10 replies to this topic

[GilsonGirl]

Dear Fellow Cloners,

Thanks for reading. I appreciate your help.
I understand there are some similar topics to this but each is slightly different so I thought I'd post my problem:::

I am trying to insert a 44bp insert into a 7.63KB commercial plasmid.

I am cutting ~ 2.0ug with XbaI and PmeI (Both NEB, in Buffer 4 @37C 1hr)
I then heat inactivate and run on 1% agarose gel, extracting the band with qiagen qiaquick column (horrible yields but much quick than phenol cholorform and no chance of loosing the pellet)

Both enzymes are working, I have run these seperately to check on a gel.

I am not dephosphorylating the plasmid. I get no colonies plating out bacteria transformed with cut plasmid so i think there is no re-ligation happening.

The oligos are IDT standard desalted custom DNA oligos. I am worried about the purification but I know it is working for others too. (I may eventually have to prep a fair few colonies and sequence to check)


I anneal the 2ug of each Oligo according to the protocol, in 50uls annealing buffer
I know that they are annealing (I have checked this on a gel)

My insert is going in according to the commercial protocol which is 50ng plasmid to 4ng insert.
I have tried ligating: (Plasmid: Insert)

50ng: 8ng
50ng: 80ng
80ng :160ng
80ng: 320ng

All to no avail.
The ligase works (I have tested it each time with singly cut plasmid I am getting about 100 colonies on single digests)
The cells are transforming very efficiently with control plasmids (3000 colonies with 0.1ng Puc1 DNA)
I am getting very low background (1 colony per plate) with no insert so the RE digests are fine.

Because I am doing miRNA target validation I will be trying to do this with a high number of inserts and continously for a fair few months (therefore keeping the costs down is a little important)

If you have any ideas then please let me know.

Thanks so much.

Edited by GilsonGirl, 06 March 2009 - 09:55 AM.

[Dr Teeth]

This is a very small insert. Therefore, 8 ng of insert is a very large number of molecules of insert compared to vector. I use a 3 molar ratio of insert to vector (based on the famed Sambrook et al. Molecular Cloning laboratory manual), which works very well.

Use the equation,

100 ng vector * X kb insert
_______________________ * 3 = N (ng insert to use for ligation)
y kb vector

For you, this means using 100 ng of your vector with only 1.73 ng of insert for a 7.63 kb vector. Therefore, using 8 and especially 320 ng of insert is way too much and you likely are getting insert/insert ligation favored over insert/vector, though I am surprised that you have not gotten clones of vector with multiple self-ligated inserts.

Edited by Dr Teeth, 06 March 2009 - 10:27 AM.

[stardust]

Hi there,

I do a similar thing. I anneal 1 µg per oligo in a 20 µl reaction and then take 100 ng of vector and 2 µl from the annealing reaction with really good results...I hope your oligodesign is right...coud you post them here for others to check? you bought them with the compatible overhangs for your enzymes right?

Stardust

[scolix]

How are you annealing the oligos? What buffer are you using?

[GilsonGirl]

Thanks for the suggestions.

I have double and triple checked the oligo sequences, and got others to do this for me also (including tech support). The oligos are ordered with the RE overhangs.

I anneal the oligos with by Heating at 90°C for 3 minutes, then transfer to a 37°C for 15 minutes. This is as per a commercial protocol and the buffer is "Oligo Annealing Buffer" whatever that may be.

Maybe I am using far too much insert as Dr Teeth suggested....

Stardust, do you mind me asking which vector you are using and who makes your oligos???

Many thanks

[GilsonGirl]

stardust, on Mar 6 2009, 09:54 PM, said:

Hi there,

I do a similar thing. I anneal 1 µg per oligo in a 20 µl reaction and then take 100 ng of vector and 2 µl from the annealing reaction with really good results...I hope your oligodesign is right...coud you post them here for others to check? you bought them with the compatible overhangs for your enzymes right?

Stardust

I am 99% sure the Oligo designs are right, I have triple checked them and had colleagues and tech support check too.

Can I ask which plasmid you are using, and where you buy the oligos from? Do you use any special purification for the Oligos? Do you have to do anything special to get them to ligate?

I guess your ration is 1:2 vector:insert then

[stardust]

Hi,

I'm cloning into a plasmid made by me, into pSuppressorNeo (Imgenex) and pcDNA6.2/GW-miRNA or something like that from invitrogen. My Oligos are from biomers (Germany)

As I said, I always use 100 ng vector and 2 µl of the 20 µl annealing reaction with 1 µg per oligo.

My annealing is different as well, 95°C for 5 min in pcr block, then I let it cool down to RT over about 3 hours in the pcr block and the I immediately use it for ligation.

My annealing buffer (10x) is 0.1 M Tris (pH 7.5), 1.0 M NaCl, 10 mM EDTA.

I have no idea unless you oligos are wrong since you seem to have check everything.

Stardust

[biotechnica]

Hi GilsonGirl,

No vector colonies means ur vector is digested properly and does not need more work. Try the following protocol:
Add the oligos in equimolar ratios - I would suggest about 10ul of each at 10-100pmoles/ul. Dilute with another 20ul MolBio Grade WATER. Heat denature this mix at 80 (or 90-94 if you feel that is better)*C for about 2 minutes followed by SLOW cooling to room temperature. Slow cooling is specially important if your oligos are GC rich. You can either do this in a thermal cycler (let the program bring it down to 4*C after annealing) or you could heat up a beaker full of water, boil for 2 mins and let the beaker (plus water and sample) get to RT over the next half hour (basically just leave it on ur bench after boiling). You can then ligate this according to your ligation protocol. An excess of oligo is better than less oligo to drive the reaction.

If electroporating, do not forget to Qiaclean the ligation mix.

Hope this helps. Let me know how it goes- good luck! I just finished two clonings like that.

[phage434]

Some things to check:
* Are you trashing the DNA with UV during gel extraction? Test by gel extracting single cut vector and then ligating to get good yield of religated vector.
* The oligos you have ordered have no 5' phosphate, so this will definitely not work with dephosphorylated vector, and would be more likely to work with kinased oligos. Consider treating your annealed fragment with kinase prior to ligation.
* As mentioned above, you have far too much of the insert. Instead of adding more insert in your tests, add far less. The lack of the 5' phosphate is preventing concatamers, which is good. The high concentrations may mean that different fragments ligate to each of the vector ends, which can then not be ligated together. Less is more. Aim for equimolar amounts of vector and annealed oligos.

[allynspear]

Are your oligos phosphorylated? I know that you technically don't need them to be since your vector still contains it's phosphates, but I have tried this both ways and I get a much higher cloning efficiency when my oligos are phosphorylated.

Best of Luck.

[Toyitta]

Dear GilsonGirl:

I don't know if you solve your problem or not, but if you still with troubles, last year I have a very similar problem with an insert of 125 pb... the ds oligos that I use was from IDT to. Check this, maybe help:
a) Because the plasmid is to big is better that the ds oligos are phosphorilated and the plasmid dephosphorilated. That works for me.
because I forgot ask the ds-oligos phosporilated I liogate this into another plasmid (pGEMT), very easy a quick and then digest from this and ligate into my vector.
c) this is another kind of solution, for insert to short like yours, her we do PCR reaction with the insert in the tail of the primer, maybe you need 2 different PCR reaction for get that insert complete, but in time is more quickly and easy.

Good luck