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 subcloning chemically competent 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.
Miniprep DNA Extraction using Qiagen kit
1) Centrifuge the vial with bacterial cells for 5 minutes at 9000 rpm to form a pellet of cells at the bottom. REMEMBER TO BALANCE CENTRIFUGE! If you are using a previously frozen pellet, ignore this step.
2) Pour out the supernatant (the liquid surrounding the pellet) into a liquid waste container. Do this carefully so that the pellet remains at the bottom of the tube.
3) Add 250uL of Buffer P1 (KEPT IN THE FRIDGE-> PUT BACK IN THE FRIDGE) to the tube containing the pellet, then pipet up and down until the pellet is completely resuspended. Make sure beforehand that the P1 bottle’s top has a checkmark in the appropriate box (RNase). When you can no longer see any trace of the pellet, expel all liquid from the pipet back into the tube and proceed.
4) If the cells aren’t already in a microcentrifuge tube, transfer them to one now.
5) Add 250uL of Buffer P2.
6) With the lid closed flip the tube upside down, then back to normal, 4-6 times to mix. Wait 1-2 minutes before proceeding.
7) Add 350uL of Buffer N3, then mix again, flipping it 4-6 times.
8) Centrifuge for 10 minutes at 13,000 rpm. REMEMBER TO BALANCE.
9) The kit comes with its own blue open top tubes with filters in them. Get one of these. There are two parts, the inner part with a filter and the outer part that’s just a tube. Label both of these parts to avoid any mix ups.
10) This is a critical step. Read this whole thing very carefully or you will likely have to repeat this whole procedure. Transfer as much of the supernatant as you can (usually between 800uL and 1000uL) from the tube you just centrifuged onto the filter of the tube from the kit. TWO IMPORTANT NOTES: 1) Do not touch the gunk at the bottom of the tube you just centrifuged, try and avoid sucking any up into the pipet by placing the pipet tip on the tube’s wall a little above where the gunk is. It is better to get a little less than 800uL and no gunk, than 800 uL and gunk. 2) DO NOT touch the filter of the tube from the kit with the pipet tip. Hold the tip just above the filter and drip the contents onto the filter. Do it slowly so that the filter has time to absorb it and you don’t overflow and lose anything.
11) Centrifuge the new tube with the filter in it for 60 seconds at 13,000 rpm. BALANCE.
12) Detach the bottom/outer part of the tube, dump its contents but keep the tube itself. It’s ok if you don’t get all of it out, just get the tube as empty as possible.
13) Put the inner part with the filter back inside the now empty outer part. Drip 500uL of Buffer PB onto the filter, again taking care not to touch the filter.
14) Centrifuge for 60 seconds.
15) Detach the outer part and discard the flow-through again, then put the inner part back inside.
16) Drip 750uL of Buffer PE into the filter, again taking care not to touch the filter. Make sure the bottle’s top has a check mark in the appropriate box.
17) Centrifuge for 60 seconds. Discard the flow-through.
18) Centrifuge again for 60 seconds.
19) Discard the whole outer part of the tube, placing the inner part in a new, clean, and labeled normal tube.
20) Drip 50uL of Buffer EB onto the filter, again taking care not to touch the filter.
21) Let it stand for 1-2 minutes.
22) Centrifuge for 60 seconds.
23) Throw out the inner portion of the tube. KEEP THE FLOW-THROUGH!!! THIS IS WHERE THE DNA IS NOW.
24) Store the DNA at 20C.
Restriction Digest
Restriction digests allow you to cut DNA at sites specific to the restriction enzyme used. Use the sequence of the DNA you’re working with and the goals of the experiment you’re working on to determine which restriction enzyme you need to use. Different enzymes require different buffers to work. You can use the New England Biolabs website to decide which buffers to use and what temperatures to heat inactivate your enzymes at. Restriction enzymes are incredibly expensive, so please check with a senior team member before doing a restriction digest to confirm you’re using the correct enzymes and buffers. KEEP THE RESTRICTION ENZYME ON ICE AT ALL TIMES!!! KEEP IN THE FREEZER AS MUCH AS POSSIBLE!
1) In a PCR tube, add 1ug of the DNA you want to digest. This is a measurement of weight, so you need to do a little math based on the concentration of the vector DNA to figure out how many uL this translates to.
2) Add 5uL of the 10X NEB Buffer you selected to match the restriction enzyme you’re using.
3) Add 1uL of the restriction enzyme.
4) Add ddH2O until total volume is 50uL
5) Close the tube, then flick it a few times gently to mix the contents.
6) Incubate for 1 hour at the temperature specified by the restriction enzyme you’re using.
7) If the restriction enzyme you’re using says to heat inactivate, heat the reaction at the required temperature.
Ligation
Ligation lets you take a strand of DNA that has compatible sticky ends (usually from a restriction digest) and insert it into a vector (usually a plasmid).
1) Label a microcentrifuge tube to perform the reaction in.
2) Add 2uL of T4 DNA Ligase Buffer to the tube.
3) Add 50ng of the vector DNA to the tube. This is a measurement of weight, so you need to do a little math based on the concentration of the vector DNA to figure out how many uL this translates to.
4) Add 37.5ng of the insert DNA to the tube. This is also a measurement of weight, so you need to do a little math based on the concentration of the vector DNA to figure out how many uL this translates to.
5) Add nuclease free water to the tube so that the total volume is 19uL. This amount will vary based on the volumes of DNA you added.
6) Add 1uL of T4 DNA Ligase.
7) Gently mix the reaction by pipetting up and down a few time.
8) Incubate at 16C overnight OR 10 minutes at room temperature.
9) Put in 65C hot water bath for 10 minutes.
10) Chill on ice, then either transform into cells or store at -20C.
PCR
PCR allows for the rapid copying of a very small amount of DNA. It is also useful because it can only amplify specific regions you want it to, letting you isolate segments of DNA that you care about. Use the New England Biolabs website to calculate the program that is optimal for your particular PCR and primers.
KEEP ALL REAGENTS AND THE REACTION ITSELF ON ICE THROUGHOUT THIS PROCEDURE! Also, take extra care to avoid any contamination. PCRs are notoriously finicky and even a tiny bit of DNA contamination will get amplified and can ruin the reaction. It is far better to use an extra pipet tip here and there than to repeat a PCR reaction because you didn’t.
1) Determine the number of PCR reactions you will be performing, then add 2.
2) Multiply all the following amounts by this new number to create the “master mix” for the PCR. In a microcentrifuge tube add
25uL Q5 2X high fidelity master mix (NEB), 0.625uL 40uM forward primer, 0.625uL 40uM reverse primer, 23.75uL Nuclease Free Water.
3) Mix the master mix by vortexing for a few seconds.
4) In small PCR tubes, on ice, add 2uL of the template DNA from each sample.
5) Add 48uL of the master mix to each PCR tube. CHANGE TIPS FOR EACH TUBE.
6) Load the PCR tubes onto the heat cycler.
7) Heat cycle through the heating program recommended for your primer and sample by the NEB website.
Proximity dependent ligation assay #1
1) Dilute all probe oligos to 100uM and all connector oligos to 200uM in nuclease free water.
2) In PCR tubes mix the following 6 reactions:
-----A) 2uL 10X DNA Ligase buffer, 2uL NEB DNA ligase, 2uL probe 1, 2uL probe 2, 1uL connector 1C, 11uL nuclease free water
-----B) 2uL 10X DNA Ligase buffer, 2uL NEB DNA ligase, 2uL probe 1, 2uL probe 2, 1uL connector 2C, 11uL nuclease free water
-----C) 2uL 10X DNA Ligase buffer, 2uL NEB DNA ligase, 2uL probe 1, 2uL probe 2, 1uL connector 3C, 11uL nuclease free water
-----D) 2uL 10X DNA Ligase buffer, 2uL NEB DNA ligase, 2uL probe 1, 2uL probe 2, 1uL connector 1C, 0.979uL Thrombin, 10.02uL nuclease free water
-----E) 2uL 10X DNA Ligase buffer, 2uL NEB DNA ligase, 2uL probe 1, 2uL probe 2, 1uL connector 2C, 0.979uL Thrombin, 10.02uL nuclease free water
-----F) 2uL 10X DNA Ligase buffer, 2uL NEB DNA ligase, 2uL probe 1, 2uL probe 2, 1uL connector 3C, 0.979uL Thrombin, 10.02uL nuclease free water
3) Incubate reactions at 37C for 2 hours, then heat inactivate at 70C for 10 minutes
4) Add 6X loading dye to each sample.
5) Load the 6 reactions into a 1.75% agarose gel, along with 1kb ladder and 105.5ng of probe 1 in its own lane as a control.
6) Run the gel.
Proximity dependent ligation assay #2
Same as previous protocol, but incubate reactions at 35C for 9 hours.
Proximity dependent ligation assay #3
Same as previous protocol, but incubate reactions at 25C for 4 hours and use 2% agasose gel.
Proximity dependent ligation assay #4
Same as previous protocol, but make two sets of the reactions. Incubate one set of reactions at 20C and the other set at 37C for 4 hours and use 2% agasose gel.
NEB Q5 Site-directed Mutagenesis
1) Assemble the following reagents in a thin-walled PCR tube: 12.5uL Q5 Hot Start High-Fidelity 2X Master Mix, 1.25uL 10 μM Forward Primer, 1.25uL 10 μM Reverse Primer, 1uL Template DNA (1-25 ng/uL), 9uL nuclease free water.
2) Mix reagents completely, then transfer to a thermocycler.
3) Perform the following cycling conditions: 30s @ 98C, 25x[10s @98C, 10-30s @ 50-72C (use NEB annealing calculator), 25s/kb @72C], 2mins @72C, hold at 4C.
4) Assemble the following reagents: 1uL PCR product, 5uL 2X KLD Reaction Buffer, 1uL 10X KLD Enzyme Mix, 3ul Nuclease-free Water.
5) Mix well by pipetting up and down and incubate at room temperature for 5 minutes.
6) Transform into chemically competent cells.
