Work in a lab & there are some “boring” experiments you have to do A LOT in order to do the “exciting” experiments that require more thought. Like “minipreps” (aka ALKALINE LYSIS) in which we separate and purify plasmid DNA we put into bacteria from all the stuff that was already in the bacteria. So we can then do things like check that the plasmid has the gene you think you put in there. And, if so, stick the plasmid into cells that’ll make protein from it. When you have a lot to clone, this is a technique you’re gonna hone!
The first times you do an experiment it’s always kinda stressful & it’s hard to imagine it ever seeming “boring” but, for things you end up having to do a lot that don’t require a lot of thought (at least “brain thought”) thanks to muscle memory, you can get bored – if you let yourself. But if you imagine what the molecules are doing as you do it, it makes it a lot more fun. And it helps you really appreciate why you’re doing what – and this is especially important when you’re working from a kit. So let the bumbling biochemist help you beat boredom & bring some pep to the MINIPREP!
Aka ALKALINE LYSIS, “minipreps” are experiments in which we separate and purify the plasmid DNA we put into bacteria from all the stuff that was already in the bacteria. You can’t just break the cells open (lyse them) & pull out all the DNA because the bacteria has its own DNA you aren’t interested in. But some cool chemistry comes to the rescue!
Say you want to study a protein. You can use molecular cloning to stick the gene for that protein into a circular piece of DNA called a VECTOR PLASMID & use transformation to stick that plasmid into bacterial cells. Not all of the cells will take in the plasmid, but only those that do will be able to grow on agar plates with food spiked with an antibiotic that the plasmid provides resistance to.
Each cell that took in the plasmid will start growing and dividing, leading to globby dots on the plate we call colonies. Each colony represents 1 original cell taking in the plasmid and it’s replicating asexually (just copying its DNA & splitting itself in 2) so each cell within a colony should have the same plasmid (though the cells might have different numbers of copies of it) but cells in different colonies might have different plasmids
If all went well, all of the plasmids taken in will be “perfect” – the gene really got in there ok, there aren’t any typos etc. But all doesn’t always go perfectly so we want to check.
But we’ll need more DNA. So we choose a few colonies and stick a bit of them in some liquid media (typically 5mL LB) and let them grow overnight (at 37°C with shaking). Then we can isolate the plasmid DNA and send it for sequencing to make sure all is A-OK before moving on.
But the plasmid is in bacterial cells that have their own genomic DNA (gDNA) as well as a bunch of proteins and stuff. So how do we isolate the plasmid DNA (pDNA)?
Clearly, we’ll need to get the plasmid DNA (pDNA) out of the cells, which we can do by breaking the cells open (LYSIS). But before you break the cells open, you want to make sure they have an ideal environment to come out into. The cell membrane provides a nice natural barrier to allow you to swap out the external environment.
Right now, the cells are in the liquid media you grew them in – usually LB with an antibiotic for selection – so that media has salts, peptides (from the tryptone), vitamins etc. (from the yeast extract), all sorts of stuff the bacteria want/need to survive and thrive (and make lots of copies of your plasmid)
If our goal is to purify the pDNA, might as well get rid of as much of that stuff we don’t want now, while the cell membrane provides a convenient natural barrier (especially because some of that stuff could interfere with future steps). Also, we want to minimize the volume we’re working with. Right now, the cells are “spread out” in excess media but if we “spin them down” by centrifugation, we can separate the cells (heavy) from the media (light).
- glycerol – osmotic balancer – keeps the cells from bursting before you’re ready – we’ll burst them with detergent not pressure differences
- RNaseA – this degrades RNA (and is why P1 has to be stored in the fridge) – we don’t want the RNA so might as well break it down so it doesn’t clog things up or try to tag along
- Tris – acts as a pH buffer – we’re not ready to change the pH yet
- EDTA – chelating agent (metal-biter) -> binds divalent cations (molecules with a +2 charge) like Mg2+, Ca2+ that DNases need – prevents DNases from degrading the plasmid (RNaseA doesn’t need cation cofactors so it can still work and chew away the RNA)
- EDTA also helps destabilize the cell membrane -> the hydrophilic heads of phospholipids have negatively-charged phosphate groups that bind to positively-charged ions that helps keep them from repelling one another
Once you’ve resuspended it in this “safer” & “cleaner” environment, you can go ahead and break the cells open – LYSIS
LYSIS BUFFER has a detergent called Sodium Dodecyl Sulfate (SDS) – this is the same detergent we use to denature proteins (unfold them and keep them unfolded) when running SDS-PAGE gels to separate proteins by size. And it’ll denature the proteins in the cells, but before it does that, it has to get into the cells. And when it does, it breaks down the door so everything comes spilling out.
The “doors” of e. coli are cell membranes made up of sandwiches of phospholipids (and lipopolysaccharides (LPS’s) in the outer outer membrane (these have sugar chains sticking off them), with a peptidoglycan wall in between (thankfully for us e. coli’s Gram negative so this wall isn’t very thick). SDS knocks down the door by being sneaky. Like other detergents, it’s amphiphilic -> it has a water-loving (Hydrophilic) part and a water-avoiding (hydrophobic) part
The phospholipid & LPS molecules making up the cell membrane are also amphiphilic, so the SDS can kinda “join their club” -> disrupt the membrane, breaking the cell open (it also denatures the proteins embedded in the membrane
Alkaline is another name for “basic” and is the opposite of acidic. In alkaline conditions, there are more hydroxide ions (OH-) than protons (H+) and the pH is higher. In acidic conditions, there are more H+ than OH- and the pH is lower. The reason we care is that molecules can give or take protons depending on how many there are around, and this changes the giver or taker’s characteristics.
The “alkaline” part of alkaline lysis comes from sodium hydroxide (NaOH). In addition to helping disrupt the cell wall, it disrupts base pairing between strands of DNA. Both the plasmid and the bacterial genomic DNA (gDNA) are double-stranded and NaOH breaks up the hydrogen bonding (H-bonding) between the 2 strands, “unzipping them”
NaOH is a “base” and a base (at least in one definition of the term) is a molecule that “steals” protons (H+) – it can steal protons from DNA “bases” (the unique parts of DNA letters that stick off the individual strands and form hydrogen bonds with bases on the other strand)
These H-bonds rely on having hydrogen, so if NaOH steals that hydrogen, base pairing is disrupted and the strands come apart. This isn’t the final product we want (we want double-stranded pDNA) so we’ll have to let the pDNA strands come back together but there is a point to this step, don’t worry – at the same time we’re unzipping the pDNA we’re also unzipping the gDNA and we won’t let that come back together. And that will let us discriminate between the pDNA (which we want) and the gDNA (which we DON’T want)
You want the lysis buffer to be able to reach all the cells (so hopefully you resuspended the pellet well in P1) You could be vigorous there, because the DNA was still protected by the cell membranes, but once you break the cells open you need to be more careful. When you add P2 (lysis buffer) you want to mix well, BUT gently -> you don’t want to mix too vigorously or you’ll “shear” the gDNA -> break it into pieces that will stay soluble. So mix by inverting the tube not by pipetting or (definitely not) vortexing.
NEUTRALIZATION – This is where we get the pDNA to come back together by “undoing” the proton stealing. Instead of trying to get the NaOH to give it back, we add a fresh source – acidic potassium acetate.
The potassium acetate acts as an acid, donating protons, so the DNA bases act as bases and take the protons -> now they can form base pairs again. If they can find their binding partners.
The plasmid DNA is small (relatively speaking), circular, and supercoiled (really twisted up), so even though the strands unzip under the alkaline conditions, they stay near each other – their partners are right next door, so they can rezip readily.
The gDNA’s a lot bigger, (e. coli’s genome’s ~4.7 million basepairs (letters) whereas a plasmid’s usually less than ~200 thousand) so this is a lot harder especially since, when it got denatured, hydrophobic bases were exposed that latched onto lipids and proteins to form a kind of tangled up mess
And the proteins and lipids in that mess are bound to SDS. And when you added potassium acetate you didn’t just add acid, you also added potassium. As you might have found out the hard way if you’ve run SDS-PAGE on proteins purified in a buffer with high KCl, unlike sodium dodecyl sulfate (the sodium salt of dodecyl sulfate), the potassium version is insoluble.
So that big web of stuff bound to the SDS (all that protein, lipids, – and gDNA!) will precipitate (come out of solution) as a kinda fluffy white precipitate.
You spin this down again and the precipitate will separate from the liquid supernatant containing your plasmid. But this liquid doesn’t just have your plasmid – you’ve gotten rid of gDNA and (most) proteins & cellular debris, but you still have a bunch of salts, EDTA, RNase, etc. to get rid of
Clean up time! At this point, things are basically like a PCR purification (“clean-up”), which I go over here: http://bit.ly/2Cd8vS9
Here’s the gist: We apply the solution to a spin column – the DNA binds the column -> we let everything else flow through. We wash it. We unstick it and let it flow through.To help with the sticking, N3 also has guanidinium chloride, a chaotropic salts which disrupts the water coat around the DNA and lets it bind the membrane instead. Once we’ve washed off the gunk we can redissolve the pDNA which is now pure!
In the QIAprep kit you have the option of adding “LyseBlue reagent” (thymolphthalein dissolved in ethanol) to P1 It serves as a pH indicator to help you see whether you’ve mixed well (but gently, remember) in the lysis step and whether you’ve really neutralized in the neutralization step.
Thymolphthalein deprotonates at a pH ~10 which makes it go from colorless to blue (more on pH indicators here: http://bit.ly/2C9bA5J )
P1 has a pH of 8 (thanks Tris!) so it’ll be colorless at this point. But when you add P2 you’re raising the pH, so it’ll turn blue. And you want it to turn blue everywhere, evenly. When you add the neutralization buffer (N3), you lower the pH, so there are more free protons, so it re-protonates and goes colorless again.