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No differences between cut and uncut, that depends on the size of the removed fragments and the kind of gel (and time) that you are running it. A difference of 100bp on short gel for a 7,4kb vector is really small. So if you really want to be sure, you might want to play with agarose concentration and running time to improve separation.
please post th photo on the forum for everyone to see
Buffers shoould not interfere with the extraction because after running on the gel they have long been diluted and moved in the gel, so it is not likely your problem.
One thing that can lead to poor extraction efficiency in all extraction kit (they are all based on the same technique: the binding of DNA to a silica matrix): what are you collecting your DNA with in the last step? the kits usually say use our solution or water. But the problem is that elution of DNA happens only at a pH slightly above 7, and milliQ/double distilled water is usually acidic...
Science has proof without any certainty. Creationists have certainty without
any proof. (Ashley Montague)
I have been eluting my digested vector with EB buffer (pH of Qiagen kit. I've also used preheated water previously to compare the yield of one vs the other. But I didnt get anything back. Now I dont know whether my nanodrop is messed up (which I dont think is true) or something else. The spectra on nanodrop looks like blank but on a gel looks alright..but then I dont get any colonies.
oh ya, one more thing, I had also digested my 'ligation mix' with third enzyme (eco R1 - whose sites are absent from the insert as well as deleted from mcs during cloning) as suggested by the manual, and yet I got colonies on my control vector - which is simply doubly digested vector, with NO insert and ligase added and then again digested with EcoR1 to remove any background parental vectors. I got colonies even on those..and when I screened for them, they were running so much higher compared to where a religated vector or an uncut vector runs..
so I am totally just losing my mind as of now!
I am attaching the gel photo of my uncut and digested and then excised/purified vector. Sorry for replying this late.
Thanks for all the help!
Except for the first gel looking weird and the one double-digest being over-digested for some reason, I don't see anything egregiously wrong. I'll have to think about it some more. I'm still inclined just to start again with fresh everything, if possible: fresh vector DNA, fresh buffers, fresh enzymes...
I used to do clinical chemistry with an instrument that had a reputation for being dependable, but quirky. Occasionally, it would fail to work properly and for the life of you there didn't seem to be anything wrong with the machine--and yet it wasn't working. The standard solution was to take it apart and put it back together again. That "fixed" a surprising number of problems.
Thanks for all your suggestions!
As you recommended I have restared everything from the scratch. That is PCR of inserts, dbl dig of inserts and vectors etc. A few questions I had I was hoping someone/you can asnwer:
(1) How much necessary is it to CIP treat your dbl digested vector?
(2) Can you set up dbl dig on a vector (without running it on a gel) and straight purify it either by et-oh ppt OR by PCR clean up kit?? If so, will you have a lot of negative colonies on ur ligation transformation due to circular/uncut vector?
(3) What is the correct way to reduce the background of selfligated vector (negative colonies) after transformation of ligation?? -- I heard that you can set up a digestion with RE whose site is not present in the insert and also deleted from the MCS of the vector. This would linearize any uncut vector; However, I dont know whether I should add 10X RE buffer and REnzyme straight to the ligation mix or not.
(4) Should I heat inactivate the ligase/RE enzyme OR would it inhibit the transformation efficiency?
(5) And lastly (for now), do I need to purify (either by Et-OH ppt) my ligation-and-digestion reaction before transformation??
So sorry for these many questions, but I have been trying this for several months, and want it to work really well.
p.s. Regarding my issues with GEL purifying dbl digested vector earlier, I purified my dbl digested vector with Promega PCR clean up kit and was finally able to get a concentration 10 ng/uL and 23 ng/UL in ~45 ul of water. Also, do we lose more DNA if it's ET-OH ppt vs PCR cleaned up??
Many thanks in advance,
(1) Sorry can't help
(2) Probably, if you trust the efficiency of your digestion. Or if you ran some of it on a gel and saw it did have an almost perfect efficiency.
(3)I would ethanol precipitate the ligation product and do the digest in the correct RE Buffer.
(4) I never did
(5) I don't think it is useful. I never digested after ligation, but I have always transformed with the ligation mix. I don't think ligation and restriction reaction are that different.
(ps) I have never compared efficiency, but I know that kits do not claim 100% efficiency. And I have heard that greater efficiency in DNA precipitation is achieved with isopropanol or (supposedly even better) butanol at room temperature.
Science has proof without any certainty. Creationists have certainty without
any proof. (Ashley Montague)
1) If the ends of your dd’d vector are incompatible, you don’t have to worry too much about CIPing the ends. The reason for the phosphatase treatment is to reduce the efficiency of self-ligation, just as, in reverse, you phophorylate the insert to enhance its ligation efficiency. Incompatible or non-sticky ends shouldn’t ligate whether or not they’ve been dephosphorylated. CIP can be partly inactivated by heat, but it is better to phenol extract after CIPing prior to ligation. I do my cloning these days by LIC, which doesn’t require using CIP/Kinase or REs.
2) You don’t have to gel purify the vector, but it’s recommended. If your vector ends are incompatible, I wouldn’t worry so much about self-ligation, and there shouldn’t be any undigested vector left, but I would be concerned about interference by the small fragment chopped out of the vector during RE digestion. Phenol/chloroform extraction followed by EtOH precipitation should reduce the amount of contamination, but gel purification should eliminate it altogether.
3) The correct way to reduce the background? Don’t have one. I’ve never resorted to RE digestion to remove unwanted DNA. The Stratagene site-directed mutagenesis kits take advantage of methylated vs non-methylated DNA to selectively digest non-mutated DNAs with DpnI which greatly enriches the products for mutants, but that’s the closest I’ve seen to what you’re suggesting to do. I’ve not seen it done, and I wouldn’t think it would be necessary. If you really want to do the extra digestion, then I would at least heat-inactivate the ligase (and I would probably phenol-extract followed by EtOH pptn) before adding the other RE in its correct buffer. I confess, I’m not too fond of the idea, but if you insist, go ahead and try.
4) You don’t need to inactivate the ligase before transformation. I probably would inactivate any RE either by heat or phenol extraction/EtOH pptn or both.
5) You shouldn’t have to do anything to your ligation mix before transformation.
Essentially what canalon said, only with more words.
The only other thing to mention is the ratio of ends of vector to ends of insert in the ligation mix. You want to have an excess of insert over vector, but you don’t want to overdo it either. I would follow the recommendations in the manufacturer’s information for the ligase—or check out the Maniatis bible on protocols in molecular biology for some direction. I haven’t heard too many good things about the Intein system, I’m afraid, but most (if not all) the complaints I’ve heard about are issues with the expression of protein, not with the cloning.
Thanks a lot for your comments/suggestions!! They are really really helpful. Unfortunately, my vector produces sticky ends (with NdeI and Xma). The reason why I was asking about Et-OH was because I am not able to gel purify the vector. Yesterday I did transformation of all my ligations that had different ratios of I:V (in range of 3:1 to 48:1)with a control plate that had digested vector + NO INSERT + ligase.
All the ligations were treated with a third enzyme to remove any background and were NOT heat inactivated prior to transformation ( I read somewhere else that heat inactivation prior to transformation lowers the efficiency of transformation). And today when I checked all 9 plates, I DID NOT get a single colony on any of them including control plates. I can only come up with 2 possible explanations:
(1)All the dbl digested vector must have either religated or not cut at all, and thus must have been removed by third enzyme
(2)Not heat inactivating the enzyme might have cause transformation to fail.
I probably should not have changed all these at once. Next time I will keep your suggestions in mind and proceed with changing one thing at a time.
I am beginning to lose my mind!
Thanks for alll your help!!
NdeI and Xma (I or III?) have compatible ends?? I didn't think so, unless you're blunting them with T4 polymerase or something like that. Presumably, your transfection positive control worked and you know your competent cells are still in good shape. When you designed your primers for pcr , did you add a few (3-4) extra junk bases at the ends? Some REs are fussy about being too close to the end of a piece of DNA sequence--and I don't know if that is the case for either of your enzymes. I think the appendices in the NEB catalog has a table for cutting efficiencies as a function of the length of the extension off the end. If you didn't allow for that in your primer design, and one or more of your enzymes is affected by that sort of thing, then you may not be cutting your insert to completion, or even, possibly, very well.
When I say that NdeI and Xma() don't have compatible ends, I mean that they can't ligate to each other. The vector should not self-ligate with high frequency because the NdeI overhang does not match the Xma() overhang; they will not hybridize with each other. The NdeI overhang is supposed to be sticky with the complementary NdeI-cut insert DNA, and so is the Xma() site sticky for its counterpart on the insert. Its not that the digestion product produces overhangs that determines whether the dd'd ends are compatible. Say you were using XhoI and SalI as your enzymes, These ends are compatible and you could expect a certain amount of self-ligation of the vector. I don't think self-ligation is your problem. I suspect the problem is somewhere in the ligation--assuming you know your competent cells are OK.
Depending on which table you want to use, NdeI requires 7 extra bases to cut a small oligonucleotide, or at least 6 extra bases to be able to efficiently cut an NdeI site near the end of a linear fragment. XmaI isn't nearly so sensitive to being near an end--it only needs 2 extra bases to be able to cut to >98%. This is coming from the appendicies in the NEB catalog, "Cleavage Close to the End of DNA Fragments". If your pcr primers don't have at least 6 extra bases before the NdeI site, I recommend making new primers with extra bases. I used to add something like GATTGG to each end for good measure. It doesn't matter too much what the extra sequence is. They are only there to enhance the cutting of the ends after the pcr reaction.
Sorry for taking so long to reply and thank you for your suggestions. I use Nde1 and Xma1 enxymes to digest my vector as well as my inserts; and my primers include GGTGGT extra residues before Nde1 and Xma1 sites.
So here's another problem I've been running into - not getting positive clones!!!
Here's what I've been doing!!
(1) dbl digesting my vector and inserts (nde and xma) (37 C for 3.5 hrs and heat kill for 20min at 65C) - both are new enzyme (nde 1 is from promega and xma is from NED - these two have worked before with my 2 other inserrts)
(2) CIP treating dbl dig pTYB2 vector (7.4kb)
(3) Et-OH/Chloro extract the dbl dig vector only (and gel purify DNA inserts ~200 bp) [I do NOT gel purify the digested vector b/c for whatever reason I am not getting any DNA back. By doing et-oh ppt i get plenty of DNA back - offcourse it carries circular/undig/supercoied along with it.]
(4) Using dig vector and insert, I set up ligation with diff ratios of 3:1, 6:1, 12:1,20:1 and 40:1..My controls are (1) digested vector (without insert) + ligase (2) dig vector (w/o inserrt) and w/o ligase and (3) uncut vector.
(5) Got plenty of colonies on all ratios of I:V; 6 colonies on control dig vector (w/ ligase) and A LOT on dig vector w/o ligase. This does not make any sense how the dbl dig vector without ligase would have more colonies.
(6) Moving on., i selected 3 colonies per plate and did screens for total of 34 colonies (all same constructs different ratios of I:V) and all came to be negative; bands same as mcs of uncut vector.
(7) So I used all these ligations and set up another digest with an enxyme (EcoR1) whose site is absent in the insert and is deleted from the mcs upon cloning. This would reduce any background!!
(8) transformed them and did not get any colonies!! this prolly suggest tht all the colonies tht I had contained either uncut plasmid or religated plasmid - which sucks!!
Has anyone had similar probs? what should i do?
Thanks in advance
I confess, your problem baffles me. It seems that all you recover is vector. In the early days of recombinant DNA, some fragments from mammalian cells or viruses would not clone into pBR322-related vectors. I seem to recall references to so-called poison sequences (distinct from things like ccdB which is a bacterial “poison” gene) and that altering or removing such sequences was required for successful cloing of those particular fragments or genes. I don’t remember what the sequence(s) is, though. You’d have to go back to the literature from the mid-eighties, I would guess. I’ve never had that kind of a problem so far as I was aware, but it could explain (maybe) the failure (apparently) of any vector carrying your 200 bp insert to survive selection—namely, that there is some kind of selection against your insert. But this is sheer speculation. If this were the problem, and everything else were fine, I would expect to see low yields in your ligations, and you don’t see that, or so it seems. You say you get a reasonable number of colonies at all ratios—no differenes at all in the yield no matter what the ratio of insert to vector? That’s kind of suspicious, too.
Are you sure you didn’t accidently switch the two controls? It makes no sense for there to be more clones in the no-ligase control. I can’t explain it as anything but a fluke.
I don’t understand why you can’t recover vector DNA from a gel. Clearly, you can purify your fragment, so it’s not that you can’t purify anything from gels. Extraction and precipitation may not remove all of the mcs fragment that enzyme digestion drops out of the vector. These little pieces of mcs will compete with your insert for vector ends during ligation and a) reduce the yield of recombinants, and b) increase the background of “undigested” vector. There shouldn’t be much uncut vector remaining in your double-digest.
The long and short of it is that I don’t know what to suggest. If anything strikes me, I’ll post it, but for now I’m as stumped as you are. You’re sure it’s TBY2 you have, and not TBY1 which doesn’t have the SmaI/XmaI site. Eh, most likely you’ve checked that, or maybe you don’t even have the other vector in the lab.
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