Team:Michigan/Experiments
Experimental Flow
Below we have outlined the path our project took, from the initial cloning through experimentation and submission.
Isolated LacZ from iGem part BBa_K564012
1) Rehydrated DNA from distribution kit
2) Transformed into DH5a chemically competent cells
3) Extracted plasmid
4) PCR mutagenesis to engineer desired cut sites around LacZ
Transferring LacZ into pET28
1) Digested pET28 and new version of BBa_K564012 (with added cut sites)
2) Ligated together
3) Submitted for sequencing to confirm
4) Transformed into new DH5a chemically competent cells
Creation of delta-M15 mutant
1) Deletion of segment including alpha fragment of LacZ using NEB Q5 site directed mutagenesis kit
2) Submitted for sequencing to confirm successful deletion
Proximity dependent ligation assay
1) Probe segments and bridge segments mixed with T4 ligase in presence or absence of thrombin
2) Gel electropheresis to detect ligation
Cloned delta-M15 mutant into submission vector
1) Digested linearized pSB1C3 with our mutant
2) Ligated together
3) Transformed into new DH5a chemically competent cells
5) Extracted DNA
6) Dried, packaged, shipped
3) Transformed into new DH5a chemically competent cells
Protocols
Below are the protocols we used for al stages of our project. We wrote them to be as easy to follow as possible, to accommodate new team members with little lab experience. We hope future teams find these highly detailed versions of common protocols useful.
Micropipetting
1) Choose the right pipet for the amount of liquid you want to transfer. You should choose the smallest pipet capable of transferring the amount of liquid you desire. (If you want to transfer 185 uL you should use the pipet with a 200 uL maximum volume NOT the pipet with a 1000 uL maximum volume).
2) Set the desired amount on the side of the pipet by twisting the knob on the top of the pipet. Be careful where the decimal point is located on the display as this varies depending on the size of the pipet. The decimal point between mL and uL or between uL and nL will always be marked somehow (usually with a line, sometimes by change between red numbers vs. black numbers). DO NOT EVER TWIST THE KNOB PAST THE UPPER AND LOWER LIMITS FOR THE PIPET!
3) Slide the bottom end of the pipet into a clean, autoclaved tip of the appropriate size. You can tell if a tip box is autoclaved because it will have a stip of autoclave tape with black stripes. Press down firmly so that when you pull the pipet back up, the tip remains fixed to the end of the pipet. You should keep tip boxes closed when you are not working with them to prevent contamination.
4) Push the plunger (knob on top) down with your thumb until you feel some resistance. This is the pipet’s “first stop.” Do not push past this stop yet.
5) With the plunger pushed to the first stop, insert the tip into the liquid you want to pipet, just below the liquid’s surface. (Minimizing the tip’s contact with the liquid helps prevent contamination). Do not allow the sides of the pipet to touch the inside of the container! Remember that only the disposable plastic tip is sterile
6) Slowly lift up your thumb to allow the plunger to rise back to its starting position. If you do it too fast you will create air bubbles that can throw off the measurements. As you let the plunger back to its starting position, the pipet will suck up the desired amount of liquid. If you are pipetting from a small container such as a 1.5 mL tube you may need to move the pipette downward as you do this in order to keep the tip under the surface of the liquid.
7) Remove the tip from the liquid. Keep the pipet completely upright at all times when there is liquid in the tip. Turning the pipet upside down or sideways will cause liquid to fall into the body of the pipet, ruining it.
8) Look at the liquid in the tip to make sure you actually pulled it up! There should not be any bubbles.
9) Lightly touch the pipet tip to the side of the new container just above the level of the liquid. Again, do not allow the sides of the pipet to touch the walls of the container! Only the disposable plastic tip is sterile
10) Slowly press down on the plunger to expel the liquid. This time, push past the first stop, all the way until the second stop, where you can push no more.
11) With the plunger still pushed all the way, remove the pipet tip from the container, scraping the tip on the wall to help get every last bit of liquid out of the tip.
12) Once clear of the new container, release the plunger, and eject the tip into a biohazardous solid waste bin (will usually just look like a small plastic bin full of used tips on the lab bench). Most pipets have a button near the plunger that will eject the tip.
Gel Electropheresis
1) Fill a beaker with 50ml of 1X TAE buffer.
2) Weigh out and add 0.75g of agarose. (to make 1.5% gel)
3) Microwave for 3 minutes, then run the beaker under cold water until the glass has cooled so you can touch it.
4) Add 2uL of Gel Red
5) Pour the beaker’s contents into the gel mold.
6) Add plastic “combs” to the mold to make wells.
7) Push any large bubbles to the side walls if possible using a spare comb or pipet tip.
8) Let the gel cool and solidify.
9) Once the gel is solid, move it into the electrophoresis device. Make sure the ‘top’ end of the gel (with the wells) is on the same side as the negative (black) terminal.
10) Pour 1X TAE buffer to the fill line.
11) Slowly and gently remove the plastic combs from the wells by wiggling and pulling them. Take care not to rip and of the walls separating the wells.
12) Select a ladder to use based on what kind of samples you’re running.
13) Load 10uL of the ladder into the leftmost well on each row you intend to use. Change pipet tips each time to avoid contaminating the ladder stock. When loading the ladder into the gel, insert the pipet tip below the surface of the buffer, and slightly inside the well. This is tricky to do and takes some practice. Take care not to push too far and puncture the bottom of the well with the pipet tip. A good way to check that you’re in the right spot is to wiggle the pipet tip every so slightly, and see if you can feel the walls of the well. When you’re confident you’re in the right spot, slowly eject the ladder into the well. The ladder is heavier than the buffer and will sink into the well. Try to avoid bubbles.
14) For each sample, in a small PCR tube, add 10uL of the DNA sample and 2uL of 6X Purple loading dye
15) Load each sample into a well in the same way you loaded the ladder. Use a new pipet tip for each sample.
16) Once everything is loaded, double check the gel is oriented in the correct direction.
17) Triple check the gel is oriented in the correct direction. (Waiting an hour only to find out your gel ran off the wrong edge and all your samples are lost is tragic.)
18) Plug the electrophoresis device into the power source. Make sure the lid is closed. If you touch the buffer while the power source is on, you will be dangerously shocked.
19) Change the power source settings to 90 volts.
20) Turn the power source on and run the gel until the dye gets close to the end but dont let it run off the end. Usually an hour to 90 mins.
21) When the gel is done, turn off the power supply, then unplug it before opening the gel box and removing the gel.
22) Look at the gel using UV light source and take a picture.
Transforming DH5a strain E.coli Cells
USE STERILE TECHNIQUE THROUGHOUT
1) Warm the water bath to 42 C
2) Thaw the plasmid DNA on ice.
3) Thaw one 50 uL vial of DH5-Alpha cells on ice.
4) Pipet 1-5 uL of the plasmid into the vial of cells, mix by gently tapping. DO NOT MIX BY PIPETTING UP AND DOWN!
5) Store the extra plasmid at -20 C.
6) Put the vial in ice for 30 minutes.
7) Make sure the hot water bath is at 42 C.
8) Put the vial in the 42 C water bath for EXACTLY 30 seconds. Do not mix or shake.
9) Put the vial back on ice for 5 min.
10) Add 250-800 uL of room temperature SOC medium to the vial. SOC is a super rich media so take extra care that nothing unsterile gets in as contamination is really easy.
11) Tape the vial to a shaking incubator, and shake/incubate at 37 C and 225 rpm for exactly an hour.
12) Transfer 50 uL, 500 uL, and 1000 uL, respectively, of the diluted culture onto three petri dishes.
13) Spread evenly using glass spreader. STERILIZE THE SPREADER BETWEEN USES!
14) Incubate the plates overnight at 37 C.
Cloned delta-M15 mutant into submission vector
1) Digested linearized pSB1C3 with our mutant
2) Ligated together
3) Transformed into new DH5a chemically competent cells
5) Extracted DNA
6) Dried, packaged, shipped
3) Transformed into new DH5a chemically competent cells
Proximity dependent ligation assay
1) Probe segments and bridge segments mixed with T4 ligase in presence or absence of thrombin
2) Gel electropheresis to detect ligation
Cloned delta-M15 mutant into submission vector
1) Digested linearized pSB1C3 with our mutant
2) Ligated together
3) Transformed into new DH5a chemically competent cells
5) Extracted DNA
6) Dried, packaged, shipped
3) Transformed into new DH5a chemically competent cells
Proximity dependent ligation assay
1) Probe segments and bridge segments mixed with T4 ligase in presence or absence of thrombin
2) Gel electropheresis to detect ligation
Cloned delta-M15 mutant into submission vector
1) Digested linearized pSB1C3 with our mutant
2) Ligated together
3) Transformed into new DH5a chemically competent cells
5) Extracted DNA
6) Dried, packaged, shipped
3) Transformed into new DH5a chemically competent cells
