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Frequently Asked
Questions About Fluorometric Chlorophyll Analysis
Q: Why measure chlorophyll?
A: All plant life contains the primary photosynthetic pigment chlorophyll
a. Microscopic, planktonic plants, or phytoplankton, occupy the
lit zone of all water bodies. With over 70% of the surface of the earth
covered in water, phytoplankton and photosynthetic bacteria are responsible
for almost ½ of the planets primary production while their total
biomass comprises less then 1% of the total plant biomass. These extraordinarily
efficient plants also act as the single largest CO2 sink on
earth. For these reasons alone it should be clear that there is an interest
in measuring concentrations of phytoplankton. Chlorophyll a fluorescence
is the most versatile, sensitive and easy way to measure the concentrations
of phytoplankton in water.
The quantitation, through extracted analysis,
or estimation, through in vivo analysis, of chlorophyll a concentration
supplies information on the abundance of phytoplankton present in all
aquatic environments. Since chlorophyll-containing organisms are the first
step in most food chains, the health and /or abundance of these primary
producers will have cascading effects to all higher organisms. Therefore,
the determination of chlorophyll concentration is one of the key indices
in monitoring the health of any natural system.
Chlorophyll measurements are also used
to directly monitor phytoplankton populations. Examples include, but are
not limited to, the monitoring of chlorophyll in natural marine and freshwater
environments, reservoirs, water and sewage treatment plants, and aquacultural
systems.
Q: How do fluorometers detect and quantify chlorophyll a in
water?
A: Fluorescence is the phenomena of some compounds to absorb specific
wavelengths of light and almost instantaneously emit longer wavelengths
of light. Chlorophyll a naturally absorbs blue light and emits,
or fluoresces, red light. Fluorometers detect chlorophyll a by
transmitting an excitation beam of light in the blue range (440nm for
extracted analysis and 460nm for in vivo analysis) and by detecting
the light fluoresced by cells or chlorophyll in a sample at 685nm (red).
Generally, this fluorescence is directly proportional to the concentration
of the material in question.
Q: What is the difference between
in vivo, in vitro , and extracted chlorophyll analysis?
A: In vitro (meaning 'in glass' and referring to 'in an artificial
environment or outside the living organism') chlorophyll analysis is another
term for extracted analysis. It entails the concentration of chlorophyll
containing cells onto a filter followed by the extraction of the chlorophyll
a from the cells. In vivo (meaning 'within a living organism')
chlorophyll analysis simply refers to the analysis of chlorophyll in the
natural environment or, in our case, in the living algal cells.
Q: What is in vivo chlorophyll analysis?
A: In vivo chlorophyll analysis is the fluorescent detection
of chlorophyll a in living algal and cyanobacterial cells in water.
In this technique, the excitation light from the fluorometer passes through
the untreated sample water and excites chlorophyll within the living cells
of the algae present. There are several factors that make in vivo
analysis a semi-quantitative measure at best. Environmental parameters,
physiology, morphology, light history and the presence of interfering
compounds all play a role in altering the relationship between fluorescence
and the concentrations of chlorophyll a. Examples of interfering
materials include other plant pigments, degradation products, dissolved
organic matter, and turbidity. In vivo fluorescence data supplies
information on the relative distribution of chlorophyll concentrations
and usually correlate well with extracted chlorophyll a samples.
In vivo detection has several very
useful applications. An example is the monitoring of general trends in
chlorophyll concentrations in real time. It is very easy to obtain large
amounts of data using in vivo instrumentation and is an excellent means
of following trends and estimating chlorophyll concentration. With the
introduction of the SCUFA®
submersible fluorometer, verticals profiling and mooring applications
are now possible. Other examples of in vivo applications include
continuous monitoring along a ship's track using the 10-AU
configured with a flowcell and discrete sampling used to monitor algal
concentrations in natural or laboratory phytoplankton populations. Examples
of discrete sampling applications include aquaculture and hatchery systems,
water treatment facilities, reservoir monitoring, and aquatic research
(see the Aquafluor™).
If water samples are taken, the in
vivo data can be correlated to extracted chlorophyll a data
to estimate actual concentrations. Otherwise, the in vivo data
can be used as a relative measurement to identify trends and patterns.
Q: How do you calibrate a fluorometer
for extracted chlorophyll a analysis?
A: Allow the fluorometer to warm-up for the time specified in the
User's Manual. Measure the fluorescence of each standard at sensitivity
settings that provide mid-scale readings (refer to your User's Manual
for proper calibration procedures). Follow directions under section 10.0
from E.P.A. Method 445.0
(Revision 1.2) for the calibration and standardization procedure using
the traditional acidification technique or the non-acidification method.
E.P.A. Method 445.0 calls for filtering onto glass fiber filters (GFF)
filters and grinding of the filters. This step may not be necessary in
some systems and tests should be run to compare extractions with and without
grinding. Non-grinding techniques can use either GFF or membrane filters
that will dissolve in the solvent.
Q: What environmental factors interfere with in vivo chlorophyll
analysis?
A: Light, temperature, water quality, and dissolved components can
all have significant effects on fluorescent readings independent of the
chlorophyll concentration. However, all of these factors can be controlled
and/or corrected to a degree if the user is aware of their effects.
Temperature has an inverse relationship
with fluorescence. For example, in a vertical profile, as the temperature
decreases, the fluorescence will increase independent of chlorophyll concentration.
The in vivo chlorophyll fluorescence response changes at a rate
of 1.4% per °C. A temperature drop of 10 °C in a vertical profile
would result in a 14% overestimation of chlorophyll at the coldest point.
Turner Designs' field and submersible
instruments have the capability to automatically compensate for temperature
effects.
Light history can have significant affects
on the fluorescence in algal cells. For example, at low light levels,
algal cells can optimize the light uptake by pushing chloroplasts to the
outer edge of the cell or by producing more chlorophyll per cell. Both
of these responses can result in data increases the fluorescence signal
while the algal biomass may be unchanged. To lessen the effects, opaque
hose should always be used when sampling natural waters with a field fluorometer.
The transport time of the water in the hose will dark-adapt cells to an
extent, significantly reducing fluorescence error caused by variations
in the light history of the cells.
Dissolved organic matter (DOM), chlorophyll
degradation products (pheophytins), chlorophyll b & c
and turbidity can also falsely increase the chlorophyll a fluorescence
signal. If the dissolved interfering compounds are suspected to be significant,
it is worth conducting a quick study to look at the effects by comparing
the fluorescence from filtered and non-filtered water samples or from
noting the fluorescence signal below the photic zone and using this value
as a blank.
Q: What is the effect of varying species on fluorometric chlorophyll
analysis?
A: Different species of phytoplankton have varied morphologies (cell
packaging), physiological states, size, and chlorophyll a : carbon
ratios. All of these factors can affect the fluorescence emitted from
a cell under a specific excitation light intensity. Luckily, in natural
environments, the phytoplankton assemblage is diverse enough that much
of the variation in fluorescence resulting from the factors above balance
eachother out so the net effect on the fluorescence reading is reduced.
Nevertheless, the researcher needs to be aware of the potential for error
in readings caused by these factors to better interpret in vivo
data.
Q: How does other chlorophylls and degradation
products affect extracted chlorophyll analysis?
A: All chlorophyll pigments and their degradation products (pheophytins)
have their own unique excitation and emission spectra. Unfortunately,
these spectra overlap significantly with the fluorescence spectra of chlorophyll
a due to the similar chemical structure (see Figure
2). Due to the overlap in fluorescence spectra, the presence of one
can result in an interference in the measurement of another, resulting
in an under or overestimation of the pigment in question.
In the case of chlorophyll a, interfering
pigments to be aware of are chlorophyll b, chlorophyll c, and pheophytin
a (see figure 3). Chlorophyll b causes
the most interference in freshwater systems with high concentrations of
chlorophytes and/or prasinophytes and in marine systems with high concentrations
of prochlorophytes. In chlorophyll extraction, the interference results
during the acidification step of the traditional extraction technique.
Chlorophyll b undergoes a wavelength shift when acidified, resulting
in an underestimation of chlorophyll a and an overestimation of
pheophytin. In environments with high chlorophyll b concentrations,
we strongly recommend using the Welschmeyer (non-acidification) method.
High concentrations of chlorophyll c
can result in a slight overestimation of chlorophyll a and an underestimation
of pheophytin a sometimes even resulting in negative pheophytin readings.
It has been reported that a chl a : chl b ratio of 1 : 1
, which is the highest ratio which could occur in nature, would result
in a chl a overestimation of 10%.
High concentrations of any of the interfering
compounds will result in an increase to in vivo chlorophyll readings.
The magnitude of the increase will depend on the instrumentation used.
Instruments using filters with a wider bandpass will be more effected
by interfering compounds than instrument with narrower bandpass filters.
Q: What is a secondary standard?
A: A secondary standard is used as an alternative to a primary calibration
standard. It is often used when primary standards are expensive, difficult
to obtain, or unstable. In the case of chlorophyll a, a secondary
standard can be used the majority of time for calibration because liquid
chlorophyll a standards are expensive, time-consuming, and photosensitive.
To properly use a secondary standard, you must first calibrate with a
primary liquid standard of the fluorophore of interest. You can then obtain
the equivalent value of the secondary standard. Henceforth, you may calibrate
using the secondary standard using the value you obtained for it initially.
An occasional calibration using a primary standard to recheck the stability
is recommended.
In the past, secondary standards have been
more stable than the primary standards they mimic, but have still required
special storage and handling conditions with relatively short lifetimes.
Examples of these include coproporphrin and fluorescent dyes such as Rhodamine
WT.
Turner Designs has developed a solid
secondary standard that is stable under ambient light and temperatures
with no special treatment or storage required. This new secondary standard
will greatly reduce time, cost, and trouble in fluorescent chlorophyll
analysis procedure.
Q: What is the best solvent and procedure
for the extraction of chlorophyll a?
A: The most commonly used extraction solvent is a 90% acetone 10%
DI water solution. Other solvents, such as methanol, ethanol and acetone/DMSO
mixtures are also commonly used and can improve extraction efficiency
with specific phytoplankton or may be found useful for the extraction
of sediment samples.
There is no 'best' solvent or procedure
for chlorophyll extraction. Several work well and have their own pros
and cons. The E.P.A.
Method 445.0 describes the recommended step-by-step process for analysis
using 90% acetone. There are many factors in the extraction process that
can lead to different results. Several examples of these factors will
be discussed below in hopes reducing some of the variability.
An excellent resource for sample collection,
storage and extraction methods is the UNESCO publication, Phytoplankton
Pigments in Oceanography.
Water collection, storage, and filtration:
Because pigment is being extracted from living cells, it is critical to
use consistent techniques of obtaining water, filtering and storing filters.
The living cell is sensitive to changes in the environment such as temperature
and light. Conditions leading to cell death or damage will affect chlorophyll
concentrations.
In the collection of water samples, it
is important to make certain that the collection containers are clean
of all chemicals. They should be rinsed several times in the sample water.
Once collected, if samples cannot be filtered immediately, they should
be stored quickly on ice in the dark. The time between collection and
filtration should be as brief as possible and should not exceed 4 hrs.
Specifics on the recommended filtration
and storage procedure can be found in E.P.A.
Method 445.0. An excellent resource for sample collection, storage
and extraction methods is the UNESCO publication, Phytoplankton Pigments
in Oceanography.
Q: How do I take and store discrete water samples in the field?
A: For discrete in vivo analysis, water samples should be measured
as soon as possible after collection. The same time constraints should
be placed on the filtering of water samples that are to be used for extracted
analysis. From the time of collection to measurement, the samples should
be stored in the dark on ice. Remember that the cells are living and significant
time in a container will alter the physiological state of the algal cells,
resulting in misrepresented chlorophyll data compared to the natural situation.
Discrete samples need to be kept at the
same temperature. This is most easily accomplished through the use of
a water bath. The bath should be covered from direct light. When the samples
are being measured in the fluorometer, a 'time in the instrument' must
be established. Use the discrete sample averaging function on the 10-AU
or TD-700 Fluorometer or wait for the reading to stabilize (~10 seconds)
and record the fluorescence. If this time is not monitored the heat and
light in the instrument will cause fluorescence to change.
When developing you own sample collection
and storage procedures, it is recommended to run your own experiment to
test change in fluorescence over a given transport time by analyzing samples
from a given sample at hour or half-hour intervals.
Q: How do I calculate actual chlorophyll
a concentrations from my fluorometric data?
A: ACIDIFICATION METHOD
Prior to running sample on the fluorometer,
the instrument must be calibrated with a pure chlorophyll a standard
and the maximum acid ratio must be determined by measuring the fluorescence
of the standard before and after acidification. If a fluorometer other
than a digital Turner Designs instrument is being used, the fluorometer
sensitivity coefficient may also need to be determined. For further information
please refer to EPA
Method 445.0.
chl a =K (Fm/ Fm
-1) x (Fb-Fa) x (v/V)
pheo a =K (Fm/ Fm -1) x [(Fm x Fa
- Fb)] x (v/V)
If necessary, the result can be multiplied
by a dilution factor.
where:
K= sensitivity coefficient, equal to 1 on 10-AU
Fm = max acid ratio Fb/Fa of pure chlorophyll a standard
Fb = fluorescence before acidification
Fa = fluorescence after acidification
Fo = fluorescence signal of sample
v = extract volume (L)
V= volume filtered (L)
NON-ACIDIFICATION METHOD (Chlorophyll a
concentration only)
1) Collect fluorescence data (one number/sample)
** DO NOT ACIDIFY**
2) Plug data into following equation:
chl a = (Fo x v)/ V
where:
Fo = fluorescence signal of sample
v = extract volume (L)
V= volume filtered (L)
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