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. E., Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemistry+and+the+worm:+Caenorhabditis+elegans+as+a+platform+for+integrating+chemical+and+biological+research.">Chemistry and the worm: Caenorhabditis elegans as a platform for integrating chemical and biological research.</a> Angew Chem Int Edit. 50 (21), 4774-4807 (2011).</li><li>Khanna, A., Johnson, D. L., Curran, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiological+roles+for+mafr-1+in+reproduction+and+lipid+homeostasis.">Physiological roles for mafr-1 in reproduction and lipid homeostasis.</a> Cell Rep. 9 (6), 2180-2191 (2014).</li><li>Lynn, D. A., Dalton, H. M., Sowa, J. N., Wang, M. C., Soukas, A. A., Curran, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Omega-3+and-6+fatty+acids+allocate+somatic+and+germline+lipids+to+ensure+fitness+during+nutrient+and+oxidative+stress+in+Caenorhabditis+elegans.">Omega-3 and-6 fatty acids allocate somatic and germline lipids to ensure fitness during nutrient and oxidative stress in Caenorhabditis elegans.</a> P Natl Acad Sci USA. 112 (50), 15378-15383 (2015).</li><li>Pang, S., Lynn, D. A., Lo, J. Y., Paek, J., Curran, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SKN-1+and+Nrf2+couples+proline+catabolism+with+lipid+metabolism+during+nutrient+deprivation.">SKN-1 and Nrf2 couples proline catabolism with lipid metabolism during nutrient deprivation.</a> Nat Commun. 5, (2014).</li><li>Saltiel, A. R., Kahn, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insulin+signalling+and+the+regulation+of+glucose+and+lipid+metabolism.">Insulin signalling and the regulation of glucose and lipid metabolism.</a> Nature. 414 (6865), 799-806 (2001).</li><li>Witting, M., Schmitt-Kopplin, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Caenorhabditis+elegans+lipidome:+A+primer+for+lipid+analysis+in+Caenorhabditis+elegans.">The Caenorhabditis elegans lipidome: A primer for lipid analysis in Caenorhabditis elegans.</a> Arch Biochem Biophys. 589, 27-37 (2016).</li><li>Yen, K., Le, T. T., Bansal, A., Narasimhan, S. D., Cheng, J. X., Tissenbaum, 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=A+comparative+study+of+fat+storage+quantitation+in+nematode+Caenorhabditis+elegans+using+label+and+label-free+methods.">A comparative study of fat storage quantitation in nematode Caenorhabditis elegans using label and label-free methods.</a> PloS One. 5 (9), e12810 (2010).</li><li>Zhang, Y., Zou, X., Ding, Y., Wang, H., Wu, X., Liang, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+genomics+and+functional+study+of+lipid+metabolic+genes+in+Caenorhabditis+elegans.">Comparative genomics and functional study of lipid metabolic genes in Caenorhabditis elegans.</a> BMC Genomics. 14 (1), 164 (2013).</li><li>Folick, A., Min, W., Wang, M. 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. E., Ruvkun, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=C.+elegans+major+fats+are+stored+in+vesicles+distinct+from+lysosome-related+organelles.">C. elegans major fats are stored in vesicles distinct from lysosome-related organelles.</a> Cell Metab. 10 (5), 430-435 (2009).</li></ol>

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 &#181;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>K2HPO4</td>
			<td>Sigma-Aldrich</td>
			<td>7758-11-4</td>
			<td>NGM</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Sigma-Aldrich</td>
			<td>7778-77-0</td>
			<td>NGM</td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>Alfa Aesar</td>
			<td>7786-30-3</td>
			<td>NGM</td>
		</tr>
		<tr>
			<td>CaCl2</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/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/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

20.0K Views.  University of Southern California. 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.

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

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and hermaphrodites. In the earlier phase of gametogenesis, the germ cells of hermaphrodites differentiate into limited number of sperm - around 300 - and are stored in a small 'bag' called the spermatheca. Later on, hermaphrodites continually produce oocytes1. In contrast, males produce exclusively sperm throughout their adulthood. The males produce so much sperm that it accounts for &gt;50% of the total cells in a typical adult worm2. Therefore, isolating sperm from males is easier than from that of hermaphrodites.
Only a small proportion of males are naturally generated due to spontaneous non-disjunction of X chromosome3. Crossing hermaphrodites with males or more conveniently, the introduction of mutations to give rise to Him (High Incidence of Males) phenotype are some of strategies through which one can enrich the male population3.
Males can be easily distinguished from hermaphrodites by observing the tail morphology4. Hermaphrodite's tail is pointed, whereas male tail is rounded with mating structures.
Cutting the tail releases vast number of spermatids stored inside the male reproductive tract. Dissection is performed under a stereo microscope using 27 gauge needles. Since spermatids are not physically connected with any other cells, hydraulic pressure expels internal contents of male body, including spermatids2.
Males are directly dissected on a small drop of 'Sperm Medium'. Spermatids are sensitive to alteration in the pH. Hence, HEPES, a compound with good buffering capacity is used in sperm media. Glucose and other salts present in sperm media help maintain osmotic pressure to maintain the integrity of sperm.
Post-meiotic differentiation of spermatids into spermatozoa is termed spermiogenesis or sperm activation. Shakes5, and Nelson6 previously showed that round spermatids can be induced to differentiate into spermatozoa by adding various activating compounds including Pronase E. Here we demonstrate in vitro spermiogenesis of C. elegans spermatids using Pronase E.
Successful spermiogenesis is pre-requisite for fertility and hence the mutants defective in spermiogenesis are sterile. Hitherto several mutants have been shown to be defective specifically in spermiogenesis process7. Abnormality found during in vitro activation of novel Spe (Spermatogenesis defective) mutants would help us discover additional players participating in this event.

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

A protocol for isolating and activating spermatids from male C. elegans is described here. Cutting the posterior end of male releases spermatids. The spermatids can be activated by addition of protease.

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has since been used extensively as a model organism 1. C. elegans possesses key attributes such as simplicity, transparency and short life cycle that have made it a suitable experimental system for fundamental biological studies for many years 2. Discoveries in this nematode have broad implications because many cellular and molecular processes that control animal development are evolutionary conserved 3.
C. elegans life cycle goes through an embryonic stage and four larval stages before animals reach adulthood. Development can take 2 to 4 days depending on the temperature. In each of the stages several characteristic traits can be observed. The knowledge of its complete cell lineage 4,5 together with the deep annotation of its genome turn this nematode into a great model in fields as diverse as the neurobiology 6, aging 7,8, stem cell biology 9 and germ line biology 10. 
An additional feature that makes C. elegans an attractive model to work with is the possibility of obtaining populations of worms synchronized at a specific stage through a relatively easy protocol. The ease of maintaining and propagating this nematode added to the possibility of synchronization provide a powerful tool to obtain large amounts of worms, which can be used for a wide variety of small or high-throughput experiments such as RNAi screens, microarrays, massive sequencing, immunoblot or in situ hybridization, among others. 
Because of its transparency, C. elegans structures can be distinguished under the microscope using Differential Interference Contrast microscopy, also known as Nomarski microscopy. The use of a fluorescent DNA binder, DAPI (4',6-diamidino-2-phenylindole), for instance, can lead to the specific identification and localization of individual cells, as well as subcellular structures/defects associated to them.

Basic Caenorhabditis elegans Methods: Synchronization and Observation

The easiness of maintaining and propagating the nematode C. elegans make it a nice model organism to work with. The possibility of synchronizing worms allows the work with a significant amount of subjects at the same developmental stage, what facilitates the study of one particular process in many animals.

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has ...

Prostaglandin Extraction and Analysis in Caenorhabditis elegans

Caenorhabditis elegans is emerging as a powerful animal model to study the biology of lipids1-9. Prostaglandins are an important class of eicosanoids, which are lipid signals derived from polyunsaturated fatty acids (PUFAs)10-14. These signalling molecules are difficult to study because of their low abundance and reactive nature. The characteristic feature of prostaglandins is a cyclopentane ring structure located within the fatty acid backbone. In mammals, prostaglandins can be formed through cyclooxygenase enzyme-dependent and -independent pathways10,15. C. elegans synthesizes a wide array of prostaglandins independent of cyclooxygenases6,16,17. A large class of F-series prostaglandins has been identified, but the study of eicosanoids is at an early stage with ample room for new discoveries. Here we describe a procedure for extracting and analyzing prostaglandins and other eicosanoids. Charged lipids are extracted from mass worm cultures using a liquid-liquid extraction technique and analyzed by liquid chromatography coupled to electrospray ionization tandem mass spectrometry (LC-ESI-MS/MS). The inclusion of deuterated analogs of prostaglandins, such as PGF2 &alpha;-d4 as an internal standard is recommended for quantitative analysis. Multiple reaction monitoring or MRM can be used to quantify and compare specific prostaglandin types between wild-type and mutant animals. Collision-induced decomposition or MS/MS can be used to obtain information on important structural features. Liquid chromatography mass spectrometry (LC-MS) survey scans of a selected mass range, such as m/z 315-360 can be used to evaluate global changes in prostaglandin levels. We provide examples of all three analyses. These methods will provide researchers with a toolset for discovering novel eicosanoids and delineating their metabolic pathways.

Prostaglandin Extraction and Analysis in Caenorhabditis elegans

In this paper, we describe an optimized procedure for extracting and analyzing prostaglandins and other eicosanoids from C. elegans using LC-MS/MS.

Caenorhabditis elegans is emerging as a powerful animal model to study the biology of lipids1-9. Prostaglandins are an important class of eicosanoids, ...

Measurement of Energy Metabolism in Explanted Retinal Tissue Using Extracellular Flux Analysis

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while maintaining transparency of the visual axis. Perturbations to this delicate balance cause blinding illnesses, such as diabetic retinopathy. Therefore, the understanding of energy metabolism changes in the retina during disease is imperative to the development of rational therapies for various causes of vison loss. The recent advent of commercially-available extracellular flux analyzers has made the study of retinal energy metabolism more accessible. This protocol describes the use of such an analyzer to measure contributions to retinal energy supply through its two principle arms - oxidative phosphorylation and glycolysis - by quantifying changes in oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) as proxies for these pathways. This technique is readily performed in explanted retinal tissue, facilitating assessment of responses to multiple pharmacologic agents in a single experiment. Metabolic signatures in retinas from animals lacking rod photoreceptor signaling are compared to wild-type controls using this method. A major limitation in this technique is the lack of ability to discriminate between light-adapted and dark-adapted energy utilization, an important physiologic consideration in retinal tissue.

This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer.

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while ...

Experimental Approach to Examine Leptin Signaling in the Carotid Bodies and its Effects on Control of Breathing

An adipocyte-produced hormone leptin is a potent respiratory stimulant, which may play an important role in defending respiratory function in obesity. The carotid bodies (CB), a key organ of peripheral hypoxic sensitivity, express the long functional isoform of leptin receptor (LepRb) but the role of leptin signaling in control of breathing has not been fully elucidated. We examined the hypoxic ventilatory response (HVR) (1) in C57BL/6J mice before and after leptin infusion at baseline and after CB denervation; (2) in LepRb-deficient obese db/db mice at baseline and after LepRb overexpression in CBs. In C57BL/6J mice, leptin increased HVR and effects of leptin on HVR were abolished by CB denervation. In db/db mice, LepRb expression in CB augmented the HVR. Therefore, we conclude that leptin acts in CB to augment responses to hypoxia.

Our study focuses on the effects of leptin signaling in carotid body (CB) on the hypoxic ventilatory response (HVR). We performed &#39;loss of function&#39; experiments measuring the effect of leptin on HVR after CB denervation and &#39;gain of function&#39; experiments measuring HVR after overexpression of the leptin receptor in CB.

An adipocyte-produced hormone leptin is a potent respiratory stimulant, which may play an important role in defending respiratory function in obesity. The ...

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 ...

Isolation and Characterization of Adult Cardiac Fibroblasts and Myofibroblasts

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that mitigate fibrosis. However, fibroblast research has been hindered due to the lack of universally acceptable fibroblast markers to identify quiescent as well as activated fibroblasts. Fibroblasts are a heterogenous cell population, making them difficult to isolate and characterize. The presented protocol describes three different methods to enrich fibroblasts and myofibroblasts from uninjured and injured mouse hearts. Using a standard and reliable protocol to isolate fibroblasts will enable the study of their roles in homeostasis as well as fibrosis modulation.

Obtaining a pure population of fibroblasts is crucial to studying their role in wound repair and fibrosis. Described here is a detailed method to isolate fibroblasts and myofibroblasts from uninjured and injured mouse hearts followed by characterization of their purity and functionality by immunofluorescence, RTPCR, fluorescence-assisted cell sorting, and collagen gel contraction.

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that ...

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 ...