Lena- these two restriction enzymes have some special features that could be causing you problems. Sap1 cuts away from the recognition site in a region that is not sequence specific, so not all SapI digested DNA is complementary. You need to make sure the overhangs have complementarity (not simply check that both have the SapI binding site). Second, NdeI is extremely salt-sensitive, and in my experience does not cut well near the end of PCR fragments. Make sure that your PCR products have at 3-4 bp of flanking nucleotides 5' to the restriction site. Finally, it is possible that your ligation is being inhibited by a contaminant. Try a control ligation with just insert alone (100-200ng) and run it on an agarose gel. This will let you know whether your inserts are ligation-competent.

The fact that your single digested vector is not re-ligating suggests to me that it may be either over-digested (damaged ends), or a contaminant such as phenol or a guandinium salt used in the gel isolation is sneaking through the protocol and killing the ligase activity. Qiagen's Qiaex II resin works better than the column-based technologies for low amounts of DNA, and might help fix this problem. Alternatively, you could try to clean up your gel isolated material by ethanol precipitation. Good luck!

Did you phosphorylate the plasmid before ligation?

Hi Lena,

this sounds really mysterious. My guess would be that your ligation is the problem, since even the single cleaved vector doesn't work. Trying a different ligase and ligating a little bit longer (over night or at least for a few hours) sounds like a good idea to me. Maybe your ligation buffer is not working anymore (too many freeze/thaw cycles)? Try a fresh batch of ligation buffer or supplement it with some ATP. Maybe your competent bacteria are also not competent enough, so that you see enough colonies with an uncleaved control, but no colonies after an (inefficient) ligation.

I hope you find a solution! Keep us up to date!

Good luck,

Eva

How big is your insert & did you CIP your vector? Also, are these home grown competent bacteria or purchased?

Hi Alberto, thank you for you answer. No, i didnt use any phosphatase, because it did help when i had a lot of false positives before, but it reduced the of true positives as well, and, as this seemed like a difficult cloning... I just wanted to see some colonies, you know the feeling?

And Eva, thank you for your suggestions. I also think it is the ligation, or thought. This is why I did supplement an old buffer with DTT and ATP, and used a new buffer/enzyme and a new quick ligase in parallel when i tested ligation with the re-assembly of the 1kd ladder.

And yet, it is possible that the competent cells are just not helthy enough, i did so many trafos that i was running out of fresh ones.

I hope to know more on monday (cant bear coming on the weekend to see some more empty plates)

Thank you

lena

Whenever I had problems like this I just got fresh ligase buffer as Eva already pointed out. Just get a fresh batch, and then aliquot it in 5-10 µl aliquots, so you use a fresh tube every time you clone something. Since I started using aliquots I never had any problems with my clonings....

Lena- these two restriction enzymes have some special features that could be causing you problems. Sap1 cuts away from the recognition site in a region that is not sequence specific, so not all SapI digested DNA is complementary. You need to make sure the overhangs have complementarity (not simply check that both have the SapI binding site). Second, NdeI is extremely salt-sensitive, and in my experience does not cut well near the end of PCR fragments. Make sure that your PCR products have at 3-4 bp of flanking nucleotides 5' to the restriction site. Finally, it is possible that your ligation is being inhibited by a contaminant. Try a control ligation with just insert alone (100-200ng) and run it on an agarose gel. This will let you know whether your inserts are ligation-competent.

The fact that your single digested vector is not re-ligating suggests to me that it may be either over-digested (damaged ends), or a contaminant such as phenol or a guandinium salt used in the gel isolation is sneaking through the protocol and killing the ligase activity. Qiagen's Qiaex II resin works better than the column-based technologies for low amounts of DNA, and might help fix this problem. Alternatively, you could try to clean up your gel isolated material by ethanol precipitation. Good luck!

Ah cloning the bane of many a scientists life! I had a few issues not too long ago and it turned out to be due to the fact that I had used TE buffer in the final gel purification stepped - when I changed this to ultra pure water most of my issues disappeared. So what was your insert in? also did you dephosphorylate your vector backbone? if so make sure you don't do that too long as that also causes cloning problems. finally you need really good competent cells as you may only get a few ligation colonies compared with your positive control.

Good luck!

Daniel, excellent suggestions.

Lena, you might also try blunt-end ligation by doing a T4 fill-in if Daniel is correct about Sap1. Also, how much total DNA is in your ligation reaction? I usually aim for 200 ng in 20 up of reaction volume.

I have a really simple question. Have you check your ligated solution by PCR to see if your ligation worked or not?

This may sounds odd, but what kind of dNTPs did you use for PCR?

Hi again.

Joshua - My inserts are 250-970bp in length, the plasmid is 6.6kbp.

Daniel - The NdeI site carrying plasmid has 10 nucleotides 5' of the restriction recognition site. The SapI primer is has a complementary sequence at the cleavage site. These things have been taken into account by the people who produced these primers before. And i have incorporated them into my primer design. However, you could be definitely right about the contamination in the gel extractions. I have looked at the DNA on nanodrop, and there is a lot of salt there - not surprising to see a high salt peak when the concentrations are below 50ng/ul, but still.

Can i ask you for a detail: The control ligation you are suggesting: Insert alone? Or did you mean the plasmid. Because i did do that, and did get a band shift - it looked like it worked.

Again - thank you

Lena sapI is not sequence specific restriction enzyme so end generated by this enzyme, in insert or vector may be different and due to that they are not compatible to each other and they can't ligate with each other. If single digested product is not ligating than please change your ligase enzyme buffer or add 1ul of 5mM ATP in 20ul reaction. Please take care when you thaw buffer of Ligase enzyme, ions present in that should be properly dissolved.

All the Best.

Everything was eluted in water. And yes, the competent cells could be an issue.

The T4 fill in is a great idea. I will have to look into it, it's been a while since i have done this. i had less DNA in my reaction, about 100ng. I didnt check my ligation by PCR, i was assuming the gel was enough, because similar inserts with the same overhangs worked so many times before.

The dNTPs were a part of the Phusion master mix, a polymerase that never gave me a headache.

Its hard to keep up here with the answers. i have heard some great suggestions. I will:

try and precipitate my DNA, see whether i can get it cleaner, and maybe more concentrated to have more DNA for the reaction

Look into the filler option

And get back to you when i know more

Kind regards,

L.

The simplest reason - the version from Eva Wenzel (Kohler). Iver ligase or ligation buffer is dead. I would pu an attention to the answer from Daniel Cohen, but SapI is not sensitive to overtiming and doesn't have a star activity. So, most likely, ligase/ligation buffer. Try a fresh one.

Hi!

I am experienced in cloning and noticed that gel-purified PCR products is very hard to clone.Sometimes there are no colonies at all. Better use PCR purification kits.

Another source for the problem may be in activity of proteins that are encoded by the fragments to clone. It is known that transcription factors, DNA repair proteins or DNA modification enzymes may be toxic to bacteria and cause their death. Check your inserts for the presence of such proteins.

well if the single plasmid cannot be religated after gel purification it is likely that the problem is due to either:

agarose quality not good enough (use best quality agarose like Top vision from Fermentas or other cloning certified agarose)

gel extraction not good enough (I use only quigen minElute )

t

DNA was degraded when you stained and imaged the gel, I switched a long time ago to sybr green or sybrsafe (molecular probe) which is not imaged on UV lamp (can be and bands visibility is the same on long and short UV contrary to ethidium bromide, which usually is harder to see with long UV) but on a blue lamp.

When I double digest the plasmid for cloning I usually just use column purification, if I want to check it, I check only an aliquot.

It is enough to run aliquots of preped fragments and plasmid and use simple rule to do the ligation. The amount of ends is what counts, so do not add excessive amount of fragments, it tends to create artifacts that prevent the ligation of the products you want together, in addition there is the danger that you create a construct with additional pieces, that will make you waste a lot of time.

Hi Lena, you already got a lot of nice suggestions above. Maybe just one more in case you did not pay attention to this: if you are purifying the insert, as well as the vector in agarose gels, you should really pay attention to illuminate the DNA as little as possible with UV light, as it can induce T-T dimers, which cannot be repared by the bacteria if there are too many of them. It can really be a killer in cloning. For this reason, I was always putting some aluminium paper under the zone of the gel where I expected the DNA to be isolated, so that only some UV light from the side could reach the DNA, and just had a quick look to locate the DNA band, then swhitched off the light during the time I cut it from the gel, and switched on the light after to check that all the DNA had been taken from the gel. Also, for the vector, as it is a long DNA fragment (and so higher probability to find a lote ot T-T dimers), I always prefered not to purifiy it on a gel: i.e. just let all the digested DNA in the tube, purify to remove restriction enzymes, and engage all + your insert in the ligation fragment. Just a little bit more clones to screen at the end to find the desired one, but at least you get clones to screen! Good luck!

Hello

Last year, I moved to another laboratory. before, I never had problems with ligations.

In this new lab, nothing seemed to work. I purchased all reagents used before, and tested everything, including water!!

at the end of 3 months, I spoke with a friend who told me that in the past had the same problem. The vector that I normally use is about 4kb, and wanted to ligated an insert of 1.5 kb. With smaller inserts, also did not work.

This friend advised me to use a different image system, with less intense UV. I searched through the faculty and I found a UV system, which has the option of low / high UV. I started doing DNA extraction from the gel with low UV.

Interestingly, it works. Strange!! But now I am happy again!!!!

Fernando

All these comments could be your solution..I would try the fill-in reaction to make both ends blunt. Of course you'll need the CIP the vector afterwards (to prevent closure). Also, since this will be a blunt-end reaction, make sure you don't have toom much ATP in the reaction, since "normal" levels used for a sticky-end ligation can inhibit blunt end ligations. I've found NEB to be hit-or-miss....I used EpiCentre's Ligation kit...works almost every time.

Hi Lena,

What size is your insert compared to the vector? - if the vector is smaller then treating the vector as the insert and insert as the vector for calculating the ligation ratios is a good way to get round this. If this isn't the case then I would tend to agree with Daniel regarding the enzyme choice - I've blunted the ends of SapI digested inserts and vectors before to get round the 'non compatible issue'. I've also found that problems like this can be due to primer sequence problems where the restriction site within the primer was found to be 'off' by a base or two (generated by a manufacturing error and not picked up during quality control - I've had this happen with Sigma, MWG and Eurofins) - as a result my insert was only digested at one end but the vector was fine and would accept other similarly digested products. Like Jennifer, I also never use TE to elute either inserts or vectors after digest/clean-up. Instead I would recomend ultra pure water or QIAGEN EB buffer - the EB buffer is better if you're like me and store hords of pre-cut plasmids in the freezer for frequent use. Finally, I always CIP or SAP treat vectors - I've had to many students waste weeks during cloning just cause they didn't do this step. So I'd try:

1) re-PCR insert and cut both insert and vector with SapI

2) Blunt end the insert and vector at this point

3) Cut both insert and vector with NdeI and CIP/SAP treat vector at end of the digest

4) Re-try the ligation

If that doesn't work, I'd be tempted to re-order the amplification primers (and if you can - avoid the SapI site) and ensure you have sufficient base pairs after the restriction sites to ensure optimal digest. I've now removed the SapI site from most of my vectors using mutagenesis and just replaced it with another unique site which cuts more specifically.

Hope some of this helps and if you need a protocol for blunt-ending inserts/vectors just drop me an email.

Hi Lena. I'm suggesting an insert along control to assess whether your PCR products work as a substrate for the ligase. Don't both transforming this ligation control- it is only for purposes of measuring success of the ligase. You should observe a ladder of ligation products. If, on the other hand, you see very little ligation, or only an insert dimer, then you know the insert is the problem. If only a dimer appears, then only one of the enzymes digested the insert. If little or no ligation products appear, then you have a chemical inhibitor (e.g. denaturant salt) present in your insert preparation that is killing the ligase. If this control works, then the problem is the transformation step. Either you need higher competency (>=10^8 CFU/ug) E. coli or you have excessive UV exposure problems, as suggested by Genevieve, Vincent, and Fernando.

Lena- sorry for the typo in the previous post, that should read "insert alone control"

Lena: I noticed couple things that may go wrong for you (small things but could make a difference). Did you purify the PCR products before digestion? If not, the polymerase in the PCR reaction may filled in the gap after digestion which resulted in unclonable fragments. I also don't understand why you do gel purification if you have nice clean PCR products. Gel purification tend to give me low recovery rate and give me headache in cloning.

Reamplify your PCR fragments with primers that contain overhangs homologous to the plasmid (20bp or so) and do Gibson isothermal cloning. You can put the reagents together yourself in home-made buffer according to the paper (Enzymatic assembly of DNA molecules up to several hundred kilobases.) or you can buy it from NEB now. That will eliminate all the issues you are currently encountering, even the SapI issue. This cloning is homology based, not restriction site based, so even if SapI chews a few nucleotides away, there are still nucleotides left for annealing. Although you will have to linearize the plasmid with a restriction enzyme of course. You can also engineer restriction enzymes between the PCR homology and cloning homology if you want, but that will increase the size of the primer. I generally do 20 bp cloning homology, one restriction enzyme site, and 20bp PCR homology, for both primers. Have done this hundreds of times. Never fails.

Hi, Lena

Sorry if I've missed such reply, but to check cleavage sites in your primers, the possible problem discussed by Emma, you can try to clone your PCR products in pGEMT or similar vectors (it's always very easy step) and than double-digest this plasmid. Insertion could be used on next steps. Sometimes such additional step saves a lot of time.

Regards,

Evgeny.

Dear Lena:

I think that your problem may be due to having active restriction enzymes in your ligation sample. Some enzymes do survive gel extraction and co-purify with your DNA. They will antagonyze DNA ligase and so you will not get any ligation done (this is easy to check, just run you ligations in a gel alongside a sample of your cut plasmid and insert).

Both SapI and NdeI can be heat-inactivated, so this problem is easy to solve. Once your DNAs are digested, perform the heat inactivation step, then do the gel extraction and proceed with ligation as usual. It should work.

In addition, please be aware that the cohesive ends created by SapI are not all equal and sometimes they are not compatible (as it cuts several nucleotides away from the recognition sequence) i.e., they cannot be ligated. I guess that is not the case for you because your plasmid with a single cut should ligate (or you have two problems instead of one).

Good luck and keep us posted!

Have you sequenced plasmid before cutting? Do you you know what exactly is in SapI cutting region? How close theseNdeI and SapI? Even if they can be cut at the same time, maybe sequential cut will do better?

Adding guanosine to your electrophoresis gel prevents DNA damage from UV light.

http://www.biotechniques.com/multimedia/archive/00009/96215rr02_9961a.pdf

Hi Lena,

I always put 5-6 bp extra at 5' end of primer before the the restriction site. Since the cleaved product is so small you won't be able to assay if it cleaved, just have faith. When I do cloning I do 50ul restriction digest with 4ug vector and insert plasmid for 4 hours and purify on agarose gel. I take 5ul of the 50ul and run in one lane then add the other 45ul to another lane with an empty lane in between. I then cut my gel and visualize the piece with the 5ul lane and do not expose the 45ul lane at all. I print a picture off of fuji (mine is printed actual size, do not know if all imaging systems do this) and then put printout on benchtop, clear page protector over the printout, and then match up the cut gel (easy to do if you look at the lanes) and then cut out desired section. I have not had any problems with this so far. I do not use CiAP on vector if the two cut sites are not compatible. Hope you find resolution in this matter. Good luck!

I am of the opinion that though the ligase was active, it did not carry out the ligation. Check, possibly the temperature was a crucial factor.

"Question: why did the single cleaved, gel-excised plasmid not close? A single restriction is usually the reason we get false positive clones anyway..."

Cut your plasmid with a different enzyme, purify by your std method. Ligate and transform. Still problems? Then it is your purification method, most likely over exposure to UV. If not, then it is enzyme or cut site specific issues as already mentioned.

When I run into a similar problem I also do a ligation of the double cut insert to make sure I see a trimer on the ethidium bromide stained gel that indicates both ends (and RE) are good, you can also do that with the double cut vector to get a vector trimer. Those two experiments are critical in trouble shooting the problem. It is not clear to me which one of your two positive controls worked, was it just the uncleaved plasmid that worked, or did the SapI cleaved plasmid also work? (if the SapI cleaved plasmid did not transform then either the SapI RE may be contaminated with a nuclease that chews on the sticky ends or your ligase may not be good-since no ligations would have worked). Again the control ligation of the insert and vector will help a lot. And make sure you are using molecular biology grade agarose that has been tested to make sure it does not contain nucleases.

And for your vector only control transformation; with even a double cut vector you should still see a few (10-20) colonies on the vector only transformation plate. I use the ratio of colonies on the vector plate only versus the vector + insert plate to give me an idea of how good the digestion and ligation went.

At this point, based on your results, I would get new REs/buffer and ligase/buffer/ATP and then repeat the digests/purifications and do the control ligation of the vector alone and insert alone.

I like NEB, but have gone to Fermentus enzymes (ThermoFisher) because of the universal buffer/fast digest system (one buffer for 174 RE, and typically 5 min. digestion time), it eliminates any issues with a double digest in non-optimal buffer for one or both RE and is fast. Fisher sells a Fermentus fast digest pack of 12 common RE for ~$220 (I do not work for or receive any benefits from Fisher/Fermentus).

Also, I found that purifying DNA from agarose gels using ethidium bromide results in damaged DNA that still clones/transforms "okay" but has mutations (even when using long wavelength UV to visualize). I now stain agarose gels for isolating DNA using methylene blue in water (0.0020-.02%, do not need to destain) and use white light to visualize, not as sensitive but will not damage your DNA like EtBr does (the results were verified by transformation-a lot more colonies/plate and by sequencing, with EtBr if I sequenced 4-5 clones of a 300 bp gene insert 1-2 would be good, with methylene blue all were good!).

Also, I like Sigma mini spin columns for purifying DNA from agarose gels, good yield, can use regular "molecular biology grade" agarose, and DNA is very clean (per instructions; use 65oC water to elute large plasmids/inserts >3kb).

Hi Lena,

I would suggest doing your cloning "in silico" using software like SnapGene (I believe there is a free version), just to make sure your ends are actually compatible and your cloning can work in principle. Also, make sure that in your vector your cut sites are not so close together that the enzymes have to compete for binding or there is not enough overhang after one enzyme cleaves for the other to bind and cleave. Good luck.

Hi Lena,

As a check to see if there is potentially some ligase inhibitors or exonucleases present in your ligation reaction, it would be useful to run your ligated products on an agarose gel. If there are ligase inhibitors present or if your ligase just isn't working, you should see no difference between your ligation rxn mix and your double digest mix. If there are exonucleases present your bands will likely disappear. If your ligation reaction is working properly you should see the emergence of extra bands corresponding to various ligation products. What you would really like to see is a band roughly corresponding to the plasmid and insert ligated together.

good luck

I am agree with all the above all answer, But my point of view for the solution make the vector with double digested. For make vector for ligation use each enzyme in different tube with appropriate amount of supercoiled plasmid DNA and incubate for 2 hours after that add alternate enzyme in two tube, incubated for 2 hours at 37C. Now pool the reaction mix in one tube and further incubation at 37C for 1 hour. Laod the double digested DNA on agarose and excise the band and gel purify. Use this vector for ligation reaction and kept for incubation at room temperature for overnight. For Insert preparation digest the PCR product with specific cloning enzyme for your purpose for at least 4 to 5 hours.

Good luck

The suggestions posted here are all good. I have experienced similar problems earlier and I believe the suggestion of Evgeny to use a TA cloning kit has worked great with my cloning experiments. Your PCR amplified products need not be gel purified and you can directly subclone them into a TA vector (pGEMT or any other TA vector). If you are using any proofreading polymerases for your PCR then you may have to add 3' A overhangs as the 3' to 5' exonuclease proofreading activity removes those overhangs. The overhangs can be easily added by following the manufacturers instruction. Once cloned in the TA vector, you can sequence to determine the orientation and then clone it into your pTXB1 vector. This will save you lot of time.

Best,

Stephen

Daniel Cohen covered most of what I would suggest, one thing I would add is that I found that you can increase your ligation efficiency by mixing the vector and insert and heat the mix to 50ºc for a few minutes then transfer to 37ºc then to room temp and then put them on ice this allow the ends to anneal more effectively once the vector-insert mix is on ice add the ligase buffer and enzyme and carry your ligation as usual.

Take a day break. Start fresh with everything new, all enzymes, buffers, water etc., This saves time than trouble shooting. Please add additional flanking sequence as its been suggested. I use CGGAATTC for NdeI (NEB, 1-2 hrs double cutting mostly with xhoI/SalI) and has mostly never failed, this sequence was from NEB catalog. Shuttle vectors are good idea too. Check the sequence comparability at ends. Good luck.

Hey Lena

Did you check the restriction products of these enzymes before designing your cloning strategy

This is the restriction site for Sap1

...GCTCTTCNNNNN...

...CGAGAAGNNNNN...

and the cut product is

....GCTCTTCN NNNN...

...CGAGAAGNNNN N...

Also I would have used both enzymes for positive control on single vector digest ligations, to check that both are working good.

I think review your whole cloning strategy, various materials that you are using (if have option use fresh ones, although i dont doubt components) and then check the easiest fragments (based upon size of insert).

All the Best

This may not be your case but FYI to everyone.

We just had a gel purification kit bring cloning efficiency to zero in our lab. When our always successful cloning stopped, it was a week before we realized the tech was using a Machrey Nagel kit (the best mfr. for mini and maxiprep kits in our hands) because we ran out of our Qiagen kit. Resumption of Qiagen returned the same failed ligations back to normal high efficiency insertion.

One other diagnostic - did you gel purify and extract your positive (uncleaved) plasmid control? If you were just using it fresh from a plasmid prep there may be a problem coming from the gel purification/extraction step that would give you problems with all of the cut plasmids. Some times you get ethanol carry-over from the gel extraction step that can interfere with subsequent ligations.

There are a variety of potential problems. As had already been noted SapI cuts away from the recognition site, so it really is not a good enzyme for cloning unless you are going to blunt the ends. Secondly, PCR fragments are not particularly easy to clone, although I am assuming that you have incorporated restriction sites into the primers so that you can cut them prior to cloning. Another potential problem is the fact that you are using a ColE1 based plasmid. These types of plasmids have problems maintaining certain types of DNA sequences. We never use any ColE1 plasmid for initial cloning work. The pSC101 based vectors have a much higher success rate (try one of the pWSK series of vectors). Once you have what you want, you can then transfer the sequence into whatever expression vector that you would like to use.

Sidney Kushner

These are good suggestions. However, I noticed that you said you used 20ul volume for transformation. I wonder what competent cells were you using? self-made or commercial ones? Because some commercial ones only give you cells in 50ul volume, if you load 20ul of ligation products in, it will significantly change the buffer concentration, which could be a cause for the failure. Just a thought.

In my experiences, for 50ul cells, you need limit to

Dear Lina, Simply clone the target in pGEMT vector and cut it with some other compatible restriction site. Then try to clone it in pTXB1. Possibly your restriction site are not properly generated after restriction digestion.

The suggestions proposed may help you to solve technical problems related with digestion & ligation & transformation steps. If it does not help, take into consideration that your vector provides intein-mediated expression of proteins. In addtition, if your target protein or its truncated derivatives are toxic for host cells, then even leaky expression from T7 promoter from pBR-based vectors could lead to T10 cell death after transformation. The solution is using another vector system, which provides very strong inhibition of gene expression and from single copy vector. To solve similar problem we had to use pACYC184 vector putting the gene of interest under tet promoter. A paper published in European journal of Biochemistry in 2000 is attached for you.

Bon courage

In your description it is not clear if you had the SapI restriction site and sticky-end containing overlap in one of your primers. If this is not the case it is most unlikely that you will be able to get ligation products ( only a 1 in 64 chance). Your sticky end ligating to the SapI site needs to be 5' ATG 3'. You probably thought of that anyway. The most worrying is that you get no ring closure with single cut vector. Was this with both restriction enzymes?

I was in a similar situation. Even after leaving enough bp in the primers to let the enzymes sit and cut properly and trying 2 different ligases I was not getting results. I solved it by moving the marker cassettes to a different plasmid that allowed me for gap-repair in S. cerevisiae. The new PCR products had enough sequence homology with the vector for proper recombination. It was easy and from yeast to E.coli competent cells just one step away. It would be great if you could do something similar, specially if you plan on cloning several different constructs.

Lena- several people have wisely pointed out that volumes matter during bacterial transformation. This is quite true. Try to keep the volume to no more than 2-3 microliters (20 would be much too high). Maybe you meant that you plated 20 uL of the transformation reaction? Depending on the efficiency of your cloning, and the competency of your cells, you may need to plate anywhere between 5%-100% of your competent cells mixture to get good results. Note that expected transformation efficiency with ligated DNA is at least 10-fold lower than with supercoiled DNA (relaxed DNA transforms less well, and only a fraction of your input vector with get correctly ligated).

Lena I think there may be problem in your double digestion. Make sure that you are using proper buffer for double digestion. Both NEB and Fermentas has their online tool which helps you to select appropriate buffer and conditions for double digestion.

https://www.neb.com/tools-and-resources/interactive-tools/double-digest-finder

http://www.thermoscientificbio.com/webtools/doubledigest/

I have pasted both links which may help you.

Secondly, donot over digest your PCR product by exposing it too long for restriction reaction.

Hope that will help you.

Dear Lena, I have only read some of the answers so someone else might have already said that, but here's my 2 cents on this:

1st, some people have concerns over the volume of ligation vs the volume of competent cells. I am not particularly concerned, as I usually use 10μl of ligation in 50 or 100μl of competent I made and they work great.

2nd, since your single-cut plasmid religation didn't work, it is obvious that you have a problem with the ligation step (well, unless you did something wrong in that reaction alone, that is, but we are troubleshooting assuming you did everything perfect). On the other hand, your ligase is working great, so this leaves the plasmid as the culprit.

So how could the plasmid be the guilty part? One, someone mentioned a problem with one of your enzymes. I am not familiar with them, but you should definitely check what's known about them. Two, I have seen time and again, that cut plasmids that have been gel-purified don't seem to work as well. TRY CUTTING THE PLASMID THEN DOING ONLY A DNA PRECIPITATION AND SEE IF THIS WORKS.

Now, a word of caution regarding the next step (after you solve your single-cut religation problem). How do you know that your inserts are actually cut? You said you saw them in a gel, so I am going to assume that the primers amplify well away from the restriction sites. But if this is not the case, do keep in mind that most enzymes require some distance from the end of the DNA to cut - it could be just one base, or many more. NEB catalogues have tables about these things so you can check what applies to your enzymes.

Good luck and tell us what works!

PS: Sorry for the CAPS but I realised my proposed solution was kinda buried in explanations.

Hi Lena,

as far as I remember SapI cuts far away from the recognition site, therefore you basically are not aware of the ends you get.

I know that I might sound funny, but in this case digesting the insert you might not have SapI cutting because its trying to cut "outside" the DNA.

What I would do is to redesign the primer carrying SapI site including a more convenient enzyme, I'm quite sure that if you look in the MCS of your plasmid you'll find out a more suitable restriction site to prerform a directional cloning.

Otherwise if you wish to keep your already cut insert and plasmid I'd blunt both of them, running dephosporilation just on the plasmid, it's obviously sub-efficient but it worked for me in the past.

Good luck!

IS DNA damage from UV light during the gel extractions a possibility?

Dear Lena

Have you gel-excised the fragment under UV light? this could be the reason of the poor ligation efficiency, there is massive DNA degradation under this conditions.

We usually stain the gel with methylene blue and never use EtBr and UV for band purification.

Good luck

Antonio Caruz

If you only truncating your insert at the N- or C-terminus you could do a PCR on the whole plasmid, then digest it with a single enzyme and ligate the digested PCR product, worked every time for me, e.g. your inserted your gene with BamHI/XhoI, one primer with with a XhoI overhang would sit on the gene and one primer with a complementary XhoI overhang which is on the plasmid starting at the stop codon.

If you want to delete part of your gene, you can use the same strategy, but with phosphorylated primers (bit more expensive but less expensive than wasting days) and directly ligate the PCR product.

I have given up using classical ligation techniques.

Look up a product called InfusionHD from Clontech. It is a in vitro homologous recombination system. All you need to do is extend your primers to include 15bp homology with the insert vector.

I have had great success using this system to clone quite large genes into a couple of different vectors.

I agree with most of the posts here. The main issue in my experience with classic ligations using PCR products is that the products are very difficult to cut and many require a few extra nucleotides to recognize the site. I typically add 10x more enzyme to cut PCR products than for plasmid DNA and this has worked well for me. Definitely check the SapI site - that might be an issue - and try transforming no more that 2.5ul of your ligation reactions. The pH of the reaction is inhibitory to transformation - if you want to transform more you can add 1ul of MES buffer (pH 6) per 10ul ligation reaction. This reduces the pH of the reaction and is therefore less inhibitory. When plating it is best to spin down the cells and plate them all since the efficiency of tranformation of ligated plasmids is much less than supercoiled plasmids.

Hope this helps - good luck!

I would also like to recommend In-Fusion cloning from Clonetech. Can be used with any vector and up to 8 inserts at the same time, overlapping by 15 base pairs, giving you exactly what you want without having to think about restriction sites. Has worked every time for me.

http://www.clontech.com/FR/Products/Cloning_and_Competent_Cells/Cloning_Kits/Cloning_Kits-HD-Liquid?sitex=10020:22372:US

•Avoid sub-cloning—clone any insert, into any location, within any vector

•High efficiency—over 95% efficiency with a broad range of fragment sizes from 0.5 kb to 15 kb

•Get only what you want—final constructs are seamless with no extra base pairs

•Be flexible—clone multiple DNA fragments simultaneously in a single reaction

•Perform site-directed mutagenesis

Lena stated that the cleavage sites on the primers are known to work on this plasmid, so that should eliminate any of the mentioned issues with the Sap I and Nde I restriction sites (which are all valid) if the cleavage sites on the primers include the Sap I recognition site and cleavage site. The simplest explanation is the ligase is not working, since she has no ligation reactions that worked in transforming E. coli, including a single cut plasmid.

If you see the insert laddering into trimers with new ligase but still do not get transformation, then putting the uncut plasmid DNA through the same steps as the cut plasmid (gel purification alone, and gel purification / ligation) should allow detection of a bad reagent that is inhibiting transformation.

I would like to thank everyone who took the time to read through my troubleshooting in detail. Because of the overwhelming number of answers, i cannot address everyone by name, but as far as the issues go:

Ligase is active, and the buffers are working and/or supplemented

The UV exposure was minimal

The molar radio i thought of, and tried different ones

The primer ends are fine, and are complementary after restriction to the plasmid

1.The gel purification is something to think about. I havent had much experience with just cutting, inactivating and then purifying and ligating, because i prefer my reaction pure, and keeping the clone screening to a minimum, but, as suggested, i will try this, because i really want to see those small dotts on my plates again!

2. The volume of the ligation on cells is something i havent thought about, but you are right, 20 to 50 is a lot.

The blun fill-up is something to consider, but not for the cloning today, i really need not extra bases for this one.

The homology recombination Koen and others have suggested is something i will most definitely be thinking about in the future, and will be doing, i like the idea a lot

And Emma, you are right, I consulted with someone who used the vector and SapI before, and they told me that the double digest is a big problem there, event hough with a single it looks like its working. I extended the digest time to 4h, and made them sequential. Im also working with another plasmid now (related), which has a bigger fragment in between the two sited, and i will hopefully be able to visualise the cleavage like that....

I think that was it. I think we went through all the Typical pitfalls for cloning within this discussion :). Hopefully next time i will tell you it worked!

Kind regards,

Lena

I have found that ligation of double-digested plasmids and PCR products is frequently problematic. A ligation-independent cloning method is far superior. The PIPE protocol (Klock, H. E., et al. (2008) Proteins, Struct. Func. Bioinfor. 71, 982-994) is an excellent choice for constructing truncation variants. Our protocol can be found at http://capsicum.colgate.edu/chwiki/tiki-index.php?page=Recombinant+DNA+Protocols#Polymerase_Incomplete_Primer_Extension_PIPE_Cloning_of_Deletion_Variants

Since the ligase is working it looks like your REs may be the problem. Cloning can be tricky, and issues with REs are a big part of it. In case you missed my earlier comment, Fermentus has 174 RE that use the same digestion buffer (including SapI and NdeI) and you can load the digestion reaction right on the gel (no need to add DNA loading buffer with dye) see: (http://www.thermoscientificbio.com/restriction-and-modifying-enzymes/restriction-enzymes/fastdigest/enzymes#S).

The Sigma DNA gel isolation columns have always worked fine for me (http://www.sigmaaldrich.com/catalog/product/sigma/na1111?lang=en®ion=US). Just remember to use methylene blue to stain the gel so UV is not used on the DNA, and make sure the water/TE that you elute the DNA off the column with is autoclaved, that will eliminate any nuclease contamination (DI water can be contaminated with nucleases, but not distilled). Keep us updated when you solve the problem.

And lastly, I had a problem once with cloning a DNA fragment, after trouble shooting I found the problem. The agarose gel DNA loading buffer was contaminated with a nuclease that was chewing on the ends of the DNA, the DNA fragment was a little fuzzy on the gel, but otherwise looked fine. Made fresh DNA loading dye and gel purified the same digested fragments (had some left over) and they cloned fine.

I suspect that your problem could be either

(1) Nde i digestion

Nde is a funny enzyme, it likes DTT which if the buffer is old needs supplementing. NdeI also requires a number of bases on the oligo if you are cutting a PCR product.

please see:

https://www.neb.com/~/media/NebUs/Files/Chart%20image/cleavage_olignucleotides_old.pdf

(2) DNA damage!

it does not take very long when you are cutting out fragments from an agarose gel for the DNA to be damaged by the UV light, this DNA will ligate fine but will not transform into bacteria very well.

The easy solution is to use "sunblock", run the gel in 1 x TBE buffer with 0.5-1mM Guanosine. The guanosine will protect your DNA!!!

OR cut out as fast as possible!!!

For NdeI also add extra fresh enzyme to the reaction halfway, it does not have such a long halflife time.

Use a TAE buffer instead of TBE .

I think your problem is the same as my problem with NdeI. I think your problem will be settle if you change this enzyme with something else. Sometime, this enzyme shows nibbling property which is really hard to understand on the gel electrophoresis. apparently, everything is ok, but you do not have any ligation, since this enzyme degrade sticky end of cut site. I had this problem with enzyme for 6 month and no thing resolve this problem, except using hindIII instead of NdeI. I f you are able to use the other enzyme do not hesitate to do it.

Regards

Mojtaba

Just an update: I have clones! I exchanged my overly-puristic approach with gel purification, for a simple quick and very dirty: cut with enzymes (increased cutting time, as well as sequential cutting) - inactivate said enzymes - purify - ligate. I have 4/10 clones with an insert at the right height. Thanks to everyone that contributed!

L.

Lena,

I agree with Daniel Cohen, because the restriction enzymes what you are working are very tricky. I do give one more strage suggestion is you please change all your reagents and start with fresh preparations then definetely it works. Why I am saying this is once I had the same kind of problem and you want to know the reason is my Low melting agarose what I was using was bad. Any way good luck

vathsala

Lena,

Great to hear, thanks for updating us.

Gary

Hi Lena, 

I am having almost the same problem and also performed most of the control experiments you have done in the very first place. You mentioned, "The digest also worked on the inserts, which I saw on a 4-20% TBE gel". How did you make sure the accuracy of two ends of the insert from gel? Also, I guess you inactivate the enzyme by heating after digestion and  after deactivation of enzyme you purified. How did you purify the DNA? Because very first thing you mentioned in the post that you exchanged your overly-puristic approach with gel purification for a simple quick and very dirty..Any suggestion will be appreciated. 

Hi Lena

We always purify PCR bands/ digested plasmids from methylene blue stained gels.

The intensity is not too high compared to etidum bromide/UV but the integrity of the DNA is preserved.

Best regards

Hi Dhadchi , 

Thanks for your up date and this is really helpful. I have a few question about your procedure. During sequential digestion did you use the reaction mixture for 2nd digestion directly after deactivation or you process it some way before 2nd digestion. Do you run a gel between to digestion? I guess you use heat deactivation i.e. 20 min at 65 C.  What type of cell you use for transformation? 

Also you have mentioned that you never had any issue with the cleavage of the 1kb insert. How did you clarify the complete digestion of the both end of inserts?

Thanks again for sharing your experience.

Thanks 

Arnab

Hi Dhadchi, 

Thanks for this detail information. I really appreciate it. Are you deactivating the enzyme just by incubating the reaction mixture at 65 C for 20 mins - 30 mins?

Thanks 

Arnab

hey,

is anyone has articles to explain why cloning fail??