PCR Cloning Protocol. Day 1: Isolation of MG1655 genomic DNA (adapted from Current Protocols in Mol. Biol. (1997), Supplement 27.

Transcription

1 Modifications to EXPERIMENTS 21 and 24: PCR and Molecular Cloning STUDENTS SHOULD USE A P-2 MICROPIPETTOR FOR ALL VOLUMES 2 µl. Day 1: Isolation of MG1655 genomic DNA (adapted from Current Protocols in Mol. Biol. (1997), Supplement 27.) 1. Obtain a MG1655 cell pellet originating from 1.5 ml of an overnight culture. 2. Gently resuspend the pellet in 567 µl TE buffer. Add 30 µl of 10% SDS and 3 µl of 20 mg/ml proteinase K. What are the final concentrations of proteinase K and SDS? Mix thoroughly and incubate for 1 hr. at 37 C. 3. Add 100 µl of 5 M NaCl and mix thoroughly. Why is this important? 4. Add 80 µl of CTAB/NaCl solution. Mix thoroughly and incubate for 10 min at 65 C. 5. In a hood, add an approximately equal volume of chloroform, mix thoroughly (but gently) by inverting the tube several times, and spin 4 min. at max speed in a microcentrifuge. You should see a white interface between the chloroform and aqueous layers. What is this precipitate? 6. Remove the aqueous supernatant to a fresh 1.5 ml microfuge tube, being careful to leave the white interface behind. In a hood, add an equal volume of 25:24:1 phenol:chloroform:isoamyl alcohol to the aqueous solution, mix thoroughly (but gently) by inverting the tube several times, and spin 4 min. at max speed in an microcentrifuge. 7. In a hood, carefully transfer the aqueous phase to a fresh 1.5 ml microfuge tube. Extract the aqueous phase with an equal volume of chloroform, mix thoroughly (but gently) by inverting the tube several times, and spin 4 min. at max speed in a microcentrifuge. Transfer the top aqueous to a fresh 1.5 ml microfuge tube. 8. Add 1.5 volumes of ice cold 100% ethanol. Gently invert the tube several times. A stringy white nucleic acid precipitate should be clearly visible. Place this tube on ice for 10 min., centrifuge 15 min. at 4 C, and then carefully remove the solution being sure not to disturb the nucleic acid pellet. When centrifuging, it is important to be consistent on how you place the microfuge tube into the rotor. In this way, you will always know where the nucleic acid pellet should be (in the event that it is not visible to you). 9. Rinse the nucleic acid with 1 ml of ice cold 70% ethanol by gently inverting the tube several times. Place this tube on ice for 10 min., centrifuge 15 min. at 4 C (in the same orientation as in step 8), and then carefully remove the solution being sure not to disturb the nucleic acid pellet. 10. Dry the pellet in a speedvac for 10 min. on medium heat. If the pellet is not completely dry, speedvac the sample for additional time. Resuspend the pellet in 100 µl TE buffer with very slow/gentle pipetting. Place the tube in a 42 C water bath for 30 min. Obtain the concentration of a 1

2 1:10 dilution of the nucleic acid using the NanoDrop. What solution will you use to make your dilution? What will you blank the NanoDrop with? Be sure to save the OD readings and spectra. Day 2: PCR amplification of a β-galactosidase gene. 1. Set up the following reaction twice in the specified order in labeled 0.5-ml sterile PCR tubes. Each reaction gets a different primer mix. One PCR product is destined for pet15b and the other for pet28a(+). Why can you not use the same primer mix for both PCR reactions? Instructors will add polymerase enzymes. Do not forget to dilute your MG1655 genomic DNA to the proper concentration prior to adding to the reaction mix. Amplification of β-galactosidase 41.5 µl sterile water 5 µl 10X PicoMaxx Buffer 1 µl 10 mm dntp Mix 1 µl 50 ng/µl MG1655 genomic DNA 1 µl 25 µm oligonucleotide primer mix 0.5 µl PicoMaxx enzyme 2. Place the tubes in the PCR Minicycler, which will use the BIOC455_LacZ PCR protocol. Be sure to record the program parameters 2

3 Day 3: Digestion and Gel Purification of PCR Products *Students will need to come to lab early in order to shorten the length of time in lab on Day 3 (ABA and ABB, report to Noyes 218 by noon, Wednesday, January 29. Section ABC should report to Noyes 218 by noon, Thursday, January 30). Only one group member needs to be present for these steps. Set up the following reactions (4 in total) in labeled 0.7-mL sterile microfuge tube. Instructors will add restriction enzymes. Please note that this is a digestion of the PCR product. Digestion of PCR products, pet15b and pet28a(+) 3 µl sterile water 4 µl 10X CutSmart Buffer 30 µl PCR or Vector Sample (Vectors should be at 50 ng/µl) 1.5 µl BamHI-HF restriction enzyme 1.5 µl XhoI-HF restriction enzyme 2. Incubate in a 37 C water bath for ~2 hr. 3. Reserve the uncut samples for your gel and for use on Day 4 (uncut vectors). After the gel, label and return the remaining vector DNA to your TA. 4. While the digests are incubating, set up a 0.7% agarose gel. Do NOT move the gel from the designated area. Carefully remove the cast gel from the casting tray and transfer it to the submarine unit. Pour 250 ml of running buffer (1X TBE) over the gel. The gel should be completely covered with buffer. 4. Add 8 µl of 6X loading dye to the digest reactions after their completion. 5. Make your uncut PCR gel samples by combining 10 µl of uncut PCR sample, 20 µl H 2 O and 6 µl 6X loading dye. Also make one undigested vector sample (either pet15b or pet28a(+), not both). Make sure your sample contains 150 ng of vector DNA. Record which vector you loaded and how you prepared the sample. 6. Load the entire volume of the above samples into the gel wells according to the following order. Also load 5 µl of the DNA ladder. Lane 1: DNA Ladder Lane 2: Uncut PCR destined for pet15b Lane 3: Digested PCR destined for pet15b Lane 4: Uncut PCR destined for pet28a(+) Lane 5: Digested PCR destined for pet28a(+) Lane 6: Uncut vector of your choosing Lane 7: Digested Vector Used in Lane 6 Lane 8: Digested Vector not used in Lane 6 3

4 7. Attach the negative electrode (black) to the well side of the chamber and the positive electrode (red) to the other side of the chamber. Run the gel at 120 V constant voltage until the first dye front has migrated 2/3 of the gel length (~30 minutes). 8. Turn off the power to the electrophoresis chamber, lift the gel tray out of the box, and take the gel to the EtBr supplemented water bath. Allow the gel to stain ~15 min. After staining, destain the gel for 10 min in a water bath. Then, image the gel (TA will assist you). MESSES WILL NOT BE TOLERATED, PLEASE CLEAN UP AFTER YOURSELF! 9. While the gel is on the UV box, take a picture of the gel. Gloves and goggles are mandatory when using UV light. Try to minimize the amount of time that your gel is exposed to UV light because it may cause adjacent thymine bases to dimerize, which leads to mutations in the DNA. 10. Using your picture, estimate the sizes of the DNA fragments of your two samples by comparing them to the DNA ladder. Record the estimated sizes in your laboratory notebook. Using the long-wavelength setting on a handheld UV lamp and a razor blade, excise only the bands for the lacz genes and vectors that have been digested. Try to keep the size of the gel slice as small as possible without leaving DNA behind in the gel. As a general rule of thumb, the smaller the gel slice obtained, the more efficient the gel purification will be! 11. Add the gel slice into a pre-weighed 1.5-ml microfuge tube. Weigh the tube with the gel slice in order to determine the gel mass. Ideally, the gel mass should be 300 mg. 12. Add 500 µl of DF Buffer to the gel slice and mix by vortex. 13. Incubate at 55 C for 15 min (or until the gel slice has completely dissolved). To help dissolve the gel, mix by inverting the tube every 2 3 min during the incubation. Be sure to solubilize the agarose completely. Cool the dissolved sample mixture to room temperature. 14. Place the DF Column into a 2-ml Collection Tube. To bind DNA, apply the sample to the DF Column without touching the filter and centrifuge at maximum speed for 30 sec. The maximum volume of the column reservoir is 800 µl. For sample volumes of more than 800 µl, simply load the additional volume and spin again. 15. Discard flow-through and place the DF Column back in the same collection tube. Collection tubes are reused to reduce plastic waste. 16. Add 400 µl of W1 Buffer to the DF Column and centrifuge at maximum speed for 30 sec. Discard the flow-through and place the DF Column back in the same collection tube. 17. Add 600 µl of Wash Buffer (with EtOH) to the DF Column and let stand for 1 min. Centrifuge at maximum speed for 30 sec and discard the flow-through. 18. Place the DF Column back in the collection tube and centrifuge for an additional 3 min to dry the column matrix. IMPORTANT: Residual ethanol from Wash Buffer will not be completely removed unless the flow-through is discarded before this additional centrifugation. It is important to remove the residual ethanol as it can inhibit downstream reactions. 4

5 19. Transfer the dried DF Column into a clean 1.5-ml microfuge tube. Open the tube and place at 37 C for 5 min. 20. To elute DNA, add 40 µl of 37 C prewarmed Elution Buffer (10 mm Tris-HCl, ph 8.5) to the center of the DF Column matrix and let the column stand for 2-5 min at 37 C. Centrifuge for 2 min at maximum speed to elute the purified DNA. IMPORTANT: Ensure that the elution buffer is dispensed directly onto the column matrix for complete elution of bound DNA. 21. Take an absorbance of 1 µl of each sample using the NanoDrop. What will you blank the NanoDrop with? If you see a massive peak around nm that swallows the DNA peak at 260 nm, continue on with the following steps. If there is a dip within this region leading into a pronounced 260 nm DNA peak, continue on to Step 28. What absorbs at nm? Be sure to save the OD readings and spectra. 22. In a hood, add an equal volume of 25:24:1 Phenol:Chloroform:Isoamyl Alcohol. Mix thoroughly and centrifuge at max speed, 2 min. at room temperature. 23. Carefully transfer the top aqueous phase into a fresh 0.65 ml microfuge tube. It is important not to collect any of the organic layer. Add ~40 µl Elution Buffer to each organic layer, mix thoroughly, and centrifuge at max speed, 2 min. at room temperature (back extraction). 24. Carefully transfer the top aqueous phase to the first aqueous layer. In a hood, add an approximately equal volume of chloroform to the combined aqueous layers. Mix thoroughly and centrifuge at max speed, 2 min. at room temperature. 25. Carefully transfer the top aqueous layer to a fresh 0.65 ml microfuge tube being careful not to collect any of the chloroform layer. Make the aqueous layer 2.5 M ammonium acetate (10 M stock). Subsequently, add 2 volumes of 100% ice-cold ethanol. Incubate on ice 10 min, and then recover the DNA by centrifugation (20 min. at max speed, 4 C). Remember how you place the tube into the microcentrifuge so that you know where the DNA pellet will be (it will not be visible). 26. Carefully remove the 100% ethanol solution without disturbing the DNA pellet. Rinse the pellet with 400 µl room temperature 70% ethanol. Recover the DNA by centrifugation (20 min. at max speed, 4 C). 27. Carefully remove the 70% ethanol solution without disturbing the DNA pellet. Dry the pellet in a Speedvac for ~10 min. Dissolve the pellet with 15 µl MilliQ water and heat at 42 C for 10 min. Afterward, centrifuge your tubes at room temperature for 1 min. at max speed. Obtain the concentration of 1 µl of your DNA using the Nanodrop. What will you blank with? Be sure to save the OD readings and spectra. Convert your ng/µl concentrations to fmol/µl (fmol = mol). 28. Carefully mix your ligation reactions using a 2:1 PCR insert:vector backbone ratio (50 fmol PCR insert: 25 fmol vector backbone) in sterile, 0.2 ml PCR tubes. There will be two reactions and two negative controls for a total of 4 ligations. What is the different between the negative controls and the reactions? You should not go beyond a 20 µl total volume (ideally you want a 10 µl volume). The total reaction volume should be 1x T4 DNA Ligase Buffer (10x Stock). It should also contain the necessary volumes of DNA and 1 µl T4 DNA ligase. Take the reaction volume to the 5

6 desired level using MilliQ water. The ligase should always be added last. All of these experimental details should be recorded in your notebook. 29. Incubate the reaction at 15 C overnight. Store the rest of your DNA sample (unused) at 4 C. Day 4: Transformation of DH5α E. coli Cloning Cells 1. Digest the pet28a(+) ligation and negative control reactions with SalI. Use anywhere from 0.5 to 1.0 µl of SalI to do this, making sure the new reaction volume is 1x CutSmart Buffer (10x Stock). Be certain to record how you set your reactions up. Digest using Minicycler program Restriction. Be certain to record the program parameters. 2. Obtain 2 microfuge tubes from the instructor; each containing 160 µl of freshly thawed DH5α competent cells. Keep these cells on ice at all times! 3. Very gently, mix the cells by flicking the tube gently with your finger, then add 40 µl of these cells to each of 8 pre-chilled 1.5-ml microfuge tubes. Label the tubes 1a to 4a and 1b to 4b and keep them on ice at all times. 4. To tube 1a, add 1 µl of the ligation mixture that used the pet28a(+) backbone. To tube 2a, add 1 µl of pet28a(+) vector (1 ng/µl). To tube 3, add 1 µl of pet28a(+) Induction Control (1 ng/µl). To tube 4, add 1 µl of ligation negative control reaction that used the pet28a(+) backbone. Very gently, mix the reactions by flicking the tubes as before (step 2). For tubes 1b to 4b, add the respective DNA s corresponding with the use of the pet15b vector. 5. Incubate the cells with these samples on ice for 15 min, swirling gently every 2 min. Quickly remove the tubes from the ice and place them in a 42 C water bath. Incubate the tubes at 42 C for exactly 45 sec. Quickly remove the tubes from the water bath and incubate them again on ice for 2 min. During the heat shock, the competent E. coli cells will take up the intact plasmid DNA. The cells are very compromised at this point and will die if the heat shock is too long (> 1 min). The incubation on ice prevents the cells from dying after heat shock. 6. Add 1.0 ml of LB media to each tube. Transfer to well-labeled sterile culture tubes and incubate at 37 C with gentle shaking for 1 h, during which time the cells will recover from the heat shock. 7. Obtain 8 LB-agar plates, 4 containing ampicillin (100 µg/ml) + X-gal (150 µg/ml), 4 containing kanamycin (50 µg/ml) + X-gal (150 µg/ml). Which plates will be used for the pet28a(+) and pet15b series of transformations? 8. For tubes 2a, 3a, 2b, and 3b, plate 200 µl of undiluted culture onto their respective plates. Record which plates you used for each. For tubes 1a, 4a, 1b, and 4b transfer the entire culture volume to 1.5 ml microfuge tubes. Pellet the cells at 8000 rpm for 3 min. at room temperature. Decant the media from the pellets and very gently resuspend them in 200 µl LB media. Plate the whole volume on their respective plates. Record the plates used for each. 6

7 a. Be sure to label around the edge of each plate with the name of the antibiotic, as well as the identity and dilution of the transformation mixture added. Be sure to use sterile technique (demonstrated by TA). Use a steel spreader, EtOH, and flame to spread the bacteria evenly over the surface of each plate. Allow the surface of the plates to dry for 5 min, invert the plates (agar side up), and place them in the 37 C incubator overnight. Extra Day: Growth of Transformed Cells 1. Retrieve your 8 plates from the 37 C incubator. Obtain 4 culture tubes and add 3 ml of LB broth. Supplement 2 tubes with 6 µl of 50 mg/ml ampicillin stock (final concentration is 100 µg/ml) and the other 2 tubes with 3 µl of 50 mg/ml kanamycin stock (final concentration is 50 µg/ml). Label the tubes with your initials and their respective antibiotic. 2. Using a sterile toothpick, select a portion of a single colony that is present on the plate containing cells from transformation tube 1a. Remove the cap, and drop the toothpick (bacteria end down) into culture tube 1a-1. Repeat this step for tube 1a-2. Then select a portion of a single colony that is present on the plate containing cells from the transformation tube 1b. Remove the cap, and drop the toothpick (bacteria end down) into culture tube 1b-1. Repeat this step for tube 1b Incubate the cultures at 37 C with shaking overnight. These strains will be used on Day 5 to isolate plasmid DNA. 4. Count the number of colonies on the plate containing cells from transformation tube 2a and 2b. Why was this control experiment performed? Describe what a low number of colonies on this plate would indicate, as well as possible causes for this result. 5. Count the number of colonies on the plate containing cells from transformation tube 3a and 3b. Why was this control experiment performed? 6. Count the number of colonies on the plate containing cells from transformation tube 4a and 4b. Why was this control experiment performed? Describe what a high number of colonies on this plate would indicate, as well as possible causes for this result. 7. Calculate the transformation efficiency as the number of transformants per ng of DNA, using your control plasmid transformation sample (2a and 2b). Keep in mind that each colony derives from a single cell (transformant), and that you need to determine the # of transformants per ng DNA plated. 8. Wrap each of your plates with Parafilm and store them agar-side up at 4 C (cold room). Day 5: Isolation of Plasmid DNA, Restriction Digestion, and Agarose Gel Electrophoresis 1. Remove the 4 culture tubes from the 37 C shaker. Each group will prep all of the 4 cultures inoculated the previous day. Add 1.5 ml of the four cultures to 4 separate microfuge tubes labeled 7

8 1a-1, 1a-2, 1b-1, and 1b-2. Cap each tube and centrifuge for 2 min at room temperature. Remove and discard the supernatant. 2. Add the remaining 1.5 ml of each culture to the appropriate microfuge tube. Cap each tube and centrifuge for 2 min at room temperature. Remove and discard the supernatant. 3. Thoroughly re-suspend each cell pellet in 200 µl of Resuspension Solution by vortex or pipetting. 4. Add 200 µl of Lysis Solution to each tube and mix gently by inverting the tube 10 times. DO NOT vortex to avoid shearing the DNA. Let stand at room temperature for NO MORE than 5 min. This step ensures that lysis is complete. 5. Add 350 µl of Neutralization Solution to each tube, cap the tubes, and mix immediately by inverting the tubes 10 times. DO NOT vortex. Remember, this must be added no later than 5 min after the addition of the Lysis Solution. Centrifuge at maximum speed for 5 min. 6. Place a GeneElute Miniprep Binding Column into each of four 2-ml Collection Tubes. Add 500 ul Column Preparation Solution to each tube and centrifuge at maximum speed for 30 sec. Discard the flowthrough and transfer the supernatant of each tube from Step 5 to one of the columns, leaving behind the white precipitate. Centrifuge at maximum speed for 30 sec. 7. Discard the flow-through from each of the bottom tubes and replace the column back into the Collection Tube. Add 750 µl of Wash Solution to each column. Centrifuge at maximum speed for 1 min. 8. Discard the flow-through from each of the bottom tubes and replace the column back into the Collection Tube. Centrifuge the empty columns at maximum speed for 3 min to dry the column matrix. Remember, the goal here is to remove all traces of EtOH. 9. Transfer each of the dried columns to clean microfuge tubes. Add 50 µl of Elution Buffer to the center of each column. Incubate at room temperature for 2-5 min. Centrifuge at maximum speed for 2 min, making sure the tube caps are toward the center of the rotor. Be sure to put the lid on the rotor to keep from shearing off the tube caps. 10. Set up the following restriction digests for each of your 4 plasmid samples in 0.7-ml microfuge tubes according to the table below. Be sure to label the microfuge tubes with the appropriate culture sample and digest. You should have 9 digest tubes. XhoI/BamHI-HF Double Digest HincII Digest NdeI Digest 6 µl DNA prep 6 µl DNA prep 6 µl DNA prep 2 µl 10X CutSmart Buffer 2 µl 10X Buffer 3 2 µl 10X Buffer 4 1 µl XhoI 2 µl 10X BSA 1 µl NdeI 1 µl BamHI 1 µl HincII 11 µl sterile water 10 µl sterile water 9 µl sterile water 8

9 11. Incubate the digests at 37 C for 1 hr 30 min. 12. While the digests are incubating, set up a 1% agarose gel one group will use each gel. Do NOT move the gel from the designated area. Carefully remove the cast gel from the casting tray and transfer it to the submarine unit. Pour 250 ml of running buffer (1X TBE) over the gel. The gel should be completely covered with buffer. 13. When the digests are complete, centrifuge the tubes at maximum speed for 5 sec in order to spin down any condensate. Add 4 µl of loading dye to each digest. Also set up three tubes of uncut plasmid (one of each sample) and add the following: 6 µl of undigested plasmid from your minipreps, 14 µl of sterile water and 4 µl of loading dye. 14. Load 10 µl of each sample into the gel wells according to the following order. Also load 5 µl of the DNA ladder in lane 4 (middle of the gel). Lane 1: DNA Ladder Lane 2: Uncut plasmid 1a-1 Lane 3: XhoI/BamHI double digest 1a-1 Lane 4: HincII digest 1a-1 Lane 5: NdeI digest 1a-1 Lane 6: XhoI/BamHI double digest 1a-2 Lane 7: HincII digest 1a-2 Lane 8: NdeI digest 1a-2 Lane 9: Uncut plasmid 1b-1 Lane 10: XhoI/BamHaI double digest 1b-1 Lane 11: HincII digest 1b-1 Lane 12: NdeI digest 1b-1 Lane 13: XhoI/BamHaI double digest 1b-2 Lane 14: HincII digest 1b-2 Lane 15: NdeI digest 1b Attach the negative electrode (black) to the well side of the chamber and the positive electrode (red) to the other side of the chamber. When orienting your gel, remember which electrode the DNA will migrate toward! 16. Run the gel at 120 V constant voltage until the first dye front has migrated 2/3 of the gel length. 17. Turn off the power, lift the gel tray out of the box, and EtBr stain/destain the gel as done on Day 2. Take a picture of your gel for your lab notebook. Be sure to wear UV-protective shielding while viewing your gel. Clearly label the samples in each lane and the MWs found on the ladder. 18. Estimate the sizes of the DNA fragments of your samples by comparing them to the DNA ladder. 19. Have you successfully constructed the desired pet15b/lacz or pet28a(+)/lacz recombinant plasmid? Explain your answer in terms of what size DNA fragments resulted from each digest and your knowledge of the composition of the parent plasmid, and the insert gene used in this experiment. What else might indicate if you have isolated the desired recombinant plasmids? 9

10 Prepare a restriction map of the newly constructed recombinant plasmid. The map should include the total number of base pairs in the plasmid, and the identity and position of the following restriction sites in the plasmid: XhoI, BamHI, NdeI, and HincII. 20. If you do not think that you have isolated the desired pet15b/xyl3b plasmid, prepare a map of the plasmid that you think you have isolated. Support your map with an explanation of how the digestion of this plasmid with the restriction enzymes used would produce the results that you obtained. 10