Phytoplankton samples collected using appropriate quantitative sampling methods can be analysed in the laboratory by various counting methods or by the measurement of chlorophyll-a concentrations within the samples (Box 4.5).
The chlorophyll-a concentration will provide an estimate of the standing crop or abundance of phytoplankton present in a water sample, but it will not provide any information on the composition of the phytoplankton present. To do this, you will need to identify and count each taxon (that is, each species or 'type') present using a microscope and a counting chamber. The data obtained by these means will provide an estimate of the number of cells per mL (cells. mL_1) of each taxon and can be used to describe the composition of the entire phytoplankton community, the dominance of each taxon within that community, and changes in community abundance and composition over time. However, because different species of phytoplankton have cell sizes that differ greatly from each other, total cell counts are often unreliable for describing these changes. For example, a large cell count of a very small-sized algal species
BOX 4.5 EXTRACTION AND QUANTIFICATION OF CHLOROPHYLL
Chlorophyll-a is an indirect measure of phytoplankton standing stock (crop), and represents the mass of phytoplankton per unit volume or area of water and should be reported as micrograms per litre (pg.L-1) or milligrams per cubic metre (mg.m-3) or per square metre (mg.m-2). Replicate water samples should be collected from the water column at pre-specified depths. The chlorophyll-a content is estimated in the laboratory using either the fluorescence or absor-bance techniques described in Strickland and Parsons (1972). Water samples are filtered onto 25-mm diameter Whatman GF/F fiber (0.7 pm nominal pore size) or equivalent with a gentle vacuum (of less than 100 mm Hg). The actual sample volume can range from 100 mL to 4 L, as long as you can see that the filter paper is distinctively green. You should work in a shaded room because, in this state, chlorophyll can degrade in bright light. The sample can be wrapped in foil and frozen for up to 3 months for later analysis. The filter paper is extracted into 90% acetone and the light absorbance at particular wavelengths is recorded in a spectrophotometer. Alternatively, the natural fluorescence of the extracted chlorophyll can be determined - this is a more sensitive method.
may be replaced over time by a smaller number of cells of a much larger sized species. Using just the cell count data, you may deduce that algal presence has decreased, whereas, in fact, algal biomass may have increased. It may therefore be important, depending on the objectives of your study, to also determine the biomass present of each algal taxon identified and counted within the sample. Biomass is usually initially calculated as a biovolume (mm3.L_1), which is converted to biomass by assuming that algal cells have a density similar to that of water (therefore a biovolume of 1 mm3.L_1 equals a biomass of 1 mg.L"1). Most correctly, biovolume estimates should be done by:
2. converting this to an average cell volume for this species using standard geometric formula best representing the shape of the cell (Hillebrand et al. 1999)
3. multiplying the cell count by this average cell volume to obtain a total volume for all of the cells for that species.
This is often very laborious as it needs to be repeated for each species present in the sample. Sometimes published tables of standard cell sizes for various species are used instead, if the error involved is considered acceptable in comparison with the costs of using actual measurements.
Samples are best preserved using Lugol's iodine solution for both freshwater and marine samples (although it may damage some of the small flagellates). Some laboratories will not analyse samples preserved with substances such as formaldehyde, as these are carcinogenic and represent an occupational health and safety hazard. Samples collected from a dense algal bloom can be analysed directly, but they usually need to be concentrated prior to analysis. This is usually done using a 100 mL aliquot of the sample that has already been well mixed by shaking the sample bottle prior to sub-sampling. The aliquot is poured into a 100 mL measuring cylinder and left to stand for a minimum of 24 hours. If small nanoplankton are present, a longer sedimentation time may be necessary. The Lugol's iodine preservative helps the cells sink more rapidly. After the required sedimentation period, most of the phytoplankton cells will have settled to the bottom of the measuring cylinder. The top 90 mL can then be drawn off using a suction pipette, taking care not to disturb the algal cells at the bottom of the cylinder. This gives a 10x concentration.
The identification and counting of phytoplankton cells is something that takes much patience, practice and experience to do correctly. There are a number of taxonomic guides and keys that have been published to assist in the identification of both freshwater and marine algae (see Chapters 5 and 6).
There are a number of methods available for counting algal cells in samples. The easiest method is using a Sedgwick-Rafter cell. Other methods (such as a Lund cell or an inverted microscope) are useful providing they can be used with at least as good an accuracy and precision as counts using a Sedgwick-Rafter cell (see Hotzel and Croome 1999 for a description of these methods). The Sedgwick-Rafter cell is a four-sided counting chamber that is 50 mm long by 20 mm wide by 1 mm deep, giving a bottom area of 1000 mm2, and an internal volume of 1 mL. They have a grid engraved on the bottom, with lines 1 mm apart. If correctly calibrated and filled, the volume of sample covering each grid square is 1 mm3. Both glass and plastic versions are available, with the glass cells being better, but more expensive. The cells are used on the stage of a normal compound microscope - preferably one with binocular eyepieces. Counting is done at 100x magnification, with higher power being used to identify small sized algal cells. A very thin microscope cover slip (No. 1 thickness) is required to cover the cell.
Immediately before commencing a count, the phytoplankton cells in the bottom of the measuring cylinder are resuspended into the remaining 10 mL of sample left in the measuring cylinder by swirling, and a further sub-sample of approximately 1 mL of this collected with a Pasteur pipette. This is then decanted carefully into the counting chamber of the Sedgwick-Rafter cell. The cell is full once the cover slip, which should be placed obliquely over the cell prior to filling with one corner open, just begins to float and can be rotated to completely cover the chamber. This avoids introducing air bubbles into the sample. The cell should not be overfilled. Once filled, the counting cell should be left to stand on the stage of the microscope for 15 minutes, to allow the algal cells to settle to the bottom. It is not necessary to count all the cells on the bottom of the Sedgwick-Rafter cell. However, a minimum of 30 grid squares should be counted. These should be selected randomly, as there is differential sedimentation of algal cells within the counting cell, with more algae sedimenting closer to the walls than in the centre ('edge effects'). Counting traverses across the width of the cell helps to overcome these edge effects and will cover 40 grid squares. A second requirement is that a sufficient number of algae are counted to provide a counting precision of ±30%. This involves counting at least 23 'units' for all of the most dominant algal taxa present. A 'unit' is either an algal cell, filament or colony, depending whether the species being counted is unicellular, filamentous or colonial. If counting 30 grid squares or two traverses does not yield a sufficient number of units (that is, more than 23), then additional grid squares or traverses will need to be counted. Record the number of grid squares counted as well as the number of algal units counted. If an algal unit lies across the line engraved in the base of the Sedgwick-Rafter cell to delineate a grid square, so that it falls within two squares, the simple rule is that if it lies on the right side of the grid square, include it in the count, but if it lies on the left side, exclude it. Similarly, if it falls across the top line of the square, include it, but exclude any algal units falling across the bottom line. Algal units are often smaller than the width of the lines engraved in the Sedgwick-Rafter cell, so the same applies for any algal units lying within the grid lines delineating a square.
The number of algal units present per mL within the actual water body is calculated as:
(no. of grid squares counted x concentration factor, which is typically 10)
For filamentous and colonial algae, it is then necessary to convert the count in units.mL"1 to cells.mL_1. Many green algae have a set number of cells per colony (for example 4, 8, 16, or 32), so, when this is known, it is easy to multiply the units by the cell number per colony to obtain cells.mL_1. However, many other phytoplankton species, especially cyanobacteria, have a variable number of cells per filament or colony. In this instance, it is necessary to count the number of cells in 20 to 30 randomly selected filaments or colonies, and then obtain an average number of cells per colony from these counts.
Further problems arise when samples contain large-sized colonies or tangled aggregations of filaments containing thousands of cells, where it is impossible to count all the cells in each colony or aggregation. In these situations, it is necessary to estimate a portion of the colony or aggregation -say 5% or 10% of the total colony size - and count or estimate the number of cells within that portion. Remember that the colonies or aggregations are three dimensional, with cells overlying cells, and outside of the focal plane at which you are viewing the colony. Once you have an estimate of the number of cells in 5% or 10% of the colony, multiply this by 20 or by 10, respectively, to obtain an estimate of the total cells per colony.
When you do these estimates of average cell numbers per filament or colony to obtain a count in terms of cells.mL-1, the errors can be quite large and are in addition to any statistical counting error. The need to make these estimates arises only during blooms and becomes acceptable because of immediate management needs. Methods to break up large colonies into smaller units to make counting easier (homogenisation, addition of chemicals or sonification) are often inadequate and may destroy a large proportion of the cells present.
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