Methods for DNA sequencing
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Bst DNA polymerase-catalyzed radiolabeled two-step sequencing reactions (26) are modified from those presented earlier (25) by altering the absolute amounts and the relative deoxy/dideoxynucleotide ratios in the termination mixes. Two separate termination mixes provided optimal overlap for sequence data starting in the polylinker and extending to approximately 600 bases from the priming site. This two-step format eliminated the need for the chase required in the Bst one-step reaction (25).
Each extension reaction contained 500-750 ng of Biomek isolated single-stranded DNA, reaction buffer, nucleotide extension mix, oligonucleotide primer (typically M13 (-40) universal sequencing primer, see Appendix D), either [a-32-P]dATP or [a-35-S]dATP and Bst polymerase. After the reactions are extended for 2 minutes at 67℃ and briefly centrifuged, four aliquots are removed and added to the appropriate base-specific termination mix. All nucleotide mixes contained the guanosine nucleotide analog, 7-deaza-dGTP, but differed in their deoxy/dideoxynucleotide ratios to yield fragments ranging in size from the beginning of the polylinker to greater than 300 bases from the primer, or fragments from about 150 to greater than 600 bases from the primer for "short" or "long" mixes, respectively. Following an incubation at 67℃ for 10 min and a brief centrifugation, the reactions are stopped by the addition of dye/formamide/EDTA, and incubated at 100℃. When desired, sequencing reactions are stored at -70℃ prior to the addition of loading dye.
When double-stranded pUC-based subclones are used as templates, the amount of primer is doubled and a denaturing/annealing step is added. Here, 3 μg of plasmid DNA, isolated by either the mini- or midi-prep diatomaceous earth method, is mixed with primer, placed in a boiling-water bath, and rapidly cooled by plunging into an ethanol/dry-ice bath (28). Following an incubation on ice, the remaining sequencing extension reagents (reaction buffer, nucleotide extension mix, either [a-32-P]dATP or [a-35-S]dATP, and Bst polymerase) are added. Reactions are performed as described above for single-stranded sequencing.
Protocol
For single-stranded DNA sequencing:
1. Prepare the following extension reaction in a microcentrifuge tube:
750 ng M13 template DNA
2 μl Bst reaction buffer
2 μl Bst nucleotide extension mix
1 μl oligonucleotide primer (2.5 ng/μl)
0.5-1 μl [alpha]-32-P-dATP or [alpha]-35-S-dATP
1 μl diluted Bst polymerase (0.1 U/μl)
q.s. sterile ddH2O
12 μl
[alpha]-32-P-dATP (PB 10384) and [alpha]-35-S-dATP (SJ 1304) from Amersham.
Dilute the Bst polymerase (BioRad 170-3406) in Bst dilution buffer.
2. Incubate the reactions for 2 minutes at 67℃, and briefly centrifuge to reclaim condensation.
3. Remove 2.5 μl aliquots for each reaction into the four base-specific termination mixes (either short or long), already pipetted into a V-bottomed microtiter plate (Dynatech).
4. Incubate the reactions for 10 minutes at 67℃, and briefly centrifuge to reclaim condensation. It is possible to store the reactions at -70℃ at this stage.
5. Stop the reactions by the addition of 4 μl of agarose gel loading dye and incubate for 5-7 minutes at 100℃.
For double-stranded DNA sequencing:
1. To denature the DNA and anneal the primer, incubate the following reagents in a boiling water bath for 4-5 minutes and rapidly cool the reaction by plunging into an ethanol/dry ice bath.
3 μg plasmid DNA
5 ng oligonucleotide primer
q.s. sterile ddH2O
9 μl
2. Incubate the reaction in an ice-water bath for 5 minutes, and then add the following reagents:
2 μl Bst reaction buffer
2 μl Bst nucleotide extension mix
0.5-1 μl [alpha]-32-P-dATP or [alpha]-35-S-dATP
1 μl diluted Bst polymerase (0.1 U/μl)
15 μl
[alpha]-32-P-dATP (PB 10384) and [alpha]-35-S-dATP (SJ 1304) from Amersham.
Dilute the Bst polymerase (BioRad 170-3406) in Bst dilution buffer.
3. Proceed with the sequencing reaction as described above in steps 2-5 for single-stranded templates To prepare polyacrylamide gels for DNA sequencing, the appropriate amount of urea is dissolved by heating in water and electrophoresis buffer, the respective amount of deionized acrylamide-bisacrylamide solution is added, and ammonium persulfate and TEMED are added to initiate polymerization. Immediately after the addition of the polymerizing agents, the gel solution is poured between two glass plates, taped together and separated by thin spacers corresponding to the desired thickness of the gel, taking care to avoid and eliminate air bubbles. Prior to taping, these glass plates are cleaned with Alconox detergent and hot water, are rinsed with double distilled water, and dried with a Kimwipe. Typically, the notched glass plate is treated with a silanizing reagent and then rinsed with double distilled water. After pouring, the gel immediately is laid horizontally and a well forming comb is inserted into the gel and held in place by metal clamps. The polyacrylamide gels are allowed to polymerize for at least 30 minutes prior to use. After polymerization, the comb and the tape at the bottom of the gel are removed. The vertical electrophoresis apparatus is assembled by clamping the top and bottom buffer wells onto the gel, and adding running buffer to the buffer chambers. The wells are cleaned by circulating buffer into the wells with a syringe and, immediately prior to the loading of each sample, the urea in each well is suctioned out with a mouth pipette.
Each base-specific sequencing reaction terminated with the short termination mix is loaded using a mouth pipette onto a 0.15 mm X 50 cm X 20 cm, denaturing 5% polyacrylamide gel and electrophoresed for 2.25 hours at 22 mA. The reactions terminated with the long termination mix typically are divided in half and loaded onto two 0.15 mm X 70 cm X 20 cm denaturing 4% polyacrylamide gels. One gel is electrophoresed at 15 mA for 8-9 hours and the other is electrophoresed for 20-24 hours at 15 mA. After electrophoresis, the glass plates are separated and the gel is blotted to Whatman paper, covered with plastic wrap, dried by heating on a Hoefer vacuum gel drier, and exposed to X-ray film. Depending on the intensity of the signal and whether the radiolabel is 32-P or 35-S, exposure times varied from 4 hours to several days. After exposure, the films are developed by processing in developer and fixer solutions, rinsed with water, and air dried. The autoradiogram then is placed on a light-box and the sequence is manually read and the data typed into a computer.
Protocol
1. Prepare 8 M urea, polyacrylamide gels according to the following recipe (100 ml), depending in the desired percentage:
4% 5% 6% urea 48 g 48 g 48 g 40% A & B 10 ml 12.5 ml 15 ml 10X MTBE 10 ml 10 ml 10 ml ddH2O 42 ml 39.5 ml 37 ml 15% APS 500 μl 500 μl 500 μl TEMED 50 μl 50 μl 50 μl
Urea (5505UA) is from Gibco/BRL.
2. Combine the urea, MTBE buffer, and water and incubate for 5 minutes at 55℃ and then stir to dissolve the urea.
3. Cool briefly, add the A & B, mix, and degas under vacuum for 5 minutes.
4. While stirring, add the APS and TEMED polymerization agents and then immediately pour in between two taped glass plates with 0.15 mm spacers. (Prior to taping, the notched, front glass plate should be treated with a small amount of silanizing reagent and then rinsed with ddH2O).
5. Insert the well forming comb, clamp, and allow the gel to polymerize for at least 30 minutes.
6. Prior to loading, remove the tape around the bottom of the gel and the well-forming comb. Assemble the vertical electrophoresis apparatus by clamping the upper and lower buffer chambers to the gel plates, and add 1X MTBE electrophoresis buffer to the chambers.
7. Flush the sample wells with a syringe containing running buffer, and immediately prior to loading each sample, flush the well with running buffer using gel loading tips.
9. Load 1-2 μl of sample into each well using a Pipetteman with gel-loading tips, and then electrophorese according the following guidelines (during electrophoresis, cool the gel with a fan):
termination electrophoresis
reaction polyacrylamide gel conditions
short 5%, 0.15 mm X 50 cm X 20 cm 2.25 hours at 22 mA
long 4%, 0.15 mm X 70 cm X 20 cm 8-9 hours at 15 mA
long 4%, 0.15 mm X 70 cm X 20 cm 20-24 hours at15 mA
10. After electrophoresis, remove the buffer wells, the tape, and pry the gel plates apart. The gel should adhere to back plate. Blot the gel to a 40 cm X 20 cm sheet of 3MM Whatman paper, cover with plastic wrap, and dry on a Hoefer gel dryer for 25 minutes at 80℃
11. Place the dried gel in a cassette and expose to Kodak XRP-1 film.
12. Develop the film for 1-5 minutes in Kodak GBX developer, rinse in distilled water for 30 seconds, fix in Kodak GBX fixer for 5 minutes, and then rinse again in distilled water for 30 seconds. Allow the film to air dry. Each base-specific fluorescent-labeled cycle sequencing reaction routinely included approximately 100 or 200 ng Biomek isolated single-stranded DNA for A and C or G and T reactions, respectively. Double-stranded cycle sequencing reactions similarly contained approximately 200 or 400 ng of plasmid DNA, isolated using either the standard alkaline lysis or the diatomaceous earth modified alkaline lysis procedures. All reagents except template DNA are added in one pipetting step from a premix of previously aliquotted stock solutions stored at -20℃ (see Appendix B). To prepare the reaction premixes, reaction buffer is combined with the base-specific nucleotide mixes. Prior to use, the base-specific reaction premixes are thawed and combined with diluted Taq DNA polymerase and the individual fluorescent end-labeled universal primers (see Appendix C) to yield the final reaction mixes, that are sufficient for 24 template samples.
Once the above mixes are prepared, four aliquots of single or double-stranded DNA are pipetted into the bottom of each 0.2 ml thin-walled reaction tube, corresponding to the A, C, G, and T reactions, and then an aliquot of the respective reaction mixes is added to the side of each tube. These tubes are part of a 96-tube/retainer set tray in a microtiter plate format, which fits into a Perkin Elmer Cetus Cycler 9600. Strip caps are sealed onto the tube/retainer set and the plate is centrifuged briefly. The plate then is placed in the cycler whose heat block had been preheated to 95℃, and the cycling program immediately started. The cycling protocol consisted of 15-30 cycles of seven-temperatures:
95℃ denaturation
55℃ annealing
72℃ extension
95℃ denaturation
72℃ extension
95℃ denaturation, and
72℃ extension, linked to a 4℃ final soak file.
At this stage, the reactions frequently are frozen and stored at -20℃ for up to several days. Prior to pooling and precipitation, the plate is centrifuged briefly to reclaim condensation. The primer and base-specific reactions are pooled into ethanol, and the DNA is precipitated and dried. These sequencing reactions could be stored for several days at -20℃.
Protocol
1. Pipette 1 or 2 μl of each DNA sample (100 ng/μl for M13 templates and 200 ng/μl for pUC templates) into the bottom of the 0.2 ml thin-walled reaction tubes (Robbins Scientific). Use the 1 μl sample for A and C reactions, and the 2 μl sample for G and T reactions. Meanwhile, preheat the PE Cetus Thermocycler 9600 to 95℃ (Program #2).
2. Prepare the Taq polymerase dilution. AmpliTaq polymerase (N801-0060) is from Perkin-Elmer Cetus.
30 μl AmpliTaq (5U/μl)
30 μl 5X Taq reaction buffer
130 μl ddH2O
190 μl diluted Taq for 24 clones
3. Prepare the A, C, G, and T base specific mixes by adding base-specific primer and diluted Taq to each of the base specific nucleotide/buffer premixes:
A,C/G,T
60/120 μl 5X Taq cycle sequencing mix
30/60 μl diluted Taq polymerase
30/60 μl respective fluorescent end-labeled primer
120/240 μl
4. Seal the reaction tubes carefully with the strip caps, and centrifuge briefly at 2500 rpm. Place the tube/retainer set in the 9600 Cycler, abort the soak file program, and run program #11. This program will cycle the sequencing reactions for 30 cycles of seven temperatures (30 cycles of 95℃ denaturation for 4 seconds; 55℃ annealing for 10 seconds; 72℃ extension for 1 minute; 95℃ denaturation for 4 seconds; 72℃ extension for 1 minute; 95℃ denaturation for 4 seconds; and 72℃ extension for 1 minute), and then will link to a 4℃ soak file until that program is aborted. (It is possible to freeze the reactions at -20℃ after cycling, prior to the pooling step).
5. Briefly centrifuge the plate to reclaim condensation. Pool the four base specific reactions into 250 μl of 95% ethanol.
6. Precipitate the sequencing reactions, and store the dried samples at -20℃.
One of the major problems in DNA cycle sequencing is that when fluorescent primers (1) are used the reaction conditions are such that the nested fragment set distribution is highly dependent upon the template concentration in the reaction mix. We have recently observed that the nested fragment set distribution for the DNA cycle sequencing reactions using the fluorescent labelled terminators (8) is much less sensitive to DNA concentration than that obtained with the fluorescent labelled primer reactions as described above. In addition, the fluorescent terminator reactions require only one reaction tube per template while the fluorescent labelled primer reactions require one reaction tube for each of the four terminators. This latter point allows the fluorescent labelled terminator reactions to be pipetted easily in a 96 well format. The protocol used, as described below, is easily interfaced with the 96 well template isolation and 96 well reaction clean-up procedures also described herein. By performing all three of these steps in a 96 well format, the overall procedure is highly reproducable and therefore less error prone.
Protocol
1. Place 0.5 ug of single-stranded or 1 ug of double-stranded DNA in 0.2 ml Robbins PCR tubes.
2. Add 1 μl (for single stranded templates) or 4 μl (for double-stranded templates) of 0.8 uM primer and 9.5 μl of ABI supplied premix to each tube, and bring the final volume to 20 μl with ddH2O.
3. Centrifuge briefly and cycle as usual using the terminator program as described by the manufacturer (i.e. preheat at 96℃ followed by 25 cycles of 96℃ for 15 seconds, 50℃ for 1 second, 60℃ for 4 minutes, and then link to a 4℃ hold).
4. Proceed with the spin column purification using either the Centri-Sep columns or G-50 microtiter plate procedures given below.
1. Gently tap the column to cause the gel material to settle to the bottom of the column.
2. Remove the column stopper and add 0.75 ml dH2O.
3. Stopper the column and invert it several times to mix. Allow the gel to hydrate for at least 30 minutes at room temperature. Columns can be stored for a few days at 4℃; longer storage in water is not recommended. Allow columns that have been stored at 4℃ to warm to room temperature before use. Remove any air bubbles by inverting the column and allowing the gel to settle. Remove the upper-end cap first and then remove the lower-end cap. Allow the column to drain completely, by gravity. (Note: If flow does not begin immediately apply gentle pressure to the column with a pipet bulb.)
4. Insert the column into the wash tube provided.
5. Spin in a variable-speed microcentrifuge at 1300 g for 2 minutes to remove the fluid.
6. Remove the column from the wash tube and insert it into a Sample Collection Tube.
7. Carefully remove the reaction mixture (20 ml) and load it on top of the gel material. If the samples were incubated in a cycling instrument that required overlaying with oil, carefully remove the reaction from beneath the oil. Avoid picking up oil with the sample, although small amounts of oil (<1 ml) in the sample will not affect results. Oil at the end of the pipet tip containing the sample can be removed by touching the tip carefully on a clean surface (e.g., the reaction tube). Use each column only once.
8. Spin in a variable-speed microcentrifuge with a fixed angle rotor, place the column in the same orientation as it was in for the first spin--this is important because the surface of the gel will be at an angle in the column after the spin.
9. Dry the sample in a vacuum centrifuge. Do not apply heat. Do not overdry. If desired, reactions can be ethanol precipitated.
The following protocol was developed at the C. Elegans Genome Sequencing Center at Washington University, St. Louis, Missouri, was conveyed to us by Dr. Richard Wilson, and now has been modified to take advantage of the Millipore 45 μl Column Loader (cat. # MACL 096 45). Additional information about this procedure also is available at the Millipore web site.
Preparation of Sephadex G-50 containing Microtiter Filter Plates:
1. Add dry Sephadex G-50 to the Millipore (cat.# MACL 096 45) 45 μl Column Loader.
2. Remove the excess of resin from the top of the Column Loader with the scraper supplied.
3. Place MultiScreen HV Plate (Millipore MAHVN4550) upside-down on top of the Column Loader.
4. Invert both MultiScreen HV Plate and Column Loader.
5. Tap on top of the Millipore Column Loader to release the resin.
6. Using a multi-channel pipettor, add 300 μl of ddH2O to each well to swell the resin. and let stand at room temperature for 3 hours.
7. Once the minicolumns are swollen in MultiScreen plates, they can be sealed with saran wrap and stored in the refrigerator at 4 ℃ for several weeks. A batch of plates also can be stored in the refrigerator at 4 ℃ for several weeks in a sealed plastic container with a damp towel to assure the plates are kept moist.
8. When needed, the matrix containing filter plate is taped over a microtiter plate and centrifuged for 2 minutes at 1500 RPM in a Beckman GS-6R to pack the columns and to remove any access buffer.
Packing and using the columns:
1. Sephadex settles out; therefore, you must resuspend before adding to the plate and also after filling every 8 to 10 wells.
2. Add 400 μl of mixed Sephadex G-50 to each well of microtiter filter plate (Millipore MAHVN4550).
3. Place microtiter filter plate on top of another microtiter plate to collect water and tape sides so they do not fly apart during centrifugation.
4. Spin at 1500 rpm for 2 minutes.
5. Discard water that has been collected in the microtiter plate.
6. Again place the microtiter filter plate on top of the microtiter plate to collect water and tape sides so they do not fly apart during centrifugation.
7. Add an additional 100-200 μl of Sephadex G-50 to fill the microtiter filter plate wells.
8. Spin at 1500 rpm for 2 minutes.
12. Spin at 1500 rpm for 2 minutes.
13. Dry the collected effluent in a Speed-Vac for approximately 1-2 hours.
Single-stranded dye-terminator reactions required approximately 2 ug of phenol extracted M13-based template DNA. The DNA is denatured and the primer annealed by incubating DNA, primer, and buffer at 65℃. After the reaction cooled to room temperature, alpha-thio-deoxynucleotides, fluorescent-labeled dye-terminators, and diluted Sequenase[TM] DNA polymerase are added and the mixture is incubated at 37℃. The reaction is stopped by adding ammonium acetate and ethanol, and the DNA fragments are precipitated and dried. To aid in the removal of unincorporated dye-terminators, the DNA pellet is rinsed twice with ethanol. The dried sequencing reactions could be stored up to several days at -20℃.
Double-stranded dye-terminator reactions required approximately 5 ug of diatomaceous earth modified-alkaline lysis midi-prep purified plasmid DNA. The double-stranded DNA is denatured by incubating the DNA in sodium hydroxide at 65℃, and after incubation, primer is added and the reaction is neutralized by adding an acid-buffer. Reaction buffer, alpha-thio-deoxynucleotides, fluorescent-labeled dye-terminators, and diluted Sequenase[TM] DNA polymerase then are added and the reaction is incubated at 37℃. Ammonium acetate is added to stop the reaction and the DNA fragments similarly are precipitated, rinsed, dried, and stored.
Protocol For Single-stranded reactions:
1. Add the following to a 1.5 ml microcentrifuge tube:
4 μl ss DNA (2 ug)
4 μl 0.8 uM primer
2 μl 10x MOPS buffer
2 μl 10x Mn[2+]/isocitrate buffer
12 μl
2. To denature the DNA and anneal the primer, incubate the reaction at 65-70℃ for 5 minutes. Allow the reaction to cool at room temperature for 15 minutes, and then briefly centrifuge to reclaim condensation.
3. To each reaction, add the following reagents and incubate for 10 minutes at 37℃. (For more than one reaction, a pot of the reagents should be made).
7 μl ABI terminator mix (401489)
2 μl diluted Sequenase[TM] (3.25 U/μl)
1 μl 2 mM a-S dNTPs
22 μl
The undiluted Sequenase[TM] (70775) from United States Biochemicals is 13 U/μl and should be diluted 1:4 with USB dilution buffer prior to use resulting in a working dilution of 3.25 U/μl.
4. Add 20 μl 9.5 M ammonium acetate and 100 μl 95% ethanol to stop the reaction and vortex.
5. Precipitate the DNA in an ice-water bath for 10 minutes. Centrifuge for 15 minutes at 12,000 rpm in a microcentrifuge at 4℃. Carefully decant the supernatant, and rinse the pellet by adding 300 μl of 70-80% ethanol. Vortex and centrifuge again for 15 minutes, and carefully decant the supernatant.
6. Repeat the rinse step to insure efficient removal of the unincorporated terminators. (Alternatively, after the first rinse step, droplets of supernatant can be removed by carefully absorbing them with a Q-tip cotton swab or a rolled up Kimwipe).
7. Dry the DNA for 5-10 minutes (or until dry) in the Speedy-Vac, and store the dried reactions at -20℃.
For double-stranded reactions:
1. Add the following to a 1.5 ml microcentrifuge tube:
5 μl ds DNA (5 ug)
4 μl 1 N NaOH
3 μl ddH2O
12 μl
2. Incubate the reaction at 65-70℃ for 5 minutes, and then briefly centrifuge to reclaim condensation.
3. Add the following reagents to each reaction, vortex, and briefly centrifuge:
3 μl 8 uM primer
9 μl ddH2O
4 μl MOPS-Acid buffer
28 μl
4. To each reaction, add the following reagents and incubate for 10 minutes at 37℃. (For more than one reaction, a pot of the reagents should be made).
4 μl 10X Mn[2+]/isocitrate buffer
6 μl ABI terminator mix
2 μl diluted Sequenase[TM] (3.25 U/μl)
1 μl 2 mM [alpha]-S-dNTPs
22 μl
The undiluted Sequenase[TM] from United States Biochemicals is 13 U/μl and should be diluted 1:4 with USB dilution buffer prior to use resulting in a working dilution of 3.25 U/μl.
5. Add 60 μl 8 M ammonium acetate and 300 μl 95% ethanol to stop the reaction and vortex.
6. Precipitate the DNA in an ice-water bath for 10 minutes. Centrifuge for 15 minutes at 12,000 rpm in a microcentrifuge at 4℃. Carefully decant the supernatant, and rinse the pellet by adding 300 μl of 80% ethanol. Vortex and centrifuge again for 15 minutes, and carefully decant the supernatant.
7. Repeat the rinse step to insure efficient removal of the unincorporated terminators. (Alternatively, after the first rinse step, droplets of supernatant can be removed by carefully absorbing them with a Q-tip cotton swab or a rolled up Kim-wipe).
8. Dry the DNA for 5-10 minutes (or until dry) in the Speedy-Vac. Polyacrylamide gels for fluorescent DNA sequencing are prepared as described above except that the gel mix is filtered prior to polymerization. Optically-ground, low fluorescence glass plates are carefully cleaned with hot water, distilled water, and ethanol to remove potential fluorescent contaminants prior to taping. Denaturing 6% polyacrylamide gels are poured into 0.3 mm X 89 cm X 52 cm taped plates and fitted with 36 well forming combs. After polymerization, the tape and the comb are removed from the gel and the outer surfaces of the glass plates are cleaned with hot water, and rinsed with distilled water and ethanol. The gel is assembled into an ABI sequencer, and the checked by laser-scanning. If baseline alterations are observed on the ABI-associated Macintosh computer display, the plates are recleaned. Subsequently, the buffer wells are attached, electrophoresis buffer is added, and the gel is pre-electrophoresed for 10-30 minutes at 30 W.
Prior to sample loading, the pooled and dried reaction products are resuspended in formamide/EDTA loading buffer by vortexing and then heated at 90℃. A sample sheet is created within the ABI data collection software on the Macintosh computer which indicated the number of samples loaded and the fluorescent-labeled mobility file to use for sequence data processing. After cleaning the sample wells with a syringe, the odd-numbered sequencing reactions are loaded into the respective wells using a micropipettor equipped with a flat-tipped gel-loading tip. The gel then is electrophoresed for 5 minutes before the wells are cleaned again and the even numbered samples are loaded. The filter wheel used for dye-primers and dye-terminators is specified on the ABI 373A CPU, also where electrophoresis conditions are adjusted. Typically electrophoresis and data collection are for 10 hours at 30W on the ABI 373A that is fitted with a heat-distributing aluminum plate in contact with the outer glass gel plate in the region between the laser stop and the sample loading wells (26).
After data collection, an image file is created by the ABI software which related the fluorescent signal detected to the corresponding scan number. The software then determined the sample lane positions based on the signal intensities. After the lanes are tracked, the cross-section of data for each lane are extracted and processed by baseline subtraction, mobility calculation, spectral deconvolution, and time correction. On the Macintosh computer, the collected data can be viewed in several formats. The overall graphics image of the gel can be displayed to assess the accuracy of lane tracking, and the data from each sample lane can be viewed as either a four-color raw fluorescent signal versus scan number, as a chromatogram of processed sequence data, or as a string of nucleotides. After processing, the sequence data files are transferred to a SPARCstation 2 using NFS Share.
Protocol
1. Prepare 8 M urea, 4.75% polyacrylamide gels, as described above, using a 36-well forming comb. Alternatively, the recipe can be scaled up to one liter.
2. Prior to loading, remove the tape from around the entire gel and carefully clean the outer surface of the gel plates with hot water. Rinse the glass with distilled water and then with ethanol, and allow the ethanol to evaporate.
3. Assemble the gel plates into an ABI 373A DNA Sequencer by placing the plates on the ledge in the bottom buffer well and clamping the gel into place with the black clamps attached to the laser stop.
4. Check the glass plates by closing the ABI lid and selecting "Start Pre-run" and then "Plate Check" from the ABI display. Adjust the PMT on the ABI display ("Calibration", "PMT") so that the lower scan (usually the blue) line corresponds to an intensity value of 800-1000 as displayed on the Macintosh computer data collection window. If the baseline of four-color scan lines is not flat, reclean the glass plates.
4. Attach the top buffer and the alignment brace, and fill both buffer wells with 1X MTBE electrophoresis buffer. Affix the aluminum heat distribution plate by setting it on the laser stop against the glass plates.
5. Pre-electrophorese the gel for 10-30 minutes by choosing "Start Pre-run" and "Pre-run Gel".
6. Use MakeSampleSheetOU to create a sample sheet or do this from within the ABI data collection software by entering the names and the fluorescent mobility file ("b920_21.mob" for fluorescent-labeled M13 -21 universal forward primer, "DyePrimer{M13RP1}" for fluorescent-labeled M13 universal reverse primer, "DyeTerm {any primer}" for AmpliTaq Terminators, and "DyeTerm{T7}-SetB" for Sequenase[TM] fluorescent-labeled dye terminators) to use for analysis. This Macintosh program and the related files are available from our ftp site at ftp://ftp.genome.ou.edu/ as a stuffit 1.5.1, binhexed file.
7. Prepare the samples for loading. Add 3 μl of FE to the bottom of each tube, vortex, heat at 90℃ for 3 minutes, and centrifuge to reclaim condensation.
8. Abort the pre-electrophoresis, and flush the sample wells with electrophoresis buffer with a syringe. Using flat-tipped gel loading pipette tips, load each odd-numbered sample. Pre-electrophorese the gel for at least 5 minutes, flush the wells again, and then load each even-numbered sample.
9. Begin the electrophoresis (30 W for 10 hours) run by selecting "Start Run" on the ABI display and by choosing "Begin Data Collection" from the controller box within the ABI data collection software on the Macintosh.
10. After data collection, the ABI software will automatically open the data analysis software, which will create the imaged gel file, extract the data for each sample lane, and process the data. Check the imaged gel file for sample tracking, and then transfer the results folder containing the sequence trace files to a SPARCstation 2 where the hard disk is mounted on the ether-netted Macintosh computer via NFS Share.
Sequencing double stranded DNA templates has become a common and efficient procedure (10) for rapidly obtaining sequence data while avoiding preparation of single stranded DNA. Double stranded templates of cDNAs containing long poly(A) tracts are difficult to sequence with vector primers which anneal downstream of the poly(A) tail. Sequencing with these primers results in a long poly(T) ladder followed by a sequence which is difficult to read. In an attempt to solve this problem we synthesized three primers which contain (dT)17 and either (dA) or (dC) or (dG) at the 3' end. We reasoned that the presence of these three bases at the 3' end would 'anchor' the primers at the upstream end of the poly(A) tail and allow sequencing of the region immediately upstream of the poly(A) region.
Using this protocol, over 300 bp of readable sequence could be obtained. We have applied this approach to several other poly(A)-containing cDNA clones with similar results. Sequencing of the opposite strand of these cDNAs using insert-specific primers occurred directly upstream of the poly(A) region. The ability to directly obtain sequence immediately upstream from the poly(A) tail of cDNAs should be of particular importance to large scale efforts to generate sequence-tagged sites (STSs) (11) from cDNAs (12,13).
Protocol 1. Synthesize anchored poly (dT)17 with anchors of (dA) or (dC) or (dG) at the 3' end on a DNA synthesizer and use after purification on Oligonucleotide Purification Cartridges.
2. For sequencing with anchored primers, denature 5-10 mg of plasmid DNA in a total volume of 50 ml containing 0.2 M sodium hydroxide and 0.16 mM EDTA by incubation at 65℃ for 10 minutes.
3. Add the three poly(dT) anchored primers (2 pmol of each) and immediately place the mixture on ice. Neutralize the solution by adding 5 ml of 5 M ammonium acetate pH 7.0.
4. Precipitate the DNA by adding 150 ml of cold 95% ethanol and wash the pellet twice with cold 70% ethanol.
5. Dry the pellet for 5 minutes and then resuspend in MOPS-Acid buffer.
6. Anneal the primers by heating the solution for 2 minutes at 65℃ followed by slow cooling to room temperature for 15-30 minutes.
7. Perform sequencing reactions, using modified T7 DNA polymerase and a-[32P]dATP (> 1000 Ci/mmole) using the protocol described earlier.
The following is a rapid and efficient method for sequencing cloned cDNAs based on PCR amplification (14), random shotgun cloning (1,3,15), and automated fluorescent sequencing (16). This method was developed in our laboratory because once the sequence of a genomic DNA containing cosmid is obtained and putative exons are predicted, the corresponding cDNAs should be sequenced in a timely manner. However, the presently implemented directed cDNA sequencing strategies, i.e. primer walking (17) and exonuclease III deletion (18), are both time consuming and labor intensive, while the alternative, i.e. randomly shearing the intact plasmid followed by shotgun sequencing (1,3,15), leads to a significant number of clones containing the original cDNA cloning vector rather than the desired cDNA insert.
This is a PCR-based approach where the "universal" forward and/or reverse priming sites were excluded from the resulting PCR product by choosing a primer pair that lay between the usual "universal" forward and reverse priming sites and the multiple cloning sites of the Stratagene Bluescript vector. These two PCR primers, with the sequence 5'-TCGAGGTCGACGGTATCG-3' for the forward or -16bs primer and 5'-GCCGCTCTAGAACTAG TG-3' for the reverse or +19bs primer, now have been used to amplify sufficient quantities of cDNA inserts in the 1.2 to 3.4 kb size range so that the random shotgun sequencing approach described below could be implemented.
Protocol
1. Incubate four 100 μl PCR reactions, each containing approximately 100 ng of plasmid DNA, 100 pmoles of each primer, 50 mM KCl (dilute from 1 M stock), 10 mM Tris-HCl pH 8.5 (dilute from 1 M stock), 1.5 mM MgCl2 (dilute from 1 M stock), 0.2 mM of each dNTP (dilute from 100 mM stock), and 5 units of PE-Cetus Amplitaq in 0.5 ml snap cap tubes for 25 cycles of 95℃ for 1 min., 55℃ for 1 min. and 72℃ for 2 min. in a PE-Cetus 48 tube DNA Thermal Cycler.
2. After pooling the four reactions to obtain sufficient quantities of PCR product for the subsequent steps the aqueous solution containing the PCR product is placed in an AeroMist nebulizer, brought to 2.0 ml by adding approximately 0.5 to 1.0 ml of glycerol, and equilibrated at -20℃ by placing it in either an isopropyl alcohol/dry ice or saturated aqueous NaCl/dry ice bath for 10 min.
3. The sample is nebulized at -20℃ by applying 25 - 30 psi nitrogen pressure for 2.5 min. Following ethanol precipitation to concentrate the sheared PCR product, the fragments were blunt ended and phosphorylated by incubation with the Klenow fragment of E. coli DNA polymerase and T4 polynucleotide kinase as described previously. Fragments in the 0.4 to 0.7 kb range were obtained by elution from a low melting agarose gel.
