Hey labrats!
I have a question about stripping and re-probing a western I am planning to run. Normally, I would block my membrane for an hour at RT and incubate with primary at 4 deg overnight. Next day, wash, incubate with secondary for an hour at RT, visualize.
I need to probe my membrane for two different proteins of similar weights. Again, normally I would just strip after the first visualization and repeat the protocol, but this time I'm on a time crunch as my PI would like to have this done ASAP for a manuscript we are submitting.
Would it work to block 1 hr RT, incubate with primary 1 hr RT, wash, incubate with secondary 1 hr RT, then strip, and repeat with the second antibody, all in one day? I don't see why not, but does anyone have any advice for how to get this done without the overnight incubation steps?
I had a question for our research technician today that I have been pondering for a while. She is normally a fountain of knowledge for this stuff but on this occasion she didn't know, so I am turning to Reddit.
My understanding of blocking a Western Blot membrane (e.g. with 5% skim milk solution) before incubation with primary antibody is that it causes protein to bind to remaining bare patches of membrane so the antibody doesn't simply bind to everywhere on the bare membrane. So there's protein bound all over the membrane?
Why then, when you use a total protein stain such as amido black at the end, does the whole membrane not become stained as the blocking has caused protein to bind everywhere? Total protein stains still only stain the lanes of protein from the transfer and not from blocking on the rest of the membrane.
I'm obviously not understanding part of the mechanism behind blocking or have missed something. Could someone explain?
Does anybody know if Pierceβ’ ECL Western Blotting Substrate will work even if it was left in room temperature for one day instead of 4ΛC.
One of my favorite geek stories is the naming of the different blots. western, northern and Southern. Notice the capitalization on Southern, named after Edward Southern who invented the technique with DNA. Then the cheeky bastards who created RNA blotting thought they would be funny and called that northern blotting to be funny. So of course when blotting of protein gels happened they just played along and picked western over eastern. I still hope someone figures out how and why to do lipids or something so we can finally "call the corners".
Also, look into the gene/protein named sonic, and lambda phage S-holes.
I'm attempting to optimise some primary antibodies and I keep noticing a lot of dark, non-specific bands - despite the background of the blot being pretty clean. I noticed that this has been happening for any of my blots where I used an anti-mouse secondary, as compared to ones where I used an anti-rabbit secondary. Upon closer inspection I noticed that the anti-mouse secondary vial had something fuzzy-like floating around... Now I'm under the impression that whatever contamination is in that vial is causing all the non-specific bands (which are VERY intense and saturated even when my primary dilutions are at 1:7500 or 1:10000).
Is this possible? Has anyone ever experienced something similar? I guess I'm looking for confirmation that my theory is correct, because this started happening for the last few blots I've run and I have been running out of things to optimise (i.e. blocking time, blocker type, etc.)
BTW I block overnight at 4 degrees in 5% BSA and use fresh, refrigerated 1X TBST - which both contribute to making the blots pretty clean overall.
so far i get southern and northern blotting.. . buttt can someone plss explain these techniques and the objectives of them to me like i'm 5 years old?
Iβm trying to optimize some of the immunology work we do in my lab and I was wondering whether you all have had more success with using PBS or TBS as a wash buffer. I know if your target is phosphorylated itβs best to use TBS but is there any other caveats to using one buffer vs another?
Was hoping for a clarification on western blotting. Do proteins go through purification process like column or affinity chromatography, then go through gel electrophoresis, and lastly western blot or just go through gel electrophoresis and then western blotting?
Also I have in my notes that w/ eliza we wouldn't want to use a sds/reducing b/c we need to see enzyme activity? but I thought that the gel would be before we look at enzyme activity. Also if a protein is exposed to sds/reducing conditions how does the antibody even bind in the first place after the protein has been denatured?
Thanks!
What are the reasons to use Western Blotting instead of a mass-spec workflow to detect target proteins?
LC-MS has advantages of multiplexing, throughput, quantification, detection of PTMs and doesn't require antibodies specific to each protein being measured. So what are the downsides?
Hi everyone. I'm currently answering a questionnaire for a new job and for some reason am tripping over the verbiage. Does the term 2D gel refer to the type of SDS PAGE used for western blotting and coomassie staining? I wanted to double-check. Thanks!
I'm trying to optimize my protein extraction techniques as I've been getting mediocre yields lately from my primary and immortalized neurons. I've usually used a 15x ripa stock but something crashes out of solution really easily when it's in the fridge (and won't dissolve back). It's been ok for a while but recently has given quite poor yields. My protocol involves scraping cells/detaching with trypsin-le, collecting, spinning and resuspending in 1x ripa with protease inhibitors. I keep it on ice for 30 min with vortexing every 10, followed by a 20m spin at 15000 x g.
Ive now bought a new RIPA buffer (1x) and it says I should add ripa (with inhibitors) directly to the plate after washing wells with ice cold pbs.
Do you have any tips for this method? I understand it's fairly common but I've never tried it myself. How ripa much should I add to a well in a 6 well plate? Can I combine 3 wells? Anything I should be concerned about (ie dna causing the lysates to get goopy)?
Thanks
Hello All,
I am having an interesting issue with some samples I am working with. I am attempting to assess the amount of a specific protein in a tumor xenograft via western blot. I have developed this assay in vitro with the cell line which I am using in vivo with success (I can consistently assess protein concentration via Western Blot in the cell line).
In the in vivo model I have euthanized the mice, extracted the xenografts, homogenized them in a solution of cell lysis buffer with protease inhibitor and flash frozen the homogenized tumor samples immediately after extraction. Next, I isolated the soluble proteins via 0.22uM filter and assessed protein concentration via the BCA assay. I aliquoted these tumor samples and diluted them so that I am loading an identical amount of protein onto my gels with each experiment (attempted 14 ug and 28 ug).
My issue is this, even though according to my BCA assay data (run in triplicate) I am loading the same amount of protein onto my gels, I see a substantial difference in the signal of my protein (Btk) which does not correlate with the treatment conditions and my house-keeping protein (beta-actin). The even weirder thing is in some samples I see no Western Blot (Btk nor Beta-Actin) signal even though I can stain a gel from the same samples and see bands via coomassie blue.
I am confident in my western blotting conditions because I have acquired an immense amount of quantitative data at the cellular level with these experimental conditions, but when I move to blotting proteins from my xenografts, I am seeing large fluctuations in the signal. My current hypothesis is that I am getting variable proteolysis during the extraction process, but other than this I am stuck and would appreciate any input. I hope this was enough detail, but I am happy to extrapolate further.
