Gravimetric Total Lipid Quantification Protocol

Measuring coral total lipids

Protocol for lipid extraction and quantification in coral samples. Developed by Ariana Huffmyer and Chris Wall based on protocols modified from Grottoli et al.

This protocol describes methods for analysis of 20 samples per batch using 500 µL of homogenized tissue slurry or homogenized larval/gamete samples. Note that preliminary analysis of this protocol will be required to optimize concentrations of larval samples, as this protocol was primarily developed for slurry of adult coral tissues.

I followed this protocol to quantify coral host total lipids for Pocillopora acuta (Pacu), Montipora capitata (Mcap) and Porites compressa (Pcom) for the ENCORE Hawaii TPC project.

Coral homogenate prep can be found in my previous post.

Protocol considerations and notes

Lyophilization is utilized in this protocol to increase the quality of lipid extraction, but this step is not required and instead liquid slurry or homogenized larvae could be used without lyophilization if not available.

Throughout this protocol, the term “pre-burned” indicates burning in a muffle furnace at 450°C for 4-6 h to remove organic products.

Careful pipetting and reducing organic contamination is crucial for this protocol. Wear gloves, work in a fume hood, and do not rush through this protocol.

Materials and Supplies

Materials

  • Glass scintillation vials: 20 mL vials recommended. 4 vials required per sample (two biological replicates with two vials required for each replicate in this protocol)
  • Small (~20-40mm diameter) aluminum pans. 1 per sample. These should be large enough to hold filters (described below).
  • Large (~40-80mm diameter) aluminum pans. 1 per sample. These should be large enough to hold liquid volume that will be produced in this protocol. It is recommended to have multiple sizes available so that the pan size can be optimized for your specific sample volume.
  • Culture vial, funnel, and filter apparatus with vacuum pump
  • Fiberglass (GF/F) filters: 20 mm recommended for this protocol due to small sample volumes, but this depends on the filter apparatus used. The filter size should be the recommended size on respective filter apparatus. These filters MUST be GF/F.
  • Pencils for labeling
  • P500 and P1000 pipette
  • Analytical balance
  • Muffle furnace (450°C)
  • Drying oven (60°C)
  • Lyophilizer (if available, optional but recommended)
  • Tweezers or forceps

Solvents

  • 100% chloroform
  • 100% methanol
  • 2:1 chloroform:methanol solution
  • 88% KCl solution (8.8g KCl per 1 L DI/MilliQ water)

Preparation

  1. Pre-burn supplies required for this protocol: small aluminum pans, large aluminum pans, GF/F filters, culture vials, and glass scintillation vials. Only burn glass, GF/F filters, and aluminum pans. Do NOT put any plastic or combustable material in the furnace. Burn these supplies at 450°C for 4-6 h to remove all organic material prior to this protocol.

  2. After materials are pre-burned and cooled, aluminum pans, and vials should be labeled with a PENCIL. Marker will be removed during the subsequent burning process. Label glass scintillation vials with permanent marker on both the vial and the lid. These will not be burned during the protocol.

  3. Using an analytical balance, weigh all pans and record on data sheet. This is critical as this weight will be used to determine quantity of lipids in this protocol.

Aliquot and Lyophilize Samples

  1. Thaw samples (kept at -80°C or -20°C) if required.

  2. Aliquot two 500 µL replicates of each sample into two 20 mL glass scintillation vials. If necessary, homogenize samples. Freeze samples at -80°C if lyophilizing, or proceed with thawed samples lyophilization is not used.

  3. If lyophilization is available and desired, turn on lyophilizer and ensure pressure and temperature reach targets as specified (-80°C and <1.0 atm pressure).

  4. Arrange vials with frozen sample in the lyophilizer beakers. If samples begin to thaw at any point, it is critical to re-freeze at -80°C to ensure proper lyophilization.

  5. Lyophilize samples for ~8 h until water fraction is completely removed (samples will appear to be a white powder or film). Ensure there is no leakage once lyophilization begins, if this happens, remove and refreeze samples.

  6. Store lyophilized samples at -80°C.

Lipid Extraction

  1. Remove samples from freezer and add 12 mL of 2:1 chloroform:methanol solution to each vial and gently swirl. It is recommended to only process <20 samples per batch (day) and to start with a small number of samples until comfortable with this protocol.

This volume was optimized (12 mL of solvent for 500 µL of original sample volume) based on conducted trials to ensure saturation and full lipid extraction. This volume should be optimized for either smaller or larger sample volumes.

  1. Allow to sit in a dark fridge for ~3 hours to allow for lipid extraction.

  2. Set up filter apparatus with GF/F filters.

  3. Remove samples from fridge and pour the extract through the filter apparatus (with vacuum pump) such that particulates are captured on the GF/F filter and the liquid fraction passes through the filter to be captured by a culture vial.

  4. Rinse any remaining material in the glass scintillation vial with 2:1 chloroform:methanol solvent until all material is removed and rinsed down to be captured by the filter.

  5. After pouring, use solvent to rinse down the sides of the filter apparatus to capture all lipids.

  6. Turn off the vacuum and slowly remove the filter stand. Using tweezers/forceps, gently remove the GF/F filter and place in a pre-burned, pre-labeled small aluminum pan. This pan will later be dried and burned (described below) to generate Ash Free Dry Weight for normalization.

  7. Pour all filtered liquid extract in the culture vial back into the original glass scintillation vial.

  8. Add 1 mL of 2:1 chloroform:methanol into the culture vial to rinse all material back into the glass scintillation vial.

  9. Set aside the filtered lipid extract in glass scintillation vials and the filters for each sample. Proceed with filtering all samples.

Lipid Extraction

  1. Add 6 mL KCl to the filtered lipid extract in the glass scintillation vial and GENTLY invert 4-5 times. DO NOT vortex or mix vigorously. You are looking for gentle separation of liquid phases. Allow vials to sit for 20-30 minutes to allow for separation. After separation, the vial will now contain a lower phase (yellow color) that contains lipids and an upper phase (white/cream color) that contains water soluble products.

  2. Remove the lower phase (containing lipids) into a new, labeled, pre-burned glass scintillation vial using a pipette. It is recommended to gently angle the vial and carefully pipette out as much of the lower phase as possible while avoiding taking any of the upper phase. We are aiming for complete separation of these phases to purify the lipids.

  3. Add 1 mL of 100% chloroform to the upper phase, invert, and allow to separate for 10-15 minutes. Again remove the lower phase with a pipette and add to the new vial with the lipid phase.

  4. Rinse the lower phase with 6 mL of KCl again. Gently invert and allow to separate for 10-15 minutes.

  5. Now remove the separated lower phase with a pipette and directly transfer into 1 large aluminum pan (ensure all pans are labeled with pencil and pre-weighed). This fraction is now your purified lipid!

  6. Squirt a little methanol in the pan with this purified lipid to remove any remaining water products. This will clear up cloudy liquid such that it is a clear, yellow color.

  7. Proceed with this purification for all samples.

Drying and Burning

  1. Allow lipid fraction in pans AND filters in pans to dry overnight (~ 8 h) at 60-65°C. It is recommended to do this step overnight so that burning can happen the next day. Be careful transferring liquid in pans - be careful not to spill.

  2. After drying, record the dry weight of lipid pans and filter pans in the data sheet.

  3. After weights are recorded burn the filter pan (NOT the lipid pan) for 6-8 h at 450°C in a muffle furnace. The burning will allow us to calculate the organic material on the filter. The lipid pan does NOT need to be burned - we know that fraction is purified lipids and therefore there is no need to burn.

  4. After cooling, record the burned weight of filter pans.

Calculations

Calculate mass of organic material on filter

(dry weight of pan + filter) - (burned weight of pan + filter) = organic material on filter (g)

Calculate mass of lipids

(dry weight of pan + lipids) - (pre-weight of lipid pan) = lipids (g)

Calculate AFDW that will be used for normalization for lipids

AFDW = (g lipids + g organic material on filter) / original sample volume = AFDW in g/mL

This can be used to multiply to total slurry volume to give AFDW per sample, which can later be normalized to surface area or used for other calculations.

Calculate percent lipids (lipids normalized to AFDW)

(g lipids) / (g lipids + g organic material on filter)

This provides the percent of lipids in total organic biomass. This typically ranges from 20-35% in coral adults.

Calculate total lipids in sample

(g lipids) / sample volume (0.5 mL) = g lipids per mL

This can be multiplied by total slurry volume to give total lipids in a coral sample and later normalized to surface area or other responses.

Suggested data sheet format

Columns for a data sheet that are recommended are as follows:

Date
Sample ID
Volume sample
Filter pan pre-weight (small pan)
Lipid pan pre-weight (large pan)
Time drying starts and ends
Dry weight of filter + pan
Dry weight of lipid + pan
Time burning starts and ends
Burned weight of filter + pan

Written on May 19, 2025