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galK Recombineering

1. Design galK primers with 50 bp homology to an area flanking the desired site to be modified. The 3’ end of these primers bind to the galK cassette. The primers should look as follows:

Forward: 5’------------50bp_homology-------CCTGTTGACAATTAATCATCGGCA-3’ Reverse: 5’----50bp_homology_compl._strand----TCAGCACTGTCCTGCTCCTT-3’

4. Transform a well-characterized BAC (or fosmid, in our case) into electrocompetent SW102 cells. Recover for 1 hour at 32°C, and plate on LB plates with 12.5 mg/ml chloramphenicol. (See steps 9-13 if you don't know how to prepare electrocompetent E. coli).

5. PCR amplify the galK cassette using the primers designed in step 1 and a proof-reading Taq-mix (we used Invitrogen's Platinum HiFi Taq mix). Use 1-2 ng template (the pgalK plasmid). 94°C 15 sec., 60°C 30 sec., 72°C 1 min., for 30 cycles. Add 1-2 ml DpnI per 25 ml reaction, mix, and incubate at 37°C for 1 hour. This step serves to remove any plasmid template; plasmid is methylated, PCR products are not. The digest was gel purified, then the PCR repeated using the DpnI-digested PCR product as template, and undergoing a second DpnI digestion on the result. This greatly cuts down on the non-recombinant background. From a strong PCR band, purified, and eluted in 30 ml ddH2O, we use 2.5 ul for a transformation (approx. 10-30 ng).

6. Inoculate an overnight culture of SW102 cells containing the fosmid in 5 ml low-salt LB + chloramphenicol (12.5 mg/ml). Incubate at 32°C.

7. Next day, turn on two shaking waterbaths: One at 32°C, the other at 42°C. Make an ice/water slurry and put a 50 ml tube of ddH2O in there to make sure it’s ice-cold (see later). Also ice three 15 ml round-bottomed Falcon tubes and three micorcentrifuge tubes (1.5-2 ml). Dilute 600 ml of the overnight SW102 culture containing the target BAC in 30 ml low salt LB with chloramphenicol (12.5 mg/ml) in a 50 ml baffled conical flask and incubate at 32°C in a shaking waterbath to an OD600 of approx. 0.6 (0.55-0.6). This usually takes 3-4 hours.

8. Transfer 10 ml each to baffled 50 ml conical flasks and heat shock at 42°C for exactly 15 min. in a shaking waterbath. The remaining 10 ml is left at 32°C as the uninduced control.

9. After 15 min, the three samples are briefly (~5 min) cooled in an ice/waterbath slurry and then transferred to three 15 ml Falcon tubes and pelleted using 5000 RPM at 0°C for 5 min. (We had good results spinning at 4150-our max rotor speed-but 5000 rpm is recommended.) It’s important to keep the bacteria as close to 0°C as possible in order to get good competent cells.

10. Pour off all of the supernatant and resuspend the pellet in 1 ml ice-cold ddH2O by gently swirling the tubes in the ice/waterbath slurry. No pipetting. This step may take a while. When resuspended, add another 9 ml ice-cold ddH2O and pellet the samples again.

11. Repeat step 10.

12. After the second washing and centrifugation step, remove 9 ml of supernatant and resuspend by swirling the pellet in the remaining 1 ml water. Transfer the cell suspension to the chilled microcentrifuge tubes, and spin again at 5000 rpm, 0 C, 5 min.

13. Transform the now electrocompetent SW102 cells. We use 25 ml cells for each electroporation in a 0.1 cm cuvette (BioRad) at 25 mF, 1.75 kV, and 200 ohms. After electroporation of the PCR product, the bacteria are recovered in 1 ml LB for 1 hour at 32°C.

14. After the recovery period the bacteria are washed twice in 1xM9 salts as follows: 1 ml culture is pelleted in an eppendorf tube at 13,200 RPM for 15 sec. and the supernatant removed with a pipette. The pellet is resuspended in 1 ml 1xM9 salts, and pelleted again. This washing step is repeated once more. After the second wash, the supernatant is removed and the pellet is resuspended in 1 ml 1xM9 salts before plating serial dilutions in 1xM9 (100 ml, 100 ml of a 1:10 dilution, and 100 ml 1:100) onto M63 minimal media plates with galactose, leucine, biotin, and chloramphenicol. Washing in M9 salts is necessary to remove any rich media from the bacteria prior to selection on minimal media.

15. Incubate 3 days at 32°C in a cabinet-type incubator.

16. Streak a few colonies onto MacConkey + galactose + chloramphenicol indicator plates. Streak to obtain single colonies (see appendix B). The colonies appearing after the 3 days of incubation should be Gal+, but in order to get rid of any Gal- contaminants (hitch-hikers), it is important to obtain single, bright red colonies before proceeding to the second step. Gal- colonies will be white/colorless and the Gal+ bacteria will be bright red/pink due to a pH change resulting from fermented galactose after an overnight incubation at 32°C.

17. Pick a single, bright red (Gal+) colony and inoculate a 5 ml LB + chloramphenicol overnight culture. Incubate at 32°C. There is normally no need to further characterize the clones after the first step.

18. Repeat steps 7 through 12 above to obtain electrocompetent SW102 cells (now ready for a galK <> mutation substitution). If you are going to transform a double-stranded DNA oligo, the two complementary oligos can be annealed in vitro: Mix 10 mg of each oligo in a volume of 100 ml 1x PCR buffer. Boil for 5 min. Let cool slowly to room temp (30 min.). Add 10 ml 3 M NaAc and 250 ml EtOH. Precipitate, wash once in 70% EtOH, and resuspend the final, air-dried, pellet in 100 ml ddH2O (final conc. of 200 ng/ml). Use 1 ml per transformation.

19. Transform the bacteria (25 ml of heat-shocked and 25 ml of uninduced control) with 200 ng double-stranded oligo, a PCR product, or anything containing a mutation and with homology to the area flanking the galK cassette. Recover in 10 ml LB in a 50 ml baffled conical flask by incubating in a 32°C shaking waterbath for 4.5 hours. This long recovery period serves to obtain bacteria, by “dilution”, that only contains the desired recombined BAC, and thus have lost any BAC still containing the galK cassette.

20. As in step 14, pellet 1 ml culture and wash twice in 1xM9 salts, and resuspend in 1 ml 1xM9 salts after the second wash before plating serial dilutions (100 ml, 100 ml of a 1:10 dilution, 100 ml 1:100, and 100 ml 1:1000) on M63 minimal media plates with glycerol, leucine, biotin, 2-deoxygalactose (DOG), and chloramphenicol.

21. Incubate at 32°C for three days.

22. The number of colonies may or may not be significantly different when comparing plates from uninduced and induced bacteria (range between 1:1 – 1:100). In either case, you will still be able to find true recombinants with a high frequency. Analyze, say, 10-12 colonies by SpeI digestion of BAC miniprep DNA (see appendix C). Include a SpeI digest of the parent BAC as a control. Clones with a digestion pattern like the parent are likely to have undergone the desired mutation. Background clones (DOG resistant without the desired mutation) willl have obvious deletions, and should not be analyzed further. The clones with correct digestion pattern should be analyzed by PCR and sequencing of the mutated region. If the pattern is identical to the parental digestion pattern, it means that the bacteria most likely became DOG resistant due to the desired homologous recombination event. Alternatively, any deletion that contains the region with the galK cassette, but excludes the chloramphenicol region, will also be selected in the counterselection procedure. With a high frequency of homologous recombination as in he SW102 strain, the background is not likely to be a problem. In the unlikely event that too high a background is observed, try to increase the length of the homology arms, to increase the frequency of homologous recombination.