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For more information:

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FOTODYNE Incorporated
950 Walnut Ridge Drive
Hartland, WI 53029

Educational Support

Getting Started

Equipment Suggestions

Agarose Gel Information

Frequently Asked Questions

Micropipettor Accuracy
and Precision

Recipes for Bacteriological Media

Classroom Timelines

Ask Technical Support

Frequently Asked Questions

1. What is the procedure for calibration of a micropipettor?

The first step is to check the accuracy and precision of the micropipettor using a gravimetric method. The simplest procedure measures the weight of water at a known temperature to determine the dispensed volume. See also How to Perform A Micropipettor Accuracy and Precision Check.

If you determine that the accuracy error (i.e. the difference between the actual aspirated volume and the present volume) exceeds the permissible value in the Micropipettor Technical Data chart, the pipette recalibration procedure should be carried out. For assistance with micropipettor recalibration, consult the micropipettor instruction manual or contact FOTODYNE Technical Service.

2. What is the maximum sample volume I can load on a mini-gel using the 6,8,10, or 12 well comb?

Electrophoresis Chamber	Comb	Max Sample Volume
1-1408 Mini Single Cell	6-well 8-well	35 mL 30 mL
1-1409 Mini Dual Cell	10-well 12-well	25mL 20mL

NOTE: Loading volumes are given for a 5 mm thick agarose gel. Typically, 6 mL of agarose are required per millimeter of mini-gel thickness.

3. How long should I run my agarose gel?

Agarose gel running times are quite flexible. That is, you can adapt the time of an electrophoresis run to meet your particular class schedule. For voltage and running time suggestions, consult the Voltage Selection Guide For Agarose Gels.

4. What aperture and exposure settings should I use with my Polaroid camera?

While actual exposure times may vary slightly according to band intensity after staining, suggested camera settings for white light and UV transillumination are given below.

Recommended Camera Settings

Stain	Aperture	Shutter Speed
Methylene Blue/Coomassie Blue (White Light Transillumination)	f/32	1/30 second
Ethidium Bromide (UV Transillumination)	f/5.6	1/2 second

NOTE: Recommended camera settings for both White Light and UV transilluminators using Type 667 film. These settings are recommended as a starting point, actual exposure times may vary according to band intensity after staining.

5. How do I rehydrate air-dried MolecuLab DNA?

Cut (digested) MolecuLab DNAs will be visible as a blue pellet at the bottom of the sample tube. Add the required volume of 1X Gel Loading Dye and flick, pipette, or vortex to dissolve the DNA. For best results, allow the rehydrated DNA to sit undisturbed overnight on the benchtop.

Uncut DNA samples will not be visible in the sample tube because the DNA pellets are transparent. Add the required volume of distilled water to the DNA sample and flick, pipette, or vortex to dissolve the DNA. For best results, allow the rehydrated DNA to sit undisturbed overnight on the benchtop.

MolecuLab DNAs may be rehydrated up to two weeks in advance. Store rehydrated MolecuLab DNAs in the refrigerator for best results.

MolecuLab 103 & 104:
Add 40 mL of distilled water to each tube of Uncut Lambda DNA
Add 20 mL of 1X Gel Loading Dye to each tube of cut, Lambda control DNA.

MolecuLab 105, 106, 107, & 108:
Add 20 mL of 1X Gel Loading Dye to each tube of cut MolecuLab DNA.

MolecuLab 117 & 118:
Add 15 mL of 1X gel loading dye to each tube of cut crane DNA.
Add 15 mL of distilled water to each tube of uncut crane DNA.

6. How should MolecuLab DNA be stored?

Dehydrated MolecuLab DNAs should be stored at room temperature. Dehydrated MolecuLab DNAs have a shelf life of several years when stored properly.

Once rehydrated, MolecuLab DNAs should be used in the classroom within two weeks. Rehydrated DNAs may be stored overnight at room temperature. For longer storage, rehydrated MolecuLab DNAs should be placed in the refrigerator.

7. How do I prepare my own agar plates? Is an autoclave required?

Agar plates can be prepared using a pressure cooker if an autoclave is not available. Media should be dissolved in deionized water and autoclaved or boiled for 20 minutes at 15 psi. See also Recipes for Bacteriological Media.

8. Which DNA stain should I use—Ethidium Bromide or Methylene Blue?

Ethidium bromide is the stain of choice in most research labs, and is also used in many schools. It stains rapidly and allows detection of relatively small amounts of DNA. A UV transilluminator is needed to see ethidium bromide-stained DNA fragments, which appear as bright orange bands against a dark background. Ethidium bromide, however, is a mutagen that requires caution in handling and disposal.

Methylene Blue has gained increasing popularity in the educational setting because it is relatively safe. This stain is not as sensitive as ethidium bromide and therefore requires more DNA for detection. Longer staining and de-staining times are needed. DNA fragments appear as dark blue bands against a lighter blue background and can be seen with white light transillumination.

9. Can I preserve my students' DNA or protein gels?

DNA gels stained with Methylene Blue can be stored with a small volume of buffer in plastic wrap or a resealable plastic bag. Gels will typically last for several weeks and will last longest when stored in the refrigerator.

Protein gels stained with Coomassie Blue or silver stain can be preserved by drying the gels between two sheets of cellophane. Once a gel is dried, it can be kept indefinitely. NOTE: Avoid getting bubbles between the cellophane sheets. Bubbles will cause the gel to crack as it dries.

DNA gels stained with ethidium bromide are difficult to preserve due to rapid diffusion of ethidium bromide stain and DNA-stain complexes from the gel. Photographic preservation is recommended for ethidium bromide-stained gels. For those with a limited budget, photocopies of a single gel photo can be distributed to each student or group.

10. How do I prepare TBE buffer if I want to make one liter of 1X buffer, not the entire ten liters?

TBE Buffer Mix (catalog number E1-1820) contains enough pre-mixed powder to make 10 liters of TBE at 1X concentration or 1liter at 10X concentration. To make a smaller volume of 1X buffer, dissolve 16.4 grams of TBE Buffer Mix in 800 mL of distilled water. Then add water to bring the volume to 1 liter.

11. After staining my agarose gel, I notice that the bands look strange. What could be wrong with the gel?

Punctured wells, incorrectly cast gels, altered buffer pH and other factors can alter the running of DNA fragments through an agarose gel. For help with "problem" agarose gels, check the Gel Electrophoresis Troubleshooting Guide.

12. My class meets for 50-minute periods. How should I plan my experiment?

Unlike the research laboratory, the classroom teaching laboratory is often restricted to short meeting times. However, short class periods need not limit biotechnology experiments such as electrophoresis, bacterial transformation, restriction digestion or PCR. For suggestions, consult our Classroom Biotechnology Timelines.