Simply Cloning - Supplement 2 - Cracking Gel

With the cracking gel you can screen several dozen colonies in less than an hour and know if any of them has a desired clone.
Here is the protocol for making a cracking buffer and for the cracking gel procedure.
Cracking buffer:
• 39.5 ml ddH2O
• 2.5 ml 1M NaOH
• 2.5 ml 10% SDS
• 0.5 ml 0.5M EDTA
Cracking gel protocol:
• Add 20 µl of ddH2O to a tube
• Transfer an E. coli colony with a pipette tip
• Add 20 µl of cracking buffer
• Add 20 µl of loading dye
• Run on an 0.7% agarose gel
Right here on the bench I have a plate with 18 colonies that I am going to screen by a cracking gel.
I also have a replica plate, a tube with the cracking buffer, 18 new Eppendorf tubes, double distilled water, gel loading dye with bromophenol blue, and a tube with a miniprep of parental plasmid, in this case pSAT6-MCS, that will serve as a control.
Let's move on to the cracking gel protocol itself.
I start by adding to each of the tubes 20 µl of water.
In the tube one, which will serve as a control, I will put 1 µl of parental plasmid, pSAT6-MCS.
In the rest of the tubes I will transfer the colonies by picking them with pipette tips, dipping the tips in the tubes with 20 µl of water, and leaving them in the tubes for the moment.
Once all the tips are in their tubes, I am going to shake off the cells by vortexing the tubes at low speed with the tips inside.
Then I take every tip and touch the labelled LB replica plate. I will keep this plate at 37oC overnight.
Once all the cells are in, I am going to lyse them by adding 20 µl of the cracking buffer to each of the tubes. Notice that there is no mixing involved at this step.
I immediately proceed with adding 20 µl of loading dye with bromophenol blue and glycerol.
I load contents of each of the tubes on a 0.7% agarose gel and run it for 1 hour at 105 volts.
Notice that for the cracking gel I am using the 0.7% agarose, which allows for better separation of large DNA fragments.
Let's look at the picture of this gel. Since we run whole cell lysates here, we can see three types of nucleic acids: RNA is that big smear at the bottom, at the very top is genomic DNA, and the plasmid we are looking for is in the middle.
In the control lane you see how the vector would run if it did not have an insert.
Out of the 18 colonies that we are screening here, only plasmids from colonies 6 and 14 run higher than the control, which means they should have the insert we were trying to introduce.
On the next day we can go back to the replica plate, and start the miniprep cultures with clones 6 and 14.

[@ashwinnair4816]

Thank you Andriy

[@CloningStrategies]

Yes, I do mix each sample right before loading on the gel. No mixing before adding loading dye because: - This protocol is time sensitive. Cracking buffer degrades DNA as well, so I am trying to everything as fast as possible - Adding cracking buffer to cell suspension is enough to break the cell walls even without mixing - I love taking shortcuts when it makes sense ;) Thanks for the question, it is spot on.

[@parrymavi7216]

Thanks so much for this video demonstration... I have tried this a number of times unsuccessfully. Seeing it done with tiny details has been super helpful... As another time saving step, how about cutting the positive plasmid from the gel and directly transforming it into expression host if you intend to use it for protein expression? more verification can be done in the time you get the colonies, so you can be ready for protein expression by day 3...

[@perunica1010]

Thanks so much for a quick responce!

[@CloningStrategies]

PS: sucrose could protect DNA from degradation

[@andreadurrant9380]

Will this protocol work for 5300 base pair vector

[@CloningStrategies]

- Don't know about sucrose. Try it. For me this protocol worked consistently, so I had very little incentive to test others. - Most polymerases are sensitive to buffer conditions, I would think adding a lysate with NaOH and SDS will mess them up

[@CloningStrategies]

PS: Make sure you use a thick comb when you are pouring gel, so you wells are big. It would be very hard to pull it off with thin wells

[@CloningStrategies]

2. Make sure you do 20 - 20 - 20: - 20 ul of water in your tube (plus super tiny amount of E.coli) - 20 ul of cracking buffer - 20 ul of loading dye with 60% glycerol Finally, when you are loading the gel, forfeit your greediness again and load only 20 ul You are almost there, one more go, and you will be ready to teach Cracking Gel Masterclass :)

[@CloningStrategies]

Hey! One of the first times I did this protocol I also incubated longer and at higher temperature. And I also got complete degradation of all DNA. The cracking buffer is very potent. It lyses cells instantly. Add the loading dye straight away and have a gel ready so you can load and run it immediately. Let us know how it goes

[@CloningStrategies]

Haha, all the sh...t that happened for me, is happening for you as well! That means that you are on the right path. Yes, this slimy stuff makes samples float out of the wells when you are loading the gel. Two things: 1. Don't be greedy. Take a super tiny amount of cells from colony, something you can barely see. There is enough DNA there for you to see on a gel. When you take more, you get the slime

[@perunica1010]

Thanks for the video! Please can you explain why there is no mixing before adding the loading dye? Do you mix each sample before loading on the gel, though? Thanks again!