Carlson, R.E. and J. Simpson. 1996. A Coordinator’s Guide to Volunteer Lake Monitoring Methods. North American Lake Management Society. 96 pp.
Chlorophyll is the green molecule in
plant cells that carries out the bulk of energy fixation in the
process of photosynthesis. Besides
its importance in photosynthesis, chlorophyll is probably the
most-often used estimator of algal biomass in lakes and streams, at
least in North America. Its
popularity results from several considerations;
is a measure of algal biomass that is relatively unaffected by
is a fairly accurate measure of algal weight and volume, and,
acts as an empirical link between nutrient concentration and a
number of important biological phenomena in lakes and reservoirs.
Chlorophyll is also relatively easy to
measure. This facility of
measurement contributes to its popularity, but the resulting values
are far more ambiguous than most are willing to admit.
Chlorophyll itself is actually not a
single molecule but a family of related molecules, designated
chlorophyll a, b, c, and d. Chlorophyll a is the molecule found in all plant cells
and therefore its concentration is what is reported during chlorophyll
analysis. Chlorophyll d is found only in marine red algae, but
chlorophylls b and c are common in fresh water. The molecular structure of chlorophylls a and b consists of a ring-like structure called a porphyrin and a long
organic phytol "tail." In the center of the porphyrin ring is a magnesium molecule
(Fig. 1). Chlorophyll c lacks the phytol chain. The
relative concentrations within the cell of these chlorophylls varies
with the species of algae, but chlorophyll a is dominant in all
the eukaryotic algae and the prokaryotic blue-green algae
|Figure 1. The chlorophyll a molecule, consisting of a porphyrin ring, a chelated magnesium molecule in the ring (purple), and a long hydrocarbon (phytol) "tail." Model courtesy of Botany Online, University of Hamburg (http://www.biologie.uni-hamburg.de/b-online/)|
Other pigments are also present in
algal cells. These are
the carotenes and the xanthophylls. In the cyanobacteria, water-soluble phycobiliproteins are the predominant accessory pigment, giving the group their
characteristic blue-green or red color. In addition to the algal pigments, some bacteria are also
pigmented with a series of bacteriochlorophylls.
In addition to the naturally occurring
pigments in algal cells, a filtered water sample will also contain
colored degradation products of these pigments. When algal chlorophyll degrades, it forms a series of
degradation products, the nature of which depends on what part of the
molecule that is affected. As a chlorophyll degrades, the initial step is either the loss
of the magnesium from the center of the molecule or the loss of the
phytol tail. The former
pathway results in the formation of the molecule, phaeophytin;
in the latter pathway, the resulting molecule is termed a chlorophyllide. The
degradation scheme is shown in Figure 2. Further degradation of either the phaeophytin or the
chlorophyllide produces a molecule termed a phaeophorbide: phaeophytin is degraded
by the loss of the phytol tail and a
chlorophyllide loses its magnesium ion. When a chlorophyll molecule breaks down, a number of distinct
phaeophytins, chlorophyllides, and phaeophorbides will be produced,
depending on the parent molecule. Some of these breakdown products.
|Figure 2. The degradation pathways of chlorophyll|
When a sample is filtered and extracted for chlorophyll analysis, it unfortunately contains a large number of these pigments other than chlorophyll a, the primary pigment of interest in monitoring programs. The absorbencies of these other pigments are not easily separable spectrophotometrically or fluorometrically from their parent molecule, producing falsely high absorbencies and subsequent erroneous values for chlorophyll a. Despite its seeming simplicity in the analysis of chlorophyll, the validity of its results depends on whether or not these interferences are adequately removed. Almost every choice of analytical method addresses specific interferences, yet ignores others. It therefore becomes the decision of the program coordinator to choose the analytical procedure that produces the most useful information for the program, not necessarily the most accurate estimate of chlorophyll a. It may be that some of the simpler techniques are more than adequate for the purposes of a monitoring program.
Techniques for Collection and Preservation of Chlorophyll Samples
By some means, the volunteer must
gather a sample of water, using either a hose sampler, some sort of
water sampling bottle, or by simply lowering the sample container over
the side of the boat. Once
the sample is taken, it is usually filtered and preserved until
delivered to the laboratory for analysis. An alternative to filtering, preservation, and storage would be
to immediately deliver the whole water sample to the laboratory. Herve
and Heinonen (1982) suggest that whole-water samples stored at 4oC in the dark can be kept up to 1 day without significant degradation
of chlorophyll. Weber et
al. (1986) found no change in refrigerated samples over 18 days,
but if the samples were left at room temperature (20oC),
50% of the chlorophyll was lost in 5 days.
Filtration is usually accomplished in
volunteer monitoring programs using a filtration funnel and hand-held
suction pump. This system
allows the volunteer to filter large amounts of water in a relatively
short time. The volunteer can also see how much algae are being collected
on the filter, and therefore judge when it is sufficiently green. This
filtration technique does have the problem that the volunteer must
measure the water in a separate, graduated container and must handle
the filter, both before and after the filtration.
The Ohio-NEFCO program uses a 25 mm
in-line Swinney filter holder instead of a filter funnel. The glass fiber filter is placed into the filter holder by
the coordinating laboratory prior to distribution to the volunteers.
The volunteers are given a 60 ml plastic syringe equipped with a latex
rubber hose and a 3-way valve. After
the sample is brought into the boat, the volunteer places the tube
into the sampler, draws 50 ml of sample into the syringe, and pushes
the sample gently through the filter, which is positioned on the
output end of the 3-way valve. Pulling back on the plunger switches the valve back into
input mode and a second 50 ml of sample is drawn up into the syringe. This alternation of drawing the sample and pushing it through
the filter is done until an appropriate amount of algae have been
filtered (See below). When
finished, the volunteer puts the exit end of the filter holder on the
input tube and draws any remaining water out of the sampler. The entire sampler is then wrapped in foil, labeled, and stored
in the freezer until picked up for analysis. The volunteer never needs to touch or manipulate the filter.
The amount algae filtered by the volunteer is important. There appears to be more variability in the acid ratio at low absorbencies (Fig 3) that may contribute to error in chlorophyll a values. This variability may be a result of too much acidity causing phaeophytin to be degraded to phaeophorbide (Hallegraeff, 1976) or an analytical error associated with low chlorophyll concentrations. Standard Methods (APHA, 1989) recommends an optical density of 0.1 to 1.0 at 664 nm if the trichromatic method is used.
|Figure 3. The relationship between the acid ratio and the absorbance of the extract. Note that the variability is apparently higher at low absorbencies, suggesting that the amount filtered could make a difference in the estimated phaeophytin values.|
Some programs instruct the volunteer to continue filtering
until the filter is slightly green. The Ohio-NEFCO and Wisconsin programs developed a relationship
between Secchi transparency and the amount of water filtered (Table
1). The Ohio-NEFCO table
works reasonably well except in reservoirs having high non-algal
turbidity. In that case,
the amount of algae filtered is insufficient. In Florida, the volunteer is instructed to filter a volume of
approximately 100 milliliters for each foot of the Secchi depth.
|Table 2. The relationship between Secchi depth and the amount of water filtered use etz, et al., 1992) and in the Ohio-NEFCO program|
| 1.0 - 1.5
|>1.5 - 2.25
| >2.25 - 3.25
|>3.25 - 6.0
| 2.1 - 4.0
| >6.0 - 9.75
| 4.1 - 8.0
| >9.75 - 16.5
Choice of Filters
There are differences
of opinion as to what type of filter to use. Three factors have been considered: retention of particles, efficiency of extraction, and cost.
Membrane filters such as Millipore HAÔ or GelmanÔ retain more particles (Lenz and Fritsche, 1980), but are more subject to clogging than are glass fiber filters. This can mean that the volunteer might either have to filter less water or have to use very long filtration times. It also means that if a volunteer is faced with a clogged filter, they have to make a decision as to how to proceed: either start over, continue filtering, or, as is done in the Wisconsin program, pour the sample into another container, change the filter, and resume filtering.
Glass fiber filters have
the advantage of being less expensive than membrane filters and,
during grinding, the
glass fibers aid in the homogenization of cells. The membrane filters
can be ground in a tissue grinder, but not as efficiently as glass
fiber filters. However, unless the superior extracting powers of a
solvent such as methanol is used with membrane filters (see below),
the advantages of superior retention of smaller particles by membrane
filters may be lost because of a lower amount of extracted
Phinney and Yentsch (1985) suggest that major retention differences
between membrane and glass fiber filters disappear above chlorophyll
concentrations of 1 μg/L glass fiber filters would seem to be
adequate for most inland waters.,. Prepas et al. (1988) found no difference in chlorophyll
concentrations collected on either Whatman GF/F (median retention size
of 0.7 um) and GF/C (median retention size of approx 0.2 um) glass
fiber filters in concentrations ranging from 2 - 175
μg/L. In oligotrophic waters, the choice of filter may be of more
μg/L. In oligotrophic waters, the choice of filter may be of more concern..
Marker et al. (1980) listed the following reasons for using a Whatman
GF/C glass fiber filter:
1. They are very efficient for chlorophyll retention in most situations and sometime even better than membrane filters.
2. They filter much faster and do not clog so rapidly.
3. On centrifugation there is no turbidity in acetone or methanol
4. The act as an excellent abrasive material to aid cell breakage
5. For comparisons between various
chemical variables, it is essential that the filters used for the
determinations should retain the same size fractions of particulate
material, and most investigators now use glass-fiber filters for these
determinations as well as for particulate organic carbon.
Some methods suggest the addition of
MgCO3 to retard degradation and to enhance filtration
efficiency. Some studies
have found that its addition has no significant effect (Lenz and
Fritsch, 1980), and it may absorb pigments (Daley et al., 1973;
Weber et al., 1986). If
it is used with glass fiber filters, it should be used prior to
filtration to decrease the pore size of the filter.
Preservation of the Chlorophylls
Once the chlorophylls are on the
filter, they become highly susceptible to degradation as the cells die
and decompose. They also
become increasingly light and temperature labile. Some method must be used to keep the pigments from degrading. The problem is compounded for volunteer programs because the
samples have to be transported to the laboratory for analysis. A
simple mailing of the samples would be desirable, but there is the
real possibility of degradation of the samples during the process.
The simplest method of preservation
apparently is to freeze the samples. Several authors report that frozen samples showed no
significant degradation even after 6 months (Lenz and Fritsche, 1980). Jones and Lee (1982), however, mention that they have
encountered problems with freezing and recommend that samples should
not be frozen unless investigation has shown that results from frozen
samples are comparable to those from fresh samples. A problem with freezing the filters is that apparently the
chlorophyll will begin to degrade as soon as it is unfrozen. This means that the samples must be brought to the laboratory
in a frozen state. This
would seem to preclude the use of the mails to get the samples to the
laboratory if they are not kept frozen during shipment.
Another preservation method is to
immediately submerse the filter in the solvent, seal and darken. Apparently the chlorophyll will not degrade as long as it is
kept dark. This means,
however, that the volunteer would have to be given the solvents and
that transportation would have to be of a liquid chemical.
Others have found that as long as the
filter is kept dry and in the dark, the chlorophyll will not degrade. More experimentation seems to be necessary before an adequate
preservation technique can be recommended.
Analysis for Chlorophyll
Several methods for chlorophyll
analysis are available. The
methods are carefully described in Standard Methods (APHA, 1991), and
the methods will not be discussed in detail here. However, there is a great deal of confusion about which method
should be used in limnological investigations, and this confusion has
resulted in a number of different methods being used by various
volunteer programs. Unfortunately,
although all of these methods report their results as chlorophyll a,
there is little evidence that the numbers derived by each method are
necessarily similar. Because
monitoring programs imply that the numbers generated are accurate as
well as precise, the choice of a technique is important. A little background about chlorophyll analysis might help
clarify the differences between the various chlorophyll methodologies
Choice of Solvents
Homogenization by grinding of the
filter enhances the rupture of the algal cells and increases
extraction efficiency of the solvent. Homogenization is an absolute necessity with an acetone
solvent, but some have found that other extractants such as ethanol or
methanol apparently do not need grinding to extract all the
chlorophyll (Sartory and Grobbelaar, 1984). Others, however, have found that even methanol extractants do
not extract as well without grinding. These
other solvents are more efficient than acetone at extracting pigments
from some green and blue-green algal cells. Methanol, however, is more toxic. Membrane filters can be ground but they lack the abrasiveness
to produce a good extraction, and their extraction efficiencies are
lowered (Long and Cooke, 1971).
There are three basic methods for
measuring chlorophyll. Analysis
using a spectrophotometer uses
Spectrophotometric analysis of
chlorophyll pigments were developed in the 1930's and 1940's (Weber et
al., 1986). Richards
and Thompson (1952) introduced a trichromatic technique that
was supposed to measure chlorophylls a, b, and c. Trichromatic equations attempted to remove interferences of the
other chlorophylls at the maximum absorption wavelength for each
Richards and Thompson, a number of modifications have been made to
these equations which purportedly produce better estimates of the
chlorophylls (Parsons and Strickland, 1963; UNESCO, 1966; Jeffrey and
Humphrey, 1975). When
these equations are ultimately compared with concentrations of
chlorophyll obtained using physical separation techniques such as
HPLC, paper or thin-layer chromatography, it is found that the degree
of correspondence is low. Apparently
the trichromatic equations are no substitute for physical separation
techniques. In addition,
these equations do not deal with the degradation products of
trichromatic "chlorophyll a" is better presented as
chlorophyll a minus most of the interference of other
chlorophylls but including all degradation products that have
absorbencies at the primary wavelength of chlorophyll a. These multiple chlorophyll equations have not been particularly
successful, but are still used in oceanographic research, where
degradation products are less of a problem.
Lorenzen (1967) and Moss (1967)
introduced an acidification step in a monochromatic method to
circumvent the interference by chlorophyll degradation products. When chlorophylls are acidified, the magnesium ion is lost from
the porphyrin ring, resulting in the production of a phaeophytin. Lorenzen (1967) produced equations capitalizing on the fact
that the ratio of pure chlorophyll a after acidification to
that before was 1.7. If
the sample contained pure phaeophytin, then the absorbance would not
change, and the ratio would be 1.0. Acid ratios between 1.0 and 1.7 would therefore indicate the
amount of degradation products in the sample, and the estimate of
chlorophyll could thus be corrected.
In natural waters, the acid ratio, and
therefore the resulting estimate of chlorophyll a, varies
"chlorophyll a," whether determined using
trichromatic equations, or using the pigment correction is really an
operationally defined term, whose meaning and values change with each
change in the technique. Numerous
authors emphatically state that the only method for measuring
chlorophyll a accurately is using some separation procedure
such as HPLC. Any other
method produces only an estimate of the chlorophyll concentration.
An alternative to the use of physical
separation techniques and the distress of choosing the
"proper" spectrophotometric equation is to report the amount
of total chlorophyll pigments (Golterman and Clymo, 1971). It is the estimate of all chlorophyll pigments and degradation
products that absorb at 665 nm. The
measure is a descendent of the Odum's et al. (1958) monochromatic
chlorophyll a equation. Golterman
and Clymo's equation uses the extinction coefficients of Strickland
and Parsons (1963) in 90% acetone which are probably the most popular extinction coefficients and
solvent. Their equation
where V is the volume filtered (L), v is
the volume of extract (ml), and p is the pathlength (cm). Using values for total chlorophyll pigments rather than
either the trichromatic equations or the acid-corrected equations gets
around the problem of interference by ignoring it. It is simply a measure of absorbance at 665 nm.
There are some very good reasons for
ignoring, or at least giving second place to, chlorophyll a values. Comparisons of
the total chlorophyll concentration with trichromatic chlorophyll a calculated by the Parsons and Strickland (1963) equations using data
from the Ohio-NEFCO program have correlations with the trichromatic
chlorophylls of greater that 0.99 with a slope of 1. Herve and Heinonen (1982) also reported no significant
differences between the Parsons and Strickland (1963) and the
"Proposed Norsk Standard" chlorophyll equation, which is
identical to that for total chlorophyll pigments. Canfield (personal communication) has not found sufficient
amounts of phaeo-derivatives in Florida lakes to warrant reporting
anything except total chlorophyll pigments. In most northern Ohio lakes, correlations between
acid-corrected "chlorophyll a" and total chlorophyll
have a correlation coefficient of 0.96 (n = 88). If, on most occasions, neither interference by other
chlorophylls or by -derivatives significantly interfere with the
chlorophyll a determination, the designation "chlorophyll a"
could be used, but it is avoids confusion to use the term "total
chlorophyll," which implies no correction for chlorophyll or
phaeo-derivatives but also does not perpetuate the myth that
chlorophyll a can be accurately determined in natural water by
Considerations for Spectrophotometric Determination of Chlorophyll
The bandwidth of the spectrophotometer is important because the wider the
bandwidth, the lower the absorbance that is obtained (Weber, et al. (1986). This relationship
results from the rather sharp chlorophyll peak. On instruments with large bandwidth, the value includes a
greater amount of lower absorption values than would be obtained on an
instrument with a narrow bandwidth. Standard Methods recommends using instruments with bandwidth of
0.5 - 2.0 nm.
If phaeo-pigments are to be determined
spectrophotometrically, it is necessary to acidify the sample after the
first reading. It is
important to follow the Standard Methods procedures exactly. Any deviation of procedure at this step can produce the type of erroneous
chlorophyll a results mentioned earlier. The amount of acid that has been used in the past has varied
(1968) used several drops of 4M HCl in his original technique, but
Riemann (1978) found that such a strong acid causes spectral shifts in
the carotenoid, fucoxanthin, which increases in absorbance with
acidification and therefore lowers the acid ratio. This shift also increases the value at 750 nm. A strong acid will also convert phaeophytin, and chlorophyllides
to phaeophorbide, which results in an acid ratio greater than 1.7. This results in negative phaeo-pigment values and chlorophyll a values higher than total chlorophyll.
The amount of time between the addition
of the acid and the reading of the absorbance is critical. The conversion from chlorophyll to phaeophytin is a first
order reaction, the rate of which is dependent on pH. When strong acids were added, as used in the Lorenzen
technique, conversion was
instantaneous. The 90
seconds recommended in Standard Methods is necessary to complete most of
the reaction, yet avoid the interference of degradation products of
chlorophyll b, which increase at a much slower rate than those of
When exposed to blue light, chlorophyll
molecules will fluoresce brightly in the red region of the spectrum. Fluorometry is a highly sensitive method to determine chlorophyll
sensitivity can be of value in a volunteer monitoring program because
the volunteer will not have to to filter as much of a sample than is
necessary in spectrophotometric analysis. Even multi-chromatic fluorescence equations exist (Loftus and
Aside from sensitivity, however, there
is little to recommend fluorometry over spectrophotometry. There are no independent fluorometric chlorophyll attenuation
coefficients, and each individual fluorometer must be calibrated against
spectrophotometric standards. The
acid ratio for pure chlorophyll must also be determined for each
instrument. Weber et al.
(1986) also mention problems of the quenching of chlorophyll a fluorescence by b-carotene and other accessory pigments, an algal
species dependent relationship between extract fluorescence and
chlorophyll concentration, and the dependence of chlorophyll
fluorescence on temperature. High
chlorophyll concentrations will quench the fluoresence, thus requiring
the dilution of some samples. Marker et al. (1980) also discourage the use of the fluorometric
technique in freshwaters if an acidification step is used to determine
the phaeophytin by-product of chlorophyll b has a fluorescence
that overlaps significantly with that of phaeophytin a, therefore
producing high values for phaeo-pigments. For this last reason, Standard Methods (APHA, 1989) does not
recommend the acidification step in inland waters. Total chlorophyll pigments, as discussed above, could be
Based on the theory discussed above and
on the methods, observations, and comments by existing programs, the
following recommendations are made.
Samples should be filtered as quickly
as possible. In the
interim, the water samples should be kept cool and dark. The type of filtration apparatus can be left to the discretion of
the coordinator. Providing
a complete filtration apparatus has the single advantage of allowing the
volunteer to see the color of the filter and thus judge the amount of
algae to filter. Its
disadvantages are that it is expensive and it requires considerable
amount of manipulation and care by the volunteer. The advantages of using the Swinney filter holder is that it is
relatively inexpensive, requires no manipulation by the volunteer, and
the filtration is done immediately, therefore requiring no sample bottle
or sample preservation. Its disadvantage is that the volunteer does not open the
holder, and therefore cannot judge if sufficient sample has been
filtered. If the volunteer
does use care and dry the filter, there is a greater chance that the
filter holder will contain residual water which might promote growth or
degradation of the chlorophyll.
Keeping the chlorophyll molecule intact
until analysis is a concern. Most
programs immediately freeze the sample. Freezing seems to provide adequate preservation for several
months. Although mailing of
the frozen sample to the laboratory is done, experience suggests that
unless it can be assured that the sample is delivered within 48 hours,
there is a possibility of degradation. Immersing the filter in the solvent or drying the filter are
possible alternatives that should be given consideration. The safest method is to have the samples picked up by program
A number of combinations and variations
of the chlorophyll technique exist. The important fact is that the final chlorophyll value is highly dependent on the technique used. Without
some standardization, program to program comparisons of chlorophyll
values should be held as suspect. The
concept behind Standard Methods is to provide just that, a standard set
of methods for all analysts. The
chlorophyll method described in APHA (1989) is recommended. This technique, since it uses acetone as the solvent, probably
does not provide total extraction of chlorophyll from some algal cells,
but does provide some analytical consistency with historical data. Using a different extractant will undoubtedly change the
amount of chlorophyll extracted per unit biomass. It will therefore change any empirical relationships between
chlorophyll and other limnological variables such as phosphorus, Secchi
depth, etc. Changing
extraction solvents should be done only with the knowledge that
published empirical relationships may no longer be valid.
The trichromatic equations are not
recommended: they take
longer and can provide erroneous chlorophyll a values. At best, the chlorophyll a values are equal to those for
total chlorophyll pigments. Chlorophyll a values obtained after acidification can be reported but
remember that calling the phaeo-pigment corrected value
"chlorophyll a" does not make it so. It would be better termed "magnesium-containing
pigments." This value
is dependent on the technique and would be expected to vary widely from
procedure to procedure.
It is strongly recommended that the
total chlorophyll pigment be reported in addition to chlorophyll a. This value, although flawed by interferences by other
chlorophylls, phaeo-pigments, as well as a number of other possible
interferences, is the only value that remains fairly independent of
chlorophyll methodology. Therefore,
it is the only measurement that provides historical consistency. Chlorophyll a methodologies have changed over the past
25 years, and with each change, the previous chlorophyll estimates
became obsolete and non-comparable to the new methods. If everyone had reported total chlorophyll, at least there would
be one consistent value that would allow comparison. In a monitoring program, where historical data consistency is
absolutely necessary, this value should be reported.
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