srpres

I have recently cloned a 2.0 kB insert which I thought may be a promoter into pGL3 and have discovered it has no activity compared to pGL3-Basic (actually, it has about 0.3 x compared to pGL3 basic, but this is understandible because pGL3-Basic has low background activity in some cell lines).

The way I identified the insert as a putative promoter region was by identifying the location of the transcriptional start site (TSS) through 5' RACE. Based on this result, I cloned in the 2 Kb piece, 1.5 kB upstream of the TSS and 0.5 kB downstream.

I know fo four reasons why this could be a negative result:

Possibility 1: something wrong with plasmid identity, integrity or concentration used in the transfections to test for activity.
Possibility 2: something wrong with the sequence.
Possibility 3: ATG start sites that are downstream of the transcription start site but upstream of the ATG start site for luciferase may interefere with the reading of the ATG for luciferase.
Possibility 4: the 2.0 kB insert is missing crucial regions, either A.) part of the promoter, B.) missing an enhancer.

I have ruled out possibility 1 and 2 by doing restriction digests, agarose gels, and sequencing to suggest that the plasmid I transfeceted is correct. Notably, there is a great Kozak sequence at the ATG for luciferase.

For #3, there are 3 ATG sites between the ATG of luciferase and the TSS, but they have poor Kozak sequences.

For #4. I did clone the 2.0 kB putative promoter insert into the pGL3-enhancer vector, which is the same as pGL3 basic but includes the SV40 enhancer. This did not improve activity.

Questions:
1. Is there another possibility I am not thinking about?

2. Could a strong splice donor site between the TSS and the ATG for luciferase interfere?

3.Any other ideas on how I can identify the problem rapidly?

Thanks for any consideration.

Lets say I wanted to clone a region of DNA (a 2.0 kB putative promoter region) from genomic DNA into a reporter vector, like pGL3 basic, by PCR-mediated amplification.

Should I use a high fidelity polymerase for the PCR or just regular Taq?

If I use a high fidelity polymerase, which one should I use?

Hello: I recently purchased a bottle of acid phenol/Chloform/IAA from Ambion, and on standing it separates into two phases--most of the volume is in the lower (denser) phase.

I wanted to use this for an RNA extraction. Do I:

1.) Swirl the bottle well before I use to mix the phases well, and then take out what I need, or

2.) Let the bottle stand, and take out solely from the lower phase?

Thanks for any consideration.

Hello: I am trying to clone a 2.5 kB insert with different sticky ends (Bam H1, Spe1) into a large vector (pGL3 with other fragments after the Luc sequence), the vector is 10 kB, so the total size is 12.5 kB The ligation product is being electroporated into DH10B cells, and then plated on Amp resistance plates (Amp = 100 ug/mL).

1.) I dont have blue white screening with this insert and the pGL3 plasmid. Instead, what I do is pick a colony, create a 1 mL LB overnight, test 2 uL of this with 13 uL qPCR mix with primers specific for the insert (amplicon is 300 bo). I have to check a lot of colonies (like 50 or more). From many colonies/growths I get a positive, but weak PCR amplifcation. I will miniprep this growth and see a contaminating plasmid. Is there a better way?

2.) I get few colonies as I expected, but the colonies seem all seem to be derived by a contaminating plasmid (this appears to be 2 or 3 kB) in size. Has anybody else have this problem? How can I solve it?

3.) Im guessing that my transformation effiency for this large construct (12.5 kB)  is low. Is there anyway that I can increase this?

Thanks for any consideration.