If you share the calculation and/or protocol, it will be a lot easier to give you advice.

Katie A. S. Burnette,

Hi, how are you? Thanks for your reply! I haven’t started yet because I don’t know the process. I read that some people use 80% acetone and incubate the seedlings overnight, while others use ethanol without grinding the seedlings for 48 hours and then simply measure the chlorophyll.

Measuring the chlorophyll (as well as carotenoid) contents is a pretty simple procedure, though still quite time-consuming and with a lot of small details to mind about.

The general procedure is easy enough: weight some tissue (most likely leaves), extract the chlorophyll from the tissue with a solvent, then measure the absorbance at 2 specific wavelengths (3 if you also do carotenoids), and use the formulas already established in literature to calculate the chlorophyll (and carotenoid) contents.

You can use several different solvents. Wellburn had devised multiple formulas used for different solvents in his excellent 1994 paper. I recommend you check it out:

Article The Spectral Determination of Chlorophylls a and b, as well ...

The formula will give you the chlorophyll contents in the extract you measure (as µg/mL). Multiply by the total volume of solvent (mL) you used to extract, then divide by the sample weight (mg) you took, and you get the chlorophyll contents per mg fresh weight of sample. It's recommended that you calculate the contents per dry weight, but it's up to you. If you do dry weight, take a few tissue samples from the same treatment and dry them, calculating the fresh/dry weight ratio, and convert.

Now for the technical aspect, there are a few things you must mind.

First is that chlorophyll is very easily degraded under light once extracted from the cells. Therefore, you must keep it relatively dark during and after extracting, and don't keep it too long.

Second is the ratio between the amount of tissue sample and the volume of extracting solvent. Spectrophotometers typically measure most accurately at absorbance ~ 1. Any measurement above 2 or below 0.2 is not reliable. If you use too high or too low sample/solvent ratio during extraction, you'd end up with a very concentrated or very diluted extract that's not good for measuring. Chlorophyll contents can differ wildly between different species and even between different treatment conditions. Therefore you need to test beforehand to find suitable sample/solvent ratio for each treatment.

Third is, the solvents used for extracting are usually very volatile. This can cause very significant errors if you don't seal the extract tight enough, keeping the extract for too long, using too small a volume while measuring absorbance (like with a microplate) or working too slow. These can cause the solvent to evaporate, changing the volume of the extract and causing error in calculation.

Therefore, you need to use good tubes with tight seals to do the extraction, and don't extract for too long. The protocol I learned years ago is to slice the leaves into thin slices and extract in the solvent for 3 days for all the chlorophyll to come out. But later on, I found that if you grind the leaves relatively small, extracting for 1 day is enough, and it gives better accuracy thanks to less solvent loss. Although, you can always recalibrate the volume right before measuring absorbance by adding solvent, and if you can do so easily and accurately, it's strongly recommended.

Measuring the absorbance should also be done in relative darkness, preferably with a 3 mL cuvette instead of 1 mL cuvette to minimize volume change error due to solvent evaporation. Don't use microplate as the 300 µL extract would disappear like crazy. Work fast. Rinse the cuvette with solvent when you change treatment or replication, and dry the cuvette with tissue paper thoroughly before putting new extract in.