EDTA also helps destabilize the cell membrane. This membrane is made up of a “sandwich” of phospholipids, which are kinda soap-like molecules that have long fatty tales that form the hydrophobic (water-excluded) “peanut butter” of the sandwich, and hydrophilic (water-loving) heads that form the “bread” facing the watery environments of the inside of the cell (cytoplasm) and the outside of the cell. These phospholipids have negatively-charged phosphate groups that bind to positively-charged ions that helps keep them from repelling one another. So, by stealing those cations, the EDTA sets the scene for some head-to-head repulsive action that’ll make the next step easier What’s this next step? Lysis! We didn’t want to break the cells open before, when the outside of the cell was so “dirty”, but now that we’ve resuspended it in this “safer” & “cleaner” environment, we can go ahead lyse them. And for this we use another buffer 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. bit.ly/sdspageruler And SDS will denature the proteins in the cells (which is fine (even good) because we don’t want them anyway), 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 those sandwiches of phospholipids (and lipopolysaccharides (LPS’s) in the outer outer membrane (these have sugar chains sticking off them), with a peptidoglycan wall (made up of peptide-linked sugars) in between (thankfully for us, e. coli is 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-avoided (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 thus the pH is higher. In acidic conditions, there are more H⁺ than OH⁻ and the pH is lower (yeah, it’s an inverse log scale so more protons is lower pH…). 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. bit.ly/phacidbase The “alkaline” part of alkaline lysis comes from sodium hydroxide (NaOH). 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.” (egads…) Nitrogenous bases, aka nucleobases, aka “bases” are the unique parts of DNA letters that stick off the individual strands and form specific hydrogen bonds (H-bonds) with bases on the other strand (A:T & C::G). bit.ly/nucleicacidstructure In addition to helping disrupt the cell wall, NaOH disrupts such base pairing between strands of DNA. Both the plasmid and the bacterial genomic DNA (gDNA) are double-stranded and NaOH breaks up the H-bonding between the 2 strands, “unzipping them.” These H-bonds aren’t as strong as the covalent bonds linking the letters up into chains, so you can unzip the strands without unchaining them. Here’s a brief overview of why… Those strong covalent bonds come from sharing pairs of electrons, and H-bonds can come as a consequence of unfair pair shares. Some atoms (like oxygen and nitrogen) are electron-hoggy (electronegative), so they pull the shared electrons closer to themselves, making them partly negative and their partner partly positive. And opposite charges attract, even partial ones, so you can get hydrogen atoms attached to electronegative atoms becoming partly positive and getting attracted to electronegative atoms with a lone pair of electrons and a partial negative charge. And we call this a hydrogen bond (H-bond). bit.ly/frizzandmolecularattractions 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 (for those not familiar with the arm-numbing power that is the vortex, it’s a really fast orbital shaker kinda like a small, very aggressive, electronic foot massager. 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. But 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 rezipping is a lot harder for gDNA, 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 (and a vinegar-y smell). 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 only 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: bit.ly/spincolumns Here’s the gist: We apply the solution to a spin column - these columns have membranes made of silica (amorphous (no-consistent-shape) silicon dioxide, SiO₂), which offers a variety of binding options for DNA. You pipet your sample into the “cuppy” part above the membrane and use centrifugation or vacuum to help pull the liquid through. 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 has guanidinium chloride, a chaotropic salt which disrupts (brings chaos to) the water coat around the DNA and lets it bind the membrane instead. Then comes the wash. Now that you’ve gotten DNA stuck on there tight, you want to wash off any lingering extra salts. You do this using a buffer containing ethanol (buffer PE in Qiagen’s kit). Just add the liquid the same way you added your sample, spin it through & toss it out. DNA’s not soluble in ethanol, but salts are, so the extra salts are removed with this wash step. After the wash & toss, spin it again to give any residual ethanol another chance to get pulled through. It’s really important to remove the ethanol or the DNA will have problems dissolving &/or when you go to load samples into an agarose gel (more here: bit.ly/agarosegelcompare) they’ll float up out of the well cuz ethanol’s less dense than the buffer. Once we’ve washed off the gunk we can redissolve the pDNA (which is now pure!) so it can flow on through. The Qiagen kit comes with an elution buffer (buffer EB - which is 10 mM Tris·Cl, pH 8.5) (Tris is that buffering agent we saw before) you can use or you can use plain old (but really pure, nuclease-free) water. If you use water, give it a minute to fully dissolve the DNA before you spin it through. Speaking of which -> once you redissolve it, you spin it through into a NEW TUBE - this one you want to keep! Then, take your spun-through, pure DNA over to the spectrometer like a NanoDrop & check out its absorbance spectrum (it should have a 260/280 ratio of ~1.8). More on that here: bit.ly/beerslawexplained 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: 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. Happy prepping! And, if the mini prep isn’t enough fun for you, there’s also bigger versions - maxi & mega & gigapreps! Those can be helpful if you’re trying to purify a lot of template DNA for in-vitro transcription (DNA-to-RNA copying) bit.ly/t7rnap more on molecular cloning: bit.ly/molecularcloningmovie & bit.ly/molecularcloningguide DIY instructions: openwetware.org/wiki/Qiagen_Buffers & malooflab.phytonetworks.org/wiki/Miniprep_solutions_recipes/ more on topics mentioned (& others) #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com
EDTA also helps destabilize the cell membrane. This membrane is made up of a “sandwich” of phospholipids, which are kinda soap-like molecules that have long fatty tales that form the hydrophobic (water-excluded) “peanut butter” of the sandwich, and hydrophilic (water-loving) heads that form the “bread” facing the watery environments of the inside of the cell (cytoplasm) and the outside of the cell. These phospholipids have negatively-charged phosphate groups that bind to positively-charged ions that helps keep them from repelling one another. So, by stealing those cations, the EDTA sets the scene for some head-to-head repulsive action that’ll make the next step easier
What’s this next step? Lysis! We didn’t want to break the cells open before, when the outside of the cell was so “dirty”, but now that we’ve resuspended it in this “safer” & “cleaner” environment, we can go ahead lyse them. And for this we use another buffer
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. bit.ly/sdspageruler
And SDS will denature the proteins in the cells (which is fine (even good) because we don’t want them anyway), 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 those sandwiches of phospholipids (and lipopolysaccharides (LPS’s) in the outer outer membrane (these have sugar chains sticking off them), with a peptidoglycan wall (made up of peptide-linked sugars) in between (thankfully for us, e. coli is 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-avoided (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 thus the pH is higher. In acidic conditions, there are more H⁺ than OH⁻ and the pH is lower (yeah, it’s an inverse log scale so more protons is lower pH…). 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. bit.ly/phacidbase
The “alkaline” part of alkaline lysis comes from sodium hydroxide (NaOH). 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.” (egads…) Nitrogenous bases, aka nucleobases, aka “bases” are the unique parts of DNA letters that stick off the individual strands and form specific hydrogen bonds (H-bonds) with bases on the other strand (A:T & C::G). bit.ly/nucleicacidstructure
In addition to helping disrupt the cell wall, NaOH disrupts such base pairing between strands of DNA. Both the plasmid and the bacterial genomic DNA (gDNA) are double-stranded and NaOH breaks up the H-bonding between the 2 strands, “unzipping them.” These H-bonds aren’t as strong as the covalent bonds linking the letters up into chains, so you can unzip the strands without unchaining them. Here’s a brief overview of why…
Those strong covalent bonds come from sharing pairs of electrons, and H-bonds can come as a consequence of unfair pair shares. Some atoms (like oxygen and nitrogen) are electron-hoggy (electronegative), so they pull the shared electrons closer to themselves, making them partly negative and their partner partly positive. And opposite charges attract, even partial ones, so you can get hydrogen atoms attached to electronegative atoms becoming partly positive and getting attracted to electronegative atoms with a lone pair of electrons and a partial negative charge. And we call this a hydrogen bond (H-bond). bit.ly/frizzandmolecularattractions
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 (for those not familiar with the arm-numbing power that is the vortex, it’s a really fast orbital shaker kinda like a small, very aggressive, electronic foot massager.
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.
But 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 rezipping is a lot harder for gDNA, 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 (and a vinegar-y smell). 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 only 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: bit.ly/spincolumns
Here’s the gist: We apply the solution to a spin column - these columns have membranes made of silica (amorphous (no-consistent-shape) silicon dioxide, SiO₂), which offers a variety of binding options for DNA. You pipet your sample into the “cuppy” part above the membrane and use centrifugation or vacuum to help pull the liquid through. 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 has guanidinium chloride, a chaotropic salt which disrupts (brings chaos to) the water coat around the DNA and lets it bind the membrane instead. Then comes the wash. Now that you’ve gotten DNA stuck on there tight, you want to wash off any lingering extra salts. You do this using a buffer containing ethanol (buffer PE in Qiagen’s kit). Just add the liquid the same way you added your sample, spin it through & toss it out.
DNA’s not soluble in ethanol, but salts are, so the extra salts are removed with this wash step. After the wash & toss, spin it again to give any residual ethanol another chance to get pulled through. It’s really important to remove the ethanol or the DNA will have problems dissolving &/or when you go to load samples into an agarose gel (more here: bit.ly/agarosegelcompare) they’ll float up out of the well cuz ethanol’s less dense than the buffer.
Once we’ve washed off the gunk we can redissolve the pDNA (which is now pure!) so it can flow on through. The Qiagen kit comes with an elution buffer (buffer EB - which is 10 mM Tris·Cl, pH 8.5) (Tris is that buffering agent we saw before) you can use or you can use plain old (but really pure, nuclease-free) water. If you use water, give it a minute to fully dissolve the DNA before you spin it through.
Speaking of which -> once you redissolve it, you spin it through into a NEW TUBE - this one you want to keep!
Then, take your spun-through, pure DNA over to the spectrometer like a NanoDrop & check out its absorbance spectrum (it should have a 260/280 ratio of ~1.8). More on that here: bit.ly/beerslawexplained
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: 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.
Happy prepping! And, if the mini prep isn’t enough fun for you, there’s also bigger versions - maxi & mega & gigapreps! Those can be helpful if you’re trying to purify a lot of template DNA for in-vitro transcription (DNA-to-RNA copying) bit.ly/t7rnap
more on molecular cloning: bit.ly/molecularcloningmovie & bit.ly/molecularcloningguide
DIY instructions: openwetware.org/wiki/Qiagen_Buffers & malooflab.phytonetworks.org/wiki/Miniprep_solutions_recipes/
more on topics mentioned (& others) #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com
THANK YOU SO MUCH FOR THIS OMG😭💗💗
Super informative video!