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Encyclopedia of Experiments

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


This video describes a protocol to stain lipids in whole, fixed worms using Nile Red dye.  To specifically view the neutral lipids in the core of lipid droplets, image the green emission spectra of the stained C. elegans.


This protocol is an excerpt from Escorcia et al, Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining, J. Vis. Exp. (2018).

1. Nile Red (NR) Staining of Lipids

  1. Preparation of 5 mg/mL NR stock solution
    1. In a 500 mL bottle, add 100 mg of NR powder to 200 mL of 100% acetone.
    2. Cover the bottle with aluminum foil to avoid any exposure to light.
    3. Before use, stir the solution for 2 h in the dark.
    4. 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.
  2. Preparation of NR working solution
    1. For every 1 mL of 40% isopropanol (v/v), add 6 µL of NR stock solution.
    2. Prepare 600 µL 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.
  3. Preparation of worms for NR lipid staining
    1. Grow worms to early L4 stage at 20 °C on nematode growth medium (NGM) seeded with late-log OP50 E. coli.
    2. 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.
    3. Centrifuge the worms at 560 x g for 1 min. Remove the supernatant and repeat this step until E. coli is cleared from suspension.
    4. Add 100 µL of 40% isopropanol to the worm pellet and incubate it at room temperature for 3 min.
    5. 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.
  4. Lipid staining with NR
    1. In the dark, add 600 µL of NR working solution to each sample. Invert the tubes three times and fully mix the worms in NR solution.
    2. Rotate the sample in the dark at room temperature for 2 h.
    3. Following the incubation, centrifuge the worms at 560 x g for 1 min and remove the supernatant.
    4. Add 600 µL of PBST and incubate the samples in the dark for 30 min to remove excess NR stain.
    5. Centrifuge the samples at 560 x g for 1 min and remove all but approximately 50 µL of supernatant.
      NOTE: NR incubation does not require agitation. Settling of worms is common during this step.
  5. Preparation of slides for microscope imaging
    1. Resuspend the worm pellet in remaining supernatant.
    2. Place 5 µL of worm suspension on a microscope slide and put a coverslip on carefully to avoid trapping any air bubbles.
    3. Seal the coverslip with nail polish before imaging worms.
    4. 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.
  6. Imaging of NR-stained worms
    1. Image the worms at 5X magnification to capture several animals per field of view.
    2. Switch to 10X magnification for better quantification of individual worms.
    3. Use FITC/GFP channel to image NR-stained worms and try different exposure times to determine the optimal conditions for quantification.
    4. 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.

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Name Company Catalog Number Comments
Imager.M2m Microscope Zeiss n/a Fluorescence microscope
ERC5s camera Axiocam n/a Color-capable
MRm camera Axiocam n/a Fluorescence-capable
Nile red Thermo Fisher N1142 Lipid Stain
Isopropyl Alcohol BDH BDH1133-1LP Fixative solution
Centrifuge 5430 Eppendorf 5428000015 Centrifuge
Tube Rotator VWR 10136-084 Rotator


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