Download introductory lab - University of California, Santa Cruz

INTRODUCTORY L AB
Objective
In this lab we will cover logistics and introduce techniques for successful examination,
preservation, and identification of algae.
Notebook Requirements - 2 charts & 5 drawings
1) 2 microscope calibration charts
2) Mazzaella flaccida: 2 drawings (thallus & cross section)
3) Ulva: 2drawings (thallus & cross section)
4) Unknown: 2 drawings (thallus & cross section) & steps to key out
A) Logistics
 Introduction
 Seymour Center Rules
 Lab Orientation
 Lab Safety
 Clean up
 Field Trip Waivers
 Check-out Plant Presses
 Check-out Microscopes – get both scopes out, check if they are both functional (i.e. plug in and
turn on lights – make sure they work). Keep both scopes out for microscope calibration.
B) Microscope Calibration
In this lab course you will be using a dissecting microscope and a compound microscope. The
dissecting microscope is useful for observing macroscopic parts of the algal thallus. The compound
microscope is used to view the internal structure of an alga. You will need to calib rate both
microscopes at various magnifications, so that you can measure objects in the field of view.
Dissecting Microscope
 Focus your dissecting scope on a small ruler and measure the diameter of your field of view
(use cm or mm units)
 Repeat measurements for each magnification
 Record measure ments for each magnification in your lab notebook:
Magnification
Diameter of Field of View
Compound Microscope
 The eyepiece of your compound scope has an ocular ruler with no known units and it is your
task to determine how many micrometers (µm) are equal to an ocular unit.
 Look through your ocular lens and observe the ocular ruler, a small ruler in the eyepiece.
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Get a stage micrometer from your TA and place it on the microscope stage on the lowest
objective lens (4x or 10X depending on your scope).
Now line up the first line of the ocular ruler with the first line of the stage micrometer.
Find another place farther down the micrometer where different points from each ruler line up.
(hint: the further apart the two points are, the more accurate your conversion factor will be)
Count the number of stage micrometer and ocular units that fall between the two points, and
divide those to find the ratio between them (see below).
Use the ratio of the stage micrometer (S) to ocular units (O) to find the true length of one ocular
unit at each magnification.
(Specimen Measurement) ocular units
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(S)
µm
= ? µm
(O) ocular units
In the future you will make all specimen measurements with your ocular ruler and then
multiply that value by your S/O ratio to report specimen measurements in micrometer units
(µm).
Record the chart below into your lab notebook so that you can give an accurate scale for the
objects you draw. Be sure to record which microscope you are using, as the calibration will
differ slightly for each individual microscope.
Objective Lens
Stage Micrometer (S)
Ocular Micrometer (O)
S/O
4X or 10X
C) Cross Sectioning Algae
Some algal features may only be seen in a squash or in a thin cross-section. To make a squash,
simply smash a small piece of an alga between a slide and a cover slip. To make thin sections, you will
need some tools: razor blade or scalpel, forceps, probe, glass slides, and cover slips. Ideally, thin
sections are only one or two layers of cells. Here’s how to make thin sections thin:
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First, cut off a manageable piece (1-2 cm2 ) of an alga and position it on a glass slide.
Use another slide to hold down the alga firmly.
Angle a sharp razor blade or scalpel away from the
top slide (see diagram below). Carefully pull the
razor blade along the edge of the top slide, slicing the
alga.
Angle the razor blade perpendicular to the slide and
slice the alga again.
Make a third slice, this time angling the razor blade
towards the top slide.
Move the top slide very slightly to expose the edge of
the alga. Repeat the above motions, making many
thin sections. Discard any large pieces.
You may need to turn the sections on their sides. To
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do so, add some salt water to the slide and nudge the sections onto their sides using the tip of
the razor blade, probe, or forceps.
Practice making thin cross sections of Laminaria sp.
Have your TA check to see if your thin section is indeed thin enough before you make your
drawing.
Draw the macrothallus and thin section of Laminaria sp. in your lab notebook.
razor blade
cut 1......2......3
specimen
thin sections
top slide
bottom slide
Side View
D) Staining Thin Sections
Staining is a useful tool when you need to examine objects that are difficult to resolve, such as
chloroplasts, pyrenoids, or cell walls. Different stains are used for different purposes. Your TA will
demonstrate proper staining technique.
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Prepare a thin section on a slide of the alga you would like to stain, as describ ed above.
Add ONE drop of stain.
Allow the stain to sit on the specimen for a minute or more, depending on the stain used.
Place a paper towel at one edge of the slide and let it slowly absorb the stain, drawing the stain
across the slide.
Rinse the sample, using seawater, by adding drops at one end of the slide while drawing liquid
away at the other end with the paper towel. Continue until the stain is rinsed away.
Add a few drops of fresh seawater around the edge of the cover slip. The seawater should flow
under the cover slip without making bubbles, and you can now view your stained slide.
coverglass
alga
eyedropper
with stain
absorbent paper
slide
Surface View
These images depict how stains are added to a sample.
Staining Specimens
Stain
Interacts with
Analine Blue
Cellu lose cell walls
Cytoplasm
Achromatic figures
IKI
Starch
Malachite Green
Cell walls
Cytoplasm
Nucleus
Chloroplasts
Methylene Blue
Cutinized cell walls
Nucleus
Crystal Vio let
Achromatic figures
Cutinized cell walls
Lignified cell walls
Nucleus
Plastids
E) Permanent slides
Permanent slides are useful for preserving a particularly good thin section or squash of an alga, or a
specimen that is hard to find or prepare. Because they take a while to dry, permanent slides should be
started in the beginning of the lab period.
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Using a labeled slide you’ve prepared, carefully remove the cover slip so as not to disturb the
sample.
Add 1-2 drops 10% Karo Syrup (light corn syrup), then let dry for 30 minutes to a few hours.
After the syrup has dried, add 1 drop of 50% Karo Syrup and let dry for 1 week in the lab. Dab
off the excess liquid after one week and add a drop of 100% Karo Syrup.
While the specimen is still wet, cover it with a cover slip and paint clear nail polish around the
edge of the cover slip to seal the slide. You now have a permanent slide you can keep.
F) Keying, Collecting, and Pressing Algae
Keying Out Algae
To key out your algae, you will need an algal key. In California, a preferred book is Marine
Algae of California, by Isabella A. Abbott and George J. Hollenberg. Once you have determined the
Division to which your alga belongs, use the dichotomous key to find the Genus and species. Writing
down each step as you key facilitates re-keying a misidentified alga.
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Today we will practice keying out an unknonwn alga together as a class.
Draw the marcothallus and cross section of the unknown in your lab notebook.
Write out the steps as you go through the dichotomous key (just write the step numbers – not
the words of each step).
Collecting Algae
When you collect algae, you may need certain tools, such as a knife to pry algae off of rocks,
and a bag in which to carry the algae. Also, consider whether you will need to collect the entire plant
and/or reproductive material for keying or pressing. Some intertidal algae may only be reached dur ing
very low tides, so consult tide tables beforehand. See table below for guidelines on collecting algae.
DO
Obtain a collecting permit or California fishing
license (for collecting up to 10 lbs of algae)
Try to collect reproductive material and entire
plant, including holdfast, for identification (except
Codium fragile where a single branch is fine)
Key out and label specimens before pressing
Press algae as soon as possible, writing species
name on the paper lightly, in pencil
Store algae, if needed, in a dry plastic bag in the
fridge for up to 1 week, “fluffing” daily
Collect drift algae/beach wrack when possible
DON’T
Don’t collect if there are fewer than 10 individuals
of the same species
Don’t forget to remove snails and other large
invertebrates from your algae while still in the field
Don’t store your collected specimens in water
Don’t collect Postelsia palmaeformis or
Phyllospadix seagrasses (see images below)
Don’t put Desmarestia, “acid weed,” in container
with other algae
Don’t collect at State Parks or protected areas
Postelsia palmaeformis
Images courtesy www.mbari.org
Phyllospadix seagrasses
Pressing Algae
An important part of this course is preparing a collection of pressed algae. This requires a press, which
includes:
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Wood slats and straps
Blotting paper/Newspaper
Cardboard
Herbarium paper
Herbarium paper may be cut in half for smaller specimens.
When pressing algae, you should follow the layering order shown in the diagram below.
1. Top of the Press
2. Cardboard
ABCDEFGHIJKLMNOPQRSTUVWXYZABCD 3. Newspaper, about 4 sheets
4. Blotting Paper
********************************************************** 5. Wax Paper
6. Specimen
7. Herbarium Paper
8. Blotting Paper
ABCDEFGHIJKLMNOPQRSTUVWXYZABCD 9. Newspaper
10. Cardboard
11. Bottom of the Press, or a new
stack for the next specimen,
sharing the cardboard layer
Tips for Good Pressing
 Keep a log of where and when each specimen was collected.
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Record each specimen’s data lightly, in pencil, in the lower right hand corner of the herbarium
paper.
Rinse algae with seawater before pressing.
Arrange each alga, exhibiting thallus characteristics clearly.
Dry in drying oven, about 150 ˚F (optional, but keep in a warm, dry environment).
Change blotting paper and newspapers often (daily for thicker specimens), to prevent molding
of the algae.
When specimen is dry, prepare a display label like the example below. Some blank labels at the
end of this lab can be photocopied and cut out for use with your pressed specimens.
University of California, Santa Cruz
ALGAE OF
Chlorophyta, Ulvales
Name
Ulva clathrata (Roth) C. Agardh
Location
Scott Creek, Santa Cruz, California
Habitat
found in high intertidal, growing on rocks
Collected by
Al Gee
Date
April 8, 2012
Identified by
Al Gee
Date
April 8, 2012
No.
University of California, Santa Cruz
ALGAE OF
Name
Location
Habitat
Collected by
Date
Identified by
Date
No.
University of California, Santa Cruz
ALGAE OF
Name
Location
Habitat
Collected by
Date
Identified by
Date
No.
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