Objective In this lab we will cover logistics and introduce techniques for successful examination, preservation, and identification of algae.

Transcription

1 INTRODUCTORY LAB Objective In this lab we will cover logistics and introduce techniques for successful examination, preservation, and identification of algae. Notebook Requirements - 2 charts & 4 drawings 1) 2 microscope calibration charts 2) Mazzaella sp: 2 drawings (thallus & cross section) 3) Pyropia spp: 2 drawings (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 calibrate 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 measurements 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.

2 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 (S) µm (O) ocular units =? µm 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: First, cut off a manageable piece (1-2 cm 2 ) 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 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.

3 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 specimen top slide Side View thin sections bottom slide 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. Prepare a thin section on a slide of the alga you would like to stain, as described 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 absorbent paper alga eyedropper with stain slide Surface View These images depict how stains are added to a sample. Staining Specimens Stain Analine Blue IKI Malachite Green Methylene Blue Crystal Violet Interacts with Cellulose cell walls Cytoplasm Achromatic figures Starch Cutinized cell walls Nucleus Cell walls Cytoplasm Nucleus Chloroplasts Achromatic figures Cutinized cell walls Lignified cell walls Nucleus Plastids

4 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. 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. 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 during 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 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 Phyllospadix spp, Codium fragile, Fucus spp, Constantinea simplex & Silvetia compressa. Don t put Desmarestia, acid weed, in container with other algae Don t collect at State Parks or protected areas

5 Pressing Algae An important part of this course is preparing a collection of pressed algae. This requires a press, which includes: 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. 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.

6 University of California, Santa Cruz ALGAE OF Name Location Habitat Chlorophyta, Ulvales Ulva clathrata (Roth) C. Agardh Scott Creek, Santa Cruz, California found in high intertidal, growing on rocks Collected by Al Gee Date April 8, 2012 No. 1 Identified by Al Gee Date April 8, 2012 University of California, Santa Cruz ALGAE OF Name Location Habitat Collected by Date No. Identified by Date University of California, Santa Cruz ALGAE OF Name Location Habitat Collected by Date No. Identified by Date