In general nucleic acids are relatively easy to work with. Unlike proteins which lose their enzymatic activities very easily, as long as you pay attention to a few important details, nucleic acids are stable. DNA is very stable, while RNA needs a bit more attention to detail. The primary concern is nucleases. For this reason all buffers, pipets, pipetman tips, tubes and anything else that will come into contact with the DNA or RNA should be autoclaved and subsequently handled as a sterile solution. Because fingers are an excellent source of nucleases, you should be careful not to touch anything with bare fingers. Again, you need to be more careful when working with RNA than with DNA. Samples should be kept on ice whenever possible. RNA is best stored frozen, preferably at -70^o C. DNA is usually stored frozen at -20^o C or at 4^o C. DNA should always be stored in TE (10 mM tris-HCl, pH 8.1, 1 mM EDTA). Since most nucleases require magnesium for activity, the EDTA helps inactivate them. RNA is usually stored in distilled water which has been treated with diethyl-pryocarbonate (DEP) to inactivate RNAses. Many RNAses survive autoclaving, so DEP should be used to treat solutions whenever possible.

Make solutions 0.1% DEP, shake, then autoclave the solution. While the solution is still hot -- but not while it is superheated -- shake the solution again. DEP is inactivated by water and heating. It is important to be sure that DEP has been inactivated as it will modify RNA and inactivate enzymes such as reverse transcriptase (used to make cDNA) or kinase (used to label RNA in vitro). For RNAs which will be used for cDNA synthesis it might be desirable to avoid DEP completely. DEP is also highly toxic to humans and should always be handled in a chemical hood. Tris, perhaps the most common buffer for nucleic acid biochemistry is incompatible with DEP, so the best thing for tris solutions is to use DEP treated water when making tris solutions, then autoclave the tris again after addition.

Phenol is used to denature and remove proteins from nucleic acid solutions. Phenol is also quite efficient at removing proteins from your skin, so be sure to wear gloves whenever using phenol. Do not use any tube which is clear. Phenol (and chloroform) will almost certainly melt it!

Following these treatments it is usually a good idea to ethanol precipitate the DNA or RNA.

To prepare phenol, crystalline phenol is warmed to melt it and extracted by shaking with 1 M tris pH 8.1, 20 mM EDTA. The aqueous layer is removed and replaced with fresh 1 M tris, 20 mM EDTA and the extraction repeated. After two or three such extractions, change the aqueous buffer to 20 mM tris pH 8.1, 1 mM EDTA and extract two or three more times. Store the phenol under this buffer. Add 8 hydroxy-quinoline (an antioxidant) to 1%. This also turns the phenol yellow which facilitates following the phenol phase during extractions.

Ethanol precipitation is an easy way to concentrate nucleic acids, or to transfer them into a new set of buffer and salts.

This procedure is good for cleaning up DNA samples before doing sequencing reactions.

DNA and RNA are very easily quantitated by simply measuring the absorbance of the solution at 260 and 280 nm wavelength. Because this is in the ultraviolet range, it is important to be sure that the cuvettes you use are UV transparent (usually that means they're made of quartz instead of glass or plastic). Pure nucleic acids should show a A260/A280 ratio of greater than 1.75 - 1.80. Values lower than this imply contamination with lipid or protein.

An A260 of 1.0 corresponds to a concentration of 40 m g/ml for RNA or 50 m g/ml for DNA.

Reference: Maniatis et al., Molecular Cloning p. 104

To avoid contaminating buffers and expensive enzyme stocks, it is very important that all tubes, pipette tips and solutions be autoclaved before use. Because of nucleases on your fingertips, be careful not to touch pipette tips with your fingers.

These represent 1X concentrations.

Notes:

While most enzymes work in the above buffers, it is critical to recognize that an enzyme produced from one company may not cut in the buffer of the same enzyme produced from a different company. The lab purchases enzymes from at least 4 different sources, so always be aware of which company's enzymes you are using.

Note:

Alternatively, Sephadex G-50 can be packed into a 1.0 ml tuberculin syringe.

Hybridization can be performed in our hybridization oven, or, if it is unavailable, you can use a seal-a-meal bag. The hybridization oven offers several advantages. First, you have greater control over the temperatures. Second, because the hybridization solution is constantly mixed by the rolling of the hybridization tubes in the hybridization oven problems with uneven hybridization are reduced.

Place the filter along the inside of the roller bottle being sure to avoid having large bubbles trapped between the filter and the glass walls of the roller bottle. The minimum volume of hybridization solution for the roller bottles is about 10 mls.

Place nitrocellulose filters in a seal-a-meal bag. Because the filters are very fragile, one approach is to cut the bag open on 3 sides and lay it out flat. Then position the filters on one side and then fold the other side over the filter. Seal the bag on 3 sides, close to the filter so that the buffer will be concentrated around the filter. If the filter is dry, it is usually possible to simply slide it into a bag, and then to seal around it. Pure nitrocellulose (vs nylon based membranes) tends to swell once in solution so don't cram it in.

Prepare 1 liter of wash buffer -- 2 X SSC, 0.1% SDS for DNA-DNA, and 0.1X SSC, 0.1% SDS for RNA-DNA. Remove the hybridization solution and place in the appropriate liquid radioactive waste container. Rinse the hybridization bottle two or three times with wash buffer and discard the waste to the radioactive waste container. Add 10-25 ml wash buffer and return to the hybridization oven. If using bags, cut open the hybridization bag and carefully squeeze the radioactive hybridization solution onto absorbant paper such as a blue diaper or benchcoat. Alternatively use a pasteur pipette to remove the labeled hybridization solution to the appropriate container for liquid radioactive waste. Carefully cut open the rest of the bag, taking extreme care to avoid spreading label everywhere -- work over absorbant paper and wear gloves, a lab coat and your radiation badge. Place the filter directly into a plastic box containing enough room temperature 2X SSC to cover the filter with a good layer of solution. Swish the filter around and discard the now radioactive solution into the radioactive waste. Repeat the 2X SSC rinse at room temperature. Take great care to avoid having the filter dry out even momentarily. This will help minimize problems with background.

Reference: Chirgwin, et al. 1979. Biochemistry 18, 5294.

This procedure is a very general one. It will work for virtually any cell or tissue type. Below is a description of the particular version for Dictyostelium. On the next page is a description for frozen tissue.

To make 100 mls: (Six tubes requires at least 150ml)

Often a more rapid procedure for the isolation of RNA from many samples, or from small amounts of tissues. The following represents an alternative.

The following assumes a 1.0 ml bed volume of oligo-dT. Adjust volumes accordingly.

The following have the advantage that they can be directly treated with DEP, thereby minimizing possible RNAse contamination.

Notes:

Other transfer media like genescreen may be better than nitrocellulose. Using genescreen allows for fixation of nucleic acids to the filter with UV light. Place the filter RNA side down on the UV transilluminator for 5 minutes before baking.

The amount of RNA/well may be increased.

Sometimes hybridization to slots which contain no RNA can occur. This is very strange. One should also confirm, by northern blotting, that hybridization is occurring to a single band and not to ribosomal RNA.

Reference: Rave et al., Nucl. Acids Res. 6:3559 (l979)

Note: Work with formaldehyde and run these gels in the hood. Formaldehyde is a possible carcinogen and can give you a bad headache.

Some reaction conditions may need to be altered depending upon the structure of the RNA. When running the sample on a sequencing gel, M13 can be sequenced and used as the sizing marker. It is generally a good idea to run probe alone with and without S1 as controls as well as a negative control (such as vegetative RNA for a developmental message).

This assay is used (1) to determine the 5' end of a messenger RNA species if one has a probe that spans the putative transcription start site and (2) to quantitate levels of messenger RNA in limited amounts of total RNA at a high sensitivity if one has a probe for the specific message.

The best probe to use is a short (<700bp) single-stranded antisense RNA probe. This is acheived by subcloning a small fragment of the gene of interest into a transcription vector such as bluescript in an orientation such that in vitro transcription will produce antisense transcripts. The DNA template must be linearised 3' to the insert so that run-through transcription does not occur. Alternately, single-stranded antisense oligos can be used as long as they are of sufficient length to be detected on a gel.

For quantification of message a standard curve should be generated using sense RNA transcripts. These are made by driving transcription off of the promoter opposite the one used for antisense probe. Again the plasmid template must be linearised downstream of the insert. The curve can go as low as 1 picogram of sense RNA.

Don't forget to make a control tube containing no RNA.

This method is from Stratagene and uses Bluescript vectors and T3 or T7 polymerases.

For linearized templates with 3' overhangs, it is necessary to blunt end the DNA with Klenow prior to transcription to prevent non-specific initiation. To blunt the template:

This technique generates single-stranded probes that are highly radioactive. They are good replacements for double-stranded DNA probes when you don't want both complementary strands hybridising. The DNA template should be completely linearised downstream of the insert to avoid "run-around" transcription. There is probably a maximum insert size that the RNA polymerase can transcribe without falling off and subcloning of smaller fragments may be necessary. Also, it may be necessary to polish the linearised ends of the template DNA to eliminate nonspecific initiation at 3' overhangs. USE GLOVES AT ALL TIMES!

Large amounts of unlabeled RNA are made using the same method as labeled transcripts except no isotope is included and all four ribonucleotides are used at the same concentration.

Subcloning of DNA restriction fragments utilizes most of the basic techniques required for cloning of DNA: restriction of DNA, phenol extraction to remove enzymes and protein contaminants, ethanol precipitation to concentrate DNA, ligation of DNA fragments, and the introduction of DNA into E. coli.

A typical subcloning operation begins with a large cloned DNA fragment, often from a lambda or cosmid genomic library. The goal is to isolate a particular segment of the cloned DNA from all the others. The first step is to digest both the DNA to be cloned "target DNA" and the vector into which the target DNA is to be inserted with the desired restriction enzyme or enzymes. See the protocol on restriction digestion.

The relative amounts of target and vector DNA to used in ligation needs some discussion. Intuitively it makes some sense that one can "drive" a reaction in the direction of insertion of target DNA into the vector by adding a lot of target DNA. However, there are competing reaction that must be taken into consideration. If the amount of target DNA is too high, ligation of target DNA to itself will occur, resulting in clones containing multiple DNA inserts. If the insert DNA is precious, you may not have enough of it to add large amounts of DNA. When the DNA is precious, excess vector relative to target DNA will maximize utilization of the precious material, but will also usually result in a much higher background of vector molecules which religate without inserted target DNA. If there is a strong selection for vector molecules carrying inserts this would not be a problem. Consequently the ratios of vector to insert DNA depends on the type of cloning. When one is subcloning, the amount of insert DNA available is usually not a problem and a 2 to 3 fold molar excess of insert over target can be used. It is important to realize that all that really matters in cloning is the relative numbers of molecule ends present in the reaction. A microgram of insert DNA which is 10 times larger than the vector will have only 1/10th the number of ends in a microgram of the vector DNA. So a 2:1 ratio means twice the number of ends, and not necessarily micrograms.

The other number than needs to be considered is the concentration at which the DNAs should be ligated. If you're subcloning into a plasmid or circular phage like M13, you want the ligation product to be a circular molecule. That means that after a vector molecule and insert molecule have been linked at one end, the concentration of DNA should be low enough that the odds are good one end of the molecule will be ligated to the other end of the same molecule, rather than to a different molecule.

When doing ligations it is often adventageous to dephosphorylate the vector. This prevents self ligation and significantly decreases backround. Calf intestinal alkaline phosphatase will remove the terminal phosphates. The amount of alkaline phosphatase used depends on the amount and type of 5' ends. Blunt or 5' recessed ends require as much as 100x more enzyme than 5' protruding ends. The number of 5' ends depends on the amount and size of the DNA fragment:

(grams DNA/length DNA in bp x 660) x 2 = moles of 5' ends

For blunt end subcloning use the amount of DNA required for the subcloning. For Maxim and Gilbert sequencing use enough DNA to achieve 30-50 pmoles 5' ends.

The enzyme can be added directly to restiction digests. There are two important criteria:

For blunt or 5' recessed end:

This procedure has generally been replace by electroporation (see the next protocol). It works in the event the electroporation machine is, for some reason, unavailable.

From here on there are two different protocols. One is for use when the vector is a plasmid such as pUC, pBR, bluescript, etc, and the other is for use when the cloning is being done in M13 phage.

2% X-gal is made in N,N-Dimethylformamide

CAW 7/87

ALTERNATIVELY,

1. Grow up a plate of transformants overnight as usual.

2. Lift colonies onto a NC filter by laying the filter down on top of the colonies on the agar plate, starting from the middle and releasing the edges as the filter moistens. Mark the filter and plate with asymmetric needle holes around the edges. Pull the filter back up immediately, without moving it around. The colonies will have been transferred to the filter. Process the filter.

3. Place the agar plates back in the incubator for a few hours or leave at room temp overnight. Enough bacteria are left on the plate to grow the colonies back up. Store the plates at 4 degrees as soon as the colonies have grown back up.

revised 1/95, Trivinos

1/95, Trivinos

This protocol was designed to minimize the number of ethanol precipitation steps so as to maximize the cDNA yield. It follows the Gubler/Hoffman protocol with modifications.