Biodiversity Research Project:
Identification and Collection Techniques
In today’s session, you will practice using the microscope, learn to identify and recognize some
common aquatic microorganisms, and learn some useful collection techniques. By the end of the
session, your team should have a well-designed project outlined, with an overall hypothesis,
experimental hypotheses, predictions, and methods for collecting data,
I. Identification of Aquatic Organisms
You may be wondering how the systematics and taxonomy exercises of our last session relate to
your biodiversity research project. In short: if you cannot identify organisms found in an ecological
survey, you cannot make relevant conclusions about why two habitats might differ in their biodiversity.
Identification is the first step towards determining the ecological tolerances of species present, and
linking those to the characteristics of the habitat in which they live.
Today you will practice observing common microorganisms with samples we have provided for
you. Some are pure cultures of a single species. Others are mixed samples of algae or protists, and
there are dichotomous keys in the lab that you can use to identify different species. These labeled
cultures are arranged on the back table.
A. Observing Microorganisms in Prepared Cultures
Take a concavity microscope slide (Figure 1) from the blue box on your lab table. These are the
thicker slides with circular indentations ground into the center. CONCAVITY SLIDES ARE
EXPENSIVE! PLEASE BE CAREFUL WITH THEM, AND DO NOT DISCARD AFTER USE!
Instead, rinse the slide well with tapwater, and dry with a Kimwipe before returning a used slide to its
storage proper box.
Figure 1. A concavity slide
To observe organisms, select a culture to sample, and draw up about 0.3 cc (one cc = one milliliter
(mL)) of liquid from the jar with a plastic pipet or 1cc syringe. (Note that you will find more organisms
at the bottom of the jar or associated with organic debris in the culture. Few organisms will be found
in the water column. (HINT: This also tends to be case in habitats you will sample in the field. )
Instructions:
1. Place 2-3 drops of your sample in the concavity on the slide.
2. Drop a coverslip onto the concave area, sealing in the liquid.
3. Place one small drop of methyl cellulose (available at your lab table) at the edge of the
coverslip, and allow it to diffuse under the coverslip. (This will slow down any rapidly
swimming microorganisms. Don’t drop methyl cellulose directly onto your sample in the
concavity, or you’ll just make a sludgy mess and be unable to see much.)
4. STARTING ON LOW POWER, observe your sample under the microscope. If necessary,
change to higher magnification only once you have focused your field of view on low power.
5. Begin at one corner of the coverslip, and gradually work your way across and down, in a zigzag pattern, until you have observed the entire coverslip field.
6. Whenever you find a motile (moving) organism (protist or animal), stop and identify it as
best you can by using the Identification Guide following this section and the other resources
available in the lab. You will find some excellent videos of common pond microorganisms at
these websites:
https://www.youtube.com/watch?v=Ln69k7LyTsU
http://www.microscopyu.com/moviegallery/pondscum/
http://tinyurl.com/tmhz
7. Record your results on the appropriate tally sheets provided.
8. Record the characteristics that allowed you to identify your organism in each case.
B. Organism Identification Guide
Since this is your first introduction to some of the vast diversity of Life, you might not be able to
identify the many organisms you see to the level of species. However, this guide should help you
narrow down the identification of the living organisms in your sample at least to the level of Phylum,
and possibly to an even less inclusive taxon.
Use all information sources at your disposal, from lab keys to online sources. Once you have
identified an organism as a diatom, for example, a Google image search might well yield a possible
identity for your organism. (Be careful. When it comes to protists, it sometimes takes a real expert to
tell them apart. We’ll be happy if you learn to tell a flagellate from a ciliate at this stage of the game.)
If you find something you can't identify, ask your instructor for help.
1. Protists
These are the simplest of the eukaryotic organisms, and they are a very diverse assemblage now
assigned to several different candidate kingdoms once subsumed under the now-defunct taxon
"Protista." The types you are likely to see today will be very small and usually highly motile. To see
them well, you'll have to use methyl cellulose to slow them down. Most common in daytime samples
will be diatoms and small flagellates. But the occasional ciliate or amoeba will show up.
A very nice image gallery, complete with taxonomic information and other facts about many
common protists can be found at http://megasun.bch.umontreal.ca/protists/gallery.html (Merci,
University of Montreal!)
2. Animalia, Porifera - The Sponges
The sponges are the simplest of animals, and they are found in both freshwater and marine
habitats. They are characterized by an amorphous body shape with no distinguishable head or tail
end. Lacking true tissues, these animals have an array of diversified cell types, each of which
performs a specific function. But you will not likely to be able to see individual cells in a sponge under
the microscope. It will just look like an amorphous blob with slightly greater degree of organization
than pond sludge. You can find a variety of freshwater sponge images as http://tinyurl.com/porif
3. Animalia, Cnidaria - Radially Symmetrical Diploblasts
Found in both freshwater and marine habitats, these animals are radially symmetrical (i.e., the
body is divisible into identical "pie shaped" wedges) and have two true tissue layers (endoderm and
ectoderm). The most common ones in freshwater will by hydras, and in marine habitats you may find
small medusae, the free-swimming “jellyfish” form cnidarians sometimes take. Check out some
images here:
http://tinyurl.com/pondhydra and http://tinyurl.com/fwmedusa
4. Animalia, Platyhelminthes - The Flatworms
If the body is dorsoventrally flattened (i.e., flattened from "top" to "bottom") and there is a distinct
head end that guides the animal's movements, there's a good chance you're looking at a flatworm. (If
you're not sure, call the instructor for a positive I.D.) These animals have three true tissue layers
(endoderm, ectoderm and mesodermal mesenchyme) and simple organ systems. View a flatworm in
action at http://www.youtube.com/watch?v=_jjzQrR5PLQ
5. Animalia, Rotifera - The Wheel Animalcules
These tiny animals are no bigger than a large protist, yet they have three true tissue layers and
complex organ systems. They feed by means of a cephalic (head end) corona of cilia which beats
food particles from the water into the mouth. They also use the corona for swimming; it pulls the
animal through the water like a little propeller when it decides to weigh anchor (pull up its sticky pedal
disk) and move.
Check out the dramatic video of rotifers at
http://www.youtube.com/watch?v=YF8OJt_pujc And yes, that’s the rotifers singing.
6. Animalia, Nematoda - The Roundworms
These worms are very thin, symmetrical, and tapered at both ends. There is no evidence of body
segmentation, and they move with a characteristic sinusoidal wave motion unique to this phylum.
This is because the body wall has only longitudinal muscles, another characteristic unique to this
phylum. Nothing else moves quite like a nematode, and you can see them swimming at
http://www.youtube.com/watch?v=SpgjnXEFadg
7. Animalia, Annelida - The Segmented Worms
The familiar earthworm is a member of this large, diverse phylum. You can identify a segmented
worm by the ringlike markings on its body, which delineate the body segments. Internally and
externally segmented, the body design and function is based on this characteristic metamerism,
which is found in many other more derived (i.e., not primitive) animal taxa. Paired bristles are a dead
giveaway that you’re looking at a segmented worm (either a polychaete or an oligochaete). Also,
they move with a characteristic “peristaltic” style, with alternat portions of the body constricting
Check them out at:
http://www.youtube.com/watch?v=X7O7UFOmRuk and
https://www.youtube.com/watch?x-yt-cl=85114404&x-yt-ts=1422579428&v=9Q9gh1k99rY
8. Animalia, Mollusca - The Mollusks
Closely related to the Annelids, the mollusks have secondarily lost their body segmentation,
though it is present in the larval forms you might see in your sample today. Mollusks can usually be
identified by the presence of a distinct head and a muscular foot, though if you happen to find a
bivalve, these features will be hidden inside the two shells. Find a gallery of larval mollusks here:
http://tinyurl.com/ku2hjux
9. Animalia, Tardigrada – The Water Bears
Closely allied to the arthropods, the tardigrades are microscopic, segmented, cuticle-covered cuties.
They can survive in the most extreme environments known, which makes them of great interest to the
folks at NASA. Check them out here: http://www.youtube.com/watch?v=6H0E77TdYnY
Watch the water bear meet a Paramecium at: http://www.youtube.com/watch?v=iLj4tBp00wo
10. Animalia, Arthropoda - The Arthropods
This is the most diverse of all animal phyla, with hundreds of thousands of species (The beetles
alone comprise more than 350,000 described species!). Arthropoda includes the familiar insects,
crustaceans, and spiders, as well as other less familiar forms. Like the annelids to which they are
closely related, the arthropods show distinct body segmentation. And if it has distinctly jointed
appendages, it's an arthropod. Some common ones you might find in your samples…
Copepods (adults and nauplius larvae): https://www.youtube.com/watch?v=Havd17RNo_c
Daphnia: https://www.youtube.com/watch?v=2g-04Uk0ut0
An ostracod: https://www.youtube.com/watch?v=4i4U1D89vVs
A mosquito larva: http://tinyurl.com/lkpgyle
A variety of crustacean larvae: http://tinyurl.com/mt73crz
11. Animalia, Echinodermata - The Spiny-Skinned Animals
Our closest invertebrate relatives that you might see today are the starfish and their relatives,
though you'll probably see only ciliated larval forms. Adults are pentaradially symmetrical. These
animals are strictly marine, and may not be present in either of your samples. Their lack of an
excretory system makes osmoregulation in freshwater or brackish water impossible for them, so you
will find them only in oceanic ecosystems. View a gallery of various echinoderm larvae here:
https://www.youtube.com/watch?v=p-9h2Jm2xPM
and
https://www.youtube.com/watch?v=3p37SvW9mso
12. Animalia, Chordata - The Chordates
This familiar group includes the sea squirts (Urochordata), the lancelets (Cephalochordata) and
the vertebrates (Vertebrata). All are united by the presence of a cartilaginous skeletal support rod
(the notochord) present at some time during development, a muscular, post-anal tail, segmentally
arranged muscle bundles (at least in development) and pharyngeal gill slits. The only kinds you're
likely to see today are fish or amphibians, if any. We just thought you'd like to know they're there.
IN THE COURSE OF COLLECTING SAMPLES FOR YOUR TEAM RESEARCH PROJECT, DO
NOT COLLECT ANY VERTEBRATES, SUCH AS TADPOLES OR FISH, AND BRING THEM TO
THE LAB. IF YOU FIND ONE IN YOUR SAMPLE, RECORD ITS PRESENCE IN YOUR SAMPLE
AT THE COLLECTION SITE, AND THEN GENTLY RELEASE IT.
By using the general identification guide above, other resources in the lab and online (Google
image search can be very useful!), you should be able to identify and differentiate many different
species within the listed taxonomic groups as you can when it comes time to survey the samples you
take for your research project. Today’s exercise should give you some practice at recognizing
common pond organisms.
The Secret to Success: Draw What You See!
It is often very helpful to DRAW A SIMPLE SKETCH of any and all organisms you have identified.
Bring unlined paper or a hardcover lab notebook for this purpose. Label it carefully with all pertinent
taxonomic information (and label anatomical structures as much as possible), and it will serve as a
reminder when you review your notes.
It really doesn’t matter if you feel you have no artistic ability. Just look carefully at the organism
and do your best to reproduce what you see. Use pencil so you can modify your drawing. A plastic
eraser (not the traditional pink kind) will be your friend, as it erases pencil without destroying the
paper. You’ll also be amazed at how much more detail you will notice if you try to draw an accurate
rendering of your little beastie.
II. Information for Planning Your Team Research Project
Once you and your team members have finished practicing identification of protists and using the
microscope, you should spend the rest of the lab period finalizing all the details of your research
project. This section of the manual provides some background regarding the methods you will use,
as well as a template you will provide to your Lab Instructor as a record of your plans.
A. Collection Techniques
For your research project, you will be collecting samples from aquatic habitats on or near campus.
Consider the question you will be asking, and be sure that you do not introduce unwanted variables
that could be controlled. For example, you should collect all samples at the same time of day, unless
you are studying changes in biodiversity over a diurnal cycle. And if you are studying changes in
biodiversity over something like a diurnal cycle, then you must carefully control all other variables,
such as location, volume of sample, type of sediment sampled, etc.
If there is time in today’s session, your lab instructor may take you to a local aquatic ecosystem
and demonstrate some common sampling techniques. Your group may choose to use the same
techniques, or you may devise a technique of your own. If you choose the latter, you must be certain
to keep things as constant as possible in such a complex system as an open habitat in the Great
Outdoors.
1. Recording Data
Whenever you collect a sample, you must record pertinent information about that sample, such as
1.
2.
3.
4.
5.
Locality (use Google Maps or other smart phone GPS application to obtain exact coordinates)
Time of day collected
Habitat type
Sample type (water column? Sediment?)
Other relevant details (point on transect, depth of sample, volume of sample, etc.)
Each sample will be stored in a collection vessel (see next section) that must be carefully labeled so
you can cross-reference each sample with its data. To this end, you will need a field notebook in
which to record all information about your samples. This can be as simple as sheets of paper on a
clipboard or as self-contained as a composition notebook or ring binder. But each team should
assign one member to record all data in its notebook. All members of the team should obtain
copies of all raw data, once all samples are collected.
Since field sites can be damp, pencil is often better than pen for field recording data. Keep
several handy, or at least have a pencil sharpener with you. (Do we really need to tell you this?)
Each sample cup (see below) should be labeled with an identifying code of your own design. For
example, if you are collecting sediment samples from Crandon Beach and comparing them to
sediment samples from Miami Beach, you might label your samples “CB1, CB2, CB3,” etc. and “MB1,
MB2, MB2,” etc.” Use wax pencil, Sharpie marker, or other water-proof ink to label your samples.
Each sample should have a matching entry in your field notebook with all pertinent data recorded
there. You need not record all the data directly on the cup if you have a complete record in your field
notebook. Your field notebook is as valuable as your samples: without it, your samples are useless.
2. Collection vessels and tools
We will provide your team with a set of six (6) plastic condiment cups with lids and plastic
disposable pipets (Figure 2) for each of your two localities (total of 12 cups per team). You will
take these to your two sample sites to hold your samples. Your task, over the next two weeks, will be
to collect six similar samples along a transect line (see below) in each of your two habitats.
Figure 1. Plastic condiment cup (a) and plastic pipet (b) are handy
field collection tools.
Your team will decide on the exact way to sample your sites. If you choose to survey the diversity
found in the “scum” (algal mats) on the edge of a pond, for example, you might simply collect a wad
of the algal mat by hand (use latex gloves, if possible, and wash your hands well after you have
collected your samples). If you are sampling the water column, you might use a plastic pipet (we
have these in lab for your use) or a section of a plastic drinking straw. The latter can be very useful
for sampling soft sediments.
Using a water-resistant marker, draw gradations on the straw that correspond to the depth of your
sample (increments of 1cc will work well) (Figure 2). Insert your straw to the desired depth, place
your finger on top of the straw to seal the top, and pull out your sample. To release your sample into
your collection cup, hold it over the cup and release your finger. If you wish to know exactly how
much sediment you have collected, empty the sample into a graduated cylinder first, and measure it.
Voila!
Figure 2. Mark your collection straw with gradations and use it to
sample sediment or water column (A). To release a sample into your
labeled storage cup, lift your finger from the top of the straw (B).
Pipets and straws are only two possible collection tools your team might use, and only these will
be provided for you. If your team wishes to use a different type of collection tool, then run your
methods by your Lab Instructor before you make a final decision. Ultimately, this project is of your
design, and we give you the freedom to devise techniques different from those described here if they
will work better for your particular project.
3. Sampling Techniques: Transects
Unlike research projects you may have done in the past, your survey of biodiversity will not involve
comparing treatment and control samples in which you manipulate a variable. Instead, you will be
comparing the biodiversity of two naturally occurring ecosystems that you predict will differ because
of some environmental factor (or possibly combination of factors, though further experimentation is
always necessary when more than one factor contributes to an outcome.)
Adequate sample size is critical to a good study. You will not get valid results if you take a single
sample from each of your areas of interest. Instead, you must take multiple samples from your area
of interest, in the hope of getting an estimate of its overall biodiversity. That is why we are providing
you with SIX cups for each locality.
A common method used by ecologists to sample diversity in natural habitats is to follow a
transect. This is a set path along which the investigator moves while collecting samples or counting
occurrences of a particular phenomenon (such as the occurrence of a particular species of organism).
At set intervals along the transect, a sample is taken at a measured distance from the center of the
path. If you know the length of your transect and the distance away from the line your sample was
taken, you can calculate a total area sampled, and should include this in your final report. There are
several different types of transects, but for our purposes, a line transect is probably the most useful.
A line transect (Figure 4) is a (usually) straight path marked by a measuring tape or string, along
which the investigator moves and counts particular items at known, set intervals. Because you will be
counting organisms that may not be readily visible to the naked eye, you will collect samples and
bring them back to the lab for analysis.
Figure 4. A transect line. Samples are taken at set intervals and known
distance from the line. This allows the investigator to calculate the total
area randomly sampled.
A transect line doesn’t have to be high tech. A pre-determined length of strong twine, marked with
distance intervals in indelible ink, will serve as well as a measuring tape. A meter stick can be used
to measure distance sampled from the transect line by placing it perpendicular to the transect line.
Your team will choose the distance from transect line to sample, and then collect either a water
sample or a sediment sample.
To collect along a transect line:
1. Stretch a transect tape or string along the area to be sampled.
2. Choose sampling intervals along the transect (e.g., every 0.5 meter).
3. At the first sampling point, place a meter stick perpendicular to the transect so
that you can record the exact distance from the transect your sample is collected.
4. Collect a sample with a pipet or other collecting tool. (Your instructor will
demonstrate and provide more details.)
5. Place your sample in a collection cup (we have provided very fancy plastic condiment cups
with lids) and label with all locality data. This can be obtained with a GPS application (e.g.,
GoogleMaps) on your smart phone.
6. In your field notebook, record all pertinent data, such as habitat type, type of sample (water
column? Sediment sample? Depth?), for each sample.
7. Repeat this procedure at every sampling point (for our purposes, three subsamples is probably sufficient for each measured point along the transect) until
you have completed the entire transect (Your team can decide how long the
transect should be. A total length of five to ten meters is probably adequate.)
4. Next Week: Bring Your Samples to Lab
Your team should plan to collect samples no more than a few hours before lab. Since some
organisms are more tolerant of disturbance than others, the composition of organisms in your sample
might change with time, as more sensitive organisms die. Once a sample is collected, keep its
temperature and lighting conditions as close to those at the sampling site as possible until you bring it
to lab for analysis. You will have two weeks for data analysis. If your two sites are distant from each
other, it will be most efficient to sample and analyze one locality the first week, and the second
locality the next. Remember to keep everything as constant as possible, sampling your two localities
at the same time of day, etc. If there are differences in weather or other environmental conditions
between the two localities, be sure to record those in your field notebook, in case you wish to refer to
them to help explain your results. In short, the ecologist records just about everything that can
affect the nature of collected data.
In lab, each team member will take sub-samples from each of the samples in your condiment cup
and observe them under the microscope, just as you did with the known samples in today’s lab
session. As a starting point, we suggest that each team member do at least two biodiversity counts
from each of the six samples taken at a given locality. These are not replicates! Each sample is
ONE replicate from your chosen locality. By having each team member do at least one count, you
are simply making sure that your sample is well analyzed, since the more of it you look at, the more
likely you are to find all the species hiding there. Therefore, you will pool the data from the counts
that each team member records from each of the six sample cups. At the end of your analysis, you
should have a biodiversity count for six different samples from each locality, and each of those
samples will have been subject to eight counts (two by each of four team members).
Whatever numbers your team finally decides to use, it is important that both localities are
analyzed in parallel fashion, with the same number of samples and sub-sample counts by all team
members.
A sample tally sheet that each team member can use to record species found in a sample count
can be found at the end of this chapter. You may use this form, or create one yourself, if you feel you
can make one more suitable to your team’s needs. If you are able to identify species to less inclusive
levels than those included on the tally sheet, either provide a spot for that on your own tally sheet, or
record the information in your field notebook.
III. Outlining Your Team Research Project
Now that you have explored the literature on biodiversity, learned how to classify and identify
organisms, and know some of the ways to sample an aquatic environment, your team should be able
to outline a complete protocol for the next two lab sessions.
Refer to the first chapter of your online lab manual, which contains information about
environmental factors to consider, how to measure biodiversity, etc. Linked to the lab syllabus is a
Word document that your team must complete and turn in (electronically) to your Lab
Instructor before the end of today’s session.
Before you leave, ask your Lab Instructor or your Undergraduate TA to inspect your lab station
and microscopes for cleanliness. Any team leaving before having their station inspected will suffer
the loss of 5 points from each team member! Part of good science is good lab technique, and that
starts here. Once you have completed the protocol document and cleaned up your lab station,
you’re done for the day.
Species Diversity Tally Sheet:
Team:
Location:
Sample:
Transect measurement:
Taxonomic
Group
Protist
(diatom)
species 1
species 2
Protist
(flagellate)
Protist
(amoeboid)
Protist
(ciliate)
Porifera
(sponge)
Cnidaria
(cnidarians)
Platyhelminthes
(flatworm)
Rotifera
(wheel
animalcules)
Nematoda
(roundworm)
Annelida
(segmented
worms)
Mollusca
(mollusks)
Arthropoda
(arthropods)
Chordata
Other
Total number of different species:
species 3
species 4
species 5
species 6
Total #