no insert issue - (Feb/21/2014 )
I am trying to clone a 3.8Kb gene into a 5.7Kb vector ( pFBDM ), here is my protocols:
Vector preparation:
10ul of vector was cut with 1uL of XmaI and KpnI ( NEB ) in Cutsmart buffer for 3 hrs at 37degC ( 50uL total reaction volume )
Restrictions enzymes are fresh stock purchased about 1 month ago.
Gel purify the cut plasmid. ( UV was avoided by cutting the gel at the corresponding site )
plasmid preparation:
purified PCR product was cut with 1uL of same enzymes in same buffer for 3 hrs then remove the enzymes with Qiagen kit
no dephosphorylation
ligation:
50ng of cut vector was used in ligation reaction
vector/insert ratio: 1/2
ligation O/N at 16 degC ( NEB T4 ligase )
*****Note: I included a negative control
cut vector+ ligase but NO insert added
plate on selective plate and culture O/N at 37 degC
result:
negative control plate: 3 colonies yielded
experimental plate: around 50 colonies
I picked up 8 colonies and do overnight culture. I cut the purified plasmid with two enzymes but no insert found!
Need your opinions.
Thanks
Well first comment,
How many nanograms of vector DNA is present in that 10ul?
KpnI doesn't do all that well in buffer 4. It might actually be better to do a sequential digest. Start with KpnI in buffer 1, then increase the volume by 50% and add buffer 4 and do the digest with XmaI. There is also KpnI-HF that will work in NEB buffer 4.
Did you dephosphorylate the vector? And if so what were the conditions used?
How many bp did you add around the restriction sites on the primers? Did you add at least 6bp on the 5' end? Many enzymes do not cut well if their sites are not flanked on both ends by other bp.
What do you mean by 1:2 ratio? Is this a mass ratio? Or a number of molecule ratio? The ratio of insert to vector (in terms of molecules) should in my opinion be 1:1.
As I used electroporation followed by an ethanol precipitation, 50ng is to little DNA in my opinion. You mush have ug of digested DNA. Be a bit more generous and use at least 200ng of DNA. The DNA in the ligation mix shouldn't be too diluted else the molecules will have trouble finding their partner.
How old is your ligase buffer? Does it still smell of DTT?
How old is your ligase. Has any of your other labmates reported ligation problems?
You should run some of the ligated DNA on a gel to see if you can see high molecular weight DNA bands that indicate ligation has occurred.
What method did you use to transform your DNA into bacteria? Electroporation or chemical transformation? If electroporation, you should run your desalted ligated DNA on a gel to make sure you have not lost your DNA.
50 colonies per plate is very low. You should have hundreds.
Here are more details.
Around 1ug of DNA was used in digestion.
I use Kpn-I-HF for digestion. The activity of XmaI and Kpn-1-HF are 100% in CutSmart buffer according to NEB website. Extra 4 bases were added to the primer and again, I checked NEB
RE database, the efficiency should be good.
I don't think incomplete digestion is the problem since only a few colonies in negative control ( digested vector only, plus ligase, no insert ).
The vector was not dephosphorylated.
1:2 ratio is the molar ratio.
The ligase buffer is in small aliquots thus fresh anytime. The ligase was a new fresh stock just purchased around 1 month ago.
I transform the bacteria using competent cells prepared with Inoue method with competency around 10^8 CFU/ug plasmid.
Besides, I have no electroporation device in my lab.
I would suggest checking the primer sequences again just to make sure.
Could you show me the sequence of the primers that you use? I want to make sure that the RE sequence are in the correct orientation.
What is the composition of the ligation mix that you use? ie what was the volume of the ligation reaction?
It could be that the DNA concentration in the ligation mix is too diluted. Thus the vector and insert are having trouble finding their partners. I would use at least 200ng of vector in 20ul reaction volume.
Thanks a lot for your suggestion.
My primer sequence are:
5': atcgCCCGGGatgCATCACCAT.............. ( CCCGGG is the recognition site of XmaI, atcg are extra 4 bases for efficient cutting, others are for coding sequence )
5': atcgGGTACCatag............................... ( GGTACC is the recognition site of Kpn-1-HF, atcg are extra 4 bases for efficient cutting )
The ligation mixture volume was 20ul.
The strange part of the experiment is that negative control yielded few colonies suggesting that the double digestion is complete and self ligation is minimal, but experimental plate yielded dozens of colonies. I am curious about where those colonies came from? Is it possible that bacteria ( Dh5 alpha ) delete the insert from vector?
The reason I use 50ng of vector in ligation is that NEB suggests that the total concentration of DNA should not be too high. Higher concentration might inhibit the efficiency of ligation.
Thanks again.
Possible but I feel unlikely. Cloning strains like the one you are using have low levels of homologous recombination.
Usually when a gene is toxic to the cell, you just don't get any colonies. Does the gene you are inserting express a toxic gene product?
When you say the vector is empty is it really empty and nothing has gone into the plasmid?
If there is a small fragment within the plasmid, a possible explanation is that the PCR amplified gene contains additional RE sites of KpnI or XmaI. Check to make sure that this is not the cast.
The Qiagen clean up kits. Did you add the ethanol to the PE buffer. And if added, once the column is washed with 750ul PE. Did you then dry the column with a 2min spin at 14000rpm? Residue from the PE buffer can disrupt down stream manipulations.
Can you do a quick check to see if your enzymes are cutting?
And when you do the ligation, please run some of the ligation mix on a gel to make sure your DNA is ligating. You should see high molecular weight bands if ligation is working.
Do you use a nanodrop or similar to determine the DNA concentration after digestion and clean up for both vector and insert?
50ng of vector and about 200ng insert?
So run some checks to see if all the enzymes are actually working, especially the T4 ligase. This enzyme will denature with time. Do check.
And if the enzymes all check out, we go back to basic principles. When something goes wrong it is usually what you put in. And that is the PCR product. It could be the 4bp guards. I use 6bp as my standard. I don't use less than 5bp. You could try cutting longer, or simply use longer guards.
The vector is designed for expression in sf9 cells not in bacterial cells. The gene of interest is the mammalian helicase gene.
I checked the Genbank and is pretty sure that there's no RE sites in PCR product. This is further confirmed by running a small aliquot of RE digested PCR product on
agarose. Similarly no evidence of digestion was seen. The digested and undigested product were of same size.
I added ethanol into PE buffer and I also centrifuged to clean up the
The functionality of REs were confirmed with single enzyme cutting of vector.
Thanks again
Okay. Need to test that the T4 ligase is ligating.Run some of the ligated vector and insert on a gel and see if you are getting high molecular weight bands.
In theory, cloning and ligation should work every time according to manufacturers' instructions. However, we must realize that we are inserting foreign DNA into bacteria and this can be troublesome. Some inserts because of bacterial replication and transcription machinery are resistant and can be toxic to bacterial cells. In the past I've ran into this issue of resistance when trying to clone a particular insert. Because of this I would try using different competent cells which should help alleviate the toxicity issue.
Also, I would recommend performing a colony PCR on the colonies you picked to make sure the insert is amplifying. That way it would be easier and you would know for sure your primers are amplifying the correct target as well as you wouldn't have to digest and run a gel and hope the insert is there.
Thanks a lot.
I will run the PCR and let you know the result.