Invertebrate Anatomy OnLine

Laboratory Techniques ©


Copyright 2001 by

Richard Fox

Lander University


            This is a chapter from Invertebrate Anatomy OnLine , an Internet laboratory manual for courses in Invertebrate Zoology.   Additional exercises can be accessed by clicking on the links to the left.   Terminology and phylogeny used in these exercises correspond to usage in the Invertebrate Zoology textbook by Ruppert, Fox, and Barnes (2004).  

1.   Removing Microorganisms from Culture Jars

            Use a plastic or glass Pasteur pipet with a bulb.   Squeeze the air from the bulb BEFORE immersing the tip of the pipet into the culture jar.   Position the tip appropriately.   For example, if you are looking for bottom dwellers such as Amoeba, place the tip at the bottom.   If you seek swimmers, such as Euglena or Paramecium, then the tip should be above the bottom.   Release the pressure on the bulb slightly so a few drops (only) of liquid are drawn into the tip of the pipet.   Transfer these drops to a microscope slide.  

2.   Preparation of a Wetmount

            Place one or two drops of liquid in the center of a clean glass microscope slide.   Add stain or other materials as appropriate and mix thoroughly with a teasing needle.   Do not add more than a total of 2-3 drops of liquid.  

            Carefully lower a clean coverslip onto the liquid in such a way that no air bubbles are trapped beneath the coverslip.   This is best accomplished by placing one edge of the coverslip on the slide touching the liquid.   Support the opposite edge of the coverslip with a teasing needle or forceps and slowly lower it to the surface of the slide.

            Use # 1½ or # 2 square 22 mm coverslips for ordinary microscopic examination.   Use # 0 coverslips if you plan to use oil immersion.   These are thinner and much easier to break so you must be more careful with them.

            It will probably be necessary to adjust the amount of liquid on the slide.   There should be no liquid outside the coverslip but there should be enough under the coverslip to support it so it does not stick tenaciously to the slide and crush or distort organisms.   If you have too much liquid, use a tissue to absorb the excess.   If you do not have enough, use a pipet to place a drop or two on the slide next to the edge of the coverslip.   The liquid will run beneath the coverslip if it is needed.   If it is not needed, you will have to remove it with tissue.  

            Excess liquid on the slide will run off the slide onto the stage where it will form a thin film between the slide and stage.   This film acts like glue and its presence makes it impossible to move the slide smoothly across the stage and almost impossible to move it at all.   It may also damage the microscope, especially if it is a salt solution.

3.   Simple Squeeze Preparations

            It is sometimes desirable to use the weight of the coverslip to squeeze an organism slightly.   This may be done to immobilize the specimen or to make it a little thinner so its internal parts are easier to see.  

            The simplest way to accomplish this is by removing some of the liquid from under the coverslip.   While watching the organism through the eyepiece, touch a piece of absorbent tissue paper to one edge of the coverslip.   Capillarity will wick the liquid from beneath the coverslip into the paper.   Remove the paper when you have withdrawn as much liquid as desired.  

            More complex supports are described below.

4.   Supported and Adjustable Squeeze Preparations

            A more elaborate method of making squeeze preparations employs wax feet on the corners of the coverslip and gives you control over the pressure applied to the organism.   Dig each corner of a coverslip into the surface of a piece of beeswax so that a tiny piece of wax adheres to the glass.   This is easier if the beeswax is softened somewhat.   Hold the piece of wax in your hand to warm it or, better still, simply store it in a shirt pocket next to your body when not in use.   Remember to remove it before you leave the lab. The four pieces of wax are referred to as "feet".  

            Place the coverslip, with its feet down, over the liquid on the slide and lower it as usual.   When the coverslip is in place, use the handle of a teasing needle or applicator stick to push gently, directly downward, on the glass above each foot.  This will flatten the foot and move the coverslip closer to the slide where it will trap and squeeze organisms.  

            The degree of squeezing depends on the pressure applied.   Once squeezed, the process cannot be reversed.

5.   Support of Coverslip with Sand

            It is sometimes necessary to support the coverslip so it does not crush or distort delicate organisms beneath it.   The wax feet described earlier can be used for this purpose and have the advantage of being adjustable (in one direction) but they are cumbersome to use and time-consuming to prepare.   Instead, many microscopists use 5-10 grains of fine beach sand scattered in the fluid on the slide before the coverslip is applied.   The sand grains support the coverslip and protect the organisms.  

            With this technique there is no possibility of squeezing the organism should that become desirable.  Supported wholemounts may be too thick for use with the high dry objective and will certainly be too thick for oil immersion.

6. Compound Microscope Light Adjustment

            When using the compound microscope, the light intensity must be carefully adjusted.   There is a tendency among beginning students to use too much light.   Transparent objects, such as Amoeba, will be invisible if the light intensity is too high.  

            In general, light intensity should be adjusted using the iris diaphragm, not the electrical rheostat control.   Set the rheostat control at about 75% of maximum and leave it there.   Use the iris diaphragm to adjust the light intensity.  Sometimes it will be necessary to use the rheostat, as for example, when using the scanning lens.  

7. Initial Focus

            Finding the correct level at which to focus is difficult when most of the material under the coverslip is water.   In the absence of a conspicuous object, there is nothing on which to focus.   Use the edge of the coverslip as an easily-found "object" at the correct level and focus on it first, then look for the small objects you are really interested in, secure in the knowledge that you are focused at about the right level.

8. Systematic Scan

            Finding small and/or scarce organisms on a slide is most efficient if done systematically.   First, be sure the light is adjusted properly and the microscope is focused at the correct height (i.e. between the coverslip and the slide).   Organisms are difficult or impossible to see if the light is too strong.   Using the scanning lens, find one corner of the coverslip, say the upper left.   Move the slide so you follow the upper edge of the coverslip to the upper right corner.   Move down a distance equivalent to a little less than the diameter of one field and then move back to the left side of the coverslip.   Your new field should just overlap the already scanned strip next to the edge of the coverslip.   Be sure there is no unexamined space between the two strips.   Continue moving back and forth across the slide, neglecting no part of it, until you find what you are looking for or you reach the lower right corner.  

9.   Alteration of Wetmount Without Removing Coverslip  

            It is frequently desirable to change the liquid below the coverslip of a wetmount without removing the coverslip.   This is easily accomplished and can (should) be done while you observe the organism.

            To accomplish such a change, place a drop of the new fluid on the slide next to and touching one edge of the coverslip.   Touch the edge of a piece of absorbent tissue paper to the opposite edge of the coverslip so it is in contact with the old liquid beneath the coverslip.   The old fluid will be absorbed by the paper and removed from the wetmount.   The new fluid will move under the coverslip to replace the old as it is removed.  

10.   Optical Sections

            When observing organisms with the compound microscope, keep the fine focus in nearly continual motion, focusing up and down through the object.   Each position of the fine focus gives you a crisp focused image of one level of the object, much as if you had sliced, or sectioned, the object at that level.   The resulting view is known as an optical section.   Other levels are out of focus and cannot be seen clearly.   To get an accurate impression of the entire organism you must take optical sections at all levels by continual refocusing up and down.

11.   Immobilization of Active Organisms

            Rotifers and many protozoans, especially ciliates, swim so rapidly that they cannot be observed alive unless immobilized.   This is accomplished by using a solution containing long polymers that entangle and impede the organisms.   Traditionally a 10% solution of methyl cellulose is used for this but other products are now available.   Your laboratory may be supplied with vials of homemade methyl cellulose solution or with commercially prepared products, such as Detain or Protoslo.   Prepare the wetmount as usual but use a mixture of culture and polymer.   The more polymer, the slower the organisms.  

            Squeezing the preparation slightly by removing a little fluid may also immobilize swimming cells or organisms. Another method is to place a whisp of cotton on the slide and add a drop of culture and then the coverslip.   The crossed cotton fibers create small chambers or cells in which organisms may be trapped thus limiting their motion.

12.   Oil Immersion

            To use the oil immersion lens, focus on the object of interest with the high dry lens and be sure the object is centered in the middle of the field.   Rotate the nosepiece halfway to the oil immersion (100X) objective.   Place a drop of immersion oil on the slide directly atop the center of the hole in the stage.   Slowly move the nosepiece around until the oil immersion lens clicks into position.  

            Be sure you rotate the nosepiece directly to the oil immersion lens and not to the high dry.   If you do the latter, you will get oil on the high dry lens.   Should this happen, clean it immediately with lens paper (only) and no harm will be done but do not allow oil to remain on this lens.  

            Look through the eyepiece and focus on the object very carefully using fine adjustment only.   When finished with the oil immersion lens clean it carefully with lens paper.   If you are using a prepared slide, clean it also.   Coverslips contaminated with oil are usually not reused and should be discarded.   Oil immersion is usually used without a coverslip or with a # 0 coverslip.

13.   Cleaning Lenses

            You should keep the lenses of your microscopes clean.   Make it a point to clean the ocular lenses (eyepieces) of your compound and dissecting microscopes at the start of each laboratory session.   The objectives, however, rarely need cleaning unless they have been accidentally smeared with the liquid from a wetmount.   Use nothing but fresh, unused lens paper to clean lenses and discard it after use.  

            If you notice specks or smears in your field of view, it means at least one of the many lenses in the system needs cleaning.   You can identify the offending lens using a simple protocol.  

            Look through the eyepiece so you can see the spots.   Move the slide.   If the spots move, they are on the slide, which should then be cleaned.  

            If the spots don't move with the slide, try rotating the eyepieces one at a time.   If the spots rotate, they are on one of the lenses of that particular eyepiece.   In this case they are almost always on the outer surface of the upper lens because it is the one exposed to oily eyelashes, dust, and the occasional fingerprint.  

            If the dirt did not rotate, it must be somewhere else.   Raise or lower the condenser.   If the spots go out of focus they could be either on the top of the substage lamp or the condenser lens.   Clean both.  

            If the contamination manifests itself as a blurred image that cannot be corrected by focusing, the objective is probably fouled.   Moisten a piece of lens paper with water and clean the outer surface of its lens.  

14.   Cleaning Permanent Slides

            Clean prepared slides by wiping lightly with a piece of absorbent paper or tissue.   The lower surface of the slide can be rubbed vigorously but the coverslip must be treated gently.   Do not apply pressure to the coverslip that would tend to move it (the coverslip) with respect to the slide.   The undetectable microscopic movements resulting from such handling can damage the object.   This is especially true of sections, which can be destroyed by enthusiastic rubbing.  

15.   Sealing Temporary Slides

            Mineral oil can be used to seal the edges of wetmounts to prevent evaporation and increase the life of the slide.    This is especially useful for wetmounts made with seawater or magnesium chloride since evaporation of water increases the salinity under the coverslip, making conditions intolerable for living organisms.

            After preparing a wetmount as usual, place a tiny drop of mineral oil beside the coverslip.   The oil will flow over the surface of the exposed liquid at the edge of the coverslip and will retard evaporation.

            Temporary slides can also be sealed with petroleum jelly.   Smear a thin film of jelly on the fleshy part of the heel of your hand and scrape all four edges of a coverslip across the jelly leaving a low ridge of jelly on each edge.   Use this coverslip to make a wetmount in the usual fashion.   The jelly ridges will form a seal between the coverslip and the slide.

16. Visualization of Ciliary Currents
            One of the important advantages of using living, relaxed specimens is that cilia and flagella continue to beat, making it possible to see the currents they generate. 
            Vizualization of ciliary currents usually requires the addition of fine particles or soluble dyes to the area of interest. The particles or dye should be suspended or dissolved in liquid of the same density (and temperature) as that surrounding the animal or else density currents will interfere with your study of ciliary currents. The suspensions should be made using either seawater or pondwater as appropriate and stored in small bottles for use when needed. The density of the suspension can be adjusted with salt (non-iodized) or Instant Ocean.
            Ciliary fields and tracts are usually very specific with regard to the particles they can transport. Particle size and particle density are strongly related to their transportability by a particular ciliary field. It is useful to have a collection of suspensions of particles of different sizes and weights. 
            Carmine suspended in water (seawater or pondwater as appropriate) is probably the best general-purpose tracer. 
            Chalk dust from the eraser trays of the classroom chalkboard (becoming harder and harder to find as more and more classrooms are equipped with whiteboards) can be suspended in water and used to advantage. (Chalk dust deliberately scraped from a stick of chalk is coarser, less homogenous, and less satisfactory than that found in the chalk tray.) 
            Water-based latex house paint, which consists of tiny latex beads in an aqueous carrier can also be used when very fine particles are needed. 
            India ink contains colloidal carbon and has long been a popular source of tiny particles. Its density must be adjusted for use in seawater. 
            Homogenized milk, with its density adjusted (with salt), can also be used, especially when fine particles are needed. As an alternative, powdered milk can be reconstituted using seawater of the appropriate salinity (density).
            If available, fine volcanic ash suspended in water is a useful tracer.  
            When tracing ciliary currents on surfaces, be sure the surface is horizontal. If you do not then gravity currents may be mistaken for ciliary currents.
            Common household food coloring, being non-toxic and inexpensive, is ideal for freshwater work but its density must be adjusted for use in seawater or else it will simply float on the surface of the water. 
17. Micro Teasing Needles
            Very fine needles, known by the German minuten nadeln, or the English microneedles, mounted at the ends of wooden applicator sticks (Fig 1) are essential for manipulation and dissection of small animals and are easily prepared. They are not indestructible and must be replaced periodically. 
            Split, with the grain if possible, 2-3 mm of the tip of an applicator stick with a scalpel. Do this safely by placing the stick on the surface of the table and pushing the blade of your scalpel into the side of the end of the stick (not the end of the end). Do not hold the stick in your hand while splitting it and do not cut toward yourself. 
            Insert about 2 mm of the blunt end of a 0.2 mm microneedle into the cleft, applying a drop of water-insoluble glue, such as Duco Cement, to the end of the stick around the cleft and base of the needle, and wrapping it tightly with sewing thread. Twirl the end of the stick between your thumb and forefinger to spread the glue over the thread, end of the stick, base of the needle and into the cleft. 
            Make at least two miocroneedles, one with a straight tip and the other bent at a 45 ° angle (Fig 1). Use two different colors of thread, such as yellow for the straight needle and blue (B for bent) for the bent tip, to color code your two microneedles. This makes it easy to select the desired needlewithout having to look closely at the tip. 
Figure 1. A micro teasing needle made from a minuten nadel and an applicator stick. lab5.gif
              Figure 1
             Stainless steel minuten nadeln (without sticks, listed as minutien pins) are available from Fine Science Tools ( Be sure to order 0.2 mm needles and not 0.1 mm. Be sure to get stainless rather than black anodized. Applicator sticks are available from VWR Scientific Products. 
            Be careful when working with these tiny needles.  Microneedles are tiny and are easily lost at the workbench. Lost needles have a habit of ending up later sticking into fingers or elbows.
            The delicate needles are easily bent but are almost as easily restored to their original condition. If the tip gets turned over into a tiny hook, it can be straightened by pulling it between the nail of your forefinger pinched tightly against the ball of the thumb, or between the points of your forceps.
18. Modification of Microdissection Forceps
            Watchmaker's or microdissecting forceps are essential for most of the dissections in this collection. They are available from Fine Science Tools, Carolina Biological, and Ward's. They are delicate tools that require careful handling and preparation. 
            The tips of new microdissecting forceps (especially inexpensive ones) often require modification before they are suitable for use. This is accomplished by grinding excess metal from the tips with a sharpening stone such as a Ouachita or Arkansas stone (available from Fine Science Tools or a hardware store). 
            Hold the blades of the forceps together lightly so the opposing tips touch each other but do not press firmly against each other. Look at them with the dissecting microscope. Do they meet each other in perfect alignment or do they pass each other or attempt to do so? Are the two tips the same length? Are the tips as sharp as you would like? Perhaps they are too sharp? 
            You can correct the alignment and produce the shape you want with the stone. Place the stone on the stage of the microscope and focus on its upper surface. Place a few drops of light machine oil on the stone. Hold the tips of the forceps lightly together and slide the point back and forth over the surface of the stone against the metal you want to remove. These stones will remove metal rapidly so check the tips frequently to be sure you are not removing too much. If the tips meet each other at a spot behind the tip, it will help to bend the tips a little. Never try to remove metal from the inside of the tips.  
            If one tip is longer than the other the forceps will be unusable until corrected. Hold the tips together and rub the point of the combined tips together on the stone. Check the lengths frequently and stop grinding when the two are equal.  
            For storage push the tips into the end of a small cork and keep them there when not being used. 
19. Dissecting Pans
            Standard dissecting pans available from biological supply companies are too large to be used conveniently with most invertebrates.  Most dissections will be conducted on the stage of a dissecting microscope and standard pans are too large to fit.  An assortment of small dissecting pans can be made from a variety of metal containers. 
            Dissecting pan wax can be purchased from Wards Natural Science Establishment. The wax is melted in a saucepan dedicated to that purpose (it will be ruined for other uses) and the molten wax poured to an appropriate depth into the chosen container.   
            The most useful sizes are those made from empty 56 g anchovy tins, 105 g sardine or smoked oyster tins, and 100 g kippered herring tins.   The old-fashioned tins that require a can opener are best but those with pull tops can also be used. Their disadvantage is that they have a lip around the opening. Tuna fish tins can also be used and the now-obsolete aluminum refrigerator ice trays make excellent pans for long worms. They can still be found at yard sales and flea markets. 
            The wax should be poured so there is ample freeboard above the wax to more than accommodate the thickness of the specimen and the fluid in which it will be immersed. Because it does not rust, aluminum pans are preferable to steel
20. Incident and Transmitted Light
            Dissecting microscopes are usually equipped with two light sources. The incident light shines down on the object from above the stage whereas transmitted light illuminates the object from below and passes through it. The two lights have different effects and different uses and are not interchangable. In general, incident light should be used for opaque objects and transmitted light for transparent objects. Avoid using transmitted light on opaque objects as the contrast between the bright light and the dark object makes it difficult to see the features of the object. 
21. Anesthetization
            Most marine invertebrates can be anesthetized and relaxed with isotonic magnesium chloride. The sensitivity of animals to magnesium compounds is variable. Some species succumb rapidly, almost immediately, whereas others resist its effects for long periods. In general those species that succumb rapidly will recover rapidly when returned to seawater but those that require long periods to relax often will not recover. There is a tendency for species with thick, impermeable cuticles or exoskeletons to resist relaxation by magnesium chloride whereas those with thin, permeable integuments are susceptible. 
            Magnesium chloride is usually used at a concentration having the same tonicity as that of the animal's body fluids. For marine invertebrates this is usually the same as that of the surrounding seawater. This is referred to as isotonic magnesium chloride throughout these exercises. For animals from undiluted seawater this is about 33 ‰ (parts per thousand). Recipes are given in the Supplies chapter.  
            A weak solution of non-denatured ethyl alcohol (5-10%) in seawater (or tapwater for terrestrial or freshwater species) rapidly relaxes some species that can resist magnesium chloride for hours.
            Magnesium chloride cannot be used to anesthetize freshwater animals because concentrations high enough to be effective are strongly hyperosmotic to the animal and cause its cells to plasmolyze. Other anesthetics must be used instead but it is sometimes very difficult, especially with freshwater or terrestrial molluscs, to find an anesthetic that will relax the organism in an expanded condition. Ethanol (5%), chloretone (0.2%), carbonated water, chilling, and chloroform-saturated water are the most widely used anesthetics for freshwater animals. For soft-bodied animals the anesthetic should be made up in 0.75% saline solution to avoid osmotic damage.
            Most terrestrial arthropods can be anesthetized with fumes of chloroform, ether. Carbon dioxide, either as a gas for terrestrial animals or in solution as carbonated water (Club Soda, Perrier) for freshwater animals is an excellent anesthetic.
22. Gradual Relaxation
            Some marine animals such as the sea hare, Aplysia, and most snails cannot be placed directly into anesthetic.  Aplysia will contract irregularly into an unusable and incomprehensible lump of tissues and snails simply withdraw into shell, close the operculum, and wait for things to get better. The problem can sometimes be circumvented by the slow addition of anesthetic to the water containing the animal.
            Use a strong solution of anesthetic (e.g. 70 ‰) magnesium chloride, 95% ethanol, or 5% chloretone) administered dropwise via a siphon with a carefully adjusted screw clamp. Alternatively, a strip of absorbent cloth or a string (known as a drip string) can be used as a wick to drip anesthetic slowly from the anesthetic container to the animal's into the container. This process may take several hours, often overnight. Be very careful that the animal is not disturbed until it is anesthetized sufficiently that it cannot respond. 
23. Preservative
            Preserved material, if it must be used, should always be ordered in one of the commercial holding fluids rather than formalin. Prolonged exposure of human skin to formalin often results in development of a dermal sensitivity that manifests itself much like poison ivy with weeping pustules and intense itching. Further, formaldehyde causes respiratory problems and is now on the federal list of carcinogens. 
            The developed sensitivity to formalin was once an occupational hazard of biologists, many of whom spent much of their professional lives handling preserved animals. The sensitivity develops as a result of prolonged exposure, with the amount of exposure required varying with individuals. Once acquired, the sensitivity remains for life and the response is triggered by exposure to minute amounts of formalin and aggravated by other chemicals. It is best to avoid all unnecessary exposure. 
            Fortunately it is now relatively easy to avoid formalin. All supply companies offer preserved specimens in some holding fluid other than formalin. This does not mean, however, that no formalin is present. Formalin remains the best agent for the initial fixation and hardening of tissues to be used for gross dissection. Specimens are usually fixed in formalin first and then transferred to some innocuous holding fluid that prevents decomposition. 
            Some formalin inevitably remains in the tissues. Its odor may be strong and unpleasant and there is more than enough to initiate a reaction in sensitized individuals. Formalin-sensitive people should wear rubber gloves when using materials prepared in this manner. The teaching staff should remove specimens from the holding fluid the day before the laboratory session and place them in tap water to soak overnight. At the beginning of the laboratory period the water should be drained and replaced with fresh. Preserved specimens should be dissected immersed in tapwater. Once opened, the body cavity of the specimen should be rinsed gently with tapwater. 
24. Pinning
            Most dissections of soft-bodied invertebrates require that you pin the body wall aside to reveal the body cavity and its viscera. In most instances # 1 insect pins are appropriate but for very small or delicate animals # 000 may be preferable whereas for larger specimens # 4 may be best. The pins should be stainless steel, especially if they are used in seawater or magnesium chloride. High quality stainless steel insect pins are available from Fine Science Tools (See Supplies chapter).  
            Insert the pins at 45 ° angles through the body wall and push them firmly into the wax of the dissecting pan. Think ahead before you insert the first pin and arrange the position of the specimen in the pan so it, the specimen, will be conveniently located after all pins are in place. The animal should be positioned so it will be visible with the dissecting microscope and so none of it extends beyond the edges of the pan. 
25. Care of Iridectomy Scissors
            Your laboratory may provide you with iridectomy scissors or you may be required to purchase a pair. In either case they require careful attention to remain in working order. The blades of these fine scissors are closed by the muscles of your fingers but they are opened by the elastic recoil of their spring-like handles. The blades are held together at an articulation by a pin, as are the blades of any scissors. Whereas your fingers are capable of supplying more than enough power to close the blades, the weaker springs cannot open them again if there is any resistance. Specifically, if rust accumulates in the articulation, it will prevent the blades from opening and the scissors will be useless. 
            After each session the scissors should be cleaned, dried, and greased before storing. Wash the scissors with warm soapy water. Dry with a soft cloth towel. Dip the scissors in acetone and then dry again. Unfasten the handles and open the articulation. Inspect the flat surfaces and scrape away any rust deposits you find there. Apply a tiny gobbet of petroleum jelly to the articulation and smear it over the surfaces. Close the scissors and secure the handles again. 
            All your scissors would benefit from this treatment but it is essential for iridectomy scissors. 
26. Berlese Leaf Litter Extraction
            The diverse and populous arthropod community of forest floor leaf litter is easily sampled using a Berlese funnel. A few handfuls of leaf litter are placed in a large funnel (about 20-25 cm in diameter at the large end), oriented with its large opening uppermost. A vial of water or alcohol (40% isopropanol or 80% ethanol) is attached to the small, lower end of the funnel. A 75-watt light bulb is left burning for several days over the top of the funnel. The heat from the lamp drives the animals downward in the funnel until they drop into the vial from which they can be recovered and studied. 
            The Berlese technique will yield a variety of small soil arthropods, usually including many species of mites and collembolans as well as some pseudoscorpions, spiders, millipedes, centipedes, beetles, flies, ants, thrips, and others. 

            Berlese equipment can be purchased from Carolina Biological and Ward's Natural Science (See Supplies chapter).