Wet Lab

After PCR or Gel Electrophoresis, DNA needs to be recovered and cleaned up to yield pure DNA for further application. We followed a PCR Cleanup Protocol to accomplish this. Two different paths were followed, either from an agarose gel or DNA in solution.

After Gel Electrophoresis:

1. A block heater with water in wells is preheated to 55°C.
2. After DNA is run through agarose gel and separated by size, the band of DNA that is of interest is cut out of the gel, leaving behind as much agarose as possible while conserving the DNA.
3. Gel piece containing DNA of interest is placed into a tared 1.7mL Eppendorf tube and weighed again, finding the weight of the gel. 300μL of ADB (Agarose Dissolving Buffer) added to Eppendorf for every 100mg of gel, up to 950uL in a single tube.
4. Tube placed in 55°C block heater until gel is dissolved, periodically shake mixing.
5. Once dissolved, contents loaded onto Zymo-Spin I Columns 600μL at a time.

From solution:

1. DNA samples in solution need an extra reagent to facilitate binding to the spin column.
2. DNA Binding Buffer added in 5:1 ratio of volume (e.x. 250μL of DNA Binding Buffer added to a 50μL solution containing DNA) and mixed thoroughly. Entire volume loaded onto Zymo-Spin I Columns 600μL at a time.

Protocol Coverage:

1. Loaded Zymo-Spin I Columns with a 2mL Collection Tube centrifuged at max RCF for 30-60 seconds, flowthrough discarded. If additional volume needs bound, load column again and repeat spin.
2. Once all of the sample is bound to the column, DNA must be washed. Add 3x600μL DNA Wash Buffer to the column and centrifuge at max RCF for 30-60 seconds, discarding flowthrough in between washes. After this, DNA in the column has been washed a total of three times.
3. Column and collection tube placed in centrifuge and spun at max RCF for 2-3 minutes, dry spinning the column and removing any remaining ethanol or other contaminants.
4. Following dry spin, 12-30μL of water, depending on expected yield, is added directly to the silica-based matrix, being careful not to touch the matrix. Allow the water to fully absorb into the column for 1-2 minutes.
5. Place the Zymo Column into a labeled 1.7mL Eppendorf tube and centrifuge at max RCF for 1 minute to elute purified DNA.
6. Analyze DNA by Nanodrop to find concentration and purity of sample.

To insert genes or sequences of interest into vectors, plasmids or inserts first need to be digested by restriction enzymes to create overhangs that can be ligated together with other overhangs with reverse complementary sequences. We used enzymes XbaI, SacI, EcoRI, and FseI at different steps during our project to cut DNA in specific spots.

1. Obtain target DNA that needs to be digested and find its concentration. Determine the enzymes to be used and ensure both can be used in the same buffer and temperature to work correctly.
2. Determine volume of solution of DNA to reach 1μg. Subtract this volume, along with the volumes of other reagents in the reaction, from 50μL to find the volume of nuclease-free water to bring the total volume to 50μL.
3. Add all parts of the reaction together, adding enzymes last. Single digests are also an option, but we only utilized double digests.
4. Program thermocycler for reaction conditions and run. For XbaI+SacI digestion, protocol was 37C for six hours, 65C for 20 minutes to heat inactivate enzymes, and held at 4C until further use.
5. After reaction is done, digested DNA can be purified by gel electrophoresis and PCR cleanup.

Transformation is the process of introducing new plasmid DNA into bacteria. We used this extensively to store and produce new plasmids.

1. Obtain ligated or supercoiled plasmid DNA for transformation, either from T4 ligase protocol or miniprepped plasmid. Preheat a heat block to 42C.
2. Thaw chemically competent cells for 5-10 minutes on ice. Once thawed, add 1uL PEG4000.
3. Add 0.5-3uL of plasmid. 0.5uL usually adequate for supercoiled plasmids, but more likely necessary for ligation products.
4. Leave cells on ice for 30 minutes
5. Place cells in tubes in 42C for 1min 10sec to heat shock cells and encourage pickup of plasmids.
6. When heat shock is done, immediately place cells on ice again for 2min.
7. After the 2min, add 700uL SOC media to cells and place in 37C 300RPM for 1hr to recover cells.
8. After recovery, plate cells following standard protocol.

Mini Prep is an important protocol that is used to isolate small plasmid DNA from bacterial cells while limiting the amount of contamination from genomic DNA and proteins. The end result is purified plasmid DNA. This protocol is a modified version of Omega Bio-Tek’s.

1. Grow 5 mL culture overnight in a 10 mL culture tube.
2. Centrifuge at max RPM for 5 minutes at room temperature. Aspirate and discard the culture media.
3. Add 250 μL Solution I mixed with RNase ensuring it is stored at the appropriate temperature. Vortex to mix thoroughly before transferring suspension to a new 1.5 mL microcentrifuge tube.
4. Add 250 μL Solution II ensuring the correct volume of ethanol was added and stored at the appropriate temperature. Invert gently several times until achieving a clear lysate. Let sit for 2 minutes and avoid vigorous mixing.
5. Add 350 μL Solution III ensuring the correct volume of ethanol was added and stored at the appropriate temperature. Immediately invert several times until a flocculent white precipitate forms in the tube.
6. Centrifuge at maximum RPM for 10 minutes in which a white pellet will form.
7. Insert a HiBindⓇ DNA Mini Column into a 2 mL Collection Tube.
8. Transfer the supernatant by carefully aspirating it into the HiBindⓇ DNA Mini Column.
9. Centrifuge at maximum speed for 1 minute and then discard the filtrate. Reuse the collection tube.
10. Add 500 μL HBC Buffer diluted with 100% isopropanol. Centrifuge at maximum RPM for one minute. Discard the filtrate and reuse the collection tube
11. Move HiBindⓇ DNA Mini Column to vacuum manifold. Add 700 μL of DNA Wash Buffer diluted with 100% ethanol. Turn on vacuum so that buffer passes slowly through; turn off vacuum when drained and relieve pressure. Discard the filtrate and reuse the collection tube.
12. Repeat step 11 two more times (three washes total).
13. Centrifuge the empty HiBindⓇ DNA Mini Column at maximum RPM for 2 minutes (with open caps) to dry the column.
14. Transfer the HiBindⓇ DNA Mini Column into a nuclease-free 1.5 mL microcentrifuge tube.
15. Add 30 μL sterile deionized water. Let sit at room temperature for 1 minute. Centrifuge at maximum RPM for 1 minute.
16. Store eluted DNA at -20 ℃.

Gel electrophoresis is a valuable technique for analysis and purification of DNA. When voltage is applied, the negatively charged DNA travels across the agarose gel, separating the DNA fragments by size. These bands of DNA can then be analyzed and cut out of the gel for purification.

1. Make the gel. A large gel is made with 150mL 1X TAE buffer and 1.5g agarose powder. Microwave the agarose/TAE buffer until agarose is completely dissolved, making sure not to let any liquid boil over. Allow the liquid to cool slightly before adding 3μL Gel Star stain and mix gently. Pour the gel in a mold with a comb for the wells and allow to cool completely. Make sure no bubbles are present on the comb. Gels can commonly be made from 0.5%-3% agarose, and percentage gel is dependent on the size of the expected DNA fragment.
2. Place the gel correctly in frame, ensuring the wells are on the anode side. Add 1X TAE buffer to the frame until the gel is completely covered.
3. Load the 12μL ladder into the left most well (Either a 1kb or 50bp ladder depending on DNA).
4. Add 6X gel loading dye to the samples to reach a final 1X concentration (e.x. 10μL into a 50μL sample) and mix thoroughly. Load the samples carefully into the other wells (30μL each) and note their location.
5. Close the frame and plug it in. Run the gel at 100V for 1.5-2hrs, depending on size and percent gel, to separate the DNA fragments.
6. When done, analyze the gel. Individual DNA bands can be cut out with a razor blade and saved for recovery by PCR cleanup.

Liquid cultures:

1. 5mL LB media added to 7mL centrifuge tube. Chloramphenicol added to make final concentration 35ug/mL (e.x. Add 3.5uL of 50mg/mL stock antibiotic to 5mL culture).
2. From either a cryostock or isogenic colony from an agar plate, touch bacteria with a pipette tip and place in liquid culture.
3. Place in a shaking incubator at specific conditions. We largely used 37C 300RPM for 18-20hr.

Agar Plates:

1. LB agar made fresh and autoclaved, or heated in the microwave until completely melted.
2. ~25mL of LB agar taken per plate. Once liquid is cooled slightly, antibiotic added and mixed, liquid poured into the plate. LB agar is allowed to cool and solidify under sterile conditions.
3. Once agar is cooled, bacteria can be plated. 100uL of bacteria in liquid added to plate. Spreading tool placed in 100% ethanol and placed in flame until all ethanol is burned off and spreading tool is sterilized. Use a spreading tool to spread liquid evenly across the plate. Resteralize the tool when done.
4. After all liquid is spread, the plate can be stored in an incubator upside down to allow bacterial growth.

Cryostocks:

After a liquid culture is grown overnight, it is taken out of the incubator and centrifuged to pellet cells. Supernatant discarded.
750uL of fresh LB media used to resuspend cells and transferred to cryostock tube.
750uL of sterile 50% glycerol added to the cryostock tube with the cells, making a final 25% glycerol solution. Tube closed tightly and briefly vortexed to homogenize culture.
Cryostock is placed in -80C for later use.

This protocol was used to grow more CC cells from our cryostock of NEB 5 alpha. This procedure was necessary for the later step of transformation.

1. Inoculate cryostock of cc cells into 5mL of LB media and incubate for 18 hours at 37C, 300RPM.
2. Place 300uL of culture and 100mL of LB into 500ml flask and put it a shaking incubator at 37C and 300RPM until optimal OD600 of 0.2-0.4.
3. Transfer to 50 mL falcon tubes and set on ice for 30 min
4. Spin the tubes down at 4000 rcf at 4C for 10 minutes
5. Discard Supernatant and resuspended in ~1mL of 0.125M CaCl2 and transfer to 1.7mL Eppendorf tubes
6. Spin down cells at 4000 rcf 4C for 1 minute and discard Supernatant
7. Wash 3x more with ~1mL 0.125M CaCl2
8. Combine cells and resuspend in 600uL 0.125M CaCl2 and 400uL 50% glycerol.
9. Aliquot 55uL of the cell solution into eppendorfs and placed in -80C.

Cell free TXTL assays were utilized to monitor fluorescence of samples loaded with riboswitch containing plasmids. As time goes on, mRFP1 is produced and fluorescence is monitored versus time.

1. Determine concentration of plasmid DNA in stock solution and amount that will be used in TXTL reaction. 1nM-20nM (25ng-500ng) is optimal.
2. Set up protocol in Tecan Spark. Set up constant temperature (29C) and loop. Loop contains orbital shaking at 180 RPM, followed by fluorescence measurement every 3-10 minutes, repeating for 12-16 hours.
3. Determine reagent volumes for TXTL assay from Diacel Arbor Biosciences myTXTL handbook. For a 12uL reaction, 9uL should be Master Mix. Determine necessary volume of plasmid and ligand to add to volume. If using T7 kit, ensure T7rnap plasmid is included. Reaction volume can be scaled down if necessary.
4. Remove TXTL Master Mix and necessary plasmids from -80C and -20C. Thaw at room temperature and place on ice until use. Gather all other necessary reagents.
5. Vortex Master Mix for ~20 seconds and centrifuge for 5-10 seconds. Pipette mix until no precipitate is observed in the stock solution.
6. Add master mix, plasmid, and any other necessary reagents to the PCR tube and mix thoroughly. Avoid incorporating bubbles.
7. Transfer samples to microplate in predetermined wells. Seal plate with plate cover.
8. Place microplate in microplate reader and begin fluorescence assay.
9. After the assay is complete, remove the plate from microplate reader and interpret the data.