Cloning <100 bp fragments into a vector
Posted 17 August 2010 - 06:29 PM
I have to clone an 85bp fragment and a 97bp fragment into a vector. I got these fragmens originally as single stranded oligos, so I annealed the complementary oligos and performed a double restriction digest so that the double-stranded fragments have the restriction sites that I need to perform the eventual ligation. Unfortunately, my ligation did not work. I used a 3:1 and 10:1 (insert:vector ratio) with vector+ligase (no insert) control and vector only (no ligase, no insert) control. When I transformed these into electrocompetent bugs, I only got colonies for the 3:1 reaction and nothing for the 10:1. Interestingly, colony morphology was very similar between the 3:1 reaction and vector+ligase (no insert) control. I understand that the fact that these two are similar could indicate that the vector is religating, but that seems unplausible as I had performed a double digest on the vector (using the same enzymes as I did for the oligos), run it on a gel and then cut/purify my fragment of interest. So unless there is some residual cut material, I fail to understand how the vector may religate with two different restriction sites on each end.
I have been at this for about a month now and have not been able to make any progress. If anybody can give me any suggestions/idea it will be much appreciated.
Posted 17 August 2010 - 08:47 PM
1- Each colony on a transformation plate represents a single plasmid molecule. Thus even slight contamination can allow you to see a few colonies coming through. Thus, if a transformation is actually successful, you should see hundreds of colonies per plate. If you only see a <10, it is likely to be contamination.
2-T4 DNA ligase, doesn't have all that good fidelity. Perfect base complimentarity is not required for T4 DNA ligase activity. So yes, it is possible, at low levels for vector molecules to religate.
Could we know more about the sequences of the oligos that you are ligating? Might there be a problem in the sequence structure?
Could you give us more details about the ligation. Did you dephos your vector? Where the oligos phophorylated? How was the vector digested? What where the conditions that you used to anneal the oligoes. What where the ligation conditions that you used? Are the ligase and ligase buffer still good? After ligation, it would be useful to run some of the ligation mix on a gel. If you see high molecular weight bands, it means the ligation is working.
Posted 17 August 2010 - 09:20 PM
Thank you for your reply. Please find below the information that you had asked for.
The sequence of my oligo is: "5'- ATGTGTCATGCATGCGGTACCTAGAACCCGCTGGTAGAACCCGCTGGTAGAACCCGCTGGTAGAACCCGCTGGTAGAACCCGCTGGTAGAACCCGCTGGTAGAAAAGCTTGGATCCATGTGCAAT-3'"
I also ordered the complementary strand: "5'- ATTGCACATGGATCCAAGCTTTTCTACCAGCGGGTTCTACCAGCGGGTTCTACCAGCGGGTTCTACCAGCGGGTTCTACCAGCGGGTTCTACCAGCGGGTTCTAGGTACCGCATGCATGACACAT-3'"
Once I had these I annealed them using the thermal cycler (heat till 95C, then slow cool -1C/min to bring it to 25C, followed by 4C hold), precipitated them, subjected them to a restriction digest with (KpnI + HindIII), precipitated them again and then moved on to the ligation step. I never phosphorylated my oligos nor am I aware that I need to do this.. could you please give me some more detail on this as to why it may be required.
The vector was prepped up, and digested with KpnI and HindIII. It was not dephosphorylated. It was then precipitated and run on a gel for extraction. I used QIAGEN extraction kit to for extraction of the vector.
Once I thought I have purified and digested versions of my oligos, I proceeded on to the ligation reaction. For this purpose I used promega's T4 DNA ligase (which has worked for me before). The reaction was run at 37C for 5-6 hrs.. After that I precipitated it and transformed into electrocompetent bugs. Next day, I did get colonies but as you said, it might be contamination or I think it is religated vector.. ne way there was no difference between my ligation reaction (3:1 ; insert:vector) with ligase + my control (vector only with ligase; no insert).
I hope this information will help.. please let me know if I can tell you more.
Thanks once again!!
Posted 18 August 2010 - 04:11 AM
You didn't mention how much of both vector and insert you were using. Likely you are using way too much of the insert, since it is short, and equimolar amounts weigh little.
Aim for 100 ng of vector and 5 ng of insert or so (guess) in a 10 ul ligation reaction.
Posted 18 August 2010 - 09:33 AM
1. Double-cut vector should be treated with SAP during the restriction digest. There will frequently be a small percentage of single-cut vector contaminating the double-cut prep. Simultaneous SAP treatment means you can clean up the reaction in a single step. If the number of ligated insert plus vector colonies is not > 2-fold higher than the number of vector only plus ligase colonies, you are unlikely to have any plasmids with inserts.
2. Rather than thinking about restriction digest reactions in terms of micrograms of material, it is more useful to consider the pmoles of DNA, which relates directly to the number of molecules. If 25 ug of pUC9 equals 8.8 e 12 molecules, then there are 8.8 e 12 unique restriction sites that must be cut. But 1 ug of a 100 bp PCR product contains 9.1 e 12 molecules and a corresponding 9.1 e 12 restriction sites. 80 Units of restriction enzyme will cut 25 ug of pUC9 or 1 ug of a 100 bp PCR fragment in KGB in 3.25 hours at 37°C.
3. Vectors cut with two enzymes inb the multiple cloning site can be purified by MWCO 30K cartridges. Run sample through and then wash with 300 uL of water before harvesting the clean vector. No gel purification needed, so you avoid the risk of UV damage during gel band harvesting.
4. A 5:1 ratio is sufficient if the vector and 100 bp insert are cut separately (allowing removal of the polylinker fragment via a 30K cartridge) and the concentrations are accurately determined. A 10:1 ratio is recommended if the polylinker is still present in the ligation reaction. If the insert concentration is in doubt, use both ratios or increase the ratio at the risk of double inserts. Higher ratios lead to double-inserts and should be avoided.
5. What you should actually be concerned about is having an equal number of molecules of insert and vector in the ligation reaction. If you combine 30 fmoles (1.8 e 10 molecules) of a 100 bp PCR product with 30 fmoles of pUC9, the 1.950 ng of insert and 52 ng of vector result in an optimal 1:1 ratio of molecules of insert and vector.
30 fmoles of 2686 bp pUC19 vector equals 52.4 ng and 1.8 e 10 molecules.
30 fmoles of 100 bp insert equals 1.95 ng and 1.8 e 10 molecules.
30 fmoles of 1000 bp insert equals 19.5 ng and 1.8 e 10 molecules.
30 fmoles of 3000 bp insert equals 58.5 ng and 1.8 e 10 molecules.
Posted 18 August 2010 - 05:50 PM
Posted 18 August 2010 - 07:11 PM
Thank you very much for your helpful comments.
Especially to tfitzwater. I have never actually thought about in terms of moles/no. of molecules. Always just thinking about 'ng' that I'm using.
So in my ligation reactions, I used 50ng of a 5.4kb vector and 2.36ng (3:1; insert to vector) or 7.86ng (10:1; insert:vector) in a 10ul reaction.
@tfitzwater: could you please elaborate on using the SAP with the digest. As I understand that SAP has its own buffer. So do you mean that when I'm digesting my vector in the same tube I should have SAP + its buffer => i mean template + Restriction enzyme + enzyme buffer + BSA + SAP + SAP buffer + water.
Alternatively I thought of this:
1) design primers that match the 5'end and 3'end of my oligo. These primers also contain overhangs that will be complementary to my restriction site of interest.
2) simply PCR up the double-stranded oligo, clean it up (via precipitation) and subsequently use it in a ligation reaction.
Do you think that will help?
Posted 19 August 2010 - 07:03 AM