Bacterial Transformation

Once the target DNA has been successfully ligated into the plasmid vector, the plasmid must be transferred into the host cell for replication and cloning. In order to do this, the bacterial cells must first be made “competent.” The term “competent” is to describe a cell state in which there exist gaps or openings in the cell wall which will allow the plasmid containing the target genes to enter into the cell. Several methods to make bacterial cells competent exist, such as the calcium chloride method and electroporation. The following is the method used by the USU team to insert the plasmids containing various biobricks into the cells.

Calcium chloride Method

• Ensure the necessary antibiotic agar plates have been prepared or begin their preparation now. Four plates per transformation will be necessary (two today, then two tomorrow for streaking). Also ensure that 10 ml liquid media is made up per transformation (also for tomorrow).
• If using Biobrick parts from iGEM distribution, use registry to identify appropriate will containing plasmid of interest and proceed to step 3, if using other DNA proceed to step 5.
• Add 10ul of sterile water to distribution well to dissolve DNA. Remove 10ul and place in 0.5ml bullet tube. Label tube with part number, use 2ul to transform and save the other 8ul in the BioBrick part box.
• Take competent cells (One Shot® TOP10 Chemically Competent E.coli, Invitrogen) from the -80˚C freezer and place on an ice bath.
• Add 2 μl of the DNA solution (or 4ul of ligation reaction) to the competent cells. Ensure the pipetting is done directly into the cell solution. Let cells incubate on ice for 30 minutes. Heat water bath to 42˚C.
• Heat shock cells in the 42˚C water bath for 30 seconds. Remove and place back in the ice bath for 2 minutes.
• In the hood, add 250 μl SOC media to each tube, bringing the total cell solution to 300 μl. Incubate at 37˚C for 1 hour.
• Add 200 μl of each transformed cell solution to the appropriate antibiotic plate. Use the Bunsen burner to create a “hockey stick” out of a glass pipette tip by holding over the flame until it bends. Allow to cool. Spread cell solution uniformly over the agar plate using the “hockey stick,” then before discarding, spread residual solution on the “stick” over a second plate to get more a more sparse colony distribution.
• Parafilm all plates and place in 37˚C incubator 12-14 hours, or overnight if that is not possible.

Electroporation Method

Making competent cells for electroporation

• Streak out E.coli strain to get single colonies
• After ovenight incubation, pick a single colony. Inoculate 50ml of SOB Media. Incubate overnight at 37℃.
• Subculture to 1L of SOB Media with 5ml of the overnight culture
• Grow to O.D.550 =0.2 (3-5 hours )at 37℃.
• Pellet cells at 5,000 r.p.m. for 10 minutes in the Sorvcal, GSA rotor
• Resuspend the cells in 500ml of cold WB and recentrifuge
• Resuspend the cells again in 500ml of cold WB and centrifuge again.
• Resuspend the cells in the WB remaining in the tube after pouring off the supernatant. If necessary, adjust volume up to 4ml with cold WB
• Transfer 200ul aliquots into microfuge tubes and store at -70℃.

Electroporation

• Gently thaw the cells at room temperature, then put into ice
• In a pre-chilled microcentrifuge tube, mix 40µl of cells with 1µl (3-5µl) DNA. Mix the suspension well and place on ice for ½ to 1 minute.
• Set the machine to the following parameters: 25µF, 2.5kV, 200Ω.
• Transfer the cell solution to a pre-chilled 0.2 cm cuvette. Shake the suspension to the bottom of the cuvette.
• Pulse the cells (4-5 msec)
• Remove the cuvette and immediately add 1ml of cold SOC Media and resuspend the cells with a Pasteur pipet.
• Transfer the cells to a new tube and invubate them at 37℃ for one hour.
• Dilute the cells in PBS or SS and plate them on selective media.

Streak Plates and Liquid Cultures from Transformed Colonies

After bacterial cells have been transformed, successfully transformed cells must be selected. Because 100% of the cells do not receive the desired plasmid and target gene, it is essential to select for cells that do have the target genes. The USU team uses antibiotic resistance to select for successful transformations. To do this, an antibiotic resistance gene is also added to the plasmid vector that contains the target genes. By doing so, it is possible to know that a cell was successfully transformed based on its ability to grow on an agar plate with antibiotics added. Because the cell is able to grow, the antibiotic resistance gene must be present as well as the target gene. From the agar plates containing the antibiotics, a colony is picked and transferred into a liquid culture for further analysis. The following is the method used by USU to clone the DNA and select for the successful transformation of various BioBricks in E.coli.

Method

• Prepare two 15 ml tubes per transformation, each with 5 ml media containing the appropriate antibiotic.
• Use a pipette tip to extract half of each colony and inoculate one agar plate per colony. Using a pipette with a tip, extract the other half of each colony and inoculate one liquid media tube per colony. Label all tubes and plates and place in the 37˚C incubator until the next morning.

Plasmid DNA Isolation

Following successful bacterial cloning and isolation, it is important to verify that the target gene is in the cell and that the resultant plasmid is correct. To do this, it is a common practice to sequence the plasmid DNA. To obtain enough DNA for sequencing, the bacterial clones are grown in a liquid culture. The cells are harvested by centrifugation and then prepared for DNA plasmid extraction. DNA plasmid extraction can be done several ways, and the overall purpose is to lyse the cells and separate the plasmid DNA from all other cellular proteins, DNA, and debris. The following is the method used by the USU team to isolate plasmid DNA containing the various biobricks. Method

• Prepare two water baths, one boiling and the other 68C.
• Centrifuge bactrerial cultures (3 to 5 ml) at 3K RPM for 20 min. Discard supernatant.
• Resuspend cell pellet in 200 μl of STET buffer. Transfer to 1.5 ml tubes.
• Add 10 μl of lysozyme (50 mg/ml) and incubate at room temperature for 5 min.
• Boil for 45 sec and centrifuge for 20 min at 13K RPM (or until pellet gets tight).
• Use a pipette tip or toothpick to remove the pellet.
• Add 5 μl RNase A (10 mg/ml) to supernatant and incubate at 68C for 10 minutes.
• Add 10 μl of 5% CTAB and incubate at room temperature for 3 min.
• Centrifuge for 5 min at 13K RPM, discard supernatant, and resuspend in 300 μl of 1.2 M NaCl by vortexing.
• Add 750 μl of ethanol and centrifuge for 5 min at 13K RPM.
• Discard supernatant, rinse pellet (which cannot be seen) in 80% ethanol, and let tubes dry upside down with caps open.
• Resuspend pellet in either sterile water or TE buffer.

Restriction Enzyme Digestion and Electrophoresis

Restriction enzyme digestion is the process by which an insert DNA sequence is separated from the rest of the DNA molecule. Specific knowledge of the DNA insert is needed to determine which enzyme and conditions to use during the digestion reaction. Once the DNA sequence is known and the correct enzymes have been selected, the DNA may be digested. Listed below is the procedure used by USU to digest the plasmid DNA. After enzyme digestion, electrophoresis is used to separate the plasmid from the insert. A gel is prepared and the respective reaction mixes are loaded into the gel. Using a DNA ladder, and knowing the size of the insert, the corresponding band can be seen and cut out of the gel. The insert may then be removed and isolated from the gel, thus yielding the desired DNA. The DNA from this may then be used in PCR reactions, sequencing, ligations for further experimentation, etc. Listed below are example protocols used by the USU team for a restriction enzyme digestion and subsequent agarose gel electrophoresis.

Method

• Resuspend DNA in 20 to 40 μl water, vortex, and do a brief centrifuge to get solution to the bottom of the tube.
• Add components to the digestion solution in the following order: DNA (23 μl), 10X restriction enzyme buffer (3 μl), Xba1 (2 μl), and Pst1 (2 μl). The volume and restriction enzymes can be varied, but it should be ensured that the total volume is 10X the amount of RE buffer. Tap tubes periodically and allow to digest at appropriate temperature while preparing electrophoresis gel.
• Prepare electrophoresis gel by adding 2 g agarose to 200 ml TAE (1% solution). This is best done in an Erlenmeyer flask of adequate volume as swirling will need to be done. Place in the microwave and microwave on high for 20 seconds at a time, pulling it out and swirling until solution is homogeneous again, then repeating (BE CAREFUL to watch the solution closely when swirling – it superheats and can boil over and cause severe burns). Continue until solution is seen boiling in the microwave then gently swirl again.
• Add 20 μl ethidium bromide to solution and swirl until dissolved evenly.
• Add 6 μl of 6X loading dye to each tube of digested DNA solution.
• Prepare the electrophoresis unit by orienting the basin sideways with rubber gaskets firmly against the side. Place desired well template in the basin.
• When the agarose solution is cool enough to comfortably touch the flask, pour into the basin until the solution is about ¾ of the way to the top of the well template.
• When the gel is solidified (should look somewhat cloudy), remove the well template and change basin orientation to have the wells closest to the negative pole (as the DNA will flow towards the positive pole). Pour 1X TAE buffer into both sides of the electrophoresis unit until it just covers the gel and fills the wells.
• By inserting the pipette tip below the TAE liquid and into the well, add 10 μl of DNA ladder solution to first (and last if desired) well, skip one well, then begin adding the digested DNA solutions to the wells by adding about 2 μl less than the total volume in the tubes to prevent air bubbles in the wells.
• Place the cover on the electrophoresis unit, plug into the power source, and turn on voltage to 70 V (this can be as high as 100 V if time is an issue), and press the start button. Separation should take two to three hours. The yellow dye shows the location of the smaller nucleotide lengths and the blue dye shows the location of the larger nucleotide lengths. DNA separation can be observed as time goes on by turning off the power supply then gently removing the basin from the electrophoresis unit (be careful not to let the gel slip out of the basin) and placing on the UV transilluminator to see DNA bands. The basin can then be placed back in the electrophoresis unit for further separation if desired. Take care to not have the power supply on without the lid to the unit in place.
• When the desired level of separation is obtained, the basin can be placed on the transilluminator for picture taking. Place the cone-shaped cover over the transilluminator and place the digital camera in the top hole for pictures.

  Media Preparation

  For all experimentation involving the need for bacterial biomass and experimentation, proper media is needed to grow the cells. We use Lysogeny broth media for E.coli. and BG11 for cynobacteria. The following is the media composition.

  
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