134567
NCBE, University of Reading
Debbie Eldridge
Science and Plants for Schools, Homerton College
Hills Road, Cambridge CB2 8PH UK
Immobilised algae
Immobilised algae for studying photosynthesis
Aim
To provide a stimulating introduction to the quantitative study of
photosynthesis.
Introduction
Typical school investigations of photosynthesis include the evolution
of oxygen by Canadian pond weed (Elodea canadensis) or testing for
starch in Pelargonium leaves that have been decolourised. Both
methods are difficult to quantify, and the second method often
misleads students.
In this procedure, algae, immobilised in calcium alginate, provide a
standardised amount of photosynthetic material, enabling semiquantitative experiments to be undertaken. The rate of carbon
dioxide uptake by the immobilised cells is used to measure the rate of
photosynthesis — this can be done simply by observing the colour
change of hydrogencarbonate indicator, either by eye or using a
colorimeter. The effect of varying the intensity or wavelength of the
light may be studied, as can the effect of temperature or cell density.
Different species of algae or cyanobacteria may also be tested.
Scenedesmus quadricauda, an
alga which grows in clusters of
four cells, which is ideal for this
investigation.
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Immobilised algae
Equipment and materials
Needed by each person or group
Equipment
• 10 mL plastic syringe (without a needle)
• Small beakers or disposable plastic cups, 2
• 100 mL measuring cylinder
• Tea strainer
• Glass stirring rod or small plastic stirrer
• 5–7 mL bijou bottles, with caps, ~ 8
• Bench lamp
• Ruler
• Access to a colorimeter or a prepared set of standard solutions (see
details below)
• OPTIONAL: For investigating the effect of wavelength of light,
coloured filters (see Suppliers)
Materials
• 3 % sodium alginate solution, 3 mL
• 2 % calcium chloride solution, 100 mL
• Suspension of algae, e.g., Scenedesmus quadricauda, 50 mL (which
concentrates to 3 mL)
• Hydrogencarbonate indicator solution, ~ 45 mL
Needed by each class
Hydrogencarbonate indicator
Makes 1 litre of 10 x stock solution
•
•
•
•
•
Cresol red, 0.1 g
Thymol blue, 0.2 g
Sodium hydrogencarbonate (sodium bicarbonate, NaHCO3), 0.85 g
Ethanol, 20 mL
Freshly-boiled distilled water, ~ 1 L
1�
Dissolve 0.1 g of cresol red and 0.20 g of thymol blue in 20 mL of
ethanol.
2�
Dissolve 0.85 g of sodium hydrogencarbonate in ~ 200 mL of
freshly-boiled (and therefore CO2-free) distilled water.
3�
Add the ethanolic solution of cresol red and thymol blue and
dilute to 1 L with freshly-boiled distilled water.
For use, dilute this stock solution with nine volumes of freshly-boiled
distilled water and adjust the pH to ~ 7.4. Ideally, this solution should
be fully aerated before use so that it is a bright red colour.
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Note
Hydrogencarbonate indicator is
very sensitive to changes in pH,
and it is therefore important that
all glassware, etc is rinsed with a
little of the indicator before use.
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NCBE, University of Reading
Immobilised algae
To prepare a set of standard solutions
If a colorimeter is not available, the colour change of the indicator can
be semi-quantified by comparing it to a series of coloured buffered
solutions. Solutions ranging from pH 7.6–9.2 can be made using boric
acid-borax buffer.
•
•
•
•
10 mL vials with lids, 9
Boric acid, 12.4 g
Sodium tetraborate decahydrate (Borax, Na2B4O7 . 10H2O), 19.5 g
9 mL stock hydrogencarbonate indicator solution
1�
Dissolve the boric acid in a litre of distilled or deionised water.
2�
Dissolve the borax in another litre of distilled or deionised water.
3�
To 25 mL of the boric acid solution, add the volume of borax
indicated in the table below and make up to 100 mL with distilled
or deionised water.
4�
Place 9 mL of each of the prepared solutions into each of a series
of vials.
5�
Immediately before the lesson, add 1 mL of the stock (that is,
concentrated) hydrogencarbonate indicator solution to each vial.
Students may compare their results with the colours in the tubes.
Borax
solution, mL
pH
1.00
7.6
1.55
7.8
2.45
8.0
3.60
8.2
5.70
8.4
8.70
8.6
15.00
8.8
29.50
9.0
57.50
9.2
To culture the algae
•
•
•
•
2 litre PET lemonade bottle
Aquarium air pump
Aquarium airline tubing and airstone or glass sparger
Cotton wool or foam, ideally non-absorbent, for closing the top of
the bottle
• Low-temperature lighting, e.g., 18 W energy-saving bulbs
(equivalent to 100 W), 2
• Algal enrichment medium, 1 L (see Suppliers)
1�
Add 1.5 g of the enrichment medium to 1 litre of distilled water in
a 2 litre lemonade bottle and shake to dissolve. Some residual
powder will settle out, this is normal and will be utilised by the
algae as the soluble nutrients are depleted.
2�
Inoculate the bottle with the algae.
3�
Insert an airline to aerate the culture with an air pump. This will
provide extra dissolved carbon dioxide and keep the algae
circulating.
4�
Stopper the bottle with a loose cotton wool or foam bung.
Continuously illuminate the culture with a bright light while it is
growing. Small fluorescent strip lights or 18W low-energy lamps
give good results. The best results are obtained when extraneous
light in the laboratory is minimised.
CAUTION! Do not place hot lamps near to the culture.
Please also refer to the Safety guidelines.
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Copyright © NCBE, University of Reading, 2008
NCBE, University of Reading
Immobilised algae
Procedure
1�
Prepare a concentrated suspension of algae. There are two ways
to do this:
Either
Leave the 50 mL of dark green algal suspension to settle out
(ideally, overnight) then carefully pour off the supernatant to
leave approximately 3 mL of concentrate.
Or
Centrifuge 50 mL of the suspension at low speed for 5 minutes.
Pour off the clear supernatant to leave approximately 3 mL of
concentrate.
2�
Pour ~3 mL of the algal suspension into a small beaker and add
an equal volume of sodium alginate solution. Stir gently until the
algae are evenly distributed.
3�
Draw the algae/alginate suspension into a syringe.
Fig. 1
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Fig. 2
Fig. 3
4�
Place a beaker of calcium chloride solution under the syringe and
allow the algae/alginate mixture to drip slowly from the syringe
tip into the liquid below. Swirl the calcium chloride solution
gently as this happens.
5�
Leave the beads of immobilised algae to harden in the calcium
chloride solution for 5–10 minutes. The alginate molecules will be
cross-linked by the calcium ions, trapping the cells in a matrix of
calcium alginate.
6�
Separate the beads from the calcium chloride solution using the
tea strainer and gently wash the beads with cold tap water. Give
the beads a final rinse with distilled water. If they are illuminated
and not allowed to dry out, the beads may be kept for several
weeks. Alternatively, the beads may be kept in distilled water in a
fridge for several months. Beads removed from the fridge should be
allowed to warm up for about 30 minutes before use.
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Fig. 4
Fig. 7
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Immobilised algae
Fig. 5
Fig. 6
7�
Take several small bottles and rinse them with a small volume of
hydrogencarbonate indicator solution.
8�
Add equal numbers of algal beads to each bottle, then add a
standard, measured, volume of hydrogencarbonate indicator
solution. Replace the lid of each bottle. Approximately 12–15 beads
will be required in each tube.
9�
Place the containers at different light intensities by standing
them at various distances from a bright light. Leave them for 1–2
hours until the indicator in some of the containers changes
colour.
Fig. 8
Fig. 9
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NCBE, University of Reading
Immobilised algae
10� Determine the relative carbon dioxide concentration in each tube
with algal beads. There are two ways to do this:
Either
Match the colours of the hydrogencarbonate indicator in the
tubes with algal beads with a range of standard solutions;
Or
Use a colorimeter to measure the absorbance of the solutions at
550 nm (that is, using a green filter). Note: It is important to
measure absorbance (not transmission) in these experiments since
there is a linear response between absorbance and pH of the
indicator over the range studied.
11� Plot a graph to show the absorbance at 550 nm, or the pH of the
hydrogencarbonate indicator against the relative light intensity,
that is, 1 / (distance2).
Fig. 10a
Fig. 10b
Fig. 11
Safety guidelines
Cultivating the algae
The build-up of gas within the culture vessel, especially one made of
glass, could be dangerous. Consequently you should ensure that the
container used to cultivate the algae is adequately vented.
Care should be taken to ensure that lights used to illuminate the
culture and the aquarium pump cannot come into contact with liquid
should it spill or leak from the culture vessel. It is therefore a wise
precaution to place the culture vessel in a deep tray with sufficient
capacity to hold all the liquid in the event of a leak. Any electrical
equipment should then be placed outside this tray.
Chemicals
None of the chemicals used in this protocol are considered to be
harmful when handled as directed.
Lamps
When lamps are used to illuminate the algae, care should be taken to
ensure that they are not used in such a manner that there is a risk of
electrical shock or overheating that may cause burns or fire.
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Copyright © NCBE, University of Reading, 2008
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Immobilised algae
Preparation and timing
Cultivating the algae
The algae should be grown for 3-4 weeks then allowed to sediment
out before the lesson by leaving the culture to stand overnight.
Sodium alginate solution
The sodium alginate takes some time to dissolve, so the solution is
best prepared the day before the lesson. Ideally, leave the alginate
overnight to dissolve, preferably on a magnetic stirrer. Large volumes
can be prepared in a blender. Note that excessive heating can reduce
the strength of the alginate by depolymerising it. If you wish to store
sodium alginate solution for more than a few days, it is advisable to
autoclave it. To prevent excessive depolymerisation of the alginate,
however, you should increase the pH to 7–8 before autoclaving.
The practical protocol
Immobilised algae can be prepared by students in 15–20 minutes. It
takes a further 20–30 minutes to set up the experiment, then 1–2
hours for the indicator to change colour.
Troubleshooting
Because sodium alginate is difficult to dissolve, it may help to leave
the alginate to dissolve overnight. If you wish to try additional
activities using buffer solutions, please note that any containing
phosphate, citrate or EDTA should be avoided, as these will cause the
alginate matrix to dissolve. Should you wish to recover the algae from
the alginate beads, 50 mM sodium citrate or phosphate buffer at pH 7
can be used to dissolve the calcium alginate gel.
Additional investigations
The basic procedure described above can be extended in several
ways. Examples of open-ended investigations include:
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Light intensity
This can be achieved in two ways: either vary the distance from
the lamp or cover the containers with neutral density filters (see
Suppliers).
Wavelength of light
Use coloured filters, wrapped around each bijou bottle and taped
at the side furthest away from the light source. If students are to
analyse the results well they will need to be aware that different
light sources produce light of different wavelengths and they will
need to know what wavelengths of light are transmitted by each
of the filters.
Temperature
Set up water baths under a bank of lights and float the sealed
containers in the water with light shining from above.
Number of algal cells
Make several sets of algal beads with different numbers of algae
in them. This can be achieved by shaking the algal culture initially
and then using different volumes of culture to obtain the
sediment.
Copyright © NCBE, University of Reading, 2008
NCBE, University of Reading
Immobilised algae
Other sources of information
Eldridge, D. (2004) A novel approach to photosynthesis practicals.
School Science Review 85 (312) 37–45.
This article describes experiments that were carried out with
immobilised algae in more detail.
Smidsrød, O. and Skjåk-Bræk, G. (1990) Alginate as an immobilization
matrix for cells. Trends in Biotechnology 8 (3) 71–78.
Science and Plants for Schools
The original version of this protocol can be found at:
www-saps.plantsci.cam.ac.uk/worksheets/ssheets/ssheet23.htm
Further information on light absorption and filters is available at:
www-saps.plantsci.cam.ac.uk/articles/broad_light.htm
Suppliers
NCBE/SAPS photosynthesis kit
A pratical kit for carrying out this work is available from the National
Centre for Biotechnology Education, University of Reading, Earley
Gate, Reading RG6 6AU, UK. W: www.ncbe.reading.ac.uk
Sodium alginate
This may be purchased from school chemical suppliers. It is also used
in food production, so may be available from food industry sources,
although the viscosity of the preparations vary, so some
experimentation may be required.
Colorimeter
A suitable colorimeter for pupils is the CO7500 from Biochrom Ltd, 22
Cambridge Science Park, Milton Road, Cambridge CB4 OFJ, UK.
W: www.biochrom.co.uk. This colorimeter has a digital readout, it is
simple to use, reliable, robust and the results are repeatable.
Filters
Coloured and neutral density filters for studying the effects of
different wavelengths or light intensity may be obtained from Lee
Filters, Central Way, Walworth Industrial Estate, Andover SP10 5AN,
UK. W: www.leefilters.com. The colours shown in the table are useful
for this investigation. The Lee filters web site has information
concerning the properties of each filter. Note that several layers of a
single neutral density filter can be used to reduce the amount of light.
Algae
Scenedesmus quadricauda and other algae are available from Sciento,
61 Bury Old Road, Whitefield, Manchester M45 6TB, UK. This firm also
sells enrichment medium for cultivating algae.
Coloured filters
Colour
Deep Amber
Primary Red
Dark Blue
Dark Green
Primary Green
Light Red
Violet
Mikkel Blue
Bray Blue
Product code
(104)
(106)
(119)
(124)
(139)
(182)
(344)
(716)
(722)
Neutral density filters
Colour
Product code
0.15 ND
(298)
0.3 ND
(209)
0.6 ND
(210)
0.9 ND
(211)
1.2 ND
(299)
Acknowledgements
This practical protocol was developed by Debbie Eldridge from King
Ecgbert School, Sheffield through the award of a Schoolteacher
Fellowship funded by Science and Plants for Schools (SAPS) and
Robinson College, Cambridge. SAPS is funded by the Gatsby Charitable
Foundation.
This version of the protocol was produced by the Volvox project,
which is funded under the Sixth Framework Programme of the
European Commission.
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Copyright © NCBE, University of Reading, 2008