Three major classes of photosynthetic pigments occur among the algae: chlorophylls, carotenoids (carotenes and xanthophylls) and phycobilins. The pigments are characteristic of certain algal groups as indicated below. Chlorophylls and carotenes are generally fat soluble molecules and can be extracted from thylakoid membranes with organic solvents such as acetone, methanol or DMSO. The phycobilins and peridinin, in contrast, are water soluble and can be extracted from algal tissues after the organic solvent extraction of chlorophyll in those tissues.

The rationale behind the extraction techniques is to disrupt cell integrity as much as possible, thereby removing pigment molecules from intrinsic membrane proteins. Freezing the tissue with liquid nitrogen, and grinding the still frozen tissue in with a mortar and pestle or blender, overcomes some of the problems of working with material that produces large amounts of viscous polysaccharides. “Freeze-thawing” tissue also breaks down cellular membranes, but may liberate more polysaccharides. Finely ground tissue can be then homogenized in organic solvent to further disrupt cellular membranes, and to liberate pigment molecules from the light harvesting pigment protein complexes.
Once the pigments are extracted into appropriate solvents they can be separated chromatographically by TLC or HPLC for spectral analysis and identification. Pigment concentrations in hydrocarbon solvents can be estimated; however, these formulas are predictive, and may overestimate some pigment concentrations (see Seely et al. 1972 for the development of equations). Uncoupling pigments from the pigment binding proteins can change the absorption patterns of the pigments, resulting in shifts in maxima from 10 to 50 nm, when compared with spectra measured for intact tissues.


Table 1:Pigment composition of several algal groups (after Dring 1982):




 Chlorophyta Green algae chlorophyll b
Charophyta Charophytes chlorophyll b
Euglenophyta Euglenoids chlorophyll b
Phaeophyta Brown algae chlorophyll c1 + c2, fucoxanthin
Chrysophyta Yellow-brown or golden-brown algae chlorophyll c1 + c2, fucoxanthin
Pyrrhophyta Dinoflagellates chlorophyll c2, peridinin
Cryptophyta Cryptomonads chlorophyll c2, phycobilins
Rhodophyta Red algae phycoerythrin, phycocyanin
Cyanophyta Blue-green algae phycocyanin, phycoerythrin




Pigments can be extracted from seaweeds by a variety of techniques though often, it is only through trial and error that an effective technique for a given species is found. Algae which produce large quantities of polysaccharide can be very difficult to work with. Choose your plants carefully!
It is important to note that light, heat, extremes of pH, and oxygen cause the destruction of pigment extracts. The extracts should be kept cold, wrapped in foil, and worked with in the lowest light possible throughout the procedure.

AAcetone Extraction

This technique can be used for the green, brown, and red algae as well as the seagrasses. Some of the greens and seagrasses may be extractable without grinding in liquid nitrogen; for brown and red algae more extreme measures may be necessary.
Note: Field collected tissue should be cleaned of epiphytes prior to the extraction.

  1. Grind approximately 2.0 g (fresh weight) of blotted tissue in a chilled mortar with liquid nitrogen. The mortar and pestle can be placed in the freezer prior to use, and chilled even further by adding a small amount of liquid nitrogen to it prior to adding the tissue. It may help to chop tough blades into workable pieces with a sharp razor blade before grinding.
  2. Quickly transfer powdered tissues to a ground-glass homogenizer. Add 2 to 3 ml of ice-cold 100% acetone. Grind over ice until the remaining debris is colorless.
  3. Transfer pigmented solution to centrifuge tube and spin at 1400 g for 2 min. Decant supernatant.NOTE: If there is color still remaining in the pellet, repeat steps 2 and 3 until the pellet is colorless.(Samples can be stored in the dark at 4o C at this point for a limited period of time)

B. Thin Layer Chromatography

When handling the silica gel plates, care should be taken not to touch the face of the plate or damage the gel along the edges.

  1. On a silica gel plate, draw a light pencil line above-the solvent level in the developing tank approximately 2 cm in width and approximately 2 cm from the bottom.
  2. Draw sample into capillary tube and carefully spot the pigment extract in a solid line along the pencil line until a thin, dark line appears. Each layer should be dry before the next is added; this process can be made more rapid by drying the TLC plate with a gentle current of nitrogen gas. Also, if your extract is dilute, it can be concentrated by passing nitrogen over the solution for 20-30 min. (The TLC plate can be wrapped in foil and stored at 4o for a limited period of time)
  3. Place a small of amount of petroleum ether:acetone (7:3 v:v) into the bottom of the developing tank. Dip filter paper in solvent and put it on sides of tank to saturate the atmosphere with solvent. Note: this should be done well in advance to the introduction of TLC plates into your development tank.
  4. Develop the your TLC plate in a tank containing petroleum ether:acetone (7:3 v:v) for approximately 20-30 min, or however long is necessary to move the solvent front near, but not off, the top of your plate.
  5. Mark the solvent front as soon as the strip is removed from the tank.
  6. Mark the location of the pigment bands and measure the distance from the origin to the pigments as well as the distance to the solvent front. The Rf values for the individual pigments can be determined from: 
    Rf = (distance moved by the pigment)/(distance moved by the solvent)
  7. Slight differences in the dryness of the plates and in the polarity of solvent system will affect the Rf values. To aid in an approximate identification of the different pigmented bands, the following values are reasonable: 


    Table 2: Rf values



     Chlorophyll a  

    Chlorophyll b  

    Chlorophyll c  








  8. After measuring the Rf values, scrape the pigments off of the plate with a spatula or razor blade. Collect the residue and transfer to an Eppendorf tube. Resuspend in the appropriate solvent (acetone for chl and ethanol for carotenoids). The amount of the solvent will depend upon the amount of pigment in the chromatographic band.
  9. Centrifuge for 1-2 min and decant the supernatant into the appropriate cuvette.
  10. Determine the absorption spectrum for that pigment and note the absorption maxima (wavelength maxima for several pigments are listed below).
  11. The absorption data and the chromatographic data can be used together to identify specific pigments for each alga.

    Table 3: Wavelength Maxima for Pigments in Various Solvents




    beta carotene  


    446, 474



    445, 471, 503


    chlorophyll a  

    428.5, 660.5
    diethyl ether
    chlorophyll b  

    diethyl ether
    chlorophyll c1  

    100 % acetone
    chlorophyll c2  

    90% acetone
    chlorophyll c2  

    100% acetone
    chlorophyll c2  

    90% acetone
    chlorophyll c  

    447, 533 or 449, 635
    90% acetone


    Jeffrey, S.A. 1972. Biochem. Biophys. Acta. 279:15-33.
    Jeffrey, S.A. and G. Hymphrey. 1975. Biochem. Physiol. Pflanzen. 167:191-194.

C. Determination of pigment concentration in organic solvents:

From Jeffery and Humphrey (1975).

1. Higher plants and algae having chlorophyll a and b.

In 80% acetone:

Total chlorophyll (a and b) (mg/L) = 20.2(A645) + 8.02(A663)
OR Total chlorophyll (mg/L) = A652/36

Chlorophyll a (mg/L) = 12.7(A663) – 2.69(A645)
Chlorophyll b (mg/L) = 22.9(A645) – 4.68(A663)

In 90% acetone:

Chlorophyll a (mg/L) = 11.93(A664) – 1.93(A647)
Chlorophyll b (mg/L) = 20.36(A645) – 5.50(A664)

2. Diatoms, chrysomonads, and brown algae containing chlorophylls a, c1 and c2 in actual proportions.

In 90% acetone:

Chlorophyll a (mg/L) = 11.47(A664) – 0.4(A630)
Chlorophyll c1 + c2 (mg/L) = 24.36(A630) – 3.73(A664)



A. Extraction of Pigments from Brown Algae

Extraction of pigments from brown algae, particularly the larger macrophytes, can be difficult because of the rubbery nature of the thalli, and the large amounts of polysaccharides in the tissue. The most commonly used methods for extraction of chlorophyll ac, and fucoxanthin are described below (Seely et al. 1972). The formulas presented do not include a correction for carotenoid absorption, and therefore, may overestimate the amount of fucoxanthin present.


  1. Remove epiphytes and rinse in distilled water. Blot dry and weigh tissue.
  2. Place discs in test tubes, cover with 4 volumes (mL) of DMSO per gram of tissue. Allow extraction to proceed in the dark for 15 min.NOTE: This is only a recommendation. The amount of DMSO required should be determined empirically.
  3. Remove discs from tubes, wash with 1.0 mL dH2O. Pour wash water into DMSO extract.
  4. Place discs into clean test tubes. Add 3 mL 100% acetone. Allow the pigments to be extraction for 2 hr in total darkness.
  5. Dilute acetone extract with 1 mL absolute methanol and 1 mL H2O.
  6. Measure absorbance of DMSO extract at 665, 631, 582, and 480 nm.
  7. Measure acetone extract at 664, 631, 581, and 470 nm.NOTE: Use 4:1 DMSO:H2O and 3:1:1 Acetone:Methanol:H2O as blanks.
  8. Calculate pigment concentrations as follows:DMSO

    Chl a = A665/ 73.6

    Chl c = (A631 + A581 - 0.3A664)/62.2

    Fucox.= (A480 - 0.772(A631 +A582 - .297A665) -.049A665)/130


    Chl a = A664/73.6

    Chl c = (A631 + A581 - 0.3A664)/62.2

    Fucox.= (A470 - 1.239(A631+ A581- 0.3A664) -.0275A664)/141


B. Extraction of Phycobilin Pigments

Phycobilin pigments are water soluble and therefore are not well extracted by organic solvents. Phycobilin pigments may be extracted from the pellet of an organically extracted pellet (although some loss may occur into the organic phase) or from fresh thalli using the following protocol (Evans 1988).

1. Weigh out 0.05 – 0.5 g of thallus. Transfer to mortar and grind in 5 mL 0.1M phosphate buffer, pH 6.8, with acid-washed sand.

2. Centrifuge for 10 min at 1,000 x g.

3. Transfer to 25 mL volumetric flask and bring up to volume.

4. Determine phycobilin concentration using following formula:


R-PE = [(A564- A592) - (A455 - A592)0.20]*0.12


C/R-PC = [(A618 - A645) - (A592 - A645)0.51]*0.15

OR FOR PLANTS CONTAINING C-PC (from Kursar and Alberte 1983)


C-PC = 166(A618 )- 108(A650)


APC = 200(A650) – 52.3(A618)


R-PE = 169(A498)- 8.64(A615) – 1.76(A650)




TLC and in vivo spectral analysis and pigment identification:

Owens, T.G., J.C. Gallagher, and R.S. Alberte. 1987. Photosynthetic light-harvesting function of violaxanthin in Nannochloropsis spp. (Eustimatophyceae). J. Phycol. 23:79-85.

Determination of pigment concentration in solvents:

A. Green algae and seagrasses:

Jeffrey and Humphrey. 1975. Biochem. Biophys. Phlanz. 167:191-194.

B. Brown algae and diatoms:

Duncan, M.J. and P.J. Harrison. 1982. Bot. Mar. 25:445-447.

Seely et al. 1972. Mar. Biol. 12:184-188.

C. Red algae:

O’Carra. 1965. Biochem. J. 94:171-174.

Kursar, T. and R.S. Alberte. 1983. Plant Phys. 72:409-414.

Evans, L.V. The effects of spectral composition and irradiance level on pigment levels in seaweeds. In: Experimental Phycology. Lobban, C.S., D.J. Chapman and B.P Kremer. Eds. New York. pp 123-134.

Algal classification:

Dring, M.J. 1982. The Biology of Marine Plants. Edward Arnold. pp 1-8.



A. Solutions and chemicals:

100 % Acetone (reagent grade)

Petroleum ether:Acetone (7:3 v:v) for 1 L : 700 ml petroleum ether and 300 ml acetone. This solution is highly flammable, use with caution.

Liquid nitrogen
Nitrogen gas

B. Materials

  • Mortar and pestle (place into freezer well in advance of the use)
  • Tissue homogenizers
  • Pipets
  • Silica gel plates
  • Developing Tanks
  • Centrifuge tubes
  • Micropipettes
  • Eppendorf tubes
  • Filter paper
  • Razor blades
  • Sand