Curcumin rocks! There’s so much potential in that lil guy. Did you know it naturally fluoresces? Rock on. 

90% of my lab life right now is Westerns haha. I have the smell of B-ME stuck in my nose. Anyway, it depends what protocol you’re using, such as what membrane you use (nitrocellulose vs pvdf) and what kind of imaging (fluorescence vs luminescence) since they use different reagents and different instruments. Also depends what problems you’re having (bands not showing up? too much background?)

But in general…

• Make sure you have enough protein. I think 20 - 30 ug/lane works the best for me. 
• Don’t let the gel run so long your protein runs off the bottom. Keep an eye on the pre-stained ladder. Since my proteins are fairly large (>40 kDa) I stop the gel once the loading dye in all the lanes reaches the bottom of the gel. 
  
• Slice off the top right corner of both your gel and membrane so you can easily orient them correctly during the transfer. 
• And don’t let the transfer buffer get too hot! I put an ice-pack in there with a stir-bar, and some people put the entire thing on ice, or even throw it into a 4 degree room. Also I think 250 mAmps for 1 hr might be the maximum it can run at or it’ll overheat. Check with your protocol though. 
• Do all the washes in TBS-T stringently. As in, for the full 5 minutes, and for 5 full washes (or per your protocol)(but 5 seems to be the standard). Not one 30 min wash. No cheatin in the wild wild west yo.
• You may need to optimize the dilution of your antibodies. Having too much or too little can ablate your signal. See what the manufacturer recommends, and then maybe you’ll need to do a curve of different concentrations for each antibody. Like for me, my anti-TGFBRI is at 1:500 but anti-B-actin is 1:2000. 
• Be careful that your antibodies don’t cross-react with stuff. Not so much a big deal in Westerns because we know what we’re looking for, but sometimes it might result in bright non-specific signals that’ll “wash” out your protein of interest. (Here’s a good (and depressing) Nature article regarding the faulty reproducibility of antibodies for your spare reading). 
• Also check the expiration dates of the antibodies. If they’re from way back when it might be time to buy new ones. 
• I dilute my primary antibody in TBS-T + 5% BSA, but my secondaries in TBS-T + 5% milk. Probably took some optimizing to figure that out. Maybe you need to fiddle around with the diluent? 
• If your bands are faint, ramp up the exposure time to 30 min or an hour, and for the machine to take a picture every 5 - 10 minutes. If your ladder doesn’t even show up, time to do some troubleshooting. Maybe you oriented the transfer slots the wrong way? (definitely happened to me before ooops) Bad secondary antibody?

I use those beige BioRad spaceship-gel-imaging-thingies to image my gels because I use chemiluminescence. I’ve also used a fancy smancy fluorescence machine that almost looks like a scanner. Those are nice because your primary and b-actin lights up in different colors, so you can probe them at the same time. I’ve also used the old-fashioned film exposure thing in the dark room, but that was years ago, and hopefully will be a time I won’t ever have to revisit. 

So for analyses on my chemiluminescence images I just use ImageJ (free from NIH). If you’d like the protocol for analyzing bands using ImageJ I can totes email it to you (or screenshot it and post it here). 

I hope that kinda helped! Not sure how much you already know since I don’t know what specific probs you’re having, but feel free to hit me up with them! My inbox is definitely always open if you have any questions or just want to chat :) ResearchGate also has a great community for QA’s and help on troubleshooting. 

Thank you for the follow (on both blogs wow!)! You’re too kind :D Best of luck on your research and I hope to see you on some Nature papers in the future yeah? :D

Thanks so much for all the advice! I actually use fluorescence so I borrow another lab group’s fancy-schmancy scanner. I’ve been having problems mainly with the background being too high or basically there being too much background on one side of my membrane as opposed to the other sides. We recently switched from using pvdf membranes to nitrocellulose and started using a LI-COR’s blocking buffer which actually helped reduce most of the background. 

In terms of using 20-30ug of protein, what protein quantification assay do you use? I’ve been using a BCA box from Thermo Scientific. 

Also, I have been doing my washes in PBS and then PBS-T once I incubate it with antibodies. Is there a big difference between PBS-T and TBS-T? 

And I would really appreciate a protocol for analyzing the bands! (I’ll message you with my school email :) )

Again, you are really great help and your advice is so amazing and thorough! Good luck with your research! I’ll definitely be aiming for Nature papers and I’m sure I’ll see you on many Nature papers too!

Hmm usually high background just means insufficient washing and blocking, or poor handling of the membrane (always use tweezers and never fingers). This article talks about all the other causes of high background (both uniform and blotchy). Worth a read and some pondering. 

For protein quant I use this Bio-Rad kit to measure absorbance and compare it against BSA standards. It then takes some analysis on GraphPad Prism to determine the ug/ml concentration. I’ve also tried using a nanodrop but that didn’t work out too well :( But those nanodrops do have the option of measuring protein conc. I just didn’t spend too much time trying to optimize it.

PBS is usually for cell work, and may actually interfere with the immuno-reactions between proteins and antibodies (especially phosphorylated ones). TBS is recommended for gel work. I would try switching over to TBS for all your washes and see if it makes a difference. (Also TBS is cheaper than PBS so yay!)

And for sure! I just got your message with your email address so I shall be sending that to you shortly :) 

Haha thanks! The dream… 

Anyway, keep me updated on your Western adventures! I’d love to hear how they go, and we can totes keep bouncing around troubleshooting ideas :D