Fluffy

No-no: I said: put both on agar plate and the overnight preculture, but when you make your big culture, put only ampicilin: aka the 1-6 L of LB which you induce with IPTG should contain only one antibiotic because 2 of them will be too stressful.

Andreea

In addition: also the preculture should contain both antibiotics. However for the main expression I would use only ampicillin because 2 antibiotics at once during expression make to much stress on the cells.

Andreea

If your cells have a plasmid with a Chloramphenicol resistance marker in it, you'll want to put your transformations on plates with both Ampicillin (I definitely prefer Carbenicillin, as I get fewer satellite colonies) and Chloramphenicol.  The RIPL plasmid in your BL21 cells is there to help with expressing proteins with codons that are rare in E. coli, so if you're expressing a eukaryotic protein, you'll definitely want to select for cells that keep that plasmid too.

We usually use 30 µg/ml Chloramphenicol and 100 µg/ml Carbenicillin for LB plates.

HI Fluffy,
by using less I meant the total amount of DNA. You always have to adjust your protocol to your DNA concentrations.
Helpful therefore is: use vector concentrations of 20-50 ng; a molar ratio of insert: vector = 4:1; final buffer concentrations has to be 1x, use 0.5 - 1 µl ligase.
The final volume of you ligation mix does not have to be 10 µl. You can vary it in a way you need it. You may also read the recommendations of the company you got the ligase from.

So this time it would be better to use less vector DNA because you will need more insert DNA.
Try this:  2 µl vector (~ 40 ng)
  11 µl insert (~ 16.5-22 ng)
  1.5 µl 10xbuffer (= 1x)
  0.5 µl ligase
If you still have only 5x ligase buffer, then: 4 µl of 5x buffer, 2.5 µl of vector, 13 µl insert and 0.5 µl ligase.

You also can vary the volume for your transformation. If you want to make sure you get some colonies, do two or three transformation at the same time using, let's say 1 µl, 2µl and 3µl of DNA. When I had difficulties with my cloning (usually when I had to use some old plasmids) I used to transform up to 10 µl (for 200 µl competent cells). When I remember right, the volume of your competent cells was 50 µl. So do not add more than 5 µl (which would be 10 % of the total volume).

Hi,

so it can work with such low amounts,I did it once with ~ 1 ng/µl. Usually I work with amounts of 4 - 15 ng/µl and it always works fine. Just do the same protocol as before, maybe with less vector concentration (20-30 is thoroughly enough)...and using therefore more in the transformation mix.

But in parallel I would repeat the digestion using more template and set up two or three reactions at the same time if you have enough template...otherwise prep new vector. Then you would have enough for ligation.

In the words of Trof, anything measured by nanodrop under 10 ng/uL is a number randomly generated. I would say it is too little to trust what you have there.

I'd recommend you read the NdeI FAQ.  Note what it says about DNA contaminants, and also the comment on long digestiosn.  It is a good practice to get into the habit of reading these FAQs when using a new enzyme.  Or to use a protocol that never requires you to use new enzymes.
http://www.neb.com/n...tR0111.asp#2089

Hi Fluffy,

negative control means you are adding water instead of template to the reaction. If a PCR band occurs you know that there is a condamination. And, often those bands are not that bright as the real positive ones...

So, if you are doing a EcoRI digestion with some other clones, wait for the result and then chose the clone that has the right size, either one of the Eco-digestion or No. two of the picture you have posted here.

I agree with Papaver.  Try cutting with another enzyme, one that cuts your vector backbone.  The bands look uncut.  Are you expecting the NdeI and XhoI sites to be present in the original vector, or are they on your insert?  It might be time for sequencing if you can't figure out what is happening.  Dead enzyme would also explain this, so check that you can cut something else (or show that the DNA can be cut with a different enzyme).

So, I would say there is no insert.

Which clone(s) have you checked by digestion? Based on your PCR-Gel, I would have chosen samples 6 and 12.
One question considering your PCR...have you run a negative control? All bands besides lane 6 and 12 seem to me like they could be false positive.

Have you tried another enzyme just for checking your constructs? Are you sure that neither NdeI or XhoI cut in your insert?

You should be adding water to your restriction digest.   I'd recommend this reaction setup:

5 ul of a 10x buffer (use the right one, probably buffer 4 if you use NEB enzymes)
0.5 ul 100x BSA
1 ul each enzyme
1 ug of DNA (less than 15 ul total volume)
sufficient water to bring the volume to 50 ul (more than 27 ul)

This does several things:
1) correct final buffer concentration (1x)
2) dilutes impurities in your DNA prep that can cause problems (primarily ethanol and Gu-HCl)
3) reduces the concentration of glycerol in the reaction (REs are in 50% glycerol, and you need to keep the final concentration below 5%)

Your reaction will be mostly done in 30 minutes of incubation at 37, and if you choose your enzymes, you can heat kill it at 80 for 20 minutes and use it directly in a ligation.  It's easiest to do this in a PCR cycler.

It is a mistake to try to maintain a high concentration of DNA in these reactions.  You don't need it for ligation, and trying to do it leads to many problems.  Load 20 ul of the cut product on a gel,  use 2 ul in a ligation, and keep the rest in the freezer if you want.

Why are you adding only 3ul of 10x buffer?     According to your recipe, the final concentration of your restriction buffer is 0.4x  which is a 60% lower than it should be, so yes there is a problem with your reactions, as your buffering conditions will not be the optimal for the enzymes.

You need to dilute your 10x buffer to a final 1x buffer concentration, not just add 3µl per reaction (I don't understand where the 3ul come from).  If you need to add that much volume of DNA, you can prepare the reactions in a final volume of 100ul, add 10ul of buffer, keep your DNA and enzyme volumes as they are, and then top up with water.

Just my 2 cents.

on the other hand, if there's an actual reason to have the buffer at 0.4x concentration just ignore my message

I think that you have to digest more template to make sure you cann see your 600 bp fragment in the gel. Also the staining is not good. You can barely see the ladder, especially the small sizes.
I don't know the expected sizes and the shown one, but what could also have happened is that your digestion was not complete. It seems a little bit that only one enzyme has cut (which I cannot exactly say because of missing sizes).Also showing the undigested vector might help.
Just give it another try. You can check with another enzymes (e.g. EcoRI) at the same time to make sure your construct is ok. It once happened to me that the fragement was double ligated into the vector...

I agree, your insert is in the direction amplified by your forward primer.  The weird bands are trash products resulting from long extention times and low annealing temperatures priming to who knows what.

Hi Fluffy

since you have cloned your fragment into the vector using two different enzymes, the direction of the insert is given by the restriction sites. I suppose you have added the Nde site to the 5 prime of your insert and the Xho I site at the 3 prime end. Anything else would not make sense to me and that's why I would check a few plasmids isolated from your clones by restriction digest phage 434 has already mentioned. Besides this I would also sequence your construct to make sure your insert (or protein) is in frame.