Is there a faster method to screen for positive colonies besides picking single colonies, growing them and then sequence/pcr them for the insert?
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There have been 12 items by Wek (Search limited from 18-January 19)
Sorry to hijack this thread but I rather not make another thread about Infusion.
I am trying to use Infusion for the first time to do some cloning. I made primers for my insert with the Infusion online tool, did the PCR and verified it by running it on a gel. Is it possible to do the infusion step with the PCR product as is? I gel purified my insert but I ended up recovering <300ng total and there's a lot of salt in it (low 260/230 but good 260/280).
I linearized my vector tonight and deactivate the RE. Can I skip the gel purification step and use whatever amount of vector I need after I nanodrop it again? I am sure Ill lose a lot of the vector and will and up with a shitty 260/230 ratio.
Btw, anyone knows approximately how much DNA you get from a 30-34 cycles reaction?It should be a few micrograms right?
I would like to create a fibroblast cell line but I am unsure about some stuff. I want to keep things simple so I plan to obtain MEFs from mouse embryos and transduce them with SV40. Can I use pBABE-puro SV40 LT as my SV40 source? I dont want to buy an immortalization kit since they are crazy expensive.
My MEF isolation protocol uses MEM + supplements for culturing these cells. Could I use DMEM instead? I plan to package the virus with phoenix cells and they are grown in DMEM. I would like to grow both cell lines with the same medium but I dont know if the extra glucose would have a negative effect on the MEFs.
Once immortalized I plan to transduce these MEFs again.
I would appreciate if someone with experience can offer some advice and answer these questions.
IIRC you just need the LT antigen to cause immortalization, small t antigen probably helps as well, you could just express those from a plasmid and then select.
So any plasmid that expresses the LT antigen will be sufficient to immortalize cells?
Also I was told long ago that when purifying low nucleic acids concentrations you can get a low 260/230 ratio due to salts eluting and not much can be done about it which makes some sense. Does anyone what's the lowest concentration of nucleus acid I should purify when working with columns? I am trying to digest 1ug.
After doing a gel purification with a Qiagen kit my insert/plasmid has a very low 260/230 ratio (<1) but a great 260/280 ratio. I have been told to ignore it and continue with my cloning. Does this happen to anyone else? I know for a fact it's not ethanol and most likely the salts from the solutions.
I have been trying to transduce a few murine cell lines with a with a lentivirus for the past month and I just dont know where could be the problem. My lentivirus is IRES-GFP so I know that my transfection of 293T cells is good (GFP+ cells after 24 hours). Here's the protocol I follow:
Day 0: plate 2e5 293T on a 6 well plate.
Day 1: add 1ug or lentivirus, 1ug of each packaging plasmids along with Fugene HD with a 3:1 ratio (9ul of Fugene). I transfect my 293T in 1ml of media complete with antibiotics.
Day 2: (24hrs after) add 2ml of media for a total of 3ml.
Day 3 (48hrs after) collect the 3ml of old media with virus and add 2ml of fresh complete mdia. Put the old meida in 4C.
Day 4: Collect the 2ml of media and pool with old media, filter through a .45um filter and add to my cell lines.
I add 1ml of the media containing virus and 8ug/ml to my cells lines (5e4 cells/well on 24 well plate). Spin infect at 37C for 99mins. The next day I see no GFP.
Any help would be appreciated.
I am trying to design a set of primers with EcoRI ends for my gene of interest. I want to PCR the gene out of a plasmid and ligate it to 2 other different plamids with the EcoRI ends but I am a complete novice in molecular biology. Can anyone verify that what I have so far is right?
Fwd Primer: 5' TAAGCAGAATTCATGXXXXXXXXXXXXXXX 3'
Rvs Primer: 5' TGCTTAGAATTCTCAXXXXXXXXXXXXXXX 3'
I will do the PCR with a high fidelity enzyme, purify the pcr product and sequence it to make sure it is right.
One more question. I purchased a plasmid, grew it out in Ecoli, purified it and sequenced it. When I align it/blast it I get >90% identity but there are 2-3 Ns and 2 dashes within my sequence. Should I worry about the Ns and the dashes? How can I make sure the frame has not shifted and created a premature stop codon (the sequence is about 1kb and my gene is 1.4kb)?
I am having a weird problem with my minipreps. I sucessfully cloned my GOI into pQCIX-puro by infusion and T4 ligation. I picked ten colonies total from both preparations and culturered them overnight. Qiagen Miniprep went well and the yield and quality of the DNA was good. I sent out the DNA for seqeuncing and they all failed to prime with the 5' primer and had homopolymeric region repeats when using a primer that binds to the middle of my GOI.
I ran about 100-200ng of the plasmid on a gel (image 2006.jpg) and there are two bands: a thick band near the top of the wells that I think it's gDNA and a very faint band below it that could be my plasmid. Its hard to tell the size of it but it seems to be within the ladder range. My plasmid should be around 8.3kbp and I am using a 10kbp ladder. I also did a normal PCR with the the primer that binds to the middle of my GOI and the 3' primer that I used to PCR my GOI for cloning (image 1005.jpg). The product should be about 362bp, which is what I got.
Based on these results I think that the cloning went well but I somehow messed up the minipreps. I didnt do anything different from the protocol. The lysis step went slightly over 5mins (max ~30-60sec over) but I dont think this could have been the problem.
Does anyone have any suggetions or advice?
I am trying to ligate my insert of 1kbp into a vector of 4-5kbp but I am having some trouble. Here are the steps that I follow:
For the insert:
Using NotI I digest my insert out of another plasmid, run the digested product on a gel and I see two bands where expected. I cut the band with my insert, gel purified (Qiagen) and put in -20C until needed.
For the plasmid:
Using NotI I digested the plasmid, run the digested product on a gel, an see only one fairly clean band where expected. There are no smears only a tiny "tail" on each edge of the band (probably due to too much DNA?). I cut the band, gel purified (Qiagen), and put in -20C.
The first time I did this I did not dephosphorylate my vector and none of the colonies I screened had the insert.
This time I couldn't dephosphorylate after gel purification due to time constraints so I threw it into -20C hoping it would not re-ligate if frozen. I plan to treat my vector with CIP tomorrow morning. Would I encounter problems during ligation because I waited 12-18hrs to dephosphorylate my vector?
During my first try, both my insert and vector after gel purification had a 260/280 of 1.8-1.9 but the 260/230 was <1.0 (I think mostly likely due to salts and not ethanol). I was told to ignore the 260/230 ratio and go ahead with the ligation. Could these salts cause problem? I know I end up diluting the salt with H2O and ligation buffer but I have no experience with cloning and this is becoming frustrating.
Also, would it be worth using CIP/AP/SAP that has expired? I think the SAP I found expired a decade ago, AP expired 4 years ago and there's no expiration date for CIP.
I have been having a series of problems transducing some cell lines with a lentivirus over the past few months.
I finally managed to transduce 2 of 3 cell lines but with a very low efficiency (<1%) and sort GFP positive cells (90% purity using IRES-GFP as the reporter). The day after the sort my cells were still green, however, after moving them to a larger vessel the GFP disappeared. The cells are growing at 37C with no problem but they are not green. I put a plate of cells in 32C and one in 37C and compared the GFP expression level, if any. The cells at 32C are starting to express GFP although at a low level but there is nothing in the 37C plate. Has anyone encountered this problem before? If so, any advice?
Another question, while working on the lentivirus transduction I started another transfection/transduction experiment with a different gene/vector. This time I am working with a retrovirus (MIGR1). My actual retroviral plasmid has no florescent reporter so I am using an empty MIGR1 that contains GFP as my positive control. Transfection with 293T cells in a 10cm dish went well (10ug of plasmid, 10ug of pcleco and 6:1 Fugene in a final volume of 8mls). I collected the sup after 3 days, spun it, and filtered. I used 1.5ml of the viral sup, 0.5ml of fresh media, and 5ug/ml of polybrene to transduced my cells in a 6well plate. After 24hrs of transduction I was able to see green cells in my control but it was about 30-40% of GFP positive cells (need higher percent because these are tumor cells). Then I changed the old media and this is where I have a problem. For the transduced cells (including the positive control), all the cells came off in one big layer. My two negative controls (regular media and media+5ug of polybrene) are fine. I tried to add the media slowly but it didn't help. These are adherent cells so something went wrong somewhere and I am not sure where. Any help is appreciated.
Btw I did performed a spinfection.
I understand the science behind tetramer preparation but I am not sure how to calculate the specific volume of antibody to be used. I have read the NIH preparation method but they do not have calculations/formulas for their guidelines. Is there a good rule of thumb formula/ratio of antibody to monomer to use (like using 1ug of antibody to 4ug of monomers)?
EDIT: I see they say "The actual amount of streptavidin to be added depends both on the molecular weight of the monomer as well as the percent biotinylation. It is necessary to divide this amount by ten to arrive at the amount to be added per time interval."
How do you find this information? Are class I monomers produced with different MW, differing from batch to batch?