GENERAL INFORMATION
Note fire route, fire extinguisher, first aid kit and eye wash station.
Zoology 351 (Aquatic Invertebrates of Alberta) is a "hands-on" course. You are mainly on your own and will get out of the course what you put in it. Essentially everyone goes their own way, but all roads lead to the lab exam.
Each person is issued:
1. A key to the lab (see Equipment - Safety).
2. A key to a microscope (storage) cabinet. The storage cabinet is for scopes and lamps, and there is a limited amount space in the cabinets for samples. But you might want to obtain a locker (center wing, 3rd floor) to store nets and samples (see Equipment and Safety).
3. A compound microscope.
4. A dissecting microscope and lamp.
5. A dip net.
6. A sorting (white enamel) pan.
7. Various preservatives and chemicals will be available in the lab.
You then go out and collect aquatic invertebrates. Put the invertebrates or samples containing the invertebrates in jars; add preservative or not, and bring the material back to the lab. Pick out the invertebrates; put them in vials or jars containing preservative, and then identify the invertebrates using the keys in the text-lab manual Aquatic Invertebrates of Alberta. The number of points awarded depends on the taxon. These are indicated in the Points section.
Collection
At the end of the term (see Class Schedule ) you hand in an identified collection with a list of taxa indicating the number of points you expect to receive (see the section: Final Preparation of Collection). More will be said about the collection as the term progresses.
Your collection should consist of specimens that you collected. Of course, if you collect with other people, the specimens can be divided the way you want, but two or more people can not share a specimen.
Again, except under special circumstances agreed to by the instructor, only specimens collected in the last week of August of 2004 or later can be included in the collection. This rule, which will be strictly adhered to, is needed so that people who could not collect during the summer are not penalized.
The collection is worth 40% of the final mark. Unlike some collection-oriented courses, there is no upper limit to the number of specimens handed in (except that you cannot collect more taxa than are actually present in Alberta). The number of points obtainable varies with the year - determined mainly by the fall weather and the number of field trips that can be held. For example in 1994, the top collection was at 1,900, while in 2002, it was only 1,367. Thus you are marked relative to other members of the class in the current year.
There is a strong correlation between a good collection and doing well on the lab exam (see Scatter Diagram in appendix). But keep in mind the above warning, and don't use this diagram as a yardstick for what is a good collection. The points allocation for certain taxa can change (as it has for water mites this year), and in some years a sample that can count in the collection has been handed out to each student thereby inflating the average points..
Laboratory Examination
The only way to get a good mark on the lab exam is to collect invertebrates and identify them throughout the term. This can't be overemphasized. You can't pass the lab exam by an intense session or two of eyeballing pictures in the text or other sources or even coming in and identifying invertebrates only during class hours. It doesn't work; it's been tried. You must collect and identify invertebrates throughout the term (which is why you are issued a key to the lab - to work on your collection at times other than during class hours).
The lab exam consists of 50 - 80 stations. You are given 1 - 1.5 minutes at each station and asked to identify the invertebrate to a particular taxonomic level, depending on the taxon in question. You use the classification scheme listed in the collection portion. You will also be asked to answer a short question on some aspect of the biology of the taxon in question that will be discussed in lecture. More will be said about the lab exam as the term progresses.
Formal Meeting
We meet for lecture and quiz only one day a week. This has been scheduled for Thursday except for 18 Nov (see Class Schedule). Since we meet formally only once a week, everyone should come and ON TIME. This is particularly important for the quiz, which starts at the beginning of the lecture period. The teaching assistant and instructor will always be in the lab on Tuesdays, 9:30- 11:00am. Depending on popular demand, we may appear for another few hours per week. If you are struggling with a specimen outside of this time, feel free to come to our offices for help. Of course, neither of us may be in; this is why it's a good idea to save up the 'hard ones' and make an appointment.
Quizzes
There are twelve quizzes that add up to 20% of the course marks. Each quiz takes place at the beginning of the Thursday lecture slot starting on 16 Sept and will be based on readings from the text Clifford (1991) and on the previous week's lecuture. Although the quizzes are brief (about 15 minutes) and relatively easy (5-8 written questions), they do require some initiative. You should prepare for the quiz by reading the chapters indicated in the class schedule (e.g. for the lecture and quiz on 11 Sept, you should have read Chapters 1 and 2 in Clifford, 1991). Subjects of quizzes usually overlap with the subject matter in the lecture that immediately follows the quiz. By the way, terms in the Glossary (pp. 449-460) are also fair game.
Textbook
Aquatic Invertebrates of Alberta is used for the quizes of course; but more importantly the keys in the book are used to identify the specimens. The pictorial keys are easy to use and when used in conjunction with the whole specimen drawings (habitus drawings) and colored photos should provide a good indication of the name of the taxon in question. We will provide you with additional keys for certain keys for certain taxa throughout the course.
Please spend time browsing through the book to become familiar with its organization. I would appreciate knowing of errors or other comments. (See appendix under Collection section for known errors, and mark you book accordingly. If you do not you may make major errors.)
Field Trips
In addition to collecting on your own, at the beginning of the term we have a few class field trips. These are NOT compulsory. Three field trips are scheduled for this year; on Saturdays or Sundays in September and October (see class schedule). You will have to sign a release form in order to attend.
More information will be given in class, for example, where to meet, the time, what to bring (see also the handout on Equipment - Safety). Don't forget to bring a pair of forceps and obtain as many jars as possible; we provide a few jars but never enough. In addition to preservative available in the lab, there will be preservative in the vans. If you take alcohol out of the lab, which almost everyone eventually will do, please mark EtOH POISONOUS on the container.
The class field trips begin on Sept. 14, but you should begin collecting as soon as possible. We realize that many have no personal transportation. This has seldom been a problem because people who do are usually willing to take others with them. Therefore, if you are going on a collecting trip and have space, please announce this to the class, e.g., a note on the bulletin board, or tell the instructor and it will announced in class.
A map showing the aquatic areas that can be sampled in the Edmonton area, by bus in some cases, is posted in the in the main lab (Z217). A good place to collect within walking distance of the building is the pond at William Hawrelak Park or the North Saskatchewan near the LRT bridge and Emily Murphy Park. But hurry, the pond is usually drained in early October.
Water Safety
Please read the section on Equipment - Safety. Be prudent and use common sense when sampling. There is no need to sample deep water with a dip net. You will collect many more invertebrates in shallow water. If sampling from a boat, you must wear a life jacket. Also, it's a good idea to sample with someone. Later this autumn, the water will be cold and there is always a risk of hypothermia if you fall in. Complaints about feeling cold when you do not have a hat or gloves, will not be greeted with much sympathy! Finally, though this has more to do with comfort than safety, wear long pants and socks when heading out on a field trip. Hip waders can do a lot of damage to bare legs.
Closing Points
We have a multitude of fascinating aquatic invertebrates in Alberta. They can affect us directly (e.g. swimmer's itch)or indirectly in many ways. Initially you will find the animals bewildering and will have difficulty recognizing orders. But eventually you will see a strange critter and say "it's a beetle larva, but I don't know what family". Being able to recognize and therefore appreciate our aquatic invertebrates takes a little work. But once you have acquired an aptitude for it, it opens up all sorts of rewarding avenues to enjoyment and employment.
EQUIPMENT - SAFETY.
Field Equipment (*supplied)
*Plankton nets (shared)
*Pond nets
*Glass jars (some will be supplied, but obtain others)
*Preservatives (denatured ethyl alcohol, ca. 80-90% (do not even remotely consider using this for parties, it has been doctored with formalin). Please mark containers of these "POISONOUS."
*Plastic bags
*Sorting trays (enamel pans)
*Fine Forceps (Don't forget to bring these on the field trips.)
Waterproof footwear. Waders or hip-boots recommended. The Department has a few pairs of mainly leaky hip boots.
Magnifying glass (at least a good idea)
Pencils/paper (for field labels)
Sunglasses and sun screen lotion.
Laboratory Equipment (*supplied)
*Key to the laboratory (Z-217)
*Dissecting scope
*Compound scope
*Scope lamp
*Storage cabinet and key to same. This cabinet is to store the scopes and lamp; there is a limited amount of space in the cabinets for samples.
It's a very good idea to obtain a locker to store nets and samples. These are located on the third floor, center wing, near the Biological Student Services Office. Information on how to obtain a locker will be discussed.
*Various chemicals (lab instructors will discuss these)
*Slides and coverslips
*Insect pins and wooden handles (to make fine probes)
*Plastic vials. These are cheap "pill vials," and only a limited number will be available. Good snap cap glass vials must be purchased.
*Bottles and vials of various sizes and shapes.
*Petri dishes
*Pipettes
*Slide box
*Various specialized keys to certain groups of invertebrates.
Materials for Final Preparation of Collection
Boxes (do not use shoe boxes). Boxes
designed commercially for new vials, with their cardboard retainers, are best.
Perhaps we will have enough to pass one out to each student.
Pill vials (see above). These are acceptable, but within 2 or 3 years the alcohol
will have evaporated. If you plan to keep your collection, screw cap glass vials
(scintillation vials), 1-4 drams, are available from the BioSciences storeroom.
These vials are superior to pill vials for permanently storing specimens, but
they are expensive.
Water Safety
Exercise common sense about water safety when collecting aquatic invertebrates. You must wear a life jacket when sampling from a boat. In fact, it is a good idea to wear a life jacket when sampling deep water of lakes (and streams) from the shore, and, if possible, sample with someone. Also be aware that a good dunking, even in September in certain waters, might result in hypothermia. More will be said about this in class.
Instructions on Laboratory Safety and Operations
1. Let the instructor, TA
or lab coordinator know if you have any allegeries.
2. Leave all books, coats and bags at the coat rack, not at your bench.
3. Do not wear contact lenses in the laboratory. They can trap
dangerous fumes and liquids against your eye.
4. KEEP THE LABORATORY CLEAN. THAT MEANS CLEAN UP YOUR OWN MESS. DO NOT
LEAVE JARS OF PRESERVED MATERIAL IN LAB BENCH DRAWERS OR COUNTERS
FOR SOMEONE ELSE TO CLEAN UP. In the interests of good housekeeping, clean off
the top of your bench when you have completed each laboratory session ñ
especially if you are using any type of chemical. Although you may not be effected,
the individual using that laboratory space may encounter severe burns, etc,
by being exposed to your mess. Return all chemicals and other materials to the
fume hoods and areas from where you obtained them.
5. ABSOLUTELY NO EATING OR DRINKING IN THE LAB. This includes party use
of lab alcohol (making cocktails for consumption); this is a cheap grade of
ethanol with many nasty residues after distillation. It is also denatured with
formalin for better preservation. ALL PRESERVATIVES ARE DESIGNED TO DO ONE
THING- KILL AND FIX. Use goggles (or perscription glassses), latex gloves,
and a lab coat at all times when handling any type of chemical. Chemicals other
than alcohol will be available ONLY during regular lab periods or under
supervision by the TA, instructor, or lab coordinator.
6. Read the Materials Safety Data Sheets posted in the fume hood. They
are for your safety and you should be aware that almost all the chemicals you
will be using are harmful in one way or another. If you spill any chemical,
use the spill mix kit located in the fume hood to soak it up. Report the spill.
7. Report an accident of any kind to the instructor, TA or lab coordinator.
8. SLIDE MAKING WITH THE MOUNTING MEDIUM PVA (polyvinyl alcohol). THIS
WILL ONLY BE ALLOWED ON TUESDAYS (9:30 ñ 11:00 AM) OR UNDER SUPERVISION.
GOGGLES, LATEX GLOVES, AND A LAB COAT ARE MANDITORY WHILE MAKING SLIDES ñ
OTHERWISE YOU WILL BE ASKED TO LEAVE THE LAB. THE MOUNTING MEDIUM PVA (actually
polyvinyllactophenol) CONTAINS PHENOL, WHICH CAUSES SEVERE INSTANT BURNS.
Slide making protocols safety precautions will be discussed after your first
formal field trip.
9. All discarded liquid
or solid waste material must be placed in the containers provided and designated
as Liquid (Ethanol) or Solid Waste, not in the sinks. A separate
container is provided for broken glass and other sharp objects, such as razor
blades. Please follow the intructorís, lab co-ordinatorís, or
T.A.'s instructions in this regard. Again, if there is a chemical spill of any
kind, pour the Spill Mix mixture provided to absorb it, and then contact an
authority.
10. Be sure that all taps,
distilled water, sink, vacuum, gas and air lines are closed after each use.
Most of the chemicals used to fix, narcotize, and preserve invertebrates or
mount them on slides can be used with the usual precautions of handling any
reagent. However, acids and strong bases, e.g. sodium and potassium hydroxide,
can be corrosive, and formalin has been implicated as a possible carcinogen.
If in doubt about the vapors of any chemical, use the fume hood. The course
coordinator will give additional information about laboratory safety. Also,
the Glossary of Aquatic Invertebrates of Alberta gives information about
certain chemicals, and please read the last paragraph of page eight.
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More Lab Safety. The appropriate protocol for the amount of spill and spill mix to use, who to call and what to do in an emergency is posted on the fume hood door and at the front of the class room. Please be aware of them in the case of an emergency.
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HANDLING SPECIMENS, MOUNTING, NARCOTIZTION AND PRESERVATION METHODS
General Information
Making Slides.
All small (<5mm) specimens for this course can be mounted in PVA (polyvinyl alcohol), which is supplied. However, use judicious quantities as gallons of this mounting medium is not available. PVA will only be available on Tuesdays of each week or under supervision of the instructor, TA or lab coordinator. Water mites (Hydrachnidia) require special methods for the best mounts. See notes under Hydrachnidia).
1. Specimens should be in 80% ethanol (transfer fluid) prior to mounting. Place 1 or 2 drops of PVA on a microscope slide to allow for shrinkage after drying.
2.Transfer your specimen(s) from the ethanol into PVA using a pipette, or specimen is large enough, using fine forceps. The specimen should be contained in the smallest volume of transfer liquid as possible. The transfer process should be also be tracked with the aid of a dissecting microscope to insure your pipette contains the specimen and that you deliver it into the PVA.
3.Orient the specimen with a fine probe and allow to restfor about 15mins. Before orienting, look at the identification key to decide what parts of the specimen need to be seen for recognition.
4. Then add another drop of PVA to the medium containing your specimen and gently place a coverslip over the specimen. While watching using a dissecting microscope, gnetly press down on either side of the specimen using forceps to orient and flatten the specimen. Don't press to hard!
5. With a marking pen, circle the location of your specimen on the underside of the microscope slide. PVA also acts a clearing agent, so this will help you find your "invisible" specimen later.
6. Allow the slides to dry for several days, or better, 1-2 weeks, in a horizontal (flat) position in the cabinet located in the fume hood. Then seal the edges with a thick coat of fingernail polish. If there is excess PVA around the edges of the coverslip, scrape this away with a razor blade before sealing.
7. When mounting larger specimens (mites, copepods, other microcrustacea), coverslip supports should be used. These can be made by breaking a coverslip into small pieces. Three or four small pieces can be placed around the specimen before applying an entire coverslip over the specimen. Alternatively, plasticine legs can be used, where the plasticine is scraped onto each corner of the cover-slip. The legs should shallow, not too large, and an additional drop or two of PVA should be used to allow for shrinkage.
8. Wipe up any mounting medium that is spilled on the bench top with a paper towel and discard in the solid waste container.
General Narcotization
Many animals (such as rotifers, ectoprocts, oligocheates, and microcrustaceans like copepods, water fleas, seed shrimp, etc) can be relaxed with chloroform. One or two drops of chloroform is applied to a cotton swab attached to the inside of a bell jar. The bell jar is then inverted over a vessel containing the specimen (which is immersed in water). The vessel must be a glass petri dish or watch glass or it will dissolve.
Cnidaria
These animals should not be fixed in the field. In the lab., place specimens in water (in a small petri dish in water from which they had been taken) until they are fully extended. Fix by quickly pouring off the water and flooding with hot Bouin's fluid. Heat the Bouin's fluid prior to use by immersing the container in a bath of boiling water for about 5min (Boil water in the kettle provided and then pour into the vessel containing the bottle of Bouins). Use a pipette to flood the petri dish. After several minutes replace the hot Bouin's fluid with cold Bouin's and leave for 12 hours. The organisms should then be rinsed with clean water, and preserved in 80% EtOH (if the ethanol turns yellow, replace it with fresh EtOH) or mounted on a microscope slide in PVA. If specimens are distinctly green when alive, transfer them to 10% formalin instead of EtOH before slide mounting.
CAUTION: Bouin's fluid is toxic (contains formalin) and when dried out can be explosive. So clean up any spills with a wet paper towel and discard in the organic waste bucket. Discard fluid into waste EtOH container.
Turbellaria
These animals should not be fixed in the field. In the lab., place several specimens in small petri dish in a minimal amount of water (preferably the water from which they came) until they are fully expanded. Narcotize them by adding Steimann's fluid, a few drops at a time, until the animals no longer respond to touch. Preserve the organisms in 80% EtOH or mount on a slide in PVA. An alternative method of killing these animals in an extended state, is by adding boiling water to a small petri dish containing several individuals.
Rotifera
These animals are small and can more readily be found when alive and moving in fresh samples of water that contain some substrate, i.e. a small amount of sand, mud or vegetation. Rotifers can also be found in moss samples soaked in a small petri dish with a minimal amount of water. After 5-15 minutes, shake the moss vigorously, remove and observe the water with a dissecting microscope.
a) Loricate rotifers: kill with 10% formalin and mount on a slide in PVA.
b) Non-loricate rotifers: place animals in small GLASS dish and narcotize under bell jar containing chloroform saturated cotton ball. Then fix in EtOH and mount on a slide in PVA. Do not use a plastic petri dish as the chloroform will dissolve it.
Ectoprocta
These animals should not be preserved in the field. In the lab., place in pond water in a small glass dish until specimens have protracted their lophophores. Narcotize under bell jar containing chloroform saturated cotton ball for about 15mins. or until specimens no longer respond to touch. Preserve in 80% EtOH; specimens can also be mounted on a slide in PVA. Statoblasts should be mounted on a microscope slide in PVA for identification.
Oligochaeta
Place specimens in a glass dish containing just enough water to cover the bottom of the dish. Wait until the specimens are elongated: then narcotize by slowly adding chloroform, drop by drop. When they no longer respond to touch, preserve in 80% EtOH. Small specimens can also be mounted on a slide in PVA after ethanol fixation.
Hirudinea
These animals should not be preserved in the field. Place leech in a petri dish containing just enough water to cover the specimen. Narcotize by adding small amounts of ENO (an antacid). After the leech has relaxed, sandwich it between two microscope slides; a piece of paper towel should be applied to one of the slides prior to this process in order to hold the specimen in place. Clamp both ends with rubber bands and immerse in 80% EtOH. Do not clamp the ends so tight that the specimen becomes 2-dimensional, making it difficult to identify.
Crustaceans
Can be killed and preserved in the field with 80% EtOH. Copepods, Ostracoda, and Conchostraca, however, need to be mounted in PVA for further identification. The specimens must be dissected before mounting; both the specimen and the dissected parts should be mounted on the same slide. Should you have live specimens, narcotize them prior to fixation in a small glass dish under a bell jar containing chloroform saturated cotton ball for about 15mins. This will expose the limbs better for easier identification. For Ostracoda and Conchostraca, the valves need to separated; use fine insect pin probes.
Porifera
Preserve in the field with 80% EtOH. For identification in the laboratory, take a small piece of sponge tissue and place in a small petri dish. Transfer as little water as possible. Separate out the gemmules (small round densely colored spheres) and pipette into a different petri dish . Add a few drops of bleach; dilute the bleach with water after the tissues disappear. With the aid of a dissecting microscope, transfer the spicules and gemmules (either by pipette or forceps) to separate microscope slides. Be sure to hand in slides of spicules and gemmules. Then add PVA and cover slip each slide.
Tardigrada
Tardigrades are very small and require careful observation with the aid of a dissecting microscope in order to locate them. Tardigrades are sometimes found associated with duckweed, in moss or the substratum the moss is attached to. For hydrated moss, dehyrate sample in a zip-lock bag. Rehydrate for about 2 days in a container; add an equal volume (volume includes the moss sample) of 20% EtOH to the vessel for narcotization. Ring out the moss, and seive residue through a fine net. For hydrated moss, the moss should be placed in small petri dish with just enough water to cover the bottom, and then shaken vigorously. Be sure to dislodge some of the substratum which the moss is attached to, which is usually sand. If the water is too murky, take a subsample and dilute with clean water. Fix in 80% EtOH and mount in PVA.
Nematoda, Nematomorpha, Insecta, Araneae
Kill and preserve in the field with 80% EtOH.
Mollusca
Kill and preserve in the field with 80% EtOH. Large bivalves (Family Unionidae) should be brought back alive and dissected for identification. This may be done by boiling the specimen in water until the shell opens, or by cutting the adductor muscles and then preserving them. Snails can be narcotized by adding menthol crystals to a small volume of water if so desired. Change EtOH if it becomes cloudy or discoloured, as this may indicate that the animal is rotting.
Hydrachnidia
Kill in the field or lab using Koenike's Fluid (45% water, 45% glycerol, 10% glacial acetic acid). Keep in Koenike's for at least one full day before clearing in 8-10% potassium hydroxide (KOH). Large, soft-bodied mites should be punctured with a fine pin to aid in clearing internal tissues. Check specimens every 15 minutes and remove when translucent. Leaving mites too long in KOH can render them brittle. Large ites can often be identified to family or genus without mounting. Simply return them to Koenike's Fluid for storage. Small mites may require mounting in PVA. First, soak mites in water for 10-15minutes to remove KOH. Mount as described above. It is helpful to mount the mites ventral side up, and to remove one palp and lay it sideways, prior to putting on the coverslip.
USE OF THE MICROSCOPE
Microscopes are delicate instruments and should be handled with care. Always pick up your microscope with one hand under the base and one hand around the neck and carry it in this manner. Never carry a microscope by the neck alone. Before inserting or removing a slide from the stage of a compound, insure that the lowest power lens (or the blank space) is pointing toward the stage. There is a real possibility of scratching the lens if there is insufficient space to move slides in and out. Every dissecting and compound microscope comes with a transformer to regulate light intensity. Before plugging it in or unplugging it, ensure that the dial is in the minimum position. In this laboratory there will never be occasion to use the transformer at maximum power. Following these two simple rules will greatly extend the life of the microscope bulbs. Always return the microscopes to your numbered place in the cupboard at the end of each lab. If there is something wrong with a microscope, do not replace it or scavenge parts, leave the microscope out on the back bench so that it may be repaired, and then tell one of the instructors in the course.
Dissecting Microscope
1. Obtain a wet-mount specimen or a prepared slide and place it on the stage.2. Push-click the transformer on. The intensity setting should not be at maximum. A mid-range is sufficient and can be used for epi-illumination. Maximum voltage of the transformer reduces the longevity of the lighting source and is not necessary.
3. Use the lowest power lens. With one eye closed, find and focus the object as sharply as possible with the focus knob. Close this eye and open the other; focus the object as sharply as possible, using the focus ring on the eyepiece. Adjust the eyepieces for the width of your eyes by pushing them together or moving them apart and keeping both eyes open. Repeat the focusing protocol if the image is blurred.
4. Keep both eyes open when viewing an object.
Compound Microscope
1. Obtain a prepared slide and place it on the stage.2. Turn the transformer on. Normally it should be used at an intensity setting of not more than 6 or 7 (or a mid-range for microscopes with no intensity numbers). Click-stop the 10X objective into place.
3. Find and focus (as sharply as possible) the object using the course and fine adjustment knob. Then focus each eyepiece according to your eyes as follows: close one eye and focus that eyepiece, using the focus ring on the eyepiece. Repeat this for the other eye. Focus on the object again and repeat this procedure if the image is blurred. Adjust the eyepieces for the width of your eyes keeping both eyes open.
4. Optimal use of the voltage regulator:
- The goal is to produce an evenly illuminated field of view with the substage condenser and its iris diaphragm adjusted so that approx. 7/10 of the back aperture of the objective lens is filled with light. This gives you the best image for this type of microscope.
- To do this, position the condenser to its maximum upper limit using the condenser pinion or knob. Remove an eyepiece and the back-aperture of the objective lens appears as a halo. Find the iris diaphragm lever and adjust it to fill about 7/10 of the aperture (it should appear as hexagon or octagon in the field of view). If the iris diaphragm is not centered then adjust it with the setscrews on the sides of condenser.
- The substage diaphragm should never be used to control the brightness of the image; altering the voltage regulator should perform this function.
- When you switch objectives (i.e., 4X, 10X, 40X) the procedure should be repeated. If your microscope is equipped with an auxiliary condenser lens, then this should be clicked into place when using the 40X objective or higher power prior to this procedure.
5. Common difficulties encountered:
- if you cannot obtain a sharp image:
- perhaps you have not focused your eyepieces individually
- the fine adjustment has either reached a stop
- the object is touching the specimen before the sharp image
- you have dirty slide, coverslip, or objective (clean only with lens paper or kim wipe)- If you have an oval field of view instead of a round view, either the objective has not fully engaged its click-stop position or perhaps the filter holder below the condenser is not swung fully to the stop.
- Keep both eyes open when viewing objects with the microscope.
- Never focus downward while looking into the microscope, especially at higher power; use the coarse adjustment while viewing from the side
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