The amplification of DNA fragments using the polymerase chain reaction (33) is performed in either the Perkin-Elmer Cetus DNA Thermal Cycler or the Perkin-Elmer Cetus Cycler 9600, by adding the following reagents to either a 0.2 ml thin-walled tube or a 1.5 ml tube, respectively: a small amount of the template DNA molecule (typically cosmid, plasmid, or genomic DNA), the two primers flanking the region to be amplified, nucleotides, buffer, and Taq DNA polymerase. The cycling protocol consisted of 25-30 cycles of three-temperatures: strand denaturation at 95degC, primer annealing at 55degC, and primer extension at 72deg C, typically 30 seconds, 30 seconds, and 60 seconds for the DNA Thermal Cycler and 4 seconds, 10 seconds, and 60 seconds for the Thermal Cycler 9600, respectively. For reactions performed in the DNA Thermal Cycler, the reaction mixtures are overlaid with two drops of mineral oil prior to temperature cycling to eliminate liquid evaporation and condensation. This is not necessary for the Thermal Cycler 9600, which is equipped with a heated lid, maintained at 100degC, that closely contacted the sample tube caps and eliminated liquid evaporation and condensation. After PCR, aliquots of the mixture typically are loaded onto an agarose gel and electrophoresed to detect amplified product. In some instances where the yield from a single PCR is insufficient, the reaction is ethanol precipitated, resuspended, and an aliquot is used as template for a second round of PCR amplification.
1. Add the following reagents to a 0.5 ml flat-topped microcentrifuge tube :
1 ul target DNA (10-20 ng) 2.5 ul each primer (40 uM ) 1 ul AmpliTaq DNA Polymerase (5 U) 10 ul 2 mM dNTPs (2 mM each dNTP) 10 ul 10X PCR buffer 73 ul ddH20 100 ul2. Cover the reaction with 2 drops of mineral oil, add a drop of oil to the heat-block well to ensure good contact between the heat-block and the tube, and place the tube in the wells of a Perkin-Elmer Cetus DNA Thermal Cycler which has been pre-heated to 95degC (Soak file #19).
3. Abort the soak file program and begin thermal-cycle program #43 for the amplification. This program has 25 cycles of a three temperature program and is linked to a 4degC soak file, which will hold indefinitely:
95degC for 1 minute4. Analyze a 10 ul aliquot on a 1-2% agarose gel.
55degC for 1 minute
72degC for 2 minutes
After an aliquot of the PCR mixture is analyzed on an agarose gel, the remainder of the reaction is concentrated by ethanol precipitation, resuspended in buffer, and subjected to a simultaneous fill-in/kinase reaction with the Klenow fragment of E. coli DNA polymerase and T4 polynucleotide kinase, the four deoxynucleotides and rATP (34). The reaction then is loaded onto a preparative 1, 1.5, or 2% low-melting temperature agarose gel, depending on the size(s) of the fragment(s) as determined above, and after minimal separation is achieved between the product(s) and the excess primers, the DNA fragments are excised and eluted. After concentration by ethanol precipitation, amplified DNA fragments are ligated into blunt-ended cloning vector, such as SmaI-linearized, dephosphorylated double-stranded M13 replicative form or pUC.
It should be noted that several other methods to purify the DNA fragments for cloning have been investigated. These included standard ethanol-acetate precipitation (1), a 50% ethanol precipitation (35), spin-column purification (36), and precipitation with polyethylene glycol (PEG) (37). The first three methods did not remove sufficient unincorporated primer, and, during the subsequent ligation of the DNA fragment, the primers apparently competed for the blunt ended vector during ligation because the efficiency of ligation was significantly lower and the vast majority of recombinant clones contained only primer derived inserts. Precipitation by polyethylene glycol resulted in only an extremely small DNA pellet, removing the PEG supernatant is difficult, and yields of PCR product were variable.
1. Ethanol precipitate the PCR reaction by adding 2.5 volumes of 95% ethanol containing 0.12 M sodium acetate, pH 4.8.
2. Perform the combined fillin-kinase reactions previously described. 3. Add 5 ul of agarose gel loading dye and load the reaction into a well of a 1.0% low-melting temperature agarose gel. Electrophorese for 30-60 minutes at 100-120 mA, and then excise the desired band visualized under UV light with a clean razor blade.
4. Elute the DNA from the gel by standard freeze-thaw methods, followed by a phenol extraction and concentrate by ethanol precipitation.
5. Resuspend the dried DNA in 10 ul of 10:0.1 TE buffer. Use this DNA in a standard blunt-ended ligation reaction. Typically, use 2-3 ul of this DNA in a 10 ul ligation reaction with 20 ng of pUC 18/SmaI-CIAP, although this will depend on the yield of amplified DNA from the PCR reaction.
Several small batches of M13RF double stranded DNA are mixed with buffer, SmaI restriction endonuclease, and calf intestinal alkaline phosphatase (CIAP) and incubated for 2-4 hours at 37degC. The reactions then are pooled, ethanol precipitated, and resuspended in buffer to yield a final concentration of about 10 ng/ul. After characterization to determine the optimal concentration for shotgun cloning ligations and to assess the efficiency of SmaI digestion and CIAP dephosphorylation, aliquots of linearized vector are stored frozen at -70degC.
1. Prepare 10-20 tubes with the following:
5 ug M13RF 2 ul NEB Buffer #4 4.5 ul SmaI (16 U/ul) 3 ul Calf Intestinal Alkaline Phosphatase (CIAP) q.s. ddH2O 20 ulSmaI from New England Biolabs (141L) and CIAP from Boehringer Mannheim (1097 075). NEB Buffer #4 (500 mM potassium acetate, 200 mM Tris-acetate, 100 mM magnesium acetate, 10 mM dithiothreitol, pH 7.9) included with SmaI from New England Biolabs.
2. Incubate at 37degC for 2-4 hours.
3. Pool the reactions, phenol extract, ethanol precipitate, and resuspend the dried DNA in 10:0.1 TE buffer to a final concentration of 10 ng/ul.
Alternatively: Preparation of M13 vector DNA for ligation M13 vectors should be digested with restriction enzymes as follows:
M13 DNA (1ug/ml) 1 µl 10x Assay buffer 1 µl ddH2O 7 µl Hinc II 1 µl total volume 10 µl (vector concentration of 100 ng/µl)Incubate at 37¡ C for 2 to 4 hours. Inactivate restriction enzyme by heating at 75¡ C for 10 minutes. If dephosphorylation of the M13 vector DNA is desired, it may be performed at this point (see dephosphorylation procedure, above). Add 90 µl water to give final concentration of 10 ng./µl; use 5 µl (50 ng.) per ligation. The linearized vector may be stored at -20¡ C.
Oligonucleotide primers, either to close gaps in a cosmid or plasmid sequencing project or for PCR, are chosen by manual observation using the following criteria:
1. A relatively even base distribution (about 50% GC) is desired, with no obvious repeated motifs.
2. When possible the 3' end of the primer contained either a G or a C residue.
3. To determine if the sequence chosen is unique, the sequence is compared to the available cosmid or plasmid sequence using Findpatterns.
Alternatively, several computer programs, the SPARCstation-based ospX , the Macintosh-based HYPER PCR and Primer, or the VAX-based Primer programs, could be employed and yield similar results.
Oligonucleotides are synthesized according to manufacturer's procedures on either the Beckman Oligo 1000 (38) or the ABI 392 synthesizer (39) using the phosphoramidite chemistry (40). The desired oligonucleotide sequence is entered into the CPU affixed to the respective synthesizer, the reagent bottles are attached, and the column is inserted which contains the respective 3' nucleotide base-specific linked by the 3-OH group to the solid support, controlled-pore-glass (CPG) silica beads (41,42). The 5'-OH group of the base is blocked with a dimethyloxytrityl (DMT) group. Typically, the columns are purchased for 30 nmoles of primer. For large scale synthesis, for lab stocks of universal primers, columns for 1 umole synthesis are obtained, and can only be used on the ABI 392. The synthesizer automatically performs a cycle of base addition which consists of 5-detritylation to remove the 5-DMT blocking group with trichloroacetic acid and dimethylchloride, activating the phosphoramidite nucleoside monomer with tetrazole and coupling the activated phosphoramidite nucleoside to the column, capping chains that were not coupled during the previous step by acylation of the 5-OH end with acetic anhydride and 1-methylimidazole (43,44), and oxidizing the internucleotide phosphate linkage from the phosphite ester to the more stable phosphotriester with iodine and water (39). After each step in the synthesis, the column is washed with acetonitrile. For the synthesis of fluorescent 5'-end-labeled oligonucleotides, the last base added has an aminolink on the 5' end (45). After synthesis, the oligonucleotide is removed from the solid support, the protecting groups are removed, and the primer is used directly after concentration by butanol precipitation (46).
1. Synthesize the oligonucleotide on either the ABI 392 or the Beckman Oligo 1000 automated DNA synthesizers according to manufacturers' instructions (38,39).
2a. After the cycles of base addition are complete, the ABI 392 automatically detaches the synthesized oligonucleotide from the solid support by adding of ammonium hydroxide and transfers the mixture into a 1.5 ml screw cap microcentrifuge tube.
2b. For the Beckman Oligo 1000, to detach the oligonucleotide from the solid support remove the column and affix it to a screw cap tube containing 1 ml of concentrated ammonium hydroxide, and with a syringe attached to the fluted end of the column, draw the ammonium hydroxide into the column. Allow this assembly to remain at room temperature for at least one hour, no longer than two hours, and after about 30 minutes mix the solution with the syringe. Push the liquid out of the column with the syringe into the tube.
3. Incubate the oligonucleotide in ammonium hydroxide at 70deg C for at least 2 hours (for 1 umole synthesis on the ABI 392, incubate overnight) to deprotect the bases.
4. Transfer 100 ul aliquots of the mixture into 9 microcentrifuge tubes, add 1.25 ml of n-butanol (46), vortex twice for 10 seconds, and centrifuge at 4deg C for 10 minutes at 13,000 rpm.
5. Decant and dry in the Speed-Vac until dry (at least 2 hours or overnight).
6. Add 115 ul of 10:0.1 TE buffer into the first tube, resuspend by pipetting up and down, and then transfer into the second tube, etc, until the dried oligonucleotide in all nine tubes is contained in one tube in about 100 ul.
7. Remove 10 ul of oligonucleotide and dilute with 990 ul of ddH2O and read the A260 in the 1 ml cuvette. The amount of oligonucleotide in the solution in the cuvette is 100 X A260, but the amount of oligonucleotide in the remaining 100 ul of solution is 10 X A260.
For synthesis of fluorescent 5'-end-labeled oligonucleotides:
1. Primers were synthesized on an ABI 392 DNA Synthesizer  at the 1uM synthesis scale with the final detritylation and the end/cleave program DMT ON, AUTO (END-CE) employed (i.e. no final detritylation step, automatic cleaving of the oligo from the column delivering a final volume of 2 ml conc. ammonium hydroxide). The final base to be added is the Aminolink-2 diamine used to couple the fluorescent dye to the oligonucleotide primer. The procedure employed for the synthesis and purification of the aminolinked-primers is as follows:
a. Edit the primer sequence into the synthesizer the final 5' end base being the Aminolink-2 e.g.. 5' 5CA GGA AAC AGC TAT GAC C 3', the 5 representing the Aminolink-2 reagent placed in the bottle 5 position.
b. Dissolve the Aminolink-2 in 3.3 ml dry acetonitrile and place it in the bottle 5 position.
c. Place fresh conc. (30%) ammonium hydroxide solution in the ammonium hydroxide bottle (bottle 10) if needed.
d. START SYNTHESIS on the ABI selecting the relevant sequences to be synthesized, DMT ON, AUTO (END CE) and EXECUTE ABI BEGIN -- YES if synthesizer has not been used in the last 6 hrs.
e. Remember to place a 2 dram vial at the outlet port for collection of the synthesized oligo.
f. When the synthesis and cleaving steps are complete remove the vial, which should contain 2 ml of solution, cap it tightly and leave at 70deg C for 12 hrs. to remove the base protecting groups.
g. Precipitate the DNA using the method of Sawadogo and Van Dyke in which to 1 part DNA/ammonium hydroxide solution 10 parts of n-butanol is added. This is best achieved by aliquoting the DNA solution into 20x1.5ml ependorf tubes of about 100 ul each and adding 1.25 ml n-butanol to each tube, vortex each for 10 sec then centrifuge for 20 min at 13K and 40degC.
h. Pour off the supernatant is poured off, drain the tube on a paper towel for 5 minutes then dry the pellet in a Savant Speed-Vac for at least 2 hrs (overnight preferable to be certain that no ammonium hydroxide or n-butanol remains).
i. To pool together and desalt the following procedure was employed:
i Combine two samples into one (therefore the 20 microcentrifuge tubes are combined resulting in 10 tubes) by dissolving one of the two samples in 40 ml of 1M NaCl and transferring it into the second tube. Further wash the first tube with 32 ml of 1M NaCl and again transfer to the second tube giving a total volume of 72 ml.
ii Then add 84 ml of 95% Ethanol and briefly vortex.
iii Add a further 84 ml 95% Ethanol, vortex and then leave at -20degC for 30 minutes to precipitate.
vi Centrifuge the precipitated samples for 20 minutes at 13K and 4degC, pour off the supernatant and dry the samples in a Savant Speed-Vac for 15 minutes.
j. The aminolink-primer oligonucleotide is now ready for coupling with the four fluorescent dyes and may be stored at -20degC until ready to couple.
2. The next step in the synthesis of the labeled primers is the coupling reaction between the fluorescent dye and the aminolinked-oligonucleotide primer followed by its eventual purification. The four dyes named FAM, JOE, ROX and TAMRA come from ABI, each as a solution in 60 ml DMSO. These dyes (which once diluted must all be used) are further diluted with 440 ml DMF to give a final volume of 500 ml (50 ml/reaction) and are therefore enough for 10 reactions each. Therefore it is best to prepare 4 x 1uM columns worth of aminolinked-oligonucleotide primers as described above which then will be distributed into 40 x 1.5 ml microcentrifuge tubes, 10 reactions for each dye. The following procedure was therefore employed to prepare the fluorescently labeled primers from the aminolinked-oligonucleotides:-
a. First prepare 4 x 1uM columns worth of aminolinked-oligonucleotide as described above yielding 40 x 1.5 ml microcentrifuge tubes of oligonucleotide ready for coupling.
b. Dissolve each of the samples in 50 ml of double distilled water and 50 ml of a 0.5M NaHCO3/Na2CO3 pH=9.
c. Add to each of the four dyes 440 ml DMF (to give a final volume of 500 ml), then briefly vortex and centrifuge.
d. From each dye add 50 ml to each of 10 predissolved aminolinked-oligonucleotide reactions and leave overnight at room temperature in the dark (covered with aluminum foil is sufficient).
e. Pool each of the 10 samples together and elute through a G-25 column to remove any unreacted dye as follows:
i Prepare four G-25 columns by eluting with 100 ml 0.1M NH4OAc (dilute from 8.0M stock).
ii Apply each sample to the column and elute with 0.1M NH4OAc collecting the leading colored band, the second colored band being the unreacted dye.
iii Aliquot the primer fractions into approximately 400 ml lots and precipitate with 1 ml of ethanol/acetate at -20degC for 2 hrs.
iv Centrifuge the samples for 20 minutes at 4degC and 13K, pour off the supernatant and dry the samples in a Savant Speed-Vac.
f. To purify the dye labeled primers from the unlabeled primers the samples were electrophoresed on a 20% polyacrylamide gel as follows:-
i For each of the three primers JOE, ROX and TAMRA prepare one, and for FAM prepare two 0.3 mm/20% polyacrylamide gel with 10 wells each capable of holding a minimum of 25 ml.
ii Pool each of the four sets of primers together in 200 ml deionized formamide/50 ml double distilled water.
. iii Apply 25 ml of dye solution to each of the 10 wells of the respective gels and electrophorese at 2500 V, 22-25 mA for 2.5 hrs.
iv Pry the gels apart and with a razor blade, cut away the colored material, place in a large falcon tube with 2 ml 1X TAE (dilute from 20X stock) and leave overnight at 37degC.
v Remove the solution and aliquot evenly into two of approximately 1 ml each and then wash the residue with 2 ml 0.1M NH4OAc (dilute from 20X stock).
vi Desalt each of the two samples for each of the four dye labeled primers by eluting with 0.1M NH4OAc through a G-25 column (prepared as above) collecting again the colored band.
vii Pool the fractions, measure the A260 and Ax (FAM, 494 nm; JOE, 527 nm; ROX, 586 nm; TAMRA, 558 nm) (the A260 in the 0.1 cm cuvette with the UV lamp and the Ax in the 1 cm cell with the VIS lamp) and then aliquot the solutions into 1.5 ml microcentrifuge tubes, 300 ml per tube. Record the total volume to calculate the OD (see appendix).
viii Dry the samples in a Savant Speed-Vac overnight or leave in a drawer in the dark until dry and then store at -20degC.
1. The Beckman Oligo 1000 DNA synthesizer can also be used to synthesize the primers at the 1uM scale In this case the Aminolink-2 reagent is again dissolved in 3.3 ml of dry acetonitrile and then quickly transferred to the Beckman X bottle and placed on the X port.
2. On START SYNTHESIS the relevant sequence is chosen, synthesis SET SCALE(nmol) - 1000 and FINAL DETRITYLATION - NO set. On completion the aminolink- oligo is manually cleaved from the column using concentrated (30%) ammonium hydroxide 1 ml, the cleaving step taking 1 hr to complete.
3. The resulting solution is then transferred to a 2 dram vial the volume brought up to 2 ml with conc. ammonium hydroxide and the left at 70degC for 12 hr. the remaining steps are as mentioned above.
PAGE Purification of synthetic, fluorescent 5'-end-labeled oligonucleotides
1. Remove DNA collection vial from DNA synthesizer following automatic cleavage from column. Bring the total volume up to 4 ml with fresh concentrated NH4OH. Cap tightly and place at 55deg. C for 4 to 12 hours.
2. Remove vial from 55deg. C water bath and place on ice for 10 to 15 minutes. Transfer the sample to three siliconized microfuge tubes and dry under vacuum for 6-10 hours (until completely dry).
3. Dissolve the contents of one tube in 100 ul of ddH2O. Determine the concentration of the sample by measuring the absorbance at 260 nm.
4. Remove an aliquot of the sample containing approximately 2 A260 and mix with 10-20 ul of dye/formamide/EDTA mix. This will be a sufficient amount to load four 1 cm wells (i.e., 0.5 A260 per well).
5. Prepare a 20% polyacrylamide gel containing 7M urea (20 cm x 40 cm x 0.4 mm). Load the samples and electrophorese at 25mA until the slow blue dye (XC) has migrated about 14 cm from the origin (for a 17-mer).
6. Remove the top glass plate and cover the gel with Saran Wrap. Carefully lift the Saran Wrap and the gel off of the bottom glass plate. Flip the gel over and cover the other side with a second sheet of Saran Wrap. Visualize the DNA bands by UV shadowing and photograph. A 17-mer will migrate halfway between the BP and XC dye bands. (Note: BP=8 nt, XC=28 nt for a 20% gel) Outline the oligonucleotide bands with a marker.
7. Excise the oligonucleotide bands from the gel and place each gel slice in a siliconized 0.5 ml microfuge tube. Add enough TAE or water to cover the gel slice (approximately 200 ul) and place in a dry 37degC incubator overnight.
8. Pool the eluant from all tubes and desalt on a small G-25 column (1 to 2 ml bed volume). Read the absorbance at 260 nm of all fractions. Pool the peak fractions, and re-measure the absorbance. Oligonucleotides may be used directly, or diluted for sequencing or labeling reactions, frozen in small aliquots or dried.
M13 clones carrying complementary inserts (i.e.: in the opposite orientation) may be rapidly screened by clone-to-clone hybridization, followed by analysis on an agarose gel.
1. Set up hybridization reaction as follows:
M13 clone 1 1.0 ul M13 clone 2 1.0 ul 5x Hind/DTT 1.0 ul ddH2O 3.0 ul Total Vol. = 6.0 ul Incubate at 55deg. C for 30 minutes. Note: M13 DNA approximately 0.5 - 1.0 ug/ul2. Add 6 ul of 2x Ficoll/BP/XC dye mix, vortex briefly and load on 0.7% agarose gel. Electrophorese at 90 mA for 1 hour.
3. Visualize DNA bands under UV light. Positives will run slower as a duplex DNA.
Solutions 5x Hind/DTT buffer: Mix equal volumes of 10x Hind with 0.1 M DTT. Store at 4degC. 10x Hind buffer: 0.5 ml 2M Tris-HCl, pH 7.6, 0.7 ml 1M MgCl2, 0.35 g NaCl, distilled water to 10 ml. Store at -20oC 0.1 M DTT: 154 mg dithiothreitol in 10 ml distilled water. Store at -20oC.