I am new to using the imager for my western blots. I am not satisfied with the images I get. I would like to know how to get good images using a imager.
1) Do you pour the substrate on the top of the blot and image it or do you put some substrate on a parafilm and invert the blot onto the parafilm let it sit for 1 min and then tap dry it and put it on the imager? (I do the latter when using films)
2) Do you cover the western blots with a saran wrap or something to prevent it from drying?
3) Any other tips to get a good exposure?
Thanks
it depends a bit on your imager (does it autoexpose or is exposure time set manually) and on your enzyme (HRP or AP), could you add these details?
I usually pour the chemi-substrate on the membrane and incubate it for a min. Then I remove the excess substrate by simply wiggling the membrane. Cover the membrane in saran wrap, use a kimwipe to smooth the srarn wrap over the membrane this will also remove any excess substrate on the membrane. I use BioRAd Chemidoc for imaging. Exposure time that works for me the best is 15 min. But, I could see signal even at lesser time. Hope this helps.
Thanks for your replies. I am using a GE Imagequant. It has provisions for both manual and auto exposure. I use HRP conjugated Ab.
The signal starts to develop rapidly and decreases fast with the HRP and I would normally just mix reagent and add on the blot, tilt the blot a bit to distribute the liquid evenly across the blot, and then measure immediately after on auto exposure. But you may also place the blot sandwiched in between pieces of saran wrap (or overhead plast). But signal drops rapidly over time so measure soon after application.
I am also using a GE imagequant. We tend to use cut pieces of polythene tubing (the bags that you can use for heat sealing, but I don't seal them for this purpose) for incubating the HRP reagent on the membrane. I pipette 1ml of substrate over the membrane, close the bag and just incubate for a min or two.
When imaging, I shake the excess substrate from the membrane and place it in a new bag. The membrane should stay wet though throughout imaging (use the bag or cling film to cover the membrane).
I tend to image using manual exposure with 10 second or 1 minute increments. Signal depends on antibody - for very strong antibodies (e.g. actin) 10-30 seconds is fine, for some weak ones I need to expose for ~30 mins. Make sure your blot is in focus before imaging.
Just a general comment, which may be of use; take care to match the intensity of your marker bands with the intensity of your bands, or make blots with only target bands and no markers, if you use "in-blot" size markers with HRP, and autoexpose, the system may adjust image aquisition parameters so only the marker bands are visible.
Thank you all for the comments, they are definitely helpful.
@ Brodin : Without the marker, how do you usually estimate the molecular weight of the recognized band ? I am using a custom antibody and trying to figure out if it actually works/how stringent it is. So, it is really important for me to see the size(s) of bands the antibody lights up. Any comments on this ?
@ Agrotis : When you expose the membrane for 30', do you see drying of the membrane ? Mine dries out even with 15 minutes exposure. But I do not use 1 ml of the substrate, I just use 200ul on a parafilm and put the membrane facing down over the parafilm, make sure no bubbles and then tap dry it using a towel and place it on the tray and cover it with saran wrap.
I pour substrate over the membrane and it does not dry even after 30 minute. I use ~750 mcl per membrane, 8.5 cm X 6.5 cm.
No, with the way I described the membrane won't get dry for at least several hours.
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