subcloning dengue NS1 gene problems - molecular biology, cloning (Nov/09/2006 )

First, thank you to read my long letter.. I am a student who do the molecular cloning research for my thesis and have no experience before.I try to subclone dengue NS1 gene (1050bp) into pET-21d(+) (5440bp) for 6 month, but still unsuccessfull
This is what I’ve done:
▪ I PCR amplify the insert with AccuPrimeTM PFX Supermix (Invitrogen), the primer added with BamHI and XhoI sites—Vector have the same RE sites too. I run PCR product in agarose gel and purify it with Wizard® SV Gel and PCR Clean-Up System (Promega). Result: single band
▪ I digest separately vector and insert with BamHI and XhoI (double digest), both RE from Fermentas. Incubation time: 3 hr in 37° C.The compositition:
Buffer Tango 2X 4 ul
ddH20 5 ul
DNA 10 ul
BamHI (10 u/ul) 0.5 ul
XhoI (10 u/ul) 0.5 ul
Total 20 ul
▪ I inactivate RE in 80° C for 20 min (as in manual)
▪ I dephosphorylate vector with CIAP (Promega) for 1 hr in 37° C, the composition:
CIAP buffer 1 ul
DNA vector 8 ul
CIAP 1 ul
▪ I purify vector and insert.
▪ I visualize DNA by running in agarose gel electrophoresis (double digest seems OK since there’s no smear, single band). Then, I count DNA concentration with BioID program(DNA compare with lambda DNA/HindIII marker). Insert concentration: 68.95 ng/ul, vector: 49.884 ng/ul.
▪ I do ligation (16 ° C, for ON) in 1:3,1:4 and 1:5 molar ratio.Total volume in reaction : 20 ul, with 3 ul T4 DNA ligase (1 u/ul, Invitrogen) and 1 ul buffer.
▪ I transform ligation product to E.coli DH5a competent cell. I use Hanahan method, efficiency about 106-107. Then plate in SOB (amp or without)
▪ The result:
(+) control transformation (with plasmid and without): OK, many
(-) control transformation: clean
vector with CIAP, without insert & ligase added (negative control) : clean
vector without CIAP, without insert & ligase added (positive control) : about 7 colonies
ligation colony: 3-10 colonies.
▪ I verify the small colony with double digest (BamHI and XhoI) with the same condition with first digestion and visualize it in agarose gel electrophoresis, but there’s no recombinant colony (no insert, only vector).
I dont know where the mistake is. Should I digest vector and insert one by one (single digest) to make (more) sure—even in agarose gel seems perfectly digested? Or I have to try other molar ratio in ligation?
Could anyone (immediately) help me please...? It’s really confusing...
Thank you,
rithzphoenix

There could be many possible problems.

* inadequate additional bases on the 5' end of your PCR primers following the restriction site

* Incomplete removal of PCR enzymes and dNTPs from the PCR process, yielding fill in or exonuclease activity on your PCR product during/after digestion

* Damage to the ends of your vector by CIAP

* Is the ligation buffer really 20x rather than 10x? And that sounds like too much ligase.

* You can get better efficiencies for your transformations with this protocol:
http://openwetware.org/wiki/TOP10_chemically_competent_cells or electroporation


You might have better luck cloning into a TOPO-TA vector following PCR, then cutting the insert from that vector and inserting it into your final vector. This avoids the problems of cutting near the 5' end of inserts, and difficulties of cleaning up following PCR.

You can test the effectiveness of ligations and the quality of your ends by ligating enough DNA to visualize on a gel and then running it. It is important to heat kill the ligase before doing this, else the ligase binds to DNA and makes it run very slowly. If you cut your PCR product with a single enzyme and ligate, you should see a double length band (do this with both enzymes). Cutting your vector with single or double enzymes and ligating
should give you long fragments. With single cut vector, you will see some recircularization, but with double cut, the shortest fragments should be double circular.

Pardon me; I don’t understand how CIAP could damage the end of vector? So, it doesn’t ‘safe’ to use? To prevent religation of vector..how much CIAP (safe condition) should I add?
The ligation buffer is 10X..Ligase is too much? hehehe...I just think that many ligase...many vector-insert ligated... So, is 1 ul ligase enough in 20 ul total volume reaction? do you have any special tips for ligation protocol?
Thanx before

CIAP is an unfriendly enzyme. Try shrimp alkaline phosphatase or antarctic phosphatase (NEB) instead. These also have the advantage that they can be heat killed. In general, I try to avoid all of the phosphatases. You should not need to phosphatase treat your vector with a double cut ligation as you describe.

Your protocol talks of a 20 ul ligation volume, and says that you add 1 ul of ligation buffer. If it is a 10x buffer (normal) then this should be 2 ul of buffer. Note that ligation buffers have ATP in them, and that they are freeze/thaw and temperature sensitive, like an enzyme, and need much more careful attention than, for example, restriction digest buffers. 3 ul of ligase is dramatically more than you need for this reaction -- you need only fractional microliters. More is not better. Ligation is quick -- mostly over in 10 minutes at room temperature for sticky end ligations. Blunt ligations are much more difficult, but I think you are not talking of these.

How much ligation mix are you adding to your competent cells? A common mistake is to think that more is better. Use 1-2 ul ligation into 50 ul of competent cells, 5 ul as an absolute maximum. Are you letting the cells + ligase sit on ice for 30 minutes? Heat shocking at 42C for 45 seconds? Immediately adding SOC and incubating for 1 hr (amp, kan resistance) or 2 hours (tet, chloramphenicol resistance)?

You should run a no-insert ligation control to check the cutting efficiency of your vector, and verify that your vector+insert ligation produces dramatically more colonies.

There’s no shrimp alkaline phosphatase or antarctic phosphatase at my lab . I choose to use CIAP because one of my RE (XhoI) only have 50-100% digestion efficiency in Tango 2X buffer. So, how about I decrease CIAP concentration (only 0,5ul maybe)?

Before I incubate my ligation mix in 16C for 16 hr, I incubate it in room temperature for 30 min. Do the incubation time is too much? I do that step as my teacher advise.

I add all (20ul)ligation mix in 200ul competent cell, incubate on ice for 30 minutes (flicking the tube sometimes), do heat shock (42C for 90 seconds), add SOC 800 ul & incubate for 1 hr, and then I plate it... Is it correct??

I will try to subclone again this week....cheers up!!!! (please pray for my luck... )
Thank you so much...

problems that I see

Amount of CIAP and the lenght of time the dephosphorylation step was conducted was far too much and far too long for the amount of vector you have used.

My calculation say that for the time you should dephos give the amount of CIAP (1ul = 10 U) and the amount of vector (8ul = 400ng) gives a time of 7.2 secs.

This is not to say my calculations are dead on, but you get the feel of the situation. Dephosphorylate more DNA, and use fewer units of CIAP. As already mentioned by phage434, CIAP is a bad enzyme, using it is like using a flamethrower to light a cigarette. Can be done, but always at the risk of burning off your face/vector.

The rule of the thumb for CIAP dephos is 1U CIAP (not 1ul) = 1pico mol DNA = 60mins at 37 Celcius in a volume of 50ul.


3ul T4 ligase for a reaction volume of 20ul is far far to much
1- Because like any enzyme, T4 ligase comes in a keeper buffer, one loaded with glycerol. Glycerol is great for keeping stuff (cells/enzymes) but it also severely inhibits enzyme activity.

2- Next , T4 ligase is not a nice enzyme either. Not as bad as CIAP but bad... it also catalyses "weird" ligase reactions. More is not better.

For a 20ul ligation reaction I would use 0.4ul T4 ligase. (And ligase it overnight at 16Celcius)