Maintaining Adult Frogs

The most convenient method of “holding” adult frogs (Rana pipiens) until needed is to render them inactive by reducing their metabolic rate. This method makes feeding unnecessary, reduces waste build-up, and reduces the occurrence of various frog diseases. In addition this method will maintain the frog in a specific physiological state (e.g., maturation, or pre- or post-ovulation, etc.)

The metabolic rate is lowered by refrigerating the frogs. If the frogs are to be held for 1– 3 days, they may be placed in a loosely covered pan with1 –2″ of water (tap water may be used). The pan should then be placed in a refrigerator at 0° – 5°C (32° – 41°F). For longer holding periods (1– 3 weeks) slight modifications are necessary. Cool the warm frogs for several hours in the refrigerator 0° – 5°C (32° – 41°F) before unpacking. Have a container of water (tap water may be used) 6 – 12″ deep, cooling at this time. This container must be well aerated at all times through the use of an airstone and air pump. Frogs may be placed in the container as soon as the above temperatures are reached. Change water at least twice a week with fresh water that has been pre-cooled to holding temperature.

Induced Ovulation

Pituitary Extract contains the lyophilized pituitary glands of seven female Ranid frogs in addition to 6 mg progesterone. Only gravid northern Rana pipiens should be used, and not the “southern” sympatric species which is not physiologically suited for induced ovulation protocols. The addition of the hormone progesterone has been found to be more effective in producing consistent ovulation success.

Since seasonal changes in pituitary potency and female sensitivity occur, the following table will serve as a general guide on the use of either collected pituitaries or pituitary extract.

Please consult the separate product literature sheet accompanying Ward’s Pituitary Extract for re-hydration instructions. Please note that you will need a syringe with a #20 gauge needle to inject the female. Syringes and needles can be obtained at a local pharmacy with a prescription or through WARD’S.

Injection procedure for inducing ovulation is as follows: Carefully, so as not to injure the ventral blood vessel or internal organs, insert the needle just through the skin and underlying muscle wall of the lower abdomen and inject the pituitary-progesterone suspension. Be certain to choose a gravid female to inject. Females have reduced nuptial pads on the “thumbs” of the forelegs. Gravid females will be more plump than their non-gravid counterparts because the abdomen will be full of eggs.

Place injected females in water approximately 11⁄2 –2″ deep and keep animals at a temperature between 18 –21°C (64 – 70°F). Injection should be done 48 – 62 hours before scheduled artificial insemination.

After 48 hours, check the enclosure for evidence of egg release. Eggs may require longer periods to “ripen” and observation of released eggs is the best indication that ovulation is proceeding. You may also “test” for ovulation by gently squeezing the abdomen towards the cloaca. To ensure a longer yield of mature eggs, do not extract them until 24 hours after the start of ovulation. Mature R. pipiens should yield between 1,000 and 2,000 eggs for fertilization.

Artificial Insemination

To obtain active sperm, either pith a male frog or overdose the animal using WARD’S Biocalm, an amphibian anesthetizing/euthanizing agent. Remove both testes, dissect away adhering tissues, and wash testes free of blood. Macerate in 10 – 20 ml of either pond water, dechlorinated tap water, or 10% Holtfreter’s Solution*. A convenient way to do this is to force the testes through a syringe with an #18 gauge needle. Large clumps may be broken by aspiration. Wait 15 –25 minutes for full sperm activity, which can be determined by examination under a compound microscope. The sperm suspension may be diluted to a total volume of 100 ml, if necessary, for very large egg masses. However, better results (e.g., more consistent rates of fertilization) are obtained by using the sperm suspension as prepared, without further dilution. Note that polyspermy may occur in concentrated sperm solutions with Octobercaught or long-held animals

Following confirmation of sperm activity, eggs are stripped from the female by holding her and applying pressure to the abdomen with the force directed towards the cloaca, thus squeezing the eggs from the ovisacs into a dry Petri dish. This is accomplished by bringing the legs of the female forward parallel to the abdomen. This assures that pressure is not dissipated laterally. The third and fourth finger are applied firmly over the throat and thoracic region to avoid dissipating pressure anteriorly. Remaining fingers are used to “milk” the abdomen toward the cloaca. Initially, rather sharp pressure may be needed to open the cloaca sphincter muscle; subsequently, gentle pressure will suffice to aid egg release.

Eggs are expressed in a circular pattern on the dry Petri dish. Due care must be exercised so as not to cause rectal prolapse by trying to obtain every “last” egg.

Expressed eggs are inseminated by pipetting the sperm suspension over them. Make sure that each egg comes in contact with the sperm suspension, although moistening is all that is required. Application of sperm suspension must be done prior to egg jelly swelling. After 10 –15 minutes, flood the eggs with medium. Use of a dry dish is recommended because eggs stick to its surface. This facilitates changing the medium. The spiral or circular pattern prevents eggs from clumping. After an additional 15 minutes, pour off the medium plus sperm and replace with fresh medium. Set the eggs aside for 30 –50 minutes. They then may be scraped free from the Petri dish and transferred to culture bowls. A single-edged razor blade, tissue section lifter, or scalpel should be used. Do not attempt to pull eggs free. About one hour after fertilization, those eggs successfully inseminated will rotate so that the black animal hemisphere is uppermost. After several hours, remove the unfertilized eggs to prevent decay. At optimum development temperature 18 – 25°C (64 – 77°F) cleavage will begin about 21⁄2 hours after fertilization, muscular response will be observed in about 96 hours, and hatching will occur in 5 – 7 days.

* Holtfreter’s Solution: Prepare a stock solution by adding 3.5 g NaCl, 0.2 g NaHCO3, 0.1 g CaCl2, and 0.05 g KLC to 1 L of glass-distilled water. Dilute this stock solution to 10% with distilled water before use.

Raising Tadpoles

About 3 –5 days after hatching, the tadpoles are ready to feed. They should be placed in an aquarium filled with tap water that has been standing overnight (to dechlorinate) or fresh pond water. Place an airstone in the water and keep the air pump constantly running. The number of tadpoles per liter of water may be as high as 100 immediately after hatching. As metamorphosis approaches (2 –3 months) the number must be gradually reduced to about five tadpoles per liter of water.

Since tadpoles are omnivorous, many types of food have proved suitable. Prepared fish food works well as does ground dry dog food, mashed hard boiled egg, chopped liver (raw or cooked), and boiled lettuce leaves. The diet should be varied from time to time. Only those foods which the tadpoles readily consume should be given. As with any aquarium set up, overfeeding must always be avoided. If the normally clear water starts to turn cloudy the tadpoles are either being overfed or have not consumed the particular type of food they are being fed. This is a dangerous condition. Be sure that the food given is being eaten and use only that amount of food which can be consumed in a short time.

As metamorphosis proceeds, a provision must be made so that the young frogs (now air-breathing) may climb from the water. This occurs very shortly after the front legs erupt from under skin flaps that have been concealing them during their development. For the remainder of its life the frog will be basically a “meat-eater”.

Feeding of Adult Grassfrogs

At metamorphosis, R. pipiens shift from an omnivorous to a carnivorous diet comprised of food that must be moving. They differ from R. catesbeiana and R. clamitans in that they do not or cannot take food while submerged. If animals are not held in hypothermic conditions, they should be fed 10 – 12 full-grown live crickets (2 – 3 times per week). Other food that is recommended include sowbugs, beetles, moths, and earthworms. Fly maggots are not recommended, since such ingested maggots may destroy the frog.

This guide is also available in PDF format on wardsci.com.

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The large number of species available for aquarium use makes it difficult to choose which plants to have in your aquarium. The guidelines presented in this leaflet should help you create an environment in which the aquarium plants and animals can survive and grow for a long period of time with relatively little care or alteration of the environmental conditions.

Generally, plants are used to increase the oxygen concentration in water by removing carbon dioxide from the water during photosynthesis, thus permitting more oxygen to go into solution. Plants that are ideal for this situation are those that have a high photosynthetic rate and do not require that they be rooted in soil for normal growth and development. Elodea (Anacharis), Myriophyllum, and Cabomba are all good aquarium plants because they are bushy, rapid growers that do not need to be rooted and are good carbon dioxide consumers.

Plants, such as Vallisneria and Sagittaria, that can be rooted in gravel are also well suited to aquariums. When rooting these plants, they should be carefully pushed into the substratum so that the crown of the plant remains at the surface. They may be securely anchored with small lead weights, stones, or other heavy, inert material until they have secured themselves with their own root growth. Vallisneria is generally a tall plant and should be planted in the back of large and relatively deep tanks. The Corkscrew Vallisneria is a shorter form that is better suited for a smaller aquarium. Sagittaria has wider leaves that Vallisneria and needs a fair amount of space for normal growth. As such, it should be separated by several inches from other plants and the sides of the tank.

If your tank is not in direct sunlight, members of the genus Cryptocoryne will do well. They are narrow-leafed plants that thrive in temperatures between 21 – 26°C (70 – 79°F) and can endure acidic water. The most common species is C. willisi, but equally good is the dwarf plant C. becketti. If you wish to add some color to your aquarium, C. cordata is a good choice because its broad leaves are red on the underside. In addition, some species of Lubwigia, or false loosestrife, have leaves that vary from green to red in color. Other interesting species that you may wish to place in your aquarium include a species of Myriophyllum that is known as Parrot’s Feather, the water fern Ceratopteris, and the hornwort Ceratophyllum. Parrot’s Feather is a bushy, rapid-growing plant with soft green feathery shoots that are very attractive. The water fern can be either planted in gravel or remain floating and will show an interesting method or reproduction. Be sure to avoid overpopulation of this species. Hornwort will do well under almost any aquarium circumstances.

The use of algae and floating surface plants in aquaria is not recommended because they do so well that they become pests. One exception is the complex green alga Nitella. It is used to form thickets for small fish and other fragile and defenseless species and can usually be kept under satisfactory control. Lemna (duckweed) and Riccia (a type of liverwort) can be used if you wish. These plants are usually rapid growers and excess material can be readily removed if the cover becomes too thick. If both are used together, Lemna may tend to overgrow Riccia. At intervals throughout the year Lemna will appear to die out and disappear, but in a few weeks it will return to its original abundance or even be more abundant. When plants such as Cladophora and Pithophora find their way into an aquarium, it is usually necessary to start over. You will need to discard all of the contents of the old aquarium, since the algae will develop rapidly from the smallest portions that may be left behind. Other species may leave slimy scums and foul odors or materials that are toxic to other life forms in the aquarium. In these cases, the aquarium should be cleaned and the sides scraped with steel wool or a razor blade. It may also be necessary to clean the aquarium with a 2% solution of HCl for thorough sterilization. Once it has been sterilized, a thorough washing with distilled or boiled water will allow the aquarium tank to be used again. The only chemical alternative for algae control is 9.065 g of potassium permanganate added to 3.785 L of distilled water. Used with care, this method will often clear an aquarium of unwanted algae.

Finally, two types of water ferns that are commonly used are Salvinia and Azolla. However, if you choose to use these plants, you should be aware of the fact that these are plants that flourish in warm, relatively stagnant water. If they are planted in a relatively shallow aquarium with a high organic content and are given adequate light and a warm temperature, they will do well. If they are planted in a large, relatively sparsely inhabited aquarium that is well aerated and has a cooler temperature, either form will die out in a short amount of time.

Regardless of whether you prefer rooted or floating plants, your selection of aquarium foliage should be based on how you intend to use your aquarium. This consideration should include the prevailing temperature, the other possible inhabitants, and the size of the tank. In all cases, you should be sure to avoid overcrowding, poor planting, and you should maintain the tank for cleanliness. Contamination, regardless of the source, should be dealt with immediately, before conditions deteriorate to the point where a complete reconstruction of the aquarium becomes necessary.

This guide is also available in PDF format on wardsci.com.

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• Nitella
• Duckweed
• Riccia
• Cladophora
• Salvinia
• Azolla

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Introduction

Protozoa are among the most fascinating organisms that can be studied in the classroom or laboratory. The Protista kingdom has seven groups that are divided into fifteen phlya. These subdivisions show the wide range of morphology and function that demonstrate the basic properties of living matter. This diversification is one of the reasons that students seem to be instantly fascinated by the study of these organisms.

The different phyla are distinguished from one another by such features as structure, means of locomotion, and formation of spores, although the locomotor organelles are the primary distinguishing feature. There are three main locomotor organelles found in the different classes of protozoa, and they are pseudopodia, cilia, and flagella.

A pseudopodium is generally not considered to be a separate organelle, but rather is formed by the extension of protoplasm outward from the main body. The pseudopodium then anchors itself while the rest of the organism flows into the pseudopodium. Pseudopodia are also used to surround and ingest food particles. Unlike the pseudopodium, both cilia and flagella are considered to be discrete organelles. Both structures move the organism by beating in a rhythmic or random pattern, and structurally are almost the same. The main difference between these two organelles is that flagella are usually found singly or in small groups near the leading end of the body, while cilia are found in large numbers in longitudinal rows on the body or near the mouth.

Most protozoa reproduce through a process called fission, which is simply the organism dividing into two cells by mitosis. In addition to fission, some ciliates are able to reproduce sexually through conjugation. In this process two cells unite and exchange genetic material.

Collection

Collection of protozoa is possible from almost any conceivable habitat. Free-living species occur wherever there is water, while parasitic species occur in most metazoa. A hay infusion set up in the lab will yield many different types of microorganisms. Mud and vegetation collected from ponds, streams, and transient bodies of water will yield microscopic life after being kept for a short time in the laboratory.

Care and Feeding

Receiving

All packages containing live materials should be opened immediately and all jars, vials, etc. should be checked for breakage.

Loosen (do not remove) the caps on the jars and tubes containing cultures to permit air exchange. (A frozen culture is not necessarily ruined. Thaw slowly, then follow instructions below.)

Inspect all cultures microscopically in the culture container using a low-power binocular microscope (Know what you are looking for). One can also pipet a bit of the liquid which contains the protozoan onto a slide and examine under a conventional microscope.

If the organism is not visible, allow the culture to rest undisturbed at room temperature for one-half to one hour and re-examine. Also refer to the particular organisms in the section entitled Comments in the chart below.

Handling

Common sense should apply here. Most Protozoans are extremely fragile; therefore all manipulations must be performed gently. Pipetting should be done slowly; cultures should not be handled roughly.

Short-Term Maintenance

Keep loosened caps on all culture-containing jars and tubes when not in use, and be sure to keep out of direct sunlight. Most protozoans do best at a temperature of 20°C, however the temperature may be anywhere between 18°C and 22°C.

Culturing

The following chart covers most of the basic knowledge needed to work with each listed protozoan, some suggestions of methods to try in working with others, as well as links to Ward’s product pages. The pH of the media should be as close to 7 as possible and all glassware used should be clean, sterilized, and free from chemical contamination.

Media

Amoeba Medium: To 100 ml of distilled water add two rice grains.

Note: Chalkey’s medium can be used instead of distilled water as a more refined medium. This consists of 0.1 g of NaCl, 0.004 g of KCl, and 0.006 g CaCl2 in one liter of distilled water. Proceed as above.

Cereal Grass: Add 1.5 g of cereal grass powder to one liter of distilled water. Boil for five minutes. Filter while hot.

Cool and use immediately. Dilute cereal grass medium 1:3 with distilled water. Decant approximately 100 ml of medium into a bowl and add three boiled wheat grains.

Note: 1.5 g of ground oven dried lettuce leaves may be substituted, but results may not be as consistent.

Difflugia: Soil-water medium is the basis. Place 5 – 10 cm of garden or muck soil free from chemical fertilizers and pesticides in the bottom of a culture container and fill three-quarters full with distilled water. Plug and steam for one hour on two consecutive days. Filter the soil water and dilute 1:4 with conditioned tap water. To 100 ml of the dilute soil water, add a pinch of extremely fine sand and a few strands of spirogyra.

Hay medium: Put 15 g of Timothy hay into one liter of distilled water. Autoclave at 15 PSI for 15 minutes. After cooling, filter and store in refrigerator. Dilute the hay medium 1:3 with distilled water. Dispense approximately 100 ml per culture container. Then add three boiled wheat grains.

Chaos: To 100 ml of distilled water, add two rice grains. Then add Paramecium caudatum grown in hay medium. Replenish P. caudatum as needed.

Split Pea: Add forty split peas to one liter of distilled water and autoclave at 15 PSI for 15 minutes, or boil for approximately 10 minutes. Cool and use immediately.

Tetrahymena medium: Dissolve 1 g Proteose Peptone in 100 ml distilled water. Dispense, cap, and autoclave at 15 PSI for 15 minutes.

Observation

The majority of experiments done with protozoa require only a microscope and good powers of observation. It is important that all slides and coverslips be completely clean. Slides can be washed in a mild detergent solution, rinsed in distilled water, and allowed to dry before use. If new slides are not pre-cleaned, they should also be washed.

Wet Mounting

There are two techniques that provide the best viewing for organisms on a slide. For the first technique, draw an outline of a coverslip and then place the slide on top of the tracing. Draw a thin line of petroleum jelly within the tracing lines using a toothpick. Place a drop of liquid containing the organisms to be viewed in the center of the coverslip and carefully lower the slide onto the coverslip to form an airtight seal. Invert the preparation smoothly to keep the seal intact. If relatively thick objects are to be viewed, such as Chaos, a few pieces of broken coverslip should be included within the petroleum jelly border to avoid crushing the organism. (Caution should be used, since the broken glass will be sharp.)

The second technique utilizes silicone culture gum to form a chamber on the slide. Remove a small piece of the gum and roll it into a sphere in the center of a clean slide. Take a second slide that has been wetted and press it on the sphere until you have the desired thickness. The wetted slide can then be removed, since the silicone gum will not stick to it. A tight bond will be formed between the silicone gum and the dry slide. Using a sharp cork borer, cut a hole through the gum to the slide and remove the plug. Fill the cell with the culture to be observed and cover with a coverslip, excluding as much air as possible. The silicone gum will allow carbon dioxide to escape and oxygen to enter. One can keep the prepared slide many days by placing it in a humidity chamber, taking it out only to study the culture. A simple humidity chamber may be made by placing the slide in a Petri dish lined with moist paper towel.

Once the organisms have been mounted on the slide, the most common behaviors to observe are feeding and sexual reproduction. Several different organisms’ feeding behavior can be easily observed. Chaos and Didinium can be observed by placing a drop of medium containing several Chaos or Didinium on a slide, and then adding a drop of concentrated Paramecia. Observe the different methods used by the Chaos and the Didinium to capture, ingest, and digest the Paramecia. In a mixed protozoa culture, Euglena and similar green forms may be observed in the food vacuoles of larger protozoa. In the mixed protozoa culture you may also observe and compare the food gathering techniques of the Ciliophora, Mastigophora, and Sarcodina.

The mating of Paramecium bursaria can be observed by mixing equal quantities of the opposite mating types on a slide or in a watch glass. Evaporation must be prevented as the experiment takes about two days. The mixing should be done about midday, since time is an environmental factor in mating. The individuals will immediately start to form clumps. After 16 hours (depending on conditions) individual mating pairs may be observed. Large numbers of conjugants normally occur from 24 to 48 hours, after which time little conjugation occurs.

Sporozoan Demonstration

All sporozoans are parasitic, absorbing nutrients from their hosts. Plasmodium vivax is a sporozoan that causes malaria in humans. Sporozoans of the genus Gregarina are found in the guts of arthropods.

Gregarines are found in the gut of the mealworm, Tenebrio. Mealworms burrow into their food and defecate. This develops an environment in which they readily infect each other with the protozoan. A simple demonstration of the presence of gregarines utilizes the live Tenebrio larva. With a scalpel or razor cut off the head and the last somite of the mealworm. Pull out the digestive tract with a probe or fine forceps and remove any adhering tissue. Then place the gut in a drop of Frog Ringer Solution (6.5 g NaCl, 0.14 g KCl, 0.12 g CaCl2, and 0.2 g MgSO4 in one liter of distilled water) on a slide. Place a coverslip over the gut and press down gently. Scan the slide under low power. The gregarines should appear as clear to dark-colored objects, longer than they are wide. Sometime they appear as short chains. They may be flushed out of the gut with another drop of saline and examined under higher power for structural details.

Immobilization

In order to view many types of fast-moving ciliates and flagellates under a compound microscope it is necessary to slow or immobilize them. There are three basic methods of immobilization: mechanical, chemical, and narcotic. Because of the unsuitability of narcotic means for classroom use and the versatility of other methods, this method will not be discussed.

Mechanical: If debris is present in the culture, remove some with the drop of culture to be examined. The organisms will eventually tend to come to rest near the debris and can be studied.

If no debris is present, a small amount of shredded filter paper or lens tissue may be placed in the drop on the slide; use only a few fibers.

Chemical: WARD’S Detain is a water-soluble, visoelastic, nonionic resin solution used to slow motile protists. Detain is less toxic to protists than the traditional chemical slowing agents stated below. Detain will dissolve readily in fresh or salt water. Due to its high viscosity, Detain will support a coverslip, and after a few minutes will form a ring around the periphery, retarding evaporation. Protozoa have been observed to survive for periods of over 96 hours following introduction of Detain.

Methyl cellulose is an excellent agent for immobilization of protozoa. It can be used as a 1.5% liquid or as a 10% paste. To use the 1.5% liquid, add a drop to the side with the culture on it. As the methyl cellulose diffuses through the medium, the change in viscosity will slow the protozoan’s movements. To use the 10% paste, follow, the same procedure used for the petroleum jelly wetmount. As the methyl cellulose diffuses inward, it will immobilize the protozoans.

To prepare the 1.5% liquid, mix 1.5 g methyl cellulose powder with 50 ml boiling water. Cool, then stir 48.5 ml of cool water into the mixture. The liquid should have the consistency of syrup. The 10% paste is prepared in like manner. Add 10 g of methyl cellulose powder to 50 ml of water, then as another 40 ml of water. The consistency of this concentration is like solidified gelatin.

Polyvinyl alcohol is used by adding one drop of 14% PVA solution to one or two drops of the culture on the slide. It may be more effective to place the drop of PVA at the margin of the culture drop, because this causes the PVA to diffuse slowerThe immobilization is both mechanical (change in medium’s viscosity) and chemical.

To prepare the 14% PVA solution, add 14g of the PVA powder to 36mL anhydrous ethyl alcohol. It may be necessary to heat in a water bath until the PVA dissolves. Stir and add 50mL of distilled water.

Vital and Supravital Staining

Many chemical stains can be applied to living protozoans to color various parts of the organism. If the stain is not harmful in the dilution used, it is a vital stain. If it is ultimately lethal, it is called supravital staining. For use, most stains must be dissolved and diluted. Percent solutions, for example a 1% methylene blue solution, will refer to only the percentage of solute weight to volume. To make a 1% aqueous solution of a stain, place 1 g of dry powder in a graduated cylinder and add enough distilled water to total 100 ml. This method may also be used for alcohol solutions, although it is not as accurate because 1 ml of alcohol is not 1 g as in the case of water. This method, however, will be sufficient for use in most cases. All water used for dilutions should be distilled. All alcohol used should be absolute (100%) ethanol. Other alcohols may be used, but the results may not be as satisfactory as those obtained with ethanol. The stains discussed are made up in absolute ethyl alcohol.

Stains

Bismarck Brown: Will stain cytoplasmic inclusions in living protozoa. Place a drop of 0.1% Bismarck Brown alcohol solution on a slide. Allow the drop to dry. Place a drop of protozoan culture on the slide and cover with a coverslip. If this concentration disrupts the protozoan cells, try a 0.05 – 0.01% dilution. Alternatively, a 1.0% Bismarck Brown alcohol solution diluted to 0.1% or 0.01% with water may be dropped directly on a drop of culture on a slide.

Brilliant Cresyl Blue: Will stain the nucleolus of a protozoan. It is used in similar concentrations as Bismarck Brown. Bromothymol Blue: A pH indicator, which is yellow at pH 6.0 and blue at pH 7.6, it is used to study the food vacuoles of protozoans. It can be used as described for Bismarck Brown, or it may be used to stain yeast cells which are then fed to Paramecia in order to demonstrate the pH of food vacuoles.

Place about 1⁄4 cake of fresh yeast or 0.5 g of dry yeast in 10 ml of distilled water. Add 0.1 ml of 1% Bromothymol Blue Solution. Boil in a water bath 5 –10 minutes to kill the yeast cells. Cool and place a small amount of the suspension (using a toothpick) in a drop of paramecium culture on a slide. Observe the pH in the food vacuoles over a period of time.

Carmine Powder: Used to demonstrate the feeding in paramecia and other ciliates. Place a drop of the protozoan culture on a slide and dip the end of a toothpick into the Carmine powder and transfer a bit of it to the slide. Cover with a coverslip. Observe the action of the cilia in sweeping the Carmine into the “mouth”. Notice the formation of food vacuoles and the circulation and discharge of Carmine.

Janus Green B: Will stain the mitochondria a greenish blue. It will also stain the Golgi apparatus. It can be used at the same concentration as Bismarck Brown. Janus Green B may be combined with neutral red to give a multiple stain. This is done by mixing one part Janus Green B solution (fifteen parts absolute alcohol and one part 1% Janus Green B solution in absolute alcohol) and two parts Neutral Red solution (ten parts absolute alcohol and one part 1% Neural Red in absolute alcohol).

Methylene Blue: Will stain the nucleus, cytoplasmic granules, and cytoplasmic processes of protozoa. It may be used in a 0.05% alcohol solution or in lesser concentrations as used with Bismarck Brown. Methylene Blue may also be used to demonstrate the discharge of trichocysts in Paramecia. Place a drop of Paramecium culture on a slide and cover with a coverslip. Put a drop of 0.1% methylene blue solution at the edge of the coverslip and observe the trichocysts discharge as the Paramecia encounter the stain.

Neutral Red: pH indicator. It is red below pH 6.8 and yellow above 8.0. It will also stain the macronucleus slightly. It can be used in the same manner as Bismarck Brown. Observe the pH changes in the food vacuoles as they form and circulate.

Fixation and Staining of Protozoa

When permanently fixing a contractile organism for observation, it may be necessary to relax the organism in order to prevent the organism from being fixed in an abnormal shape. If direct fixation does not destroy or distort the protozoan, the relaxation is not necessary. A 3% solution of copper acetate will immobilize and relax most ciliated protozoans. Other chemicals which may be used include the following: 10% methyl alcohol, 1% hydrochloride, and 1% magnesium sulfate. These chemicals are added drop by drop until the absence of any contraction is observed when the protozoan is under stimuli. Then the protozoans are fixed.

Glutaraldehyde: Suitable for fixation of some ciliates and other protozoans. To use add one to two drops of 25% glutaraldehyde to one drop of the protozoan culture on a slide. If this method causes distortion or disintegration of the protozoan the following method using glutaraldehyde may be effective.

Place a drop of the protozoan culture on a slide and invert it over a watch glass containing a few drops of glutaraldehyde. Leave the slide in contact with the fumes approximately 30 seconds, then remove and proceed to the staining of the protozoan.

Schaudinn’s Fluid: A broad-spectrum fixative for protozoa. To use, the protozoan culture to be fixed should have as little culture fluid as possible. The fixative should then be added slowly. For use in the fixation of a drop of protozoan culture on a slide, add one part Schaudinn’s Fluid to three parts water. Then add one drop of the diluted fixative to one drop of the protozoan culture on the slide. When the turbidity clears the protozoan may be stained.

Preparation of Schaudinn’s is as follows: add two parts saturated aqueous solution of mercuric chloride to one part absolute alcohol (ethanol). Just before use add 1 ml glacial acetic acid to 99 ml of the fixative.

Staining

Toluidine Blue: used to stain cilia, cirri, flagella, and membranelles of protozoans. It may be used in a 0.1– 0.001% aqueous solution. Experience will determine the best dilution for the protozoans to be stained. To use, add one drop to the fixed material on the slide. In order to be the most effective, the stain should just tint the liquid slightly. Water mounted protozoa have many limitations when it comes to staining because it is difficult to remove the stain from the liquid. For fine work and to produce permanent slides one should research into the literature on protozoa and microtechnique.

This guide can also be found in PDF format on wardsci.com.

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