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One of the most powerful and widely used diagnostic tools of the microbiologist, this technique requires a few simple steps:
1. Thin smears of bacterial cultures should be air-dried and flame-fixed on clean glass slides. 18- to 24-hour cultures will give the best results.
2. Apply Gram crystal violet stain from eye-dropper or dropper bottle. Slide should be flooded with stain and let stand for one minute. Gently rinse slide with water from a dropper or by dipping in a beaker or cup of clean, distilled water.
3. Flood smears with Gram iodine (serves as a mordant, or dye retainer) and let stand for one minute. Rinse with clean water as above.
4. Decolorize smears with 95 percent ethyl alcohol by applying one drop at a time-just until no more color runs off-for approximately 30 seconds. It is very important not to overdecolorize in this step. Rinse with water as above.
5. Apply Gram safranin counterstain and let stand for 30-45 seconds. This step serves to stain those bacteria (Gram-negative) not holding the crystal violet after step 4. Rinse with water as above. Air or blot dry the slides and examine under oil-immersion.
Note: Do not allow any of the stains to evaporate to dryness on slides prior to rinsing. Spring-type wooden clothespins make excellent slide holders during flaming, staining and rinsing.
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TWO CULTURE MEDIAS FOR ALGAE Bristol's Modified Medium Make six stock solutions, each with one of the following salts dissolved in 400 milliliters of water:
Take 10 mL of each stock solution and add to 900 mL of distilled water. Next add one drop of 1% ferric chloride solution, 40 mL of Pringsheim's soil-water extract and autoclave. Pringsheim's Soil Water Select a rich garden soil which has not been recently fertilized. Add 300 g of the soil to a gallon jar. Fill three-fourths full with distilled water then add one gram of calcium carbonate (CaCO3). Plug or cap jar loosely and steam for one hour per day on each of three consecutive days. To culture Spirogyra, do not add calcium carbonate. To culture Euglena add one-fourth of a pea cotyledon before steaming. This will enhance growth. To culture diatoms, add 10-30 mg of sodium metasilicate (Na2SiO3 9H2O) to every liter of medium. |
Culture media made by soaking or heating fresh or dried organic (usually plant) material in water are known as "infusions." Organic materials used include hay, wheat seed, rice and oat flakes-either individually or in some combination.
The source of the water used is critical. Spring water, well water, and filtered water from unpolluted streams or ponds, are the best choices. The ratio of organic material should be approximately 7 grams (0.25 ounces) per liter.
To pre-treat the organic material, place it in boiling water for one to two minutes, remove it, and maintain it under sterile conditions until it is needed. When you are ready to set up the cultures, pour one inch of water (pre-boiled) into a culture dish (3.5" to 4.5" diameter) and add the organic material. Some specific examples are: Hay/Rice Medium-To the dish of pre-boiled water add two or three grains of rice and three small (2") pieces of timothy hay. Let stand, uncovered, 24 to 48 hours and inoculate. Recommended for all types of amoeba and ciliates.
Wheat Medium-To the dish of pre-boiled water add four to six grains of pre-treated wheat seed and let stand, uncovered, 24 to 48 hours. Inoculate with Blepharisma, all types of Paramecia, Stentor or Vorticella.
For culturing non-specific organisms, prepared infusions can be inoculated with water (a few mL) from any nearby pond, stream or puddle. See what develops!No, agar itself contains very little nutritive capacity for an organism to survive on. Additives, or nutrients, must be added to sustain the growth of organisms. A simple and inexpensive culture medium consists of 15 g of agar, 2 beef bullion cubes, and 1 L of deionized or distilled water. Heat and stir the mixture to a gentle boil. This media will support adequate growth of many bacterial species.
Cultures need to be prepared up to one week in advance of shipping date; therefore orders should be placed at least two weeks in advance of desired delivery date.
The adult flies may die in transit. However, cultures will contain developing larvae and pupae which will emerge within a few days.
Before starting any disposal procedure, please consult with local government authorities first and review the disposal guidelines in a recent Flinn Scientific Catalog/Reference Manual. Never dispose of any chemicals down the drain if the school is not connected to a sanitary sewer system.
Your specimens are probably packaged in a large bucket or pail. If not, transfer them into a pail. We want to rinse and wash away the preservative from the specimen. The room in which this process is undertaken should be well ventilated.
Place the bucket in a large sink. Attach a length of tubing to the cold water outlet and, wearing gloves, force the exit end of the tubing into the very bottom of the bucket. If possible, use a water outlet equipped with a siphon breaker to eliminate the possibility of backflow.
Turn the water on slowly. You may want to start the water flowing before you force the tubing into the bucket to better gauge and control the water flow. A very slow-but steady-flow is desirable. Allow the water to flow into the bottom of the bucket, forcing the preservative to overflow into the sink. Continue for a period of 8-12 hours.
After the wash cycle is complete, turn off the water, and drain all the remaining water from the container. Place the specimens in the garbage as you would any other organic waste.
You may have biological specimens on your premises that are preserved in formaldehyde. You are justly concerned about exposing yourself and your students to the material, yet you do not want to discard the expensive animals. It is possible to displace the present formaldehyde solution with a substitute liquid.
Your specimens were probably packaged in a large container, e.g., jar, bucket, pail, etc. Your first step is to acquire a quantity of formaldehyde substitute material sufficient to replace the volume of liquid in your jar or bucket. Flinn Scientific sells a formaldehyde substitute known as "Formalternate" (Flinn Catalog No. F0056). Formalternate contains no formaldehyde!
Formalternate is a concentrated solution. The instructions accompanying each bottle will direct you to dilute the concentrate by a ratio of 9 parts water to 1 part Formalternate concentrate.
Once the substitute material has been acquired, you are ready to displace the formaldehyde in your jar or bucket. The room in which this process is undertaken should be generously ventilated. Place the jar or bucket in a large sink. Attach a length of tubing to the cold water outlet and, wearing gloves, force the exit end of the tubing into the very bottom of the bucket. We suggest you do this by taping the exit end of the tubing to a long stick and slowly working the stick to the bottom of the container.
Turn the water on very slowly. You may want to start the water flowing before you force the tubing into the bucket to better gauge and control the water flow. A very slow, but steady, flow is desirable.
Allow the water to flow into the bottom of the vessel forcing the formaldehyde preservative to overflow into the sink. Continue overnight or for a period of 10-12 hours.
After the washing cycle is complete, turn off the water, remove the stick and tubing, and drain all the remaining water from the container. Now replace the water with the formaldehyde substitute you acquired earlier.
No, you will not have washed every vestige of formaldehyde from your specimen, but you have removed a great deal of it. When an animal or specimen is opened as part of the dissection process, the residual formaldehyde will still be evident. However, your risk has been drastically reduced. We strongly suggest that you ventilate the room, wear gloves, and continue to implement prudent laboratory practices.
If you are still concerned about the presence of formaldehyde, simply elect to discard any formaldehyde preserved materials and start with new, non-formaldehyde preserved specimens.
Let's now address the issue of museum jars or other display jars. We suggest you do not replace the formaldehyde liquid in such jars. Despite claims to the contrary, the formaldehyde substitutes will not preserve the display animals for decades and decades as is the case with formaldehyde.
To protect against formaldehyde evaporation from these kinds of containers, acquire some paraffin wax, (Flinn Catalog No. P0003). You can also buy this kind of wax at your local grocery store. It is used to seal home prepared canned goods.
To seal your container, melt enough wax in a melting vessel to enable you to completely immerse the cap or closure portion of your museum jars. Allow the hot wax to thoroughly surround the cap of the museum jar. This wax will give you a seal that will last for years and years.
If you have further questions about formaldehyde substitutes, please call or write our technical services department at (800) 452-1261. We will be glad to assist you in any way we can.
Unfortunately, there are companies who provide specimens with what we believe to be high levels of formaldehyde. Instead of removing most of the formaldehyde from the specimen and replacing it with a non-formaldehyde preservative, they simply mask the formaldehyde scent. What this means to you and your students is that you are still being exposed to formaldehyde, but the characteristic odor may not be noticeable. This practice offers a false sense of protection to both you and your students.
Flinn preserved animals have the lowest formaldehyde content of any available. Our preserved animals go through a painstaking and time-consuming process in which virtually all of the formaldehyde is removed before the specimen is transferred to the non-formaldehyde preservative. Our non-formaldehyde preservative has not only replaced the formaldehyde in the animal but also lowers the vapor pressure of any residual formaldehyde still remaining in the animal. Our preserved animals are 99.7% formaldehyde free. The end result is a preserved animal you can safely dissect without fear of being exposed to formaldehyde.
We encourage you to continue to ask questions until you are satisfied the specimens you are using are safe.
Formaldehyde, in its basic form, is a gas. Most science teachers think of formaldehyde as a liquid. The liquid is actually a mixture of formaldehyde gas and water. The most common concentration used in school science departments (particularly biology) is a 37% solution. The solution consists of 37 grams of formaldehyde gas to 100 mL of water. To prevent polymerization of formaldehyde solution about 10-15% of methyl alcohol is added. It is the addition of the methyl alcohol that causes the substance to be called formalin as opposed to formaldehyde. It is correct to use the terms formalin and formaldehyde solution interchangeably. When diluting full-strength (37%) formaldehyde solution, assume it to be 100%. A fixative-strength (10%) solution is therefore a 3.7% solution of formaldehyde gas in water.
Flinn Scientific, Inc. is very concerned about how and where our preserved animals are acquired. Flinn Scientific, Inc. will only do business with reputable and responsible companies. All preserved animals are acquired and handled in strict accordance with U.S. Department of Agriculture (USDA) regulations and guidelines.
We encourage you to have your students work in teams as much as possible in order to reduce the number of specimens you must purchase. Consider also the option of performing demonstration dissections using a color video system.
How to Set Up an Aquarium
How to Maintain a Healthy Aquarium
Some live cultures are not immediately visible upon first glance at the shipping container. Certain organisms do not evenly distribute themselves throughout the culture medium. So, a random sample from the container may turn up nothing but water! Some cultures tend to attach themselves to the bottom or sides of the jar. Amoeba will typically attach to the bottom of the jar. Stentor, Vorticella and Hydra collect on the sides and bottoms of the jars. Place the culture container on a stereo-microscope or under a strong magnifier for a closer look. They may be hard to see, but they are almost certainly there. To dislodge these creatures, gently swirl the contents of the jar and they should break free from the sides. Then pipet them out and place on a slide.
Imagine the learning opportunities you will provide your students by connecting a color video camera to a microscope. The opportunities are limited only by your imagination! Demonstrate dissections that you may not wish the entire class to perform, or point out and clarify features only once instead of doing so for each student. Record demonstrations or lectures for students who are absent. For lab tests using microscope slides, simply project the slides onto the monitor. Never again will you need to set up and check 15 microscopes for a test!
Tired of kids looking at their eyelashes, bubbles, or dust particles? A video camera attached to your microscope will enable you to show your students in advance what they should be looking for. Success is guaranteed, less time is wasted, and students will not get frustrated and lose interest.
We recommend using the Video Flex® system line of cameras. The Video Flex® system consists of an ultra-compact video camera affixed to a 25" flexible gooseneck. Video Flex® includes audio capability and all accessories needed to get it up and running. For a great, multipurpose camera, on or off the microscope, consider the Video Flex® line of cameras.
For more information click on the following link:
Compound microscope objectives in the 90X to 100X range require immersion oil.
Background
The upper limit of the resolving power of light microscopes is slightly above 1000X. Objectives of 90 to 100X, when coupled with a 10X eyepiece, approach that upper limit. Even in the range of 900 to 1000X, a clear image is only possible if every bit of available light is directed through the microscope optics to the viewer's eye. Immersion oils play an essential role in maximizing the amount of light producing the image the viewer sees.
In the airspace between the slide and the objective lens, light is refracted, scattered, and effectively lost. This happens because the refractive index of air (approximately 1.0) is very different from that of glass (approximately 1.5), and light passing through a glass/air interface is refracted (bent) to a large degree. By reducing the amount of refraction at this point, more of the light passing through the slide will be directed to the very narrow diameter lens of the high-power objective. The more light, the clearer the image. Placing a material with a refractive index equal to that of glass in the airspace between slide and objective directs more light through the objective and produces a clearer image. Immersion oils are formulated for just this purpose.
High-power objectives of 90X or higher are almost invariably intended for use with oil and will be engraved with the words, "oil", or "immersion", or "HI" (homogeneous immersion). These objectives are assembled with special sealants that prevent penetration of oil into the lens system. Applying oil to an objective not designed for immersion will ruin the objective.
Immersion oils are commonly available in two viscosities-low viscosity (Type A), and high viscosity (Type B)-and should be labeled with a refractive index of 1.515. The low viscosity oil is applied to the airspace between slide and objective, the high viscosity oil is (less commonly) applied between the condenser and the slide.
Procedure
Low viscosity oil between slide and objective:
1. With low- or medium-power objective, locate a point or area of interest on the slide and center it in the image field.
2. Rotate the objective turret so that the high power objective is just to one side of the slide. Place a single drop of immersion oil (low viscosity, Type A) on the slide (using the circle of light from below as a guide) and place a drop directly on the objective lens. Failure to apply oil to the objective will likely result in trapped air and reduced image quality.
3. Slowly rotate the high power objective into place and adjust the fine focus to fully resolve the image.
High viscosity oil between condenser and slide (optional).
Condensers with a numerical aperture (N.A.) of 1.0 and greater (usually engraved directly on the condenser) are also sealed to prevent oil penetration. Do not immerse condensers with an N.A. less than 1.0.
1. Before placing the slide on the microscope stage, rack the condenser down (using the condenser focusing mechanism) and apply a drop of oil (high viscosity, Type B) to the condenser lens.
2. Apply a drop of oil to the bottom of the slide directly below the specimen, and place the slide on the stage so that the drops will meet when the condenser is raised.
3. Raise the condenser until the drops converge. Follow the steps detailed above to oil the slide to the objective.
Cleanup
Immersion oil should be cleaned from lens and slide surfaces when observations are complete. Oil left on lens surfaces will eventually dry and be very difficult to remove.
1. Carefully wipe oil from all glass surfaces with a folded piece of clean lens paper.
2. With a second piece of lens paper, moistened with a small amount of alcohol (ethyl or isopropyl), wipe glass surfaces to remove any streaks of residual oil.
3. To remove oil that has been allowed to dry on lens surfaces, moisten a folded piece of clean lens paper with a small amount of xylene. Gently wipe lens sufaces, giving the xylene a few moments to work. Xylene may soften cements used to assemble the objective-so wipe the surfaces again with clean lens paper moistened with dilute alcohol or distilled water.
Lens Cleaning:
Most manufacturers recommend the following: Moisten a Kimwipe®* or piece of lens tissue with a small amount of alcohol (ethyl or isopropyl) and wipe the lens gently in one direction. Never apply alcohol or any other solvent directly to the lens as it may work its way into the mechanism and cause unseen damage.
Immersion oil should always be wiped from all surfaces immediately after use. In the event immersion oil is allowed to harden, moisten a wipe with a small amount of xylene and use this to redissolve and remove the hardened oil. Note, xylene may leave a film on the lens and may dissolve the cement used to seal the immersion objective. To prevent this, always moisten a second wipe with alcohol and use this to remove any residual xylene.
* (Dry, unmoistened Kimwipes are not recommended for routine lens cleaning.)
Lamp Replacement:
When replacing lamps or bulbs, avoid touching the glass with bare fingers. Fingerprints left on the bulb will actually "burn into" the glass and reduce the bulb quality and life expectancy.
Microscope Fitness:
It is a good idea, on a weekly basis, for the scope user to put the focusing mechanism through its paces. During normal use the mechanism is probably not worked through its full range of motion. Lubricating grease can build up and harden because it does not remain evenly distributed. To remedy this, rotate both coarse and fine focus knobs from end-stop to end-stop several times. Be sure to rotate the lower power objective into place before carrying out this procedure.
For more information about Cleaning and Maintaining Microscopes, click on the following links to view in-depth videos!
Simple equipment can be sterilized using a bleach or Lysol solution. Glassware can be sterilized by placing in an oven at a temperature of 160-190 °C (320-374 °F) for at least an hour. To sterilize culture media, an autoclave or pressure cooker is required.
Fluorescent bulbs produce very little heat and last 10 times longer than conventional bulbs. Their whiter, more natural light provides true color images with enhanced contrast and resolution.
Use a mixture of 92% petroleum ether and 8% acetone.
