Fluorometric Analysis of Chlorophyll a

GSI Protocol #07
Published: August 2024
Prepared by: Sarah Cole, M.S., Staff Scientist; Stephen G. Hesterberg, Ph.D., Executive Director


Water Collection

MATERIALS

  • 0.5-1.0 L HDPE dark sided bottles or aluminum foil

  • Permanent waterproof marker and chemical labels/tape

  • Cooler with ice

  • Refrigerator/Freezer (-20°C)

INSTRUCTIONS

1. Sample water in triplicate at the beginning of each site visit to avoid increased turbidity.

2. Use a horizontal water sampler or lower sealed sample bottles to desired depth. If collecting by hand, remove the sample bottle lid and fill with water. Resecure lid while the sample bottle is submerged. If using a water sampler, transfer water to a sample bottle. If necessary, wrap the sample bottles in aluminum foil. Label each sample appropriately (e.g., site, date, replicate, etc.).

3. Immediately place sample bottles on ice until they can be either filtered or frozen.

4. If filtering the next day, samples can be refrigerated overnight. If freezing samples before filtering, pour off excess water. Filter all frozen samples within 7 days of collection.


Filtration

MATERIALS

  • 100 mL graduated cylinder

  • 47 mm, <0.7 um glass fiber filter (GF/F)

  • Vacuum filtration system, including: 1 L funnel flask, 47 mm diameter two-piece funnel cup, clip, vacuum pump, diaphragm vacuum filter, air-line tubing

  • Forceps

  • Nitrile gloves

  • Deionized (DI) water

  • Aluminum foil

  • 15 mL polypropylene centrifuge tubes

  • Freezer (-20°C)


INSTRUCTIONS

1. Turn off all lights; utilize red light if necessary.

2. Set up filtration apparatus by first connecting the vacuum pump and filter flask with an airline and filter. Place the bottom portion of the funnel cup into the top of the filter flask (Figure 1a) and add one GF/F using forceps while wearing nitrile gloves. Add the top portion of the funnel and use the clip (Figure 1b) to secure the two pieces together (Figure 1c).

3. Shake sample and pour known volume into graduated cylinder. Start with 100 mL, but for waters with low Chlorophyll a concentrations (<2 ug/L), 200-500 mL may be required (GF/F filters should at minimum possess a faint color). Rinse the graduated cylinder three times with DI water and empty onto filter. Record the volume of sample that was filtered.

4. Turn on vacuum pump and filter only until the GF/F is dry; turn off vacuum pump.

5. Remove the clip and top portion of the filter funnel. Using forceps, carefully fold the GF/F filter into quarters by halving twice and place into a pre-labelled 15 mL centrifuge tube. Do not touch the portion of the filter that has sample on it with the forceps; only touch the outer edge or bottom of the GF/F.

6. Cover 15 mL centrifuge tubes in aluminum foil and place in a -20°C freezer until extraction (within three months).

7. Filter at least one ‘blank’ for every 25 samples or at least once per batch of samples. A blank consists of filtering a known volume of DI water and is treated like other samples. Filter the same volume of DI water in preceding samples.

8. Empty flask periodically and do not exceed 800 mL when filtering.

Figure 1. (A) Funnel flask with attached vacuum tube and the bottom portion of funnel cup which holds the GF/F filter. (B) Top portion of funnel cup and clip. (C) Fully set-up filtration apparatus ready to filter.

Extraction

MATERIALS

  • 99.5% Acetone

  • Deionized (DI) water

  • 1 L glass or polypropylene plastic container

  • Chemical Label/Tape

  • Lab marker

  • Nitrile gloves

  • Tissue grinder with Teflon pestle

  • Refrigerator

INSTRUCTIONS

1. Create a 1 L stock solution of 90% acetone by combining 100 mL of DI water with 900 mL ACS grade (99.5%) acetone in a well-ventilated area. Store stock solution in a 1 L reusable glass or polypropylene plastic bottle (NOT polyethylene). Label with ‘90% acetone’ and the date.

2. Turn off all lights; utilize red light if necessary.

3. Remove 15 mL centrifuge tubes from the freezer and allow to thaw completely (est. 30-60 min). Add 7 mL of 90% acetone to each centrifuge tube; record if a different volume is used.

4. Using the tissue grinder, disturb the GF/F against the side of the tube until the filter is well ground (10-15 sec) and shake. Clean the tissue grinder between samples with DI water.

5. Wrap centrifuge tubes in aluminum foil and place processed samples in the refrigerator overnight and no longer than 24 hrs.


Fluorometric Analysis

MATERIALS

  • 15 mL tube centrifuge

  • 0.1 N hydrochloric acid

  • Deionized (DI) water

  • 250 mL glass or polypropylene bottle

  • Chemical labels or tape

  • Lab marker

  • Turner Trilogy Fluorometer with CHL-A optical module

  • 3.5 mL glass cuvette

  • Solid standards

  • Transfer pipettes

  • Kim wipes

  • Polypropylene or glass waste container


INSTRUCTIONS

1. Following extraction, remove samples from the refrigerator and allow them to warm to room temperature; shake tubes vigorously before centrifugation at 350 rpm for 20 min.

2. Create a 250 mL stock solution of 0.1N HCl in a well-ventilated area and while wearing nitrile gloves. Store stock solution in a 250 mL reusable glass or polypropylene plastic bottle (NOT polyethylene). Label bottle with ‘0.1N HCl’ and the date.

3. Turn on calibrated Turner Instruments Fluorometer and turn off all lights; utilize red light if necessary.

4. Select ‘CHL-A’ module on the touch screen, open the fluorometer, and ensure that the module is inserted.

5. Verify instrument calibration by recording solid standard fluorescence in RFU mode:

a. Select the ‘Mode’ button on the bottom of the touch screen and select ‘Raw Fluorescence’.

b. Place the secondary standard in the fluorometer sample compartment with the handle towards the rear of the instrument.

c. Close lid and press the large green button on the top right of the touchscreen to record fluorescence.

d. Readings should fall within 10% of the calibration value. If the measurement is beyond the suitable range, the instrument will have to be recalibrated using direct fluorometric calibration.

e. The box holding the solid standards should be labelled with the calibration values and date of calibration.

6. Change the mode to ‘Direct Fluorescence’. To do this select ‘Calibrate’ from the bottom of the touch screen, followed by ‘Use Stored Calibration’, and then the most recent calibration of the fluorometer.

7. Prior to analysis of the first sample, record fluorescence of a blank cuvette:

a. Fill a cuvette with 90% acetone, wipe with a Kim wipe, and place in the fluorometer

b. Hit the green button on the top right of the touchscreen to read direct fluorescence.

c. The instrument will ask you the total volume filtered and volume of solvent. For the blank, enter the average volume for the subsequent samples.

d. Read the before acidification value of the blank. Remove the cuvette and acidify by placing 3 drops of 0.1 N HCl in the cuvette.

e. Return the cuvette to the instrument and take the reading after acidification.

f. Chlorophyll a concentration should read <2 ug/L. If chlorophyll a concentration is higher than 2 ug/L, repeat taking a blank measurement. If the concentration is still beyond the acceptable range, the fluorometer will need to be calibrated.

g. Empty the cuvette into an appropriate acetone waste container of either glass or polypropylene (NOT polyethylene). Label with ‘90% acetone waste’ and date.

h. Clean the cuvette by rinsing with 90% acetone twice.

8. To quantify sample chlorophyll a, pipette ~2 mL of centrifuged sample into a cuvette and record fluorescence:

a. When transferring samples, avoid the bottom of the centrifuge tube, being careful to extract liquid only from above the GF/F. The exact amount of liquid does not matter so long as the volume is greater than what the fluorometer required.

9. Wipe the sides of the cuvette with a Kim wipe and place the cuvette in the fluorometer; close the lid. Press the green button on the top right of the touchscreen to record sample fluorescence before acidification. If your sample concentration is greater than the linear range of the standard curve for the instrument, samples need to be diluted with 90% acetone to fall within the measurable range.

10. Remove the cuvette from the instrument and place three drops of 0.1 N HCl into the same sample cuvette; repeat the measurement following acidification.

11. Record both chlorophyll a and phaeophytin values.

a. Empty the cuvette into an appropriate acetone waste container. Clean the cuvette by rinsing with 90% acetone twice before starting another sample.

12. Repeat step 8 with all samples

13. Run at least one ‘blank’ for every 25 samples processed to ensure calibration of the instrument.

14. After all samples have been measured, place leftover samples into the labeled acetone waste container.



FLUOROMETRIC DIRECT CALIBRATION

MATERIALS

  • High and low range Chlorophyll a standards

  • Turner Trilogy Fluorometer with CHL-A optical module

  • 90% acetone

  • Kim wipes

  • Cuvette

  • Transfer pipettes

  • 0.1 N HCl

  • Solid standards

  • Chemical label/tape

  • Lab marker

INSTRUCTIONS

1. Turn on fluorometer and ensure that the CHL-A module is installed

2. Select the CHL-A module on the touchscreen

3. Touch the “Calibrate” button to begin the calibration

4. Select “Run New Calibration”

5. Select ug/L as the unit

6. Fill the cuvette with 90% acetone as the blank and with Kim wipe. Place the cuvette in the instrument and close the lid. Touch “OK”

7. After the blank has been measured open the lid and remove the blank. Empty the contents in the acetone waste.

8. Start to calibrate with standards of known concentration. You can use 1-5 standards. Standards must be read in order of increasing concentration.

9. Fill the cuvette with ~2 mL of the low standard. Wipe with a Kim wipe, place in the instrument and close the lid.

10. Enter the concentration for the first standard and touch “OK”

11. After the standards has been measured you will be prompted to acidify the sample.

a. Remove the cuvette and close the lid. Place 3 drops of 0.1 N HCl into the cuvette.

b. Replace the cuvette and take the after acidification reading.

12. After the standard has been measured you will be prompted to either enter more standards or proceed with the current calibration. You need at least a 2-point calibration. Select “Enter More Standards”

13. Repeat with the next standard that has higher concentration.

14. When all the standards have been measured, touch “Proceed with Current Calibration”

15. Name the calibration with the date for future use.

16. Take reading for the solid standards in RFU mode for future use in checking calibration.

a. Select the Mode button on the bottom of the touch screen and select raw fluorescence.

b. Place the secondary standard in the fluorometer sample compartment with the handle towards the rear of the instrument.

c. Close lid. Press the large green button on the top right of the touchscreen to read fluorescence.

d. Take three (3) readings of the solid standard. Reading may not be the exact same but should be in range (+/- 10%). Take the average of the readings.

e. On the box holding the solid standard record the average of RFU and the date of calibration with label.

f. Repeat for second solid standard.


Safety

• Personal Protection Equipment – Wear long pants, long sleeves/lab coat, closed-toed shoes, and goggles when processing chlorophyll a samples. Utilize a fume hood or wear a respirator when working with acetone.

• Chemicals – Review chemical SDS for hydrochloric acid (HCl) and acetone before processing chlorophyll a samples.


References

Arar, E. J. and Collins, G. B. (1997) Method 445.0: In vitro determination of chlorophyll a and pheophytin in marine and freshwater algae by fluorescence. United States Environmental Protection Agency, Office of Research and Development, National Exposure Research Laboratory, Cincinnati, OH.


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