Question

Methods – What is the procedure? What is measured? Why would it be used?

1. Southern blot analysis

2. Restriction enzyme digestion

3. Primer extension

4. Electron microscopy

5. PAGE

6. PCR

7. Western blot

8. Gel mobility shift assay

9. Immunoprecipitation

10. ChIP

11. Plasmid vector

12. Making transgenic mice

13. Microinjection

14. Electroporation

15. cDNA synthesis

16. Photolithography

Answer 1

1. southern blot analysis :-

Southern blotting was named after Edward M. Southern who developed this procedure at Edinburgh University in the 1970s. To oversimplify, DNA molecules are transferred from an agarose gel onto a membrane. Southern blotting is designed to locate a particular sequence of DNA within a complex mixture. For example, Southern Blotting could be used to locate a particular gene within an entire genome.

Procedure :-

1. Digest the DNA with an appropriate restriction enzyme.

2. Run the digest on an agarose gel.

3. Denature the DNA (usually while it is still on the gel).
For example, soak it in about 0.5M NaOH, which would separate double-stranded DNA into single-stranded DNA. Only ssDNA can transfer.

A depurination step is optional. Fragments greater than 15 kb are hard to transfer to the blotting membrane. Depurination with HCl (about 0.2M HCl for 15 minutes) takes the purines out, cutting the DNA into smaller fragments. Be aware, however, that the procedure may also be hampered by fragments that are too small.

Be sure to neutralize the acid after this step, or the base after the prior step if you don't depurinate.

4. Transfer the denatured DNA to the membrane. Traditionally, a nitrocellulose membrane is used, although nylon or a positively charged nylon membrane may be used. Nitrocellulose typically has a binding capacity of about 100µg/cm, while nylon has a binding capacity of about 500 µg/cm. Many scientists feel nylon is better since it binds more and is less fragile. Transfer is usually done by capillary action, which takes several hours. Capillary action transfer draws the buffer up by capillary action through the gel an into the membrane, which will bind ssDNA.

You may use a vacuum blot apparatus instead of capillary action. In this procedure, a vacuum sucks SSC through the membrane. This works similarly to capillary action, except more SSC goes through the gel and membrane, so it is faster (about an hour). (SSC provides the high salt level that you need to transfer DNA.)

After you transfer your DNA to the membrane, treat it with UV light. This cross links (via covalent bonds) the DNA to the membrane. (You can also bake nitrocellulose at about 80C for a couple of hours, but be aware that it is very combustible.)

5. Probe the membrane with labeled ssDNA. This is also known as hybridization.
Whatever you call it, this process relies on the ssDNA hybridizing (annealing) to the DNA on the membrane due to the binding of complementary strands.
Probing is often done with 32P labeled ATP, biotin/streptavidin or a bioluminescent probe.

A prehybridization step is required before hybridization to block non-specific sites, since you don't want your single-stranded probe binding just anywhere on the membrane.

To hybridize, use the same buffer as for prehybridization, but add your specific probe.

6. Visualize your radioactively labeled target sequence. If you used a radiolabeled 32P probe, then you would visualize by autoradiograph. Biotin/streptavidin detection is done by colorimetric methods, and bioluminescent visualization uses luminesence.

32P labeled ATP :-
Treat the dsDNA fragment that you are using as a probe with a limiting amount of Dnase, which causes double-stranded nicks in DNA. Add 32P, dATP, and other dNTPs to DNA polymerase I, which has 5' to 3' polymerase activity and 5' to 3' exonuclease activity.

2. Restriction enzyme digestion :-

Restriction Digestion is the process of cutting DNA molecules into smaller pieces with special enzymes called Restriction Endonucleases (sometimes just called Restriction Enzymes or RE's). These special enzymes recognize specific sequences in the DNA molecule (for example GATATC) wherever that sequence occurs in the DNA.
Restriction Digests begin by mixing the DNA and the RE, but it's unfortunately not quite as simple as that. Restriction Enzymes are delicate and need to be treated carefully. Because enzymes are proteins and proteins denature as the temperature is increased, RE's are always stored in a freezer until they are used. In fact, all of the ingredients in a Restriction Digest are kept on ice until it's time for the reaction to begin. The actual reaction conditions vary from one enzyme to the next, and include temperature, NaCl and/or MgCl2 concentration, pH, etc. All of these variables except temperature are optimized by mixing the enzyme and DNA with a buffer specific for the enzyme of choice.Once all the ingredients are mixed in the reaction tube, the tube is incubated at the Restriction Enzyme's optimal temperature for 1 hour or longer. Then finally when the digest has run for the appropriate amount of time, the reaction tube is put back on ice to prevent nonspecific degradation of your DNA. Once the Restriction Digest is completed, Agarose Gel Electrophoresis is performed to separate the digest fragments by size and visualize the fragments and perhaps purify them for further experiments.

3. Primer extension :-

Primer extension is another technique used to analyze RNA structure and expression. In this method, an oligonucleotide primer is annealed to RNA and extended to a cDNA copy by reverse transcriptase in the presence of labeled dNTPs. Alternately, the primer is labeled and no label is included in the extension reaction. If the RNA of interest is present, extended products will appear on a denaturing gel. Furthermore, the size of the extended product will indicate the position of the 5' end of the RNA, and, if an excess of primer is used, the amount of cDNA produced will reflect the amount of target RNA in the sample.

Primer extension provides the same type of information as S1 mapping. However, primer extension is unaffected by splice sites. In cases where only a genomic probe is available and an intervening splice site prevents S1 mapping of the start site, primer extension offers a useful alternative. Primer extension offers additional advantages over S-1 mapping. A genomic clone of the target RNA is not required; only 30 - 50 bases of sequence need be known to generate the primer. Additionally, probe preparation is easier, because the primer is single stranded. This means that no elaborate procedures are needed prior to labeling.

Procedure :-

Primer Selection and Preparation:
Select a priming site that is 30 - 50 bases long, containing no self-complementary sequences. The site should be within 150 bases of the transcriptional start site, as reverse transcriptase has a tendency to find pause/termination sites in larger transcripts.

End-label the primer using 32P ATP and T4 polynucleotide kinase. Use the buffer and protocol recommended by the enzyme supplier for best results. Labeling of 100 µg of primer should incorporate 1-5x107 cpm, or 3x105 cpm/µg.

Remove unincorporated label by 3 rounds of precipitation with 1 volume 4M ammonium acetate and 10 volumes ethanol. Precipitate for 30 minutes @ -70°C, and redissolve in 30 µl water between precipitations.

Hybridization:
Add 0.1µg (3x104 cpm) of labeled probe to 50 µg RNA sample in 100 µl. Add 0.1 volume 3M sodium acetate, and 2.5 volumes ethanol, and precipitate for 30 minutes at room temperature. Pellet, remove supernatant and allow pellet to air dry for 15 minutes. Overdrying will make redissolving the pellet difficult

Redissolve RNA/probe in 30 µl of hybridization buffer (3M NaCl, 0.4M HEPES [pH 7.6], 1 mM EDTA).

Hybridize overnight at 30 - 50°C (optimize temperature to reduce background).

Precipitate 30µl hybridization with 150µl 0.3M sodium acetate and 500µl of ethanol. Wash pellet with 70% ethanol containing 30mM sodium acetate pH 5.3. Remove supernatant and allow pellet to air dry 15 minutes.

Primer Extension Reaction:
Redissolve sample pellet in a mixture of 18 µl H2O, 2.6 µl 10X RT buffer, 3.5 µl 4mM dNTP's and 2 µl RNase inhibitor.

Add 400 units of reverse transcriptase (AMV). Allow reaction to proceed at 42°C 1.5 hours.

Stop reaction with 1 µl 500 mM EDTA.

Digest substrate RNA with 1µg (1 µl of 1mg/ml) RNase A to prevent gel distortions. Digest 1 hour at 37°C. Extract reactions with phenol and then ethanol precipitate.

Redissolve in 5 µl of water, add denaturing loading buffer and analyze 2-5 µl on a denaturing PAGE gel.

4. Electron microscopy :-

Electron microscopy (EM) is a technique for obtaining high resolution images of biological and non-biological specimens. It is used in biomedical research to investigate the detailed structure of tissues, cells, organelles and macromolecular complexes. The high resolution of EM images results from the use of electrons (which have very short wavelengths) as the source of illuminating radiation. Electron microscopy is used in conjunction with a variety of ancillary techniques (e.g. thin sectioning, immuno-labeling, negative staining) to answer specific questions. EM images provide key information on the structural basis of cell function and of cell disease.  

There are two main types of electron microscope – the transmission EM (TEM) and the scanning EM (SEM). The transmission electron microscope is used to view thin specimens (tissue sections, molecules, etc) through which electrons can pass generating a projection image. The TEM is analogous in many ways to the conventional (compound) light microscope. TEM is used, among other things, to image the interior of cells (in thin sections), the structure of protein molecules (contrasted by metal shadowing), the organization of molecules in viruses and cytoskeletal filaments (prepared by the negative staining technique), and the arrangement of protein molecules in cell membranes (by freeze-fracture).

Conventional scanning electron microscopy depends on the emission of secondary electrons from the surface of a specimen. Because of its great depth of focus, a scanning electron microscope is the EM analog of a stereo light microscope. It provides detailed images of the surfaces of cells and whole organisms that are not possible by TEM. It can also be used for particle counting and size determination, and for process control. It is termed a scanning electron microscope because the image is formed by scanning a focused electron beam onto the surface of the specimen in a raster pattern. The interaction of the primary electron beam with the atoms near the surface causes the emission of particles at each point in the raster (e.g., low energy secondary electrons, high energy back scatter electrons, X-rays and even photons). These can be collected with a variety of detectors, and their relative number translated to brightness at each equivalent point on a cathode ray tube. Because the size of the raster at the specimen is much smaller than the viewing screen of the CRT, the final picture is a magnified image of the specimen. Appropriately equipped SEMs (with secondary, backscatter and X-ray detectors) can be used to study the topography and atomic composition of specimens, and also, for example, the surface distribution of immuno-labels.