Marcantonio Lab Protocol Manual
互联网
2146
Alkali Denaturation
Alkali Denaturation of Supercoiled Plasmid DNA
1) To 20 ul of purified, concentrated DNA solution add 2 ul of freshly prepared NaOH/EDTA solution and incubate at 37 degrees for 30 minutes.
2) Neutralize the reaction by adding : 3 ul 3M NaOAc 5.2 and 7 ul H2O
3) Vortex briefly, then add 75 ul ethanol and vortex to mix.
4) Let stand on dry ice for 10 minutes to precipitate the DNA.
5) Spin for 5-10 minutes in a microcentrifuge to pellet the DNA.
The pellet is usually very small and often difficult to detect at this step. Care must be taken not to pour it off or aspirate it away in the following steps. If unsure about the status of the pellet , you may wish to evaporate the ethanol off in a Speed-Vac concentrator rather than try to remove the ethanol mechanically.
6) Remove supernatant and add 200 ul of ice-cold 70% ethanol to wash pellet. Invert tube a few times, then spin in a microcentrifuge for 5-10 minutes.
7) Carefully remove the wash as discussed above and dry pellet in a Speed-Vac concentrator.
8) Suspend pellet in a volume of H2O appropriate for the number of sequencing reactions the DNA is to be used for, typically :
<center> 10 ul for one primer reaction<br /> 15 ul for two primer reactions</center>
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> 2M NaOH / 2mM EDTA <ul> <li> 0.5M EDTA 1 ul</li> <li> 10N NaOH 50 ul</li> <li> DD-H2O 199 ul</li> </ul> </li> </ul> </td> <td> <ul> <li> 3M NaOAc 5.2<br /> see "Molecular Biology Solutions"</li> </ul> </td> </tr> </tbody> </table> </center>
Competent HB-101s
Preparation of Competent Cells (HB-101)
1) Inoculate 50 ml of L2 media in a 250 ml flask with a single colony of cells or a frozen stock of cells. Grow overnight at 37 degrees with moderate shaking (200-250 rpm).
2) Inoculate 400 ml of L2 media in a sterile 2 liter flask with 4 ml of the overnight culture. Incubate at 37 degrees with shaking, until an OD590 of 0.375 is achieved. This procedure requires that the cells be actively growing. Overgrowth of the culture can result in drastically decreased efficiency of transformation.
During this incubation period, sterile polypropylene Eppendorf tubes should be placed in a freezer storage box and cooled to -20 degrees in a freezer. These tubes will be used to store the cells until use.
The cells must be kept chilled for all subsequent steps. This can be accomplished by performing all techniques on ice or, preferably, in a 4 degree cold room.
3) Aliquot culture into 8 pre-chilled sterile 50 ml polypropylene tubes. Let stand on ice for 10 minutes.
4) Centrifuge cells for 7 minutes at 3000 rpm at 4 degrees.
5) Pour off the supernatant and gently resuspend each pellet in 10 ml of ice-cold CaCl2 solution by pipetting up and down on ice.
6) Centrifuge cells for 5 minutes at 2500 rpm at 4 degrees.
7) Pour off the supernatant and gently resuspend each pellet in 10 ml of ice-cold CaCl2 solution by pipetting up and down on ice. Let stand on ice for 30 minutes.
8) Centrifuge cells for 5 minutes at 2500 rpm at 4 degrees.
9) Pour off the supernatant and gently resuspend each pellet in 2 ml of ice-cold CaCl2 solution by pipetting up and down on ice.
10) Aliquot cells into the prechilled Eppendorf tubes prepared earlier. Immediately freeze at -70 to -80 degrees.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> L2<br /> see "Molecular Biology Solutions"</li> </ul> <ul> </ul> </td> <td> <ul> <li> CaCl2 Soln<br /> see "Molecular Biology Solutions"</li> </ul> </td> </tr> </tbody> </table> </center>
CsCl Gradient
CsCl/ Ethidium Bromide Equilibrium Centrifugation
1) Resuspend the pellet obtained in the final step of the crude lysate preparation in 4 ml of TE buffer. Transfer to a 15 ml conical tube. Add 4.4 g of CsCl (Optical grade), dissolve by shaking, and add 0.4 ml of ethidium bromide (10 mg/ml). Caution: Ethidium bromide is a potent mutagen and an environmental hazard. Always wear gloves when handling and be sure to dispose of properly.
2) Transfer the solution to a Beckman Quickseal ultracentrifuge tube. This can be done using a syringe and needle or by using the Beckman funnel. Top off the tube, if necessary, using CsCl/TE solution (1 g/1 ml). Be sure that no bubbles remain in either the sample tube or balance tube. These could result in the collapse of the tubes during the spin, which could severely damage the rotor and centrifuge.
3) Band the plasmid by centrifugation in a VTi65 rotor at 58,000 rpm for>14 hours at 20 degrees.
The different capacities of covalently closed plasmid DNA and chromosomal DNA to bind the intercalating agent ethidium bromide permits separation in a CsCl gradient. Ethidium bromide lowers the density of DNA and plasmid DNA binds less than chromosomal DNA.
4) Insert an 18-G needle gently into the top of the tube. This acts as an air inlet to allow the plasmid DNA band to be drawn out below.
5) Recover plasmid DNA by suction with a syringe and 18-G needle. The collection needle should be gently inserted, beveled side down, approx. 1 cm below the plasmid band (lower of the two bands). As the needle approaches the band, the beveled edge should be gradually rotated until it is facing up, directly below the band. Avoid contamination with the protein/EtBr complexes which pellet on the outside edge of the tube.
Do not draw hard on the syringe if the needle becomes clogged, as this may create turbulence and mix the gradient. Leave the clogged needle in the tube and insert another to try again. Drawing the plasmid DNA through a clogged needle could also result in shearing of the DNA.
6) If higher purity plasmid DNA is desired (for transfection into eukaryotic cells), add plasmid DNA band to another ultracentrifuge tube, top off with CsCl/TE solution containing 0.2 mg/ml of EtBr, and repeat steps 3,4, and 5.
7) Remove the ethidium bromide by extraction using an equal volume of TE-saturated n-butanol. Vortex the tube vigorously and remove the upper organic phase. Repeat this procedure until no more red color remains in the lower aqueous phase. This procedure generates hazardous contaminated organic waste. Dispose of properly.
8) Dilute the lower phase with two volumes of TE buffer to dilute the CsCl.
9) Precipitate the plasmid DNA using 2 volumes of 100% ethanol at room temperature or at -20 degrees for>1 hour. Centrifuge at 10,000 rpm for 10 minutes at 4 degrees to pellet the DNA.
10) Wash the pellet with 1 ml of 70% ethanol and air dry. Resuspend the pellet in 0.5 ml of TE buffer, check its concentration by UV absorbance, and store at 4 degrees.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> TE<br /> see "Molecular Biology Solutions"</li> </ul> <ul> </ul> </td> <td> <ul> <li> </li> </ul> </td> </tr> </tbody> </table> </center>
Exonuclease III Deletion
Exonuclease III Deletion and Ligation Procedures
The following procedure is based on using doubly cut, Cesium-banded plasmid DNA, where 10 time points are to be taken. The amount of each solution to be used is determined by the desired number of time points. Reaction volumes can be scaled up or down in proportion to those described below.
1) Pre-warm a water bath to 30 degrees. Pre-heat a heating block to 70 degrees. Pour a 1% agarose minigel.
2) Remove the four enzyme buffers, as well as the DTT, dNTP mix, PEG, and S1 Stop buffer from the -20 freezer to allow them to thaw.
3) Label 10 Eppendorf tubes 1-10 for the ten time points. Place these tubes on ice.
4) Dissolve the pelleted DNA in 27 ul of DD-H2O. Add 3 ul of 10x Exo III Buffer. Warm this tube to 37 degrees in the water bath.
5) Prepare S1 Nuclease Mix as follows:
- DD-H2O 86.0 ul
- S1 7.4x Buffer 13.5 ul
- S1 Nuclease 0.5 ul
6) Add 7.5 ul of S1 Nuclease Mix to each of the ten labelled tubes on ice.
7) Set a timer at five minutes, add 1.5 ul of Exo III enzyme to the warm DNA tube, mix by tapping, place the tube back in the water bath, and start the timer.
8) Remove 3 ul samples every 30 seconds into the S1 tubes on ice, pipetting up and down to mix.
9) When all the samples have been taken, move the tubes to room temperature for 30 minutes.
10) While waiting, prepare the following mixtures and keep on ice:
<center> <table> <tbody> <tr> <td> <ul> <li> Klenow Mix <ul> <li> 1x Klenow Buffer 15.0 ul</li> <li> Klenow Enzyme 0.5 ul</li> <li> 50% PEG 25.0 ul</li> <li> 100 mM DTT 2.5 ul</li> <li> T4 DNA Ligase 0.5 ul</li> </ul> </li> </ul> </td> <td> <ul> <li> Ligase Mix <ul> <li> DD-H2O 197.5 ul</li> <li> Ligase 10x Buffer 25.0 ul</li> </ul> </li> </ul> </td> </tr> </tbody> </table> </center>
11) After 30 minutes at room temperature, add 1.0 ul of S1 Stop Buffer to each tube and heat at 70 degrees for 10 minutes to inactivate the S1 nuclease.
12) Remove 5.0 ul from each tube to run on the minigel to determine the extent of digestion. While loading the gel, let the 10 tubes cool to 37 degrees in the water bath.
13) Add 1.0 ul of Klenow Mix to each tube, mix, and incubate at 37 degrees for 3 minutes.
14) Add 1.0 ul of the dNTP mix to each tube, mix, and incubate at 37 degrees for another 5 minutes.
If you must interrupt the procedure, then do it here by placing the tubes on ice or at 4 degrees.
15) Add 20.0 ul of Ligase Mix to each tube, mix well, and incubate at room temperature for at least one hour.
16) Transform HB101 cells using 10 ul from each tube according to the protocol, "Transformation of HB101 Competent Cells".
Glassing Out
"Glassing Out" DNA Using the Geneclean Kit
If you are purifying DNA that is already in aqueous solution, then proceed directly to step 3. If the DNA is being gel purified, then begin with step 1.
1) Excise DNA band from ethidium bromide-stained gel using a fresh razor blade and place into a clean Eppendorf tube. Use long-wave UV light for as short a time as possible to visualize the DNA.
2) Determine the approximate volume of the gel slice by weight. If the slice weighs more than 0.4 g, it may be necessary to divide it into two fragments and continue the procedure in two Eppendorf tubes.
3) Add 3 volumes of NaI solution (BIO 101).
If DNA is not contained in agarose, go directly to step 5.
4) If agarose is present, place the tube in a 50 degree water bath. After a minute, mix the contents of the tube and return it to the water bath. After approximately 5 minutes, the agarose should be completely dissolved. Do not proceed until the agarose is completely dissolved. Do not leave the tube in the water bath for any longer than is necessary to dissolve the agarose.
5) Vortex the Glassmilk tube for 1 full minute, holding the tube horizontally.
6) Add 5 ul of Glassmilk suspension to the solution and mix by inverting rapidly. If there is more than 5 ug of DNA present, then 10 ul of Glassmilk suspension should be used.
7) Place tube in the Labquake rotator and let rotate for 5 minutes at room temperature. The DNA binds to the silica matrix during this step.
8) Pellet the silica matrix with bound DNA by spinning in a microcentrifuge for 5 seconds at full speed (Wait for the centrifuge to come to full speed, count to five, and let it come to a halt).
9) Carefully aspirate off the supernatant and resuspend the pellet in 500 ul of NEW WASH (BIO 101) by pipetting up and down.
10) Spin for 5 seconds in a microcentrifuge and remove the supernatant. Repeat this wash procedure two more times. These washes remove the proteins and most of the RNA because they do not bind to the silica matrix, as well as removing any salts or ethidium bromide.
11) Spin the tube in a Speed-Vac concentrator to remove the last traces of NEW WASH.
12) Resuspend pellet in 25 ul of DD-H2O (or any other low salt buffer such as TE) by pipetting up and down. Incubate the tube in a 50 degree water bath for 3 minutes.\ The DNA does not bind to the silica matrix in the absence of salts and is eluted during this step.
13) Spin for 30 seconds in a microcentrifuge to make a solid pellet. Carefully remove 20 ul of the supernatant containing the cleaned DNA and place in a fresh Eppendorf tube.
If a small amount of Glassmilk is carried over, it does not interfere with subsequent use of the DNA but can be removed by briefly spinning the tube before withdrawing any of the solution.
14) Resuspend the pellet in an additional 20 ul of your low-salt buffer, incubate, spin, and transfer 20 ul of the supernatant into the tube containing the clean DNA.
Large-scale Plasmid Prep
Large-Scale Isolation of Plasmid DNA
A. Preparation of Crude Lysate
1) Inoculate 200 ml of L2 containing appropriate selective agent with a single colony or culture containing the desired plasmid. Grow at 37 degrees overnight.
2) Spin down cells at 5000 rpm for 15 minutes in Sepcor bottles.
3) Carefully pour off supernatant and aspirate off any media remaining.
4) Resuspend pellet in 4 ml of ice-cold GTE by pipetting up and down. Let stand 5 minutes at room temperature.
Be sure to completely resuspend cells. Avoid creating bubbles.
5) Transfer to 50 ml conical tube (Corning). Add 8 ml of freshly prepared NaOH/SDS solution and mix by inverting. Do not vortex. Let stand on ice for 5 minutes.
6) Add 6 ml of ice-cold KOAc solution and mix by inverting rapidly. Let stand on ice for 5 minutes.
7) Transfer to 2 Oak Ridge tubes and centrifuge at 11,000 rpm for 20 minutes at 4 degrees. The pellet formed in this step will contain most of the chromosomal DNA, SDS-protein complexes, and other cellular debris.
8) Transfer the supernatant in each tube into 2 Oak Ridge tubes, for a total of 4 tubes. Add an equal volume of phenol/chloroform to each tube and mix by vortexing. Centrifuge for 5 minutes at 10,000 rpm and transfer the upper, aqueous layer to a fresh Oak Ridge tube.
9) Add 2 volumes of ethanol, mix by vortexing, and let stand at room temperature for 5 minutes.
10) Centrifuge for 10 minutes at 10,000 rpm.
11) Pour off supernatant and wash with 1 ml of ice-cold 70% ethanol. Be careful, as the pellet often loosens from the tube upon washing.
12) Dry pellet by air and resuspend in TE w/RNase.
Note: The volume in which this pellet is resuspended in is dependent upon the intended use of the plasmid DNA. If the DNA is to be further purified by CsCl gradient centrifugation, then the DNA must be suspended in a final volume of 4 ml TE, no RNase being necessary.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> NaOH / SDS Solution <ul> <li> DD-H2O 9.3 ml</li> <li> 10N NaOH 200 ul</li> <li> 20% SDS 500 ul</li> </ul> </li> </ul> </td> <td> <ul> <li> 3M GTE<br /> see "Molecular Biology Solutions"</li> <li> KOAc Solution<br /> see "Molecular Biology Solutions"</li> <li> L2<br /> see "Molecular Biology Solutions"</li> </ul> </td> </tr> </tbody> </table> </center>
Minigel
Pouring a Minigel
1) Rinse off the minigel apparatus, the gel casting tray and the desired combs with water.
2) Weigh out 0.8 g of agarose for every 1% concentration of gel you desire. Place this in a 250 ml Ehrlenmeyer flask.
A typical minigel is 1%.
3) To <700 ml of DD-H2O, add 14 ml of 50x TAE buffer and mix.
4) Add 80 ml of your 1x TAE to the agarose.
5) Melt the agarose in a microwave oven as follows:
45 sec @ HIGH/Mix/45 sec @ HIGH/Mix/1 min @ DEFROST/Mix
If the agarose is properly dissolved you should see no globules floating in the solution when held up to the light.
6) Add 4 ul of Ethidium bromide to the solution and slowly pour the gel into the apparatus. Remove any bubbles with a pipet tip if necessary.
7) Let the gel harden, turn the casting tray 90 degrees around, and pour in the remaining buffer solution.
8) Add the proper amount of 6x loading dye to each sample, mix by pipetting up and down, and carefully load each sample into a well using a Pipetman. Also load any molecular weight markers that are needed.
9) Attatch the minigel apparatus cover and the power supply leads, turn on the power supply and adjust the current to the desired level. 100 mAmps is a typical amount of current applied to a minigel, although you may adjust this to speed up the gel or slow it down.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> Ethidium Bromide Solution<br /> see "Molecular Biology Solutions"</li> </ul> </td> <td> <ul> <li> 6x Loading Dye<br /> see "Molecular Biology Solutions"</li> </ul> </td> </tr> </tbody> </table> </center>
Phenol Equilibration
Phenol Equilibration
1) Warm the solid redistilled phenol in a 65 degree water bath until the entire contents of the container is liquified.
2) Add an equal volume of 1M Tris 8.0 to the melted phenol. Mix thoroughly by carefully shaking. Let the phases separate overnight at 4 degrees.
3) Remove the upper, aqueous layer by aspiration and replace with an equal volume of 0.1M Tris 7.5.
4) Mix carefully and allow the phases to separate as before.
5) Store at 4 degrees.
Plasmid Miniprep
Miniprep of Plasmid DNA (Alkaline Lysis Method)
1) Inoculate 2 ml of L2 media containing the appropriate antibiotic with a single bacterial colony picked using a sterile toothpick. Incubate at 37 degrees overnight with vigorous shaking.
2) Transfer 1.5 ml of culture into an Eppendorf tube. Spin for 30 seconds to pellet cells.
3) Remove the medium by pipet or aspiration, leaving the pellet as dry as possible.
4) Resuspend pellet in 100 ul of ice-cold GTE solution by gently pipetting up and down. Let stand 5 minutes at room temperature. Be sure to completely resuspend cells. Avoid creating bubbles.
5) Add 200 ul of freshly prepared NaOH/SDS solution and mix by inverting tube rapidly or by tapping. Do not vortex. Let stand on ice 5 minutes. This treatment lyses the bacteria. SDS denatures bacterial proteins and NaOH denatures chromosomal and plasmid DNA.
6) Add 150 ul of KOAc solution and vortex to mix. Let stand on ice for 5 minutes. This neutralizes the mixture, causing the plasmid DNA to reanneal rapidly. Most of the chromosomal DNA and bacterial proteins precipitate along with the SDS-potassium complex.
7) Centrifuge for 5 minutes at 4 degrees.
8) Transfer the supernatant to a fresh Eppendorf tube.
9) Add an equal volume of Phenol/chloroform and mix by vortexing. Centrifuge for 2 minutes at room temperature and transfer the upper, aqueous layer to a fresh Eppendorf tube.
10) Add 2 volumes of ethanol and let stand at room temperature for 2 minutes.
11) Centrifuge for 5 minutes at room temperature. The prior three steps further purify and concentrate the plasmid DNA.
12) Aspirate off the supernatant and dry completely in a Speed-Vac concentrator. Resuspend pellet in 30 ul of TE w/RNase.
Note: 3 ul of DNA is an appropriate volume to use in a typical analytical restriction digest reaction.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> NaOH / SDS Solution per 1 ml <ul> <li> DD-H2O 930 ul</li> <li> 10N NaOH 20 ul</li> <li> 20% SDS 50 ul</li> </ul> </li> </ul> </td> <td> <ul> <li> TE<br /> see "Molecular Biology Solutions"</li> <li> KOAc Solution<br /> see "Molecular Biology Solutions"</li> <li> L2<br /> see "Molecular Biology Solutions"</li> </ul> </td> </tr> </tbody> </table> </center>
Powdered Media
Preparation of Powdered Tissue Culture Media
1) Remove Penicillin-Streptomycin solution and histidine solution (1 tube/ 500 ml) from -20 freezer to thaw.
2) To a flask as close to the final volume of the media you wish to prepare, add approximately 850 ml of deionized, distilled H2O per liter of media with gentle stirring.
3) Add one envelope of powdered media for every liter of media. Rinse inside of envelope with DD-H2O to remove all traces of powder and add to flask.
4) Add the following reagents to the flask for each liter of media:
- 3.7 g sodium bicarbonate:
- 2 tubes of histidine solution (21 mg/ ml)
- 10 ml sodium pyruvate solution (100x) (GIBCO)
- 10 ml Penicillin/Streptomycin Solution (GIBCO)
6) Dilute the medium to the appropriate volume using a graduated cylinder and check the pH, adjusting if necessary.
7) Process the medium into sterile containers by passing through a bottle-top 0.2 um membrane filter device ( Falcon 7105 or Corning 25970). The volume in each bottle should be as near to 450 ml as possible to account for the addition of media later.
8) Seal bottle-tops using parafilm and store at 4 degrees.
Note: This media should be labelled so that any users know what has and what has not been added. Often serum or selective agents may be added later.
Protein Gel
Preparing and Running a Protein Gel (7% Polyacrylamide)
A. Preparation of Running Gel Solution
1) Add to a 50 ml cylinder:
- DD-H2O 26 ml
- 30% acrylamide stock (37.5:1 Acryl/Bis) 10.5 ml
- 2M Tris 8.8 8.4 ml
- 20% SDS 0.23 ml
B. Assembling the Gel Sandwich
1) Clean two glass plates, one small and one large, by THOROUGHLY scrubbing with a non-abrasive detergent and brush. Rinse THOROUGHLY with water and allow to air dry. Also clean and dry two side spacers and a bottom spacer. Be sure to remove all traces of ink, soap, tape, dirt, dust, sweat, lunch, and any other debris that may settle on the plates.
2) Immediately before assembling plates, rinse them with ethanol and dry with a Kimwipe.
3) Place the larger plate on lab paper. Place the side spacers along the short edges of the plate with the foam block face up. Carefully place the smaller plate over this so that the lower edges of the two plates meet and the foam spacer block is snug against the top of the smaller plate..
4) Clamp the gel sandwich together with six clamps, two on each side and two on the bottom.
C. Pouring the Running Gel
1) Before pouring the gel, pipet a small amount of molten 1.2% agarose into each lower corner of the gel sandwich to seal the corners in order to prevent leakage of the gel solution.
2) Add the following reagents to the gel solution:
- TEMED 30 ul
- 10% Ammonium Persulfate 200 ul
These reagents catalyze the polymerization of the gel solution. Depending on the precise environmental conditions, you have approximately 5 minutes to pour the gel after the addition of the catalysts.
3) Holding the plates at a slight angle so that the gel will flow down the large plate, carefully pour the gel between the glass plates
4) Pour gel solution until there is about 1.5 to 2 inches of room left at the top of the sandwich.
This is to allow room for pouring the stacking gel.
5) Add 1-2 ml of Isobutanol to the sandwich. This will smooth out the upper surface of the gel and remove any bubbles that may have formed during the mixing and pouring process.
6) Leave the gel sandwich upright to polymerize.
D. Preparation of Stacking Gel Solution
1) Add to a 50 ml beaker:
- DD-H2O 14.7 ml
- 30% acrylamide stock (37.5:1 Acryl/Bis) 2.5 ml
- 1M Tris 6.8 2.5 ml
- 20% SDS 0.1 ml
E. Pouring the Stacking Gel
1) Pour the isobutanol off the running gel.
2) Rinse the gel by pouring approximately 1/2 inch of stacking gel solution onto the gel and gently rocking the gel sandwich back and forth. Pour the rinse off as waste.
3) Repeat the rinse as above.
4) Add the following reagents to the stacking gel solution:
- TEMED 40 ul
- 10% Ammonium Persulfate 200 ul
5) Pour the stacking gel solution into the gel sandwich directly from the beaker. Pour gel solution until it reaches the top of the smaller plate.
6) Gently insert the comb into the gel sandwich, being careful not to form bubbles in the well-spaces.
As the gel polymerizes, it may contract. If this starts to happen, you should add some more stacking gel solution to the corners. Failure to do so may result in the loss of usable wells.
7) Leave the gel sandwich upright to polymerize.
F. Setting Up the Gel
1) Gently remove the comb from the gel sandwich.
2) Remove the spring clips from the bottom and sides of the gel, as well as the bottom spacer.
3) Position the gel sandwich so that the shorter plate faces the apparatus and the foam blocks form a seal with the gasket, and clamp it onto the gel apparatus using spring clips on the sides.
4) Pour 1x Running Buffer into the upper buffer chamber of the gel apparatus and check for leaks. Readjust the gel sandwich and/or spring clips if necessary. 1 liter of 1x Running Buffer will be more than enough for the two chambers of the V16 Apparatus.
5) Use a syringe with a bent needle to blow away any bubbles that formed just beneath the surface of the gel sandwich. Failure to remove these bottom bubbles will result in a real mess.
G. Sample Application and Running the Gel
1) Suspend samples in a suitable volume of gel loading buffer (<80 ul).
2) Heat samples to 95 degrees for 2 minutes just prior to loading gel.
3) Load samples into wells of gel using a pipettor.
4) Connect the leads to the gel apparatus and the power supply and adjust current to 40-50 mAmps for 4 hours OR 10 mAmps to run overnight.
H. Disassembling the Gel
1) Turn off the power supply and disconnect the leads from the apparatus.
2) Pour off the buffer from the upper and lower buffer chambers as waste.
3) Remove the spring clamps from the apparatus and remove the gel sandwich.
4) Remove the side spacers and gently pry the plates apart , being careful not to tear the gel.
I. Fixing the Gel
1) Pour 200 ml of 30% Methanol/10% Acetic Acid into a 500 ml Nalgene storage box. Place gel in this solution and shake gently on a rotator for 1 hour.
At this point, treatment of the gel will depend upon the exact application you are involved in. You may need to soak the gel in ENHANCE, Coomassie Blue, or nothing at all.
See the appropriate protocol for your application.
Quantitation of DNA
Detection of Nucleic Acids Using Absorption Spectroscopy
The absorption of the sample can be measured at several different wavelengths to assess the purity and concentration of nucleic acids, as follows:
<center> A260---Quantitative for pure nucleic acids in microgram quantities<br /> A260/A280---Indicator of nucleic acid purity<br /> A325---Indicates particulates in solution or dirty cuvettes<br /> </center> 1) Turn on the UV source on the spectrophotometer to allow it a few minutes to warm up before measuring absorbances to prevent drift.
2) Pipet 2 ul of the nucleic acid sample into an Eppendorf tube containing 498 ul of DD-H2O. Mix this and transfer it to a clean cuvette. Pipet 500 ul of DD-H2O into another cuvette that will be used as a reference sample.
3) Place the reference cuvette into the instrument, close the lid, and set the wavelength to 260 by pressing the numbered keys 2,6, and 0 followed by the l key.
4) Calibrate the instrument by pressing the Cal key. Wait until the readout indicates 0 absorbance and replace the reference cuvette with the sample cuvette.
5) Close the lid and read the absorbance.
6) Repeat steps 3, 4, and 5 for the wavelengths 280 and 325.
7) When you are finished taking measurements, turn the UV source off and clean the cuvettes using the cuvette washer.
Use the A260 to determine the amount of DNA present by the following formula:
<center> A260 x 12.5 = concentration of DNA (ug/ul)</center>
Use the A260/A280 ratio and the A325 to estimate the purity of the nucleic acid sample as follows:
<center> Ratios of 1.8 to 1.9 indicate highly purified preparations of DNA. Ratios of 1.9 to 2.0 indicate highly purified preparations of RNA.</center>
Absorbance at 325 suggests contamination by particulates/dirty cuvettes.
Purify/Concentrate-SQ
Purification and Concentration of Aqueous DNA Solutions for Plasmid DNA to be sequenced
1) Add an equal volume of Phenol/chloroform to the sample to be purified and vortex vigorously for 10 seconds.
If the volume of the DNA solution to be purified is below 100 ul, it is easier to perform the extraction step if the volume is first brought to 100 ul by adding TE buffer or H2O.
2) Spin for 15 seconds at room temperature in a microcentrifuge. If the phases are not well separated, spin for 1 minute longer.
3) Carefully remove the upper aqueous layer containing the DNA and transfer to a fresh microcentrifuge tube.
4) Add 2/3 rd volume of 5M NH4OAc to the DNA solution, mix by vortexing briefly, and add 2 volumes of ice-cold ethanol. Mix by vortexing and place in crushed dry ice for at least 5 minutes to allow the DNA to precipitate.
5) Spin for 5-10 minutes in a microcentrifuge. If the concentration of the DNA is believed to be low, or the size of the fragments is very small, more extensive centrifugation may be required.
6) Carefully pour off the supernatant or aspirate it off, being very careful not to remove pellet.
7) Add 1 ml 70% ethanol and invert several times to wash pellet. Spin as above.
8) Dry the pellet in a Speed-Vac concentrator.
9) Resuspend the pellet in an appropriate volume of H2O or TE buffer for further manipulations or storage. If this DNA is to be sequenced using the Sequenase Version 2.0 Kit, then the pellet should be resuspended in 20 ul H2O.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> TE<br /> see "Molecular Biology Solutions"</li> </ul> <ul> </ul> </td> <td> </td> </tr> </tbody> </table> </center>
Sequencing DS-DNA
Sequencing Double-Stranded DNA Using the Sequenase Version 2.0 Kit
1) Prepare the DNA for sequencing as per the protocols, "Purification and Concentration of Aqueous DNA Solutions" and "Alkali Denaturation of Supercoiled Plasmid DNA".
2) Warm a heating block up to 65 degrees.
3) For each sample to be sequenced, put the following in a 1.5 ml Eppendorf tube:
- DNA (prepared as discussed) 7 ul
- Sequencing primer 1 ul
- Reaction Buffer 2 ul
5) While cooling, prepare the following mixtures. Prepare slightly more than you actually need, in order to avoid problems during the actual procedure.
Dilute Labelling Mix: need 2 ul per sample
- 2 ul Labelling Mix (USB)
- 8 ul DD-H20
- 7 ul Enzyme Dilution Buffer (USB)
- 1 ul Sequenase 2.0 (USB)
- 0.1M DTT (USB) 1.0 ul
- (35S) dATP 0.5 ul
- Dilute Labelling Mix 2.0 ul
- Dilute Sequenase 2.0 ul
6) Also at this time, prepare the microtiter dish that will be used during the termination reactions. Transparent tape should be applied over any wells that will not be needed, in order to provide a better seal against water leaking into the sample wells. The horizontal rows should be labelled A-T-G and C, while the vertical columns should be labelled with a code for your samples. When this is complete, place 2.5 ul of the appropriate ddNTP Termination Mix into each respective well. Use the appropriate set of Termination Mix tubes for either dGTP reactions (red-capped tubes) or for dITP reactions (green-capped tubes).
Be sure the drop of Termination Mix makes it to the bottom of the well by tapping the plate upon the bench. Keep the plate loosely covered with a plate sealer until needed.
7) Once the annealed template-primer has reached 35 degrees or lower, add 5.5 ul of the Master Mix by pipetting it onto the side of the sample tube and tapping it down to mix thoroughly. Before adding the Master Mix, it is a good idea to briefly spin down the samples to recover any solution clinging to the top or sides of the tube.
Try not to contaminate the pipet tip with any sample, as it can be used to add Master Mix to all sample tubes, minimizing the amount of solid radioactive waste generated.
8) Incubate at room temperature for 1 minute to allow for primer extension and labelling.
9) Pipet the labelling reaction up and down to mix, and add 3.5 ul to each of the four wells on the microtiter plate containing the four termination mixes. One pipet tip can be used to add the reaction mix to the four wells by pipeting the sample onto the side of the well. The plate is then tapped on the bench causing the labelling reaction to mix with the termination mix. This procedure, once again, minimizes the amount of solid radioactive waste generated.
10) Once all the reactions have been added to the termination mixes in the plate, cover the plate tightly with a plate sealer and place in a water bath at 37 degrees for 5 minutes.
The water in the bath should be high enough to bubble around the wells when you look under the plate. During this step the chains are extended a little more and then terminated by the addition of a dideoxynucleotide.
11) Wipe off the water from the outside of the plate with a Kimwipe and carefully remove the plate sealer, making sure that no drops of water fall into the wells.
12) Add 4 ul of Stop Solution to each well by depositing the drop on the side of the well and tapping to mix.
13) Samples may be stored, covered, at -20 degrees for up to 1 week before running on a gel.
Sequencing Gel
Preparing and Running a Sequencing Gel (6% Polyacrylamide/Urea)
A. Preparation of Gel Solution
1) Weigh out 50 g of Urea into a clean 250 ml beaker containing a stir bar.
2) Add to the beaker:
- DD-H2O 40 ml
- 40% acrylamide stock (19:1 Acryl/Bis) 15 ml
- 20x TBE 5 ml
B. Assembling the Gel Sandwich
1) Clean two glass plates, one small and one large, by THOROUGHLY scrubbing with a non-abrasive detergent and brush. Rinse THOROUGHLY with water and allow to air dry. Be sure to remove all traces of ink, soap, tape, dirt, dust, sweat, lunch, and any other debris that may settle on the plates. Any such contamination may result in extreme difficulty when pouring the gel.
2) When plates are nearly dry, wipe with DD-H2O and dry, then wipe with ethanol and let dry.
3) Siliconize one surface of the larger plate by wiping its surface with Sigmacote applied to a Kimwipe. This procedure should be performed in a fume hood to avoid contact with the Sigmacote vapors.
Siliconizing one plate helps the completed sequencing gel adhere preferentially to the other plate when disassembling the sandwich.
4) Immediately before assembling plates, rinse them with ethanol and dry with a Kimwipe.
5) Place the larger plate, siliconized side up, on lab paper. Place the vinyl side spacers along the long edge of the plate with the foam block face up. Carefully place the smaller plate over this so that the lower edges of the two plates meet and the foam spacer block is snug against the top of the smaller plate..
6) Tape the edges of the plates to seal the sandwich. First tape the bottom edges, then the sides.
The tape should be firmly pulled and pressed against each surface of the glass plates in order to accomplish a leak-free seal. Each edge should be sealed thoroughly with two strips of tape. Special attention should be paid to sealing the lower corners of the sandwich.
C. Pouring the Gel
1) Immediately before pouring the gel, filter the gel solution through a disposable Nalgene unit and pour into a fresh, clean 250 ml beaker.
2) Add the following reagents to the filtered gel solution:
- TEMED 15 ul
- 10% Ammonium Persulfate 1 ml
Following addition of the APS use the pipet tip to stir the solution. These reagents catalyze the polymerization of the gel solution. Depending on the precise environmental conditions, you have approximately 10 minutes to pour the gel after the addition of the catalysts.
3) Holding the plates at a 45 degree angle so that the gel will flow down one of the side spacers, carefully inject the gel between the glass plates using a 50 ml syringe, maintaining an even flow to avoid the formation of air bubbles. If air bubbles form, move the plates to a more vertical position and tap the glass plates to work the bubbles out.
4) Pour gel solution until there is a reservoir of liquid extruding from between the plates when they are resting at a 5 degree angle.
5) Insert the flat edge of a sharkstooth comb between the plates to an approximate depth of 2-3 mm beneath the edge of the short plate. Insert the other comb likewise, making the edges of the two combs meet evenly.
6) Place a spring clip on both sides of the sandwich over the combs to keep them in place.
7) Leave the plates in this position to polymerize.
8) If the gel is going to be used the same day then skip to the next section of this protocol, D. Setting Up the Gel. If the gel is to used at a later time, then remove the spring clips, soak several paper towels in water and wrap them around the top part of the gel to prevent it from drying out. Then, wrap the entire top half of the sandwich in plastic wrap and reapply the spring clips.
D. Setting Up the Gel
1) Remove the spring clips from the top of the gel, as well as any paper towels and plastic wrap, if used. Remove the tape from the lower edge of the glass plates by cutting with a razor blade.
2) Position the gel sandwich so that the shorter plate faces the apparatus and the foam blocks form a seal with the gasket. Do not yet tighten the screw knobs.
3) Carefully remove the sharkstooth combs from the gel, keeping track of which comb went in which side of the gel. Rinse the combs with water and place them between the plates so that the teeth just make contact with the gel surface. Clamp the combs in place using thumb screws.
If done properly, a slight indentation will be evident where the teeth touch the gel surface. Do not puncture the gel surface or tear it by moving the inserted combs laterally.
4) Gradually tighten the screw knobs, in an alternating manner, to complete the seal. Be careful not to overtighten the screws as this may damage the plates.
5) Close the upper buffer drain valve and fill the upper and lower reservoirs with 500 ml of TBE electrophoresis buffer in each.
6) Close the upper and lower safety lids and attatch the apparatus to the power supply. (Red lead on bottom, black lead on top) Gel should be pre-electrophoresed at 30 watts, constant power, for 15-30 minutes before loading samples, although this is not necessary.
E. Sample Application
1) If the sequencing reactions were just completed, then the samples must be heated to 75 degrees for 2 minutes prior to loading. If they were stored at -20 before running the gel, then they should be allowed to warm up for a few minutes prior to loading.
2) Immediately prior to loading the samples, the wells must be washed out to remove the urea which leeches into them from the gel. Use a syringe with a bent needle to rinse the wells with buffer. If the wells are not rinsed properly, then the sample will not form a tight band as it migrates through the gel and band resolution on the autoradiograph will suffer.
3) Using the Eppendorf Ultra-Micro pipettor, load 3 ul of sample into each well, loading in the order G-A-T-C.
4) Close the upper safety lid, reattatch the black lead and switch on the power supply to provide 60 watts of constant power.
F. Disassembling the Gel
1) Turn off the power supply and disconnect the leads from the apparatus.
2) Open the upper chamber drain valve to release the contents of the upper buffer chamber into the rear portion of the lower buffer chamber.
3) Loosen the screw knobs and remove the gel sandwich from the apparatus. The buffer in the front of the lower buffer chamber contains radioactive nucleotides and care should be taken not to drip this on unprotected surfaces when moving the gel sandwich.
4) The buffer in the front of the lower buffer chamber should be carefully poured into a large beaker to be stored in the radiation hood for future disposal.
5) Remove the tape from the gel sandwich and slide out the side spacers and the combs.
6) Place the sandwich on a protected lab bench with the small plate on top. Being extremely careful not to tear the gel, carefully pry the plates apart using a spatula. The gel should stick preferentially to the large plate, which has not been siliconized. However, if it decides to adhere to the small plate instead, then go with the flow. It doesn't matter which plate it sticks to as long as it is all on one plate!
7) Cut a piece of Whatmans #3 blotting paper to approximately the size of the gel and press it firmly and evenly onto the surface of the gel, especially at the corners and edges.
8) Carefully lift a corner of the paper to see if the gel is adhering to it. If so , then continue lifting the paper away from that corner to remove the gel from the glass plate.
If the gel is not sticking to that corner, then try lifting at another corner or edge. If there is still difficulty, then wet the paper a bit using a squeeze bottle and try again.
G. Exposing the Gel
1) Place the gel on the gel dryer, backing paper down. Cover the surface of the gel with a layer of plastic wrap and cover with the rubber seal.
2) After making sure that the vacuum traps are empty and the second flask is immersed in a slurry of dry ice and ethanol, apply a vacuum and turn on the dryer. It should be set at 80 degrees for 40 minutes.
3) When the gel is finished drying, first break the vacuum by lifting the rubber seal and turning the pump off.
4) Remove the plastic wrap from the gel and expose it directly to XAR film in an autoradiograph cassette overnight at -80 degrees. No intensifying screen is needed. In fact, it can decrease the resolution of the fragments on the gel.
5) Develop the film in the X-OMAT Film Processor. If necessary, re-expose for a longer period to enhance the readability.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> 20x TBE<br /> see "Molecular Biology Solutions"</li> </ul> </td> <td> <ul> </ul> </td> </tr> </tbody> </table> </center>
Transformation
Transformation of HB-101 Competent Cells
1) Thaw tubes of competent cells on ice, occasionally inverting gently to mix.
2) Pre-cool Falcon 2059 tubes on ice.
3) Aliquot 30 ul cells into each tube.
4) Add up to 10 ul DNA in TE or ligation buffer and place on ice for 10 minutes. This allows DNA to attatch to cells.
5) Place in 42 degree water bath for 1 minute. This treatment shocks the cell membranes to allow take-up of the DNA.
6) Add 0.5 ml of L2 media and shake at 37 degrees for up to 1-2 hours. Remember: Do not use media containing selection agent, usually ampicillin, as this is the period in which antibiotic resistance develops.
7) Pipet cells onto L2 plates containing selective agent and spread using a sterile glass spreader (a bent Pasteur pipet).
8) Wait until all liquid has been absorbed into plate, then incubate overnight at 37 degrees.
<center> <h3> Solutions</h3> <table> <caption> </caption> <tbody> <tr> <td> <ul> <li> TE<br /> see "Molecular Biology Solutions"</li> </ul> </td> <td> <ul> <li> L2<br /> see "Preparation of Bacterial Media..."</li> </ul> </td> </tr> </tbody> </table> </center>
