|
Analysis of Foreign DNA in Transgenic Mice
The analytical method that you select depends upon the properties of the transgene that has been microinjected. DNA samples prepared from tail tissue may be analyzed by Southern blot, dot blot or PCR analysis. Southern blot analysis is the preferred method because the results are unambiguous. Southern blot analysis is also the only way to determine whether you have more than one site of transgene integration, and will provide an estimate of the copy number of the gene.
Condition 1. -- The transgene that is microinjected contains a reporter gene that is not normally present in the mouse. In this case, slot or dot blots using a labeled reporter gene for a probe will allow rapid screening for the presence of the transgene. However, if your probe gives a high background, the results may not be interpretable. Low copy number integration events may also not be detected.
Condition 2. -- The transgene that is microinjected contains a structural gene from another species (human vs. mouse) with regions that diverge sufficiently to allow you to distinguish between the two using the appropriate probe. Southern or dot blot analysis is recommended. If your probe gives high background, results from dot or slot blots may not be interpretable. For maintenance of the transgenic mouse line, PCR analysis may be used if primers that are specific to the transgene only are identified. It is always important to use genomic DNA for a positive control rather than plasmid DNA.
Condition 3. – The transgene is a chimeric fusion gene that does not normally exist in nature. PCR may be used to identify a unique fragment that spans the junction. The restriction pattern of the fusion gene will be different from the endogenous gene and may be analyzed by Southern blot analysis.
Condition 4. -- Your transgene is very similar to an endogenous gene. Founders must be identified by Southern blot analysis. The restriction pattern at the integration site will distinguish (Figure A). Potential complications in this situation can be avoided by engineering an identifying marker into the transgene.
Restriction Digestion Strategy The transgene will most often integrate in a head-to-tail array. Selection of an enzyme that cuts once in the transgene will yield fragments that are equivalent to the size of the transgene (Figure A). Because integration into the genome is random, the size of sequences flanking the insertion site (junction fragments) will differ from founder to founder because of the position of the endogenous restriction site. The endogenous gene should generate junction fragments of a different size (Fig. B). Use of 2 unique restriction enzymes provides confirmation. Alternatively, use of an enzyme that cuts twice within the transgene will also produce a fragment of predictable size. Methylation sensitive enzymes can be problematic and their use should be avoided.
Sometimes, double integration events occur, whereby the transgene integrates into different chromosomes. This is clearly detected in the second generation of animals, when chromosomes segregate and the size of sequences flanking the insertion appear as different sizes. This event can occur in ~20% of transgenic founder animals.
Figure A. Random Transgene Integration
Figure B. Endogenous Gene
Restriction Digestion of Genomic DNA
1. Digest 5-10 µg of genomic DNA in the appropriate restriction enzyme, using the conditions recommended by the enzyme vendor.
2. To test whether the DNA is completely digested, set up a second tube in which you take ½ µg of DNA from your original digestion reaction, and add ½ µg of lambda or Ad-2 DNA, depending upon which produces a better restriction band pattern with your enzyme. Add ½ x volume of restriction buffer to a total volume of 2x the ½ µg of genomic DNA.
3. Digest both samples overnight.
4. Run the second digest containing lambda DNA on a mini-gel. If the digestion of lambda DNA is complete, then the 1st sample is also completely digested. DNA that does not digest completely is usually contaminated with proteins. If it is partially digested, 5X excess of enzyme may be added and digestion continued for 4 to 16 hours. If the DNA is mostly undigested, you may need to re-extract with phenol to remove those contaminants, and repeat the digestion. Sometimes precipitation of the DNA with 0.5 x volume of 7.5 M NH4OAc and 2X volume of 100% ethanol will produce DNA clean enough to digest. Alternatively, addition of 4 µl of 0.1 M spermidine/ 100 µl of sample prior to the addition of restriction enzyme will give a complete digestion. Digested DNA samples may be frozen in the freezer until you are ready to run the gel. If the total volume is too large to load into the well, precipitate with 0.1 volume of 3M sodium acetate and 2.5 volumes of ethanol. Re-suspend in an appropriate volume of TE buffer (usually 20 to 35 µl, depending upon the size of the comb you use).
Gel Electrophoresis
1. Run the digested genomic DNA in 0.5% agarose prepared in 1x Tris-acetate buffer. Prior to loading the digested DNA, add 0.1 volume of a 10x gel-loading buffer.
20 X Tris-acetate buffer (TAE)
96.8 g Tris-base
27.2 g NaAc*3 H2O
7.4 g Na2-EDTA
Dissolve in 900 ml dd H2O. Titrate to pH 7.8 with glacial acetic acid (30-35 ml/l)
Adjust volume to 1 liter with ddH2O and autoclave.
Add 0.5 µg/ml ethidium bromide to 1X TAE running buffer. [IMPORTANT NOTE: Ethidium Bromide is a potent mutagen. Wear gloves when handling gels or buffers containing it. Consult with your biosafety officer for guidelines regarding inactivation and disposal.]
Gel loading buffer: Prepare in a 1.5 ml Eppendorf tube.
4 mg bromophenol blue
4 mg xylene cyanol
0.5 ml glycerol
0.5 ml dd H2O
Store at 4 °C
The gel is generally run slowly overnight. Usually, we start it late in the afternoon. Load digested DNA samples onto the gel. Also run size markers (1 kb ladder, or size markers suitable for the band which you expect to see). Run the gel 15 to 30 minutes at 40 volts to run your samples into the gel and verify that it is running in the right direction. Reduce the voltage to 20 volts and run overnight. As a rule of thumb, run the gel at 0.5 to 5 volts per cm of gel length to get the best band resolution.
Photograph the gel prior to gel transfer. It is useful to place a ruler next to the gel prior to photographing the gel so that you can measure migration from the well to determine sizes of your band.
Southern Blot Analysis
1. After photographing the gel, wash for 10 minutes in 0.2N HCl (depurination buffer). Place the gel in the gel tray in a clean glass pyrex dish or a plastic Tupperware container. Pour enough buffer over the gel to submerge the gel. Place it on a horizontal platform shaker and shake gently. If the gel does not float freely in the buffer, add more buffer. This step is particularly important if you are interested in bands larger than 10 kb. Skipping this step will prevent the transfer of high molecular weight DNA. Remove the gel in its gel tray, and pour off the depurination buffer.
2. Wash the gel for 15 minutes at room temperature in denaturation buffer (0.2N NaOH, 0.6 M NaCl). As before, add enough buffer so that the gel will float freely. Pour off the depurination buffer. Repeat.
3. Add enough Neutralization buffer (1.5 M NaCl; 0.5 M Tris-base, pH 7.4) to allow the gel to float freely. Shake on horizontal platform shaker for 15 minutes. Pour off buffer. Repeat.
4. Pre-cut nylon membrane and three sheets of Whatman paper to the size of the gel. (Any number of vendors sell membrane for DNA transfer). Pre-wet the membrane with water, then transfer into 10X SSC. Set aside.
20X SSC: 701.2 g NaCl and 352.8 g of sodium citrate into 4 liters of water. Adjust pH to 7.0. Autoclave. Store at room temperature.
5. Prepare a rectangular pyrex dish for transfer (Figure C). Invert a spare gel tray in the pyrex dish and cut a wick using Whatman paper that covers the gel tray and extends over the ends of the gel tray to the bottom of the pyrex dish. Add 10X SSC to the bottom of the pyrex dish. Allow the 10X SSC to wick up the ends of the Whatman paper. Once the Whatman paper is completely wet, carefully slide the gel onto the Whatman paper positioned over the gel tray. Using gloved fingertips, press out any bubbles between the gel and the Whatman paper.
6. Flood the gel surface with 10X SSC and overlay with pre-wetted nylon membrane. Use a clean plastic or Pasteur pipette like a rolling pin to press bubbles out between the membrane and the gel.
7. Flood the membrane surface with more 10X SSC and overlay with the pre-cut Whatman paper briefly pre-wetted with 10X SSC. Use the pipette to roll out any bubbles. Flood surface of Whatman paper with 10X SSC and overlay with a second pre-cut, pre-wetted Whatman paper. Repeat procedure a third time with the third Whatman paper.
8. Overlay with a stack of blotting pads (these can be ordered from VWR) cut to the same size as the gel. Stack several inches of blotting pad on top. Pour additional 10X SSC buffer into the pyrex dish reservoir. The DNA is transferred by capillary action as the buffer wicks up through the blotting pads. There should be a sufficient number of pads such that the pads are not saturated the following morning. Allow the DNA to transfer overnight.
9. The following morning, remove the blotting pads. Discard wet blotting pads, and keep the dry ones to use for subsequent experiments. Remove the Whatman paper, exposing the nylon membrane. Using a pencil or a permanent marker, use the gel as a guide to mark the location of the sample wells onto the membrane.
10. Place the still damp membrane onto a clean sheet of Whatman paper and UV crosslink the DNA (any number of companies now sell UV crosslinkers). The membrane may be stored in a clean plastic sealing bag or between 2 clean sheets of Whatman paper in the refrigerator until you are ready to probe.
Figure C
Radiolabeling the DNA probe
The DNA probe may be labeled using any commercially available kit. We have used both the Amersham Pharmacia Biotech Multiprime Labeling System (RPN 1601Y) and the Gibco BRL RadPrime DNA Labeling Kit (#18428-011). It is important to denature the plasmid DNA prior to labeling, and to denature again after the labeling reaction is complete.
Membrane Hybridization
There are many hybridization buffers that have been successfully utilized for Southern blot analysis. Below is the recipe for a buffer that we find works particularly well in our hands. Prepare 10 ml for each membrane. (Volume will vary depending upon whether you hybridize in a bag or use a commercially available hybridization oven.)
Working Concentration Stock Concentration
50% deionized formamide 5 ml
1% BSA 0.4 ml 25% BSA, prepared in autoclaved H2O
1 mM EDTA 0.02 ml 0.5M EDTA, autoclaved
500 mM Na2HPO4, pH 7.2 2.5 ml 2M NA2HPO4
5% SDS 2.0 ml 25%
10 ml total volume
2M Na2HPO4 134 g Na2HPO4 7H2O and 4 ml of 85% H3PO4 in 500 ml. Autoclave and store at room temperature.
1. Prehybridize the membrane in the hybridization buffer for at least 1 hour.
2. Add the denatured radiolabeled probe to fresh hybridization buffer (usually 1 to 2 x 106 cpm/ml of buffer).
3. Hybridize in a hybridization oven or in a heat-sealed plastic bag overnight at 42 °C. It is important to remove as many bubbles as possible to maximize contact between the membrane and the hybridization buffer.
4. The following day, wash the blot in:
-2X SSC, 0.1% SDS at room temperature for 5 minutes on a horizontal platform shaker in a covered tupperware container, or by rotating in the hybridization oven.
-2X SSC, 0.1% SDS at room temperature for 15 minutes. Repeat.
-0.1X SSC, 0.1% SDS at room temperature for 15 minutes. Repeat.
-Monitor the counts coming off the membrane with a handheld Geiger counter. Stop washing when the counts drop to 200 to 700 cpm. If the counts are above this, continue washing as outlined below. Be sure to check the counts between each change of wash buffer!
-2X SSC, 0.1% SDS at 65 °C for 15 minutes. Check counts. Repeat wash if necessary.
-0.1X SSC, 0.1% SDS at °C for 15 minutes. Check counts. Repeat wash if necessary. Repeat washes up to a total wash time of 1 hour if necessary. If your counts are still too high, there may be a problem with high background from your probe.
-Wrap the damp, washed membrane in plastic wrap, and place in a X-ray film cassette. In a light-tight, darkened room, place x-ray film (Kodak XAR-2) next to the wrapped membrane, and a single intensifying screen. Close the X-ray film cassette holder, making sure that it is light-tight, and store overnight to several days at –80 °C to expose the film
-Develop the exposed X-ray film in an X-ray film processor. Alternatively, if the signals are good, a phosphorimager may
|