and put a coverslip on carefully to avoid trapping any air bubbles.</li>
		<li>Seal the coverslip with nail polish before imaging worms.</li>
		<li>Prepare only a couple slides at a time. This will ensure NR imaging remains constant across samples. The quality of NR-stained images diminishes after 6 h. Discrete lipid droplets are difficult to observe and background fluorescence increases, which interferes with NR signal detection.</li>
	</ol>
	</li>
	<li>Imaging of NR-stained worms
	<ol>
		<li>Image the worms at 5X magnification to capture several animals per field of view.</li>
		<li>Switch to 10X magnification for better quantification of individual worms.</li>
		<li>Use FITC/GFP channel to image NR-stained worms and try different exposure times to determine the optimal conditions for quantification.</li>
		<li>Save files in TIF format to avoid losing data due to compression. 
		NOTE: Typical exposure times range from 100-1000 ms. Having an optimal exposure time, use it for all samples to maintain imaging consistency.</li>
	</ol>
	</li>
	<li>NR staining image quantification
	<ol>
		<li>Upload micrographs to ImageJ. In the Plugins pull-down menu, use the Bio-Formats function if the image file is not recognized.</li>
		<li>In the Image pull-down menu, under the Stacks function, use Images-to-Stack to create an image stack when several micrographs are compared.</li>
		<li>In the same pull-down menu, adjust Brightness/Contrast of the image stack and propagate changes to all images by clicking Apply.</li>
		<li>Employ the Polygon selection tool to delineate each imaged worm and use the Measure function under the Analysis pull-down menu to quantify fluorescence intensity emitted by NR.</li>
		<li>For each image, measure five background locations and calculate the average background fluorescence intensity.</li>
		<li>Subtract the background fluorescence intensity from each imaged worm using the formula N = G - (A x B), where N stands for net fluorescence, G for gross fluorescence, A for total worm area and B for average background fluorescence.</li>
		<li>To normalize the fluorescence intensity by worm size, divide each worm&#39;s net fluorescence by its total area. The results are reported as fluorescence intensity, in arbitrary units (a.u.), per pixel. 
		NOTE: Avoid exporting images in JPEG format. While compression makes files more manageable to store, data integrity is compromised by this format. Make sure that all images are modified and analyzed identically. Remember to include a scale bar to properly evaluate size at different magnifications. To better visualize differences in fluorescence intensity using ImageJ, switch image type from RGB to 8-bit and select Fire from the image lookup tables (LUT) menu. This creates a heat-map image in which increased color brightness correlates with increased fluorescence intensity.</li>
	</ol>
	</li>
</ol>
2. Oil Red O Staining (ORO) of Lipids
<ol>
	<li>Preparation of ORO stock solution
	<ol>
		<li>In a 250 mL bottle, add 500 mg of ORO powder to 100 mL of 100% isopropanol and mix it well.</li>
		<li>Once prepared, store the solution tightly sealed with no light exposure.</li>
	</ol>
	</li>
	<li>Preparation of ORO working solution
	<ol>
		<li>Dilute ORO stock solution in water (3:2) to 60% isopropanol.</li>
		<li>Before use, filter the working ORO solution through a 0.2 &#181;m cellulose acetate sterile syringe filter. 
		NOTE: For best solution quality, prepare working ORO solution the day before, and let it mix overnight. However, if solution is needed same day, dilute and mix ORO solution on a rocker for at least 2 h before use. Before rocking, wrap conical tube in paraffin tape to avoid leakage of ORO solution.</li>
	</ol>
	</li>
	<li>Preparation of worms for ORO lipid staining
	<ol>
		<li>Grow the worms at 20 &#176;C on nematode growth medium (NGM) seeded with late-log OP50 E. coli to desired life stage.</li>
		<li>Add 1 mL of PBST solution to the plate and swirl until all worms are off the plate. Tilt the plate and wash it with 1 mL of PBST. Transfer the worm suspension to a 1.5 mL microfuge tube.</li>
		<li>Centrifuge the worms at 560 x g for 1 min. Remove the supernatant without disturbing the pellet and repeat the washing step with 1 mL of PBST three times. Remove all supernatant but 100 &#181;L.</li>
		<li>Add 600 &#181;L of 40% isopropanol to the worm pellet and rock it at room temperature for 3 min.</li>
		<li>Centrifuge the worms at 560 x g for 30 s and remove all supernatant but 100 &#181;L without disrupting the worm pellet. 
		NOTE: Use higher volumes of PBST to wash worms off plates if doing high-throughput staining. Do not wash worms in PBST more than 15 min prior to sample fixation. Do additional washes if supernatant is not cleared of bacteria. Carry out incubation in 40% isopropanol using a nutator or rocker. Always monitor tubes to make sure full sample agitation is occurring.</li>
	</ol>
	</li>
	<li>Lipid staining with ORO
	<ol>
		<li>Add 600 &#181;L ORO working solution to each sample. Invert the tube three times and mix the worms well in ORO.</li>
		<li>Rotate the samples at 30 rpm for 2 h at room temperature.</li>
		<li>Centrifuge the samples at 560 x g for 1 min and eliminate all supernatant but 100 &#181;L.</li>
		<li>Resuspend the samples in 600 &#181;L of PBST and rotate the tubes at 30 rpm for 30 min to remove excess ORO stain.</li>
		<li>Centrifuge samples at 560 x g for 1 min and eliminate all but 50 &#181;L of supernatant.</li>
		<li>To better distinguish the location of the animal&#39;s germline and intestinal cells, stain nuclei by adding 1 &#181;L of (2-(4-amidinophenyl)-1H -indole-6-carboxamidine) (DAPI) for every 1 mL of ORO working solution. Carry out ORO staining for 2 h with adding DAPI prior to mounting slides for imaging. Intestinal nuclei are larger in size and the gonad is distinguished by the developing germ cells. 
		NOTE: After ORO staining, worms may adhere to the sides of the microfuge tube and fail to form a noticeable pellet. If this happens, centrifuge the worms again or allow worms to settle by gravity for at least 10 min.</li>
	</ol>
	</li>
	<li>Preparation of the slides for worm imaging
	<ol>
		<li>Resuspend the worms in the remaining supernatant and mix the solution well.</li>
		<li>Put 5 &#181;L of worm suspension on a microscope slide and place a coverslip carefully, ensuring that no air bubbles form.</li>
		<li>Seal the coverslip with nail polish and image worms. 
		NOTE: After fixing and staining, worms can be very rigid and difficult to pipette. To circumvent this, make wide-bore pipettes by cutting off their tips.</li>
		<li>Store the slides in a micro-centrifuge rack at 4 &#176;C for up to 24 h if not imaging immediately after staining.</li>
	</ol>
	</li>
	<li>Imaging of ORO-stained worms
	<ol>
		<li>Use a color-capable camera to image ORO-stained worms.</li>
		<li>Use 5X magnification to image several worms in one field of view.</li>
		<li>Switch to 10X magnification for better examination of individual worms.</li>
		<li>Export images in TIF format to avoid data loss due to compression. 
		NOTE: If staining with ORO + DAPI, switch the color-capable camera to one that can capture fluorescence.</li>
	</ol>
	</li>
	<li>Image analysis of ORO-stained worms
	<ol>
		<li>Upload micrograph(s) to ImageJ. In the Plugins pull-down menu, use the Bio-Formats function if the image file is not recognized.</li>
		<li>In the Image pull-down menu, under the Stacks function, use Images-to-Stack to create an image stack when several micrographs are compared.</li>
		<li>Adjust Brightness/Contrast under the same pull-down menu to improve the visibility of lipid droplets.</li>
		<li>Categorize the images according to lipid accumulation throughout the animal&#39;s body or in specific tissues.</li>
		<li>For better evaluation of lipid localization and for the identification of non-specific image artifacts using ImageJ, select the Color function under the Image pull-down menu and click on Stack to RGB to show the signals of each RGB component.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Greenspan, P., Mayer, E. P., Fowler, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nile+red:+a+selective+fluorescent+stain+for+intracellular+lipid+droplets.">Nile red: a selective fluorescent stain for intracellular lipid droplets.</a> J Cell Biol. 100 (3), 965-973 (1985).</li><li>Guo, Y., Cordes, K. R., Farese, R. V., Walther, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+droplets+at+a+glance.">Lipid droplets at a glance.</a> J Cell Sci. 122 (6), 749-752 (2009).</li><li>Horobin, R. W., Kiernan, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Conn's biological stains: a handbook of dyes, stains and fluorochromes for use in biology and medicine. , (2002).</li><li>Hulme, S. 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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+imaging+of+lipid+dynamics+using+Coherent+Anti-stokes+Raman+Scattering+(CARS)+and+Stimulated+Raman+Scattering+(SRS)+microscopy.">Label-free imaging of lipid dynamics using Coherent Anti-stokes Raman Scattering (CARS) and Stimulated Raman Scattering (SRS) microscopy.</a> Curr Opin Genet Dev. 21 (5), 585-590 (2011).</li><li>Pino, E. C., Webster, C. M., Carr, C. E., Soukas, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biochemical+and+high+throughput+microscopic+assessment+of+fat+mass+in+Caenorhabditis+elegans.">Biochemical and high throughput microscopic assessment of fat mass in Caenorhabditis elegans.</a> J Vis Exp. (73), (2013).</li><li>O'Rourke, E. J., Soukas, A. A., Carr, C. 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The authors declare no conflicts of interest.

The rise in obesity and metabolic disease rates makes C. elegans a suitable model to study the mechanisms that regulate fat accumulation in cells and tissues. Recent evidence suggests that the changes in lipid levels are correlated with cellular processes ranging from insulin signaling8, the activation of hormone receptors2, to reproductive output5. Compared to label-free microscopy and chromatography methods, Nile red and oil red O are relatively inexpensive dyes used to stain neutral lipids in the worm consistently and reproducibly10,12,13. The first allows for the quantification of total lipid levels, while the second facilitates the evaluation of lipid distribution among tissues such as the hypodermis, intestine, and the germ line. When used in conjunction, NR and ORO enable the researchers to determine how genotype and environmental changes affect lipid accumulation and where in the worm these changes are occurring.
These dyes, nevertheless, do not directly assess lipid accumulation in the same non-invasive manner as does CARS microscopy12. Therefore, they are prone to errors during the fixation and staining, which may compromise measurement accuracy13. Additionally, these dyes may inadvertently interact with the lipofuscin and intestinal granules, resulting in fluorescence unrelated to intracellular neutral fat stores10,14. Additionally, in cases where lipid levels appear low, NR and ORO staining cannot distinguish if the outcome results from permeability issues or reduced fat accumulation. Moreover, the sensitivity of these stains to light requires special measures during storage and active handling that limit the degradation and photo bleaching. However, if caution is exercised when performing and drawing conclusions from label-based assays, NR and ORO staining are excellent means of forward genetic screens to study lipid metabolism pathways and examine their interactions with other physiological functions. The use of daf-2 and fat-6 worms is recommended to make relative comparisons of increased and decreased lipid accumulation, respectively. The washing and fixating steps for NR and ORO staining are similar. Therefore, both can be performed on the same day. However, only the slides with ORO-stained worms can be stored for imaging later because NR staining is inconsistent after 6 h. During the washing, it is critical to include Triton 100-X not only to enhance the permeability to NR and ORO, but also to avoid the worm loss due to excessive adherence to plasticware. Additionally, the worms should be washed less than 30 min before staining to ensure maximum permeability to each stain. While NR and ORO incubation times may be reduced to 1 h and 30 min, respectively, it is encouraged to follow the time suggested in this article for more consistent staining and imaging results. Prolonged exposure of NR solution to light and failure to filter ORO solution before use will increase background fluorescence during imaging. Carrying out these staining protocols requires 3-4 h in addition to imaging time. Though stain incubation offers at most 2 h of pause between steps, it is strongly recommended that the protocols are carried out with as few interruptions as possible to reduce the many sources of error inherent to all dye-based methods of lipid quantification and examination.

Lipids are essential for life. They are integral components of membranes, act as secondary messengers and signal transducers, and have crucial functions in energy storage. When lipid metabolism is dysregulated, it leads to diseases like obesity and type II diabetes, which are pressing public health concerns9. Caenorhabditis elegans (C. elegans) is an excellent model organism in which to study lipid metabolism because it has a relatively short life cycle, a transparent body, a known cell lineage, and a fully sequenced genome. Primarily a hermaphrodite, C. elegans allows researchers to raise large numbers of isogenic animals in short periods of time to carryout high-throughput forward genetic screens to study a wide array of metabolic genes and pathways4. This approach has revealed a high degree of conservation in 273 C. elegans lipid metabolism genes among humans, mice, rats and drosophila. Furthermore, over 300 lipid genes in C. elegans have human orthologues that are associated with diseases unrelated to metabolic syndrome11. Traditionally, examination of lipid storage in C. elegans has mostly relied on dye-labeled assays, which provide robust information about lipid accumulation. Less common is a description of where lipids localize and measured differences in lipid abundance across tissues. However, recent work has revealed that lipid distribution can be as important as lipid accumulation6.
Lately, studies have begun integrating methods such as high performance liquid chromatography-mass spectrometry (HPLC-MS), gas chromatography-mass spectrometry (GC-MS), and coherent anti-stokes Raman scattering (CARS) microscopy to address the shortcomings of stain-based approaches by directly analyzing the contents of lipid extracts, specific lipid fractions, and lipid deposits, respectively10,11. Moreover, CARS microscopy has revealed that Nile red can only serve as a proxy for fat accumulation when used as a fixative dye, for its use as a vital stain leads to off-target staining of auto-fluorescent organelles10. However, the required technical expertise and costs associated with these chromatography and microscopy methods make their use untenable for many research questions. In this article, we discuss a convenient and reliable method to fixate and stain neutral lipid deposits in C. elegans using Nile red and oil red O to distinguish lipid abundance in whole animals and in specific tissues.
Nile red, 9-diethylamino-5H-benzo[&#945;]phenoxazine-5-one, is a benzophenoxazone dye that readily dissolves in various organic solvents, but is mostly insoluble in water. It is an excellent lysochrome dye used to stain neutral lipids such as triglycerides or cholesterol esters because it features a strong color, solubilizes well in lipids, has negligible interaction with surrounding tissues, and is less soluble in the solvent than in lipids. It has an excitation and emission maxima of 450-500 and 520 nm, respectively1. When Nile red-stained C. elegans is viewed for green fluorescence, discrete lipid bodies can be observed throughout the intestine and other tissues either in clusters or evenly dispersed, depending on the animal&#39;s genotype or experimental treatment7.
Oil red O is a lysochrome, fat-soluble dye used to stain triglycerides and lipoproteins. It is called an azo dye because its chemical structure contains two azo groups attached to three aromatic rings. It is difficult to ionize, which renders it highly soluble in lipids. Its stain color is red and its light absorption maximum is 518 nm 3. C. elegans stained with oil red O show red lipid droplets that stand out against the animal&#39;s transparent body, which facilitates qualitative assessment of lipid distribution among different tissues6.

<table><tbody><tr><td>Imager.M2m Microscope</td><td>Zeiss</td><td>n/a</td><td>Fluorescence microscope</td></tr><tr><td>ERC5s camera</td><td>Axiocam</td><td>n/a</td><td>Color-capable</td></tr><tr><td>MRm camera</td><td>Axiocam</td><td>n/a</td><td>Fluorescence-capable</td></tr><tr><td>Nile red</td><td>Thermo Fisher</td><td>N1142</td><td>Lipid Stain</td></tr><tr><td>Oil red O</td><td>Alfa Aesar</td><td>A12989</td><td>Lipid Stain</td></tr><tr><td>DAPI</td><td>Thermo Fisher</td><td>D1306</td><td>DNA stain</td></tr><tr><td>Isopropyl Alcohol</td><td>BDH</td><td>BDH1133-1LP</td><td>Fixative solution</td></tr><tr><td>0.2 &amp;micro;m seterile syringe filter</td><td>VWR</td><td>28145-477</td><td>Cellulose acetate filter</td></tr><tr><td>Centrifuge 5430</td><td>Eppendorf</td><td>5428000015</td><td>Centrifuge</td></tr><tr><td>Shaker Rotisserie</td><td>Lab Quake</td><td>400110Q</td><td>Shaker</td></tr><tr><td>Tube Rotator</td><td>VWR</td><td>10136-084</td><td>Rotator</td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>7758-11-4</td><td>NGM</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>7778-77-0</td><td>NGM</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Alfa Aesar</td><td>7786-30-3</td><td>NGM</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>10035-04-8</td><td>NGM</td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>7647-14-5</td><td>NGM</td></tr><tr><td>Cholesterol</td><td>Sigma-Aldrich</td><td>57-88-5</td><td>NGM</td></tr><tr><td>Peptone</td><td>BD Biosciences</td><td>211677</td><td>NGM</td></tr><tr><td>Agar</td><td>Teknova</td><td>L9110</td><td>NGM</td></tr><tr><td>LB media</td><td>Sigma-Aldrich</td><td>L3147</td><td>Bacterial growth</td></tr></tbody></table>

quantification of lipid abundance and evaluation of lipid distribution in caenorhabditis elegans by nile red and oil red o staining

Caenorhabditis elegans is an exceptional model organism in which to study lipid metabolism and energy homeostasis. Many of its lipid genes are conserved in humans and are associated with metabolic syndrome or other diseases. Examination of lipid accumulation in this organism can be carried out by fixative dyes or label-free methods. Fixative stains like Nile red and oil red O are inexpensive, reliable ways to quantitatively measure lipid levels and to qualitatively observe lipid distribution across tissues, respectively. Moreover, these stains allow for high-throughput screening of various lipid metabolism genes and pathways. Additionally, their hydrophobic nature facilitates lipid solubility, reduces interaction with surrounding tissues, and prevents dissociation into the solvent. Though these methods are effective at examining general lipid content, they do not provide detailed information about the chemical composition and diversity of lipid deposits. For these purposes, label-free methods such as GC-MS and CARS microscopy are better suited, their costs notwithstanding.

SKN-1 is a bZip, cytoprotective transcription factor that shares homology with mammalian NRF2 and has been shown to mediate fatty acid oxidation. Depending on the glucose concentration in their diet, worms with a constitutively activated skn-1 allele show different lipid levels when stained with Nile red7. Figure 1A-C shows activated skn-1 animals exposed to conditions that lead to increasing lipid levels. NR fluorescence captured using a FITC/GFP channel is prominent along the intestine, but is dimmer in the head, tail, and intestinal lumen. As lipid levels increase, discrete NR-stained particles are more difficult to discern, which may necessitate lower exposure times. Though this method uses fluorescence intensity as a quantitative proxy for lipid accumulation, it is inadequate at distinguishing unspecific staining of cellular structures that are not fat stores and is prone to signal interference from intestinal auto fluorescence. Therefore, alternative label and non-label methods must be used in parallel to unambiguously quantify neutral lipids in the animal10.
Age-dependent Somatic Depletion of Fat (Asdf), is a phenotype that occurs in aged worms whereby animals display decreased somatic cell lipids, while germ cell lipids remain unchanged6. Oil red O staining is not a method that reliably quantifies lipid levels, but is excellent for visualizing lipid localization and is useful for determining fat depletion phenotypes like Asdf in the worm. While it is easy to categorize the worms according to the presence or absence of Asdf, it may be difficult to identify animals with intermediate fat loss (Figure 2A). Compared to non-Asdf animals, which show bright red staining throughout the body with few translucent areas (Figure 2B), Asdf worms exhibit noticeable fat depletion in intestinal cells (Figure 2C). Animals in the process of developing Asdf (Figure 2D) often show translucent spots where most fat loss eventually occurs. Moreover, Asdf worms may still feature bright red staining in the head and tail regions because the phenotype is not defined by complete somatic fat loss, but by extensive fat depletion relative to non-aged (non-Asdf) animals. The use of oil red O staining, thus, allows for qualitative determination of lipid localization and the identification of phenotypes that substantially change fat deposits in the worm.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57352/57352fig1.jpg" /> 
Figure 1: Staining of lipids by Nile red. Nile red staining of activated skn-1 mutants under conditions that elicit increased lipid accumulation (A-C). The gain of function skn-1 strain (lax188) harbors an E237K amino acid substitution that renders SKN-1 constitutively active. L4-stage worms were exposed to 0, 15 and 30 J/m2 UV-light followed by a 12-h recovery period on unseeded NGM plates before NR-staining and imaging. Images shown are representative of the phenotype. Lipid quantification was performed as mentioned in section 1.7. Fire is an ImageJ look up table (LUT) used to create a thermal image that shows fluorescence intensity differences between images. Merge shows FITC/GFP and DIC images combined. Quantification of fluorescence intensity for each image is shown at the bottom of the image montage. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57352/57352fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57352/57352fig2.jpg" /> 
Figure 2: Staining of lipids by oil red O. Oil red O staining of activated skn-1 mutants (lax188) showing the presence or absence of the Asdf phenotype (A-D). A &#38; D show animals with intermediate Asdf phenotypes. B &#38; C display opposite ORO-staining. While B is non-Asdf, C shows substantial somatic fat depletion with concomitant increase in germline fat accumulation, which defines the Asdf phenotype. The worms were stained with ORO followed by imaging 144 hafter L1-synchronized animals were placed on NGM media seeded with OP50 bacteria. Images shown are representative of animals with or without the Asdf phenotype. Inset cartoons are modified from Lynn, et al.6 and represent the absence (B) or presence (C) of somatic fat depletion. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57352/57352fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining at JoVE.com

Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining

Nile red staining of fixed Caenorhabditis elegans is a method for quantitative measurement of neutral lipid deposits, while oil red O staining facilitates qualitative assessment of lipid distribution among tissues.

Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining

Caenorhabditis elegans is an exceptional model organism in which to study lipid metabolism and energy homeostasis. Many of its lipid genes are conserved ...

quantification-lipid-abundance-evaluation-lipid-distribution

Nanyang Technological University

Research

JoVE Journal

Biology

20.2K Views.  University of Southern California. Nile rode kleuring van vaste Caenorhabditis elegans is een methode voor kwantitatieve meting van neutrale lipide deposito's, terwijl kwalitatieve beoordeling van lipide verdeling over weefsels olie rood O kleuring vergemakkelijkt.

Video: Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining

A Simple and Rapid Protocol for Measuring Neutral Lipids in Algal Cells Using Fluorescence

Algae are considered excellent candidates for renewable fuel sources due to their natural lipid storage capabilities. Robust monitoring of algal fermentation processes and screening for new oil-rich strains requires a fast and reliable protocol for determination of intracellular lipid content. Current practices rely largely on gravimetric methods to determine oil content, techniques developed decades ago that are time consuming and require large sample volumes. In this paper, Nile Red, a fluorescent dye that has been used to identify the presence of lipid bodies in numerous types of organisms, is incorporated into a simple, fast, and reliable protocol for measuring the neutral lipid content of Auxenochlorella protothecoides, a green alga. The method uses ethanol, a relatively mild solvent, to permeabilize the cell membrane before staining and a 96 well micro-plate to increase sample capacity during fluorescence intensity measurements. It has been designed with the specific application of monitoring bioprocess performance. Previously dried samples or live samples from a growing culture can be used in the assay.

A simple protocol to determine the neutral lipid content of algal cells using a Nile Red staining procedure is described. This time-saving technique offers an alternative to traditional gravimetric-based lipid quantification protocols. It has been designed for the specific application of monitoring bioprocess performance.

Algae are considered excellent candidates for renewable fuel sources due to their natural lipid storage capabilities. Robust monitoring of algal ...

Chemistry

Assessing Hepatic Steatosis with 3D Fluorescence Lipid Droplet Reconstruction

Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing dysfunction caused by free fatty acids (FAs). The liver, given its critical role in the body's fat metabolism, is persistently threatened by the intracellular accumulation of LDs in the form of both microvesicular and macrovesicular hepatic steatosis. The histologic characterization of LDs is typically based on lipid-soluble diazo dyes, such as Oil Red O (ORO) staining, but a number of disadvantages consistently hamper the use of this analysis with liver specimens. More recently, lipophilic fluorophores 493/503 have become popular for visualizing and locating LDs due to their rapid uptake and accumulation into the neutral lipid droplet core. Even though most applications are well-described in cell cultures, there is less evidence demonstrating the reliable use of lipophilic fluorophore probes as an LD imaging tool in tissue samples. Herein, we propose an optimized boron dipyrromethene (BODIPY) 493/503-based protocol for the evaluation of LDs in liver specimens from an animal model of high-fat diet (HFD)-induced hepatic steatosis. This protocol covers liver sample preparation, tissue sectioning, BODIPY 493/503 staining, image acquisition, and data analysis. We demonstrate an increased number, intensity, area ratio, and diameter of hepatic LDs upon HFD feeding. Using orthogonal projections and 3D reconstructions, it was possible to observe the full content of neutral lipids in the LD core, which appeared as nearly spherical droplets. Moreover, with the fluorophore BODIPY 493/503, we were able to distinguish microvesicles (1 &#181;m &lt; d &#8804; 3 &#181;m), intermediate vesicles (3 &#181;m &lt; d &#8804; 9 &#181;m), and macrovesicles (d &gt; 9 &#181;m), allowing the successful discrimination of microvesicular and macrovesicular steatosis. Overall, this BODIPY 493/503 fluorescence-based protocol is a reliable and simple tool for hepatic LD characterization and may represent a complementary approach to the classical histological protocols.

Author Spotlight: Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment  

Herein, we demonstrate an optimized BODIPY 493/503 fluorescence-based protocol for lipid droplet characterization in liver tissue. Through the use of orthogonal projections and 3D reconstructions, the fluorophore allows for successful discrimination between microvesicular and macrovesicular steatosis and may represent a complementary approach to the classical histological protocols for hepatic steatosis assessment.

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing ...

Biochemistry

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the environment, but also determines body shape and plays a role in motility4-6. Several layers secreted by epidermal cells comprise the cuticle, including an outermost lipid layer7.
Circumferential ridges in the cuticle called annuli pattern the length of the animal and are present during all stages of development8. Alae are longitudinal ridges that are present during specific stages of development, including L1, dauer, and adult stages2,9. Mutations in genes that affect cuticular collagen organization can alter cuticular structure and animal body morphology5,6,10,11. While cuticular imaging using compound microscopy with DIC optics is possible, current methods that highlight cuticular structures include fluorescent transgene expression12, antibody staining13, and electron microscopy1. Labeled wheat germ agglutinin (WGA) has also been used to visualize cuticular glycoproteins, but is limited in resolving finer cuticular structures14. Staining of cuticular surface using fluorescent dye has been observed, but never characterized in detail15. We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. This optimized protocol for DiI staining is a simple, robust method for high resolution fluorescent visualization of annuli, alae, vulva, male tail, and hermaphrodite tail spike in C. elegans.

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. With this optimized protocol, alae and annular cuticular structures are stained by DiI and observed using compound microscopy.

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the ...

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 ...

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 ...

Worm-align and Worm_CP, Two Open-Source Pipelines for Straightening and Quantification of Fluorescence Image Data Obtained from Caenorhabditis elegans

An issue often encountered when acquiring image data from fixed or anesthetized C. elegans is that worms cross and cluster with their neighbors. This problem is aggravated with increasing density of worms and creates challenges for imaging and quantification. We developed a FIJI-based workflow, Worm-align, that can be used to generate single- or multi-channel montages of user-selected, straightened and aligned worms from raw image data of C. elegans. Worm-align is a simple and user-friendly workflow that does not require prior training of either the user or the analysis algorithm. Montages generated with Worm-align can aid the visual inspection of worms, their classification and representation. In addition, the output of Worm-align can be used for subsequent quantification of fluorescence intensity in single worms, either in FIJI directly, or in other image analysis software platforms. We demonstrate this by importing the Worm-align output into Worm_CP, a pipeline that uses the open-source CellProfiler software. CellProfiler&#8217;s flexibility enables the incorporation of additional modules for high-content screening.&#160;As a practical example, we have used the pipeline on two datasets: the first dataset are images of heat shock reporter worms that express green fluorescent protein (GFP) under the control of the promoter of a heat shock inducible gene hsp-70, and the second dataset are images obtained from fixed worms, stained for fat-stores with a fluorescent dye.

Worm-align and Worm_CP, Two Open-Source Pipelines for Straightening and Quantification of Fluorescence Image Data Obtained from Caenorhabditis elegans

Worm-align/Worm_CP is a simple FIJI/CellProfiler workflow that can be used to straighten and align Caenorhabditis elegans samples and to score whole-worm image-based assays without the need for prior training steps. We have applied Worm-align/Worm_CP to the quantification of&#160;heat-shock induced expression in live animals or lipid droplets&#160;in fixed samples.

An issue often encountered when acquiring image data from fixed or anesthetized C. elegans is that worms cross and cluster with their neighbors. This ...

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits obesity and many associated diseases including cardiovascular disorders, type II diabetes, and cancer. To advance the current understanding of lipid metabolic regulation, quantitative methods to precisely measure in vivo lipid storage levels in time and space have become increasingly important and useful. Traditional approaches to analyze lipid storage are semi-quantitative for microscopic assessment or lacking spatio-temporal information for biochemical measurement. Stimulated Raman scattering (SRS) microscopy is a label-free chemical imaging technology that enables rapid and quantitative detection of lipids in live cells with a subcellular resolution. As the contrast is exploited from intrinsic molecular vibrations, SRS microscopy also permits four-dimensional tracking of lipids in live animals. In the last decade, SRS microscopy has been widely used for small molecule imaging in biomedical research and overcome the major limitations of conventional fluorescent staining and lipid extraction methods. In the laboratory, we have combined SRS microscopy with the genetic and biochemical tools available to the powerful model organism, Caenorhabditis elegans, to investigate the distribution and heterogeneity of lipid droplets across different cells and tissues and ultimately to discover novel conserved signaling pathways that modulate lipid metabolism. Here, we present the working principles and the detailed setup of the SRS microscope and provide methods for its use in quantifying lipid storage at distinct developmental timepoints of wild-type and insulin signaling deficient mutant C. elegans.

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows selective, label-free imaging of specific chemical moieties and it has been effectively employed to image lipid molecules in vivo. Here, we provide a brief introduction to the principle of SRS microscopy and describe methods for its use in imaging lipid storage in Caenorhabditis elegans.

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits ...

Lipid Supplementation for Longevity and Gene Transcriptional Analysis in Caenorhabditis elegans

Aging is a complex process characterized by progressive physiological changes resulting from both environmental and genetic contributions. Lipids are crucial in constituting structural components of cell membranes, storing energy, and as signaling molecules. Regulation of lipid metabolism and signaling is essential to activate distinct longevity pathways. The roundworm Caenorhabditis elegans is an excellent and powerful organism to dissect the contribution of lipid metabolism and signaling in longevity regulation. Multiple research studies have described how diet supplementation of specific lipid molecules can extend C. elegans lifespan; however, minor differences in the supplementation conditions can cause reproducibility issues among scientists in different labs. Here, two detailed supplementation methods for C. elegans are reported employing lipid supplementation either with bacteria seeded on plates or bacterial suspension in liquid culture. Also provided herein are the details to perform lifespan assays with lifelong lipid supplementation and qRT-PCR analysis using a whole worm lysate or dissected tissues derived from a few worms. Using a combination of longitudinal studies and transcriptional investigations upon lipid supplementation, the feeding assays provide dependable approaches to dissect how lipids influence longevity and healthy aging. This methodology can also be adapted for various nutritional screening approaches to assess changes in a subset of transcripts using either a small number of dissected tissues or a few animals.

Lipid Supplementation for Longevity and Gene Transcriptional Analysis in Caenorhabditis elegans

The present protocol describes lipid supplementation methods in liquid and on-plate cultures for Caenorhabditis elegans, coupled with longitudinal studies and gene transcriptional analysis from bulk or a few worms and worm tissues.

Aging is a complex process characterized by progressive physiological changes resulting from both environmental and genetic contributions. Lipids are ...

Live Cell Imaging of Lipid Droplet Size and Fusion Using Oil Red O and BODIPY

Evaluation of Lipid Droplet Size and Fusion in Bovine Hepatic Cells

Lipid droplets (LDs) are organelles that play an important role in lipid metabolism and neutral lipid storage in cells. They are associated with a variety of metabolic diseases, such as obesity, fatty liver disease, and diabetes. In hepatic cells, the sizes and numbers of LDs are signs of fatty liver disease. Moreover, the oxidative stress reaction, cell autophagy, and apoptosis are often accompanied by changes in the sizes and numbers of LDs. As a result, the dimensions and quantity of LDs are the basis of the current research regarding the mechanism of LD biogenesis. Here, in fatty acid-induced bovine hepatic cells, we describe how to use oil red O to stain LDs and to investigate the sizes and numbers of LDs. The size distribution of LDs is statistically analyzed. The process of small LDs fusing into large LDs is also observed by a live cell imaging system. The current work provides a way to directly observe the size change trend of LDs under different physiological conditions.

Author Spotlight: Evaluation of Lipid Droplet Size and Fusion in Bovine Hepatic Cells  

The present protocol describes how to use oil red O to dye lipid droplets (LDs), calculate the size and number of LDs in a fatty acid-induced fatty hepatocyte model, and use BODIPY 493/503 to observe the process of small LDs fusing into large LDs by live cell imaging.

Lipid droplets (LDs) are organelles that play an important role in lipid metabolism and neutral lipid storage in cells. They are associated with a variety ...

Nile Red Staining of C. elegans: A Method to Visualize Lipid Droplets in Fixed Animals

Source: Escorcia, W., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/57352/quantification-lipid-abundance-evaluation-lipid-distribution">Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining</a>. J. Vis. Exp. (2018).
This video describes a protocol to stain lipids in whole, fixed worms using Nile Red dye.&#160; To specifically view the neutral lipids in the core of lipid droplets, image the green emission spectra of the stained C. elegans.

Source: Escorcia, W., et al. Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O ...

Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining

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