Hello Anchal,

What is the "mother vector" and the "host vector"? I can still understand the "host", but....

Normally, when you ligate something, like here in your case, your question says it is a 300 bp fragment into an approx. 5.4 Kb vector, you take the digested insert fragment and purify it (XbaI and SacI, as you mention) and ligate with the linearized digested, eluted and thus purified vector. You can do the following to further ensure ligation goes well:

1) Try to dephosphorylate the vector, when you dephosphorylate the vector, it will not re-circularize after ligation, so whatever clones you will get will only come from the ligation of the insert into the vector.

2) At the same time, you can set up, both the ligations, with dephosphorylated vector as well as normal digested vector. In case of digestion being complete, you should get colonies even with the normal vector. Set up ligations in the 1:3 and 1:1 ratio.

3) Run very small amount of completely double digested vector (say 30-50 ng), undigested vector, same amount and linearized with XbaI and another aliquot linearized with SacI on a 1% gel and run it longer, so you might see the difference between linearized and double digested vector, before venturing ahead with ligation. If the amount of DNA is more, you won't see it, so make sure, you have small amount, that is visible on the gel, but that does not form a thick band. This step will ensure that your Restriction enzymes are working fine and your digestion is complete. For the insert, you should do the same, purify it after digestion by gel elution.

6) When the digestions and everything will be fine, you should get a clone, in 10 colonies even and there won't be need to screen 100s of them. Since it is directional cloning and set up the ligation at 16-18 degrees for overnight.

7) When you transform the ligation mix, and then add 1 ml of LB to let the transformed plasmid grow and express Ampicillin for an hour, after this, when you plate it on LB Amp plates, then plate 100 microlitre, 200 microlitre, then whatever is remaining, in this case around 700 micolitre, you spin it down in the centrifuge at 8000 rpm for 1 minute and then you can see the cell pellet. Throw the supernatant, around 500 microlitre and resuspend the pellet gently in the remaining 200 microlitre and plate this amount as 'Rest' of the ligation mix.

In this way, you would have plated every ligated product you might have.

Hope it helps.

Best,

Simran

Thanks simran,  The mother plasmid is my vector. I did dephosphorylation and yet things did not work.. this is why i am worried 

What is then host plasmid? I thought it is the vector. 

I find this terminology strange. Anyways, the plasmid from where you are taking 300 bp insert is only an insert provider, so its size, as far as this ligation is concerned, does not matter, as after digestion, you only need the 300 bp fallout or insert.

The vector, where you are inserting this 300 bp fragment, its this VECTOR, whose size is of importance when you calculate vector:insert ratio.

1:9 molar ratio is also rare. Normally, 1:1, 1:3 or 1:5 is fine, depending on the size of insert and vector. Here your insert is small compared to the vector, that is around 5 kb, whichever "host or mother" it is. So, 1:3 should be fine. Or you can try 2 different ratios, to be sure, but for sure, 1:9, no, I have never ever taken this ratio.

i am sorry simran there was this typho error plzz see my question again ... neways u assumed it coorect.. n i would do my ligation in 2 ratios to see how it comes

No problem, but make sure that your vector is completely digested. Also, if you do not have much difference in your SacI and XbaI sites in the vector, then it will be a problem in getting the right clone, so make sure that your double digestion is complete. For insert, it is alright as it is going to be 300 bp and you can see that clearly on a gel after digestion from around a 5 kb backbone.

Best,

Simran

Hi Anchal,

I just wanted to say you have to digest both vectors completely to get rid of undigested plasmid. and run for a longer time on gel to separate them properly.  to check the efficiency of ligation do self ligation test for the digested plasmid which apparently gives very less or no colonies. if there are more or equal no. of colonies in self ligation plate in compare to test (vector + insert),the ligation efficiency is not good.

for example, self ligation has 10 colonies and test has 50- means good efficiency.

Dear Anchal,

like Simran, I have a bit of difficulties understanding your terminology. Anyway, I can give you the following tips for your experiments.

- ALWAYS purify EVERY fragment by carrying out agarose electrophoreses and gel extractions before any ligation experiment. It might seem like a waste of time, but you will always save a lot of it in the end. Agarose electrophoresis is not only a convenient DNA separation process, it is also a way to control that the restriction enzymes made their job properly, to control the size of your restriction fragments, ...etc.

- In your experiment, I do not really understand the purpose of dephosphorylating the vector in which you want to clone your 300 bp insert. XbaI and SacI are giving cohesive ends that are not compatible, so there is no need of dephosphorylating them. On the opposite, you loose this way one of main the advantages of making a directional cloning, which is, among others, higher yield. It might explain why you only get negative clones, especially if your are using an odd vector:insert ratio (like Simran, I have never heard of it). Removing the phosphate groups from your digested vector makes it harder for the ligase to attach compatible ends, so, if you do not need it, do not use it!

- I do not know which vector you are using to clone your insert, but check that the two restriction sites XbaI and SacI are not too close from each other in its sequence. Sometimes, carrying out a double digestion in such conditions is troublesome, and result in the proper digestion of only one of the two sites, leading to cloning failure which is difficult to troubleshoot. In this case, there is no other way than carrying sequential digestions with one enzyme after the other, preferably including a gel purification in between.

I hope this will help,

Pierre-Henri

Hi Anchal, some suggestions:

1,make sure that the vetor is completely digested.

2,optimize the molar ratio of the vector and fragment.

3,lengthen the ligation time appropriately.

4,check out the activity of your ligase and buffer.

Good luck!

hello thanks for your reply.. 

2 questions: 

1. how long can i lengthen my ligation and what temp.. is 16 degress ohk

2. how to chk activity of ligase and buffer

Hello, Anchal,

16℃ is ok, thats the temperature we used usually. You can try to keep the ligation system under 16℃ over 10 hours (overnight is ok).

Many reasons can lead to the inactivation of ligase and buffer, eg kept under improper conditions.  Sometimes we add 5mM ATP into the ligation system. And of course, you can use new ligase and buffer.

Hope useful.

Hi Anchal,

You can always perform a ligation at 4 deg C (i.e. in the fridge) overnight. That works as well for me as 16 deg C overnight.

You can check the activity of your ligase and the ligase buffer by setting up a control reaction with DNA ladder. Electrophorese both the ligase-treated and untreated ladder on an agarose gel and check that the ligase-treated ladder gives a single band corresponding to the size of the plasmid used to generate the ladder.

thankyou

Hi, Anchal

maybe the clones you got were plasmid only, before the selection restriction digestion i generally run my clones with the empty vector, i think the inserted 300bp can be distinguished by 0.8% agrose gel.

In my opinion, the ligation experiment is always failed in the double digest. so do the double digest again and verify both the two enzyme are work well. Good luck!

Hi Anchal,

I would suggest you to please check each step of your expt. first check vector digested properly by doing PCR using vector primers covering the cloning site as after digestion your vector will not amplify in the PCR reaction, while your control vector (without digestion) will amplify and give you band on agarose gel. second thing after cloning reaction and transformation you again use vector primers or your primers to check the presence of  insert in the cloning vector by colony PCR again. In this reaction you will get band size of cloned insert greater than your vector size band.third thing prepare medium with ampicillin as per protocol so that its conc. should not change.

Regards,

hi,

Use vector:insert in rario of 1:3

I think you are using too high concentration of Insert DNA.