Chlorophyll a (Welschmeyer method)

The determination of chlorophyll a is used as an estimate of phytoplankton biomass. It is not indicative of species composition nor the physiological state of the population. Chlorophyll a is extremely sensitive to light and heat. Therefore all chlorophyll determinations should be performed in subdued light and a cool room, including extracting in a refrigerator. There are various methods for determining the chlorophyll a content of phytoplankton. The method below is a modification of Welschmeyer, N.A. (1994).

Chlorophyll a and phaeopigments: The chlorophyll a content of phytoplankton is measured by vacuum filtering a water sample through a 25 mm GF/F filter, then extracting the pigment on the filter in methanol. Sample processing should begin as soon as possible or within 6 hours of collection and duplicate filters are extracted for each water sample. Chlorophyll a is extremely sensitive to light and heat. Therefore, all chlorophyll a determinations should be performed in subdued light and a cool room. Furthermore, extraction should also take place under dark, cool conditions such as in a refrigerator. There are various methods for determining the chlorophyll a content of phytoplankton. The method presented here is a modification of Welschmeyer (1994). This method measures chl a without interference from chl b and phaeopigments by the use of a different lamp and filter set than in the acidification method of Holm-Hansen and Riemann (1978). The fluorometer is calibrated at least every 6 months or as required with chlorophyll a derived from spinach (Sigma Chemical Co #C-5753). The calibration is checked before each sampling trip against a coproporphyrin standard measured at the time of calibration. Refer to the attached Standard Operating Procedure for chlorophyll a for the full method. The determination of chlorophyll a is used only as an estimate of the biomass of phytoplankton. It is not indicative of species composition nor the physiological state of the population. The amount of chlorophyll in a body of water is often a defining parameter in describing eutrophication or water quality.

1. Turner Designs model 10-AU-005 fluorometer with optical kit 10-040R
2. fluorometer accessories (spare lamps, filters, etc.)
3. 13 x 100 cuvettes
4. refrigerator
5. centrifuge
6. 15 ml centrifuge tubes (preferrably disposable polypropylene)
7. test tube rack for centrifuge tubes
8. filtering manifold equipped with 25 mm filter funnels
9. vacuum pump
10. 25 mm GF/F filters
11. large vacuum reservoir
12. vacuum lines with dessicant trap
13. vortex mixer
14. methanol
15. waste methanol container
16. waste MEOH funnel
17. MeOH squirt bottle
18. Di water squirt bottles
19. plastic funnel w/ 153 um mesh netting
20. 1-10 ml Brinkmann Dispensette
21. aluminum foil
22. parafilm
23. test tube brushes
24. assorted pipets: 25 ml, 10 ml, 5 ml, 2 ml, 1 ml
25. assorted beakers: 50, 100, 250, 500 ml
26. assorted grad cylinders: 100, 250, 500, 1000 ml
27. bulb-type filler
28. graduated cylinders cut for chl: 50 and 100 ml
29. Kimwpipes
30. pasteur pipets
31. Latex pipet bulbs
32. data sheets

1. Perform the entire procedure in subdued light and let fluorometer warm up for at least 2 hours.
2. Place a 25mm GF/F filter on each filter base and attach the funnels.
3. Prescreen approx. 150 mL of the raw water sample through 153 µm mesh screening funnel into a small container such as a 250 ml beaker.
4. Rinse either the 50 or 100 mL cut off graduated cylinders with 10-20 mL of the prescreened sample.
5. Fill the grad cylinder with the prescreened sample and pour into a filtering funnel. 50 ml or less is sufficient for most inshore or coastal water.
6. Repeat step 5 twice for each sample producing duplicates for each sample until all 6 funnels are used (3 samples x 2 reps).
7. Turn the vacuum pump on and set vacuum to approximately 10 inches Hg.
8. Open the ball valves on the manifold.
9. When the sample has completely passed through the filter, remove the funnel and use forceps to fold filters in half with the pigmented portion inside. Place filter in a numbered plastic centrifuge tube with the side of the filter holding material exposed to the inside of the tube.
10. Record tube # and sample information on the chl data sheet.
11. Dispense 10 mL of methanol into the tube using the dispensette on the methanol bottle.
12. Mix the tube well on the tube buzzer and put in a sleeve of the specially constructed PVC extraction rack.
13. Process all samples as above, and place rack in the refrigerator remembering to record the initial time.
14. Extract for 18-20 hours.
15. Remove tubes from refrig and allow to warm up for 10 minutes.
16. Mix tubes well on buzzer and centrifuge for 10 minutes at 2000-3000 rpm (setting 75 on the Dynac centrifuge).
17. Meanwhile, prepare 8 clean cuvettes in a rack by the fluorometer.
18. Rinse a clean cuvette with a small amount (<1 ml) of the sample and discard rinse into waste methanol jug.
19. Pour enough sample into the cuvette to fill 3/4 full, but not enough to disturb the pellet.
20. Carefully clean the exterior of the cuvette with a Kimwipe to remove fingerprints, dirt, etc.
21. Place cuvette in fluorometer and replace cover.
22. The fluorometer should be set to Auto-range. At this point, the fluorometer will beep as it adjusts the scale.
23. When the auto-ranging is finished, press the asterisk (*) to begin the discrete sample averaging sequence. Record the chl a concentration when “Done” appears on the screen. Actual Chl a in your sample is calculated by performing a volume correction.
24. Remove sample and discard into the waste methanol jug.
25. Rinse cuvette 3 times with methanol.
26. Put cuvette upside down in a test tube rack lined with Kimwipes to dry. It is important not to have any water in cuvette.
27. Use next cuvette for next sample.
28. While reading this batch, centrifuge the next 8 samples.