Algal Microcosms
Concepts: Designing and conducting a scientific investigation requires
introduction to the major concepts in the area being investigated, proper
equipment, safety precautions, assistance with methodological problems,
recommendations for use of technologies, clarification of ideas that guide the
inquiry, and scientific knowledge obtained from sources other than the actual
investigation .(NSES)
Objective:
1. Students will participate and apply the processes of scientific investigations
to create models and design, conduct, evaluate and communicate the results
of these investigations.
2. Students will develop oral and written presentations using clear language,
accurate data, appropriate graphs, tables, maps, and available technology.
3. Students will draw logical conclusions based on scientific knowledge and
evidence from investigations.
Summary:
Algae respond (positively and negatively) to a myriad of environmental conditions
(i.e. pH, current velocity, temperature). To introduce students to the response of
algae to the environmental variables, students will be designing an experiment in
which they will measure the growth of algae in tanks exposed to different amounts
of light. The students will keep accurate records in notebooks and utilize the data
sheet provided. Students will make several algal counts and record findings in
their notebooks. Each group will use the information collected from observations
to create a presentation of their findings.
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb
Teacher Notes:
This lesson consists of many parts and is an integral part of the module. The
students will be doing the following:
1. Setting up a notebook to record all observations and data
2. Designing an experiment to determine which environment produces the most
algae based on the amount of light to which the algae is exposed
3. Create a presentation to share the findings of their experiment with the class
4. It is important to note that this will be an integral part of the next module,
“Culturing Algae”, and the students will need to complete this experiment to
be able to continue experimenting with other variables in the module
“Culturing Algae”.
Equipment/Materials:
1. 3- 2.5 gal. aquaria per group
2. Substrate for the bottom of each aquarium (cleaned gravel or sand)
3. Distilled or RO water
4. Algal samples to stock the aquaria with
5. Sampling supplies: glass slides, Pasteur pipettes, toothbrush, rubber policemen
etc.
6. Air pump with tubing and air stones (diffusers)
7. Full spectrum fluorescent light units with timers
8. Plastic cover slips
9. Compound microscope
10.Counting chambers
11.Small glass containers (finger bowl)
12.Graduated cylinder or small beaker
13.Water bottle
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb
Useful Websites:
Activity Sheets:
Data Sheets
Student Notebooks
Engage:
1. View the Scientific Method (2:43)
http://www.youtube.com/watch?v=jPaGOHwv7mQ
2. Let students brainstorm how they will design and set up and experiment to
determine in which light conditions algae grows best.
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb
Explore:
1. Each group will take 3-2.5 gallon aquaria and fill the bottom with
approximately one-half inch of sand or gravel (easier to manipulate).
2. Use RO or Distilled water to fill the aquaria to roughly ¾ full. Tap water could
be utilized, but let it sit for 34-48 hours to make certain that the chlorine residue
has dissipated.
3. Carefully position 10 glass slides on the bottom of the tank. They should be
resting on top of the gravel/sand.
4. Place the tubing and air stone into the tank and turn on the attached air pump to
provide gentle agitation of the water. This will prevent stagnation of the water.
5. Place the class’ aquaria into four different groups and position the light units
over top, making sure aquaria from the different groups are not too close to
other groups (thereby distorting the light cycles).
6. The light timers of the four groups will be set to one of the following cycles:
a. 12 hours of light followed by 12 hours of darkness (Fall/Spring equinox)
b. 16 hours of light followed by 9 hours of darkness (Summer solstice)
c. 9 hours of light followed by 16 hours of darkness (Winter solstice)
d. Continuous light 24 hours a day
e. Complete darkness could be an alternative fifth setup; however algal
growth in this unit will be minimal (if at all). The instructor may choose
to set up a demonstration tank in total darkness for comparison with the
class.
7. Inoculate each tank with algae from a common “cocktail”. The cocktail may be
generated from the various algal cultures used in the survey laboratory OR from
the field samples collected by the students.
a. Desired algal specimens should be deposited in a large beaker
b. Agitate the solution to homogenize and create your algal cocktail. This
may be done in a low tech manner by simply mixing with a glass rod or
by using a blender.
c. Inoculate each aquarium with set volume of the cocktail (~25-50 mL
should be adequate).
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb
Explain:
1. Students explain the process they used for setting up their tanks.
2. Share how they will be able to determine the growth of algae in each tank.
3. Explain the importance of keeping accurate notebooks which include the
date, descriptions, illustrations, and data collected from each observation.
4. Go over the Data Sheets and review each section.
5. Set up notebooks.
Extend:
Algal Sampling Protocol:
1. The tanks should be visually observed with the naked eye on a daily basis.
2. Once a week the student groups should:
a. Extract one of the glass slides out of their aquarium.
b. Scrape the upper surface of the glass slide with a toothbrush or rubber
policeman.
c. Using a water bottle filled with distilled or deionized water, rinse the
upper surface of the slide and scraping device into a small container
(i.e., finger bowl).
d. Measure the volume of the rinsed sample using a graduated cylinder
or beaker.
e. Using a Pasteur pipette, extract a small portion of the scraped sample.
f. Place a drop of two of the sample into the well of a counting chamber.
3. Each group should count and identify approximately 100 algal cells from
each sample.
a. Sketches, measurement, and notes should be made of each type of
algae.
b. Some students may struggle with the classification of algae. In this
case, have them group the algae into shape and color classes (e.g.,
green unbranched filaments, unicellular golden cells). Still maintain
descriptions and measurements. They may be able to make favorable
identifications at a later point in time.
c. Advanced students may attempt to quantify the density of algae
growing on their slides by using the equations listed in the Teacher
Background section.
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb
Evaluate:
1. Collect student notebooks and check for detailed descriptions and data
sheets.
2. Final presentation* using presentation rubric from Algae Research Project
*Presentations:
Presentations for Algal Microcosms should include information collected from
tank observations recorded in student notebooks. Students should compile data
from observations and graph data collected to inform others about the effect of
light on algae growth in their tanks. Presentation should include a visual display of
the information collected (illustrations of types of algae, location of tanks, etc.) and
charts and graphs to demonstrate the type and number of algae found throughout
the investigation.
Since all of the groups have basically the same design for this experiment, only a
brief discussion is necessary. Each group should explain their unique findings, in
detail, using charts and graphs and other methods to illustrate findings.
Enrichment Activity:
Teacher Background:
Algal Counting and Quantification: Counting Algal Cells
Algae can be counted in a manner that allows the student to estimate the density of
cells (either per volume or surface area). Measure the volume of the sample in the
laboratory and pipette a small subsample into a Palmer-Maloney nanoplankton
counting chamber. The volume seen in a single microscope field of view (Vf) can
be calculated using the formula:
Vf = πr2d
Where r is the radius; f the microscope field; and d is the depth of the counting
chamber.
It is necessary for the student to count (enumerate) 100 algal cells. The density of
any specific alga can be generated with the formula:
Di = N1Vs/VcA
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb
Where:
Di is the density of cells of the ith taxon
Ni is the number of cells of the ith taxon counted in the counting chamber
Vs is the total volume of the sample
Vc is the volume of the samples counted (fields of view counted x Vf)
A is the area in cm2 of substratum sampled. A typical glass slide measures 3
inches (7.62 cm) long by 1 inch (2.54 cm) wide. This results in a total sampled
area of 3 in2 or ~19.36 cm2.
Example:
A student has finished counting 100 organisms scraped from their glass slide.
His/her results are as follows:
Chlamydomonas: 15
Euglena: 10
Spirogyra: 50
Navicula: 25
Gomphonema: 5
Field of View: 2 views under the 40x objective
1. Calculate the volume of a microscope view under 40x
a. Vf = πr2d
i. π – 3.14
ii. r2 - 0.52 mm
iii. 1.5 mm
iv. Vf = 1.1775 mm3 (0.0011775 mL)
2. Calculate the density for each taxon.
3. Di = N1Vs/VcA
a. For Chlamydomonas:
i. N1: 15
ii. Vs: 40 mL
iii. Vc: 0.0011775 * 2 = 0.0035325 mL
iv. A: 19.36 cm2
v. Di for Chlamydomonas on the slide: ~3,761 individuals per cm2
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb
Counting chambers are expensive, but homemade versions can be constructed at
minimal cost. A hole can be punched through a plastic cover slip (industrial hole
punch or a cork borer will work). The cover slip can then be attached to a glass
slide using an appropriate adhesive paste. The depth of the well and radius can be
calculated using precision calipers.
© 2010, Ohio Northern University: Dr. Kenneth Reid, Dr. Debra Gallagher and Dr. Robert Verb