Because of excessive loading. Load less and you ll be fine.

Pre-stained markers tends to fluoresce with ECL+ substrate, but this is not a problem, since you can visualise them!

Syed A Ali, thank you for the answer! But can you please explain the possible mechanism of changes in antibodies binding and level of chemiluminescence when I load too much?

Claude Alban, thank you! I myself think that it's handy to see the marker bands on the photo, but isn't it unspecific signal? So I'm afraid that I do something wrong and there may be some false-positive bars in other lanes. Our marker isn't supposed to be stained. (A few days ago I've seen an advertisement of protein mass marker specially designed to be seen on the photo, and it's seen only because it contains fluorescent dyes).

Hi Ekaterina, are you using nitrocellulose membrane? It normally gives more background problems... Anyway, my impression is that your marker sample is sort of dirty or partially degraded, so possibly with debris from the marker proteins themselves, and you are creating a peptide smear in your marker lane. If your blocking is really as low as you say (0.5%, typically is 10 times higher... so maybe it's just a writing confusion), and you are using a low dilution of your secondary Ab (say 1:1000 to 1:2000) you might be unspecifically staining the lane except where your marker protein has covered the background, obtaining a negative fingerprint of the marker on the membrane... if this description is clear...!

In practice you could load less marker as Syed suggested to reduce background (or buy a new aliquot of marker and store it better: we store it at -20 °C until we start using it, then keep it at 4 °C, otherwise continuous freeze-thaw cycles ruin it), blot on PVDF that normally is less sensitive to background if you can, anyway use 5% milk for the blocking, make sure the pH of your TBS-Tween or PBS-Tween is above 8 to increase Ab affinity to the antigen, dilute 1:5000 to 1:20000 your secondary Abs, and you should get a clean blot.

Good luck!

Pietro Pilo Boyl, thanks for your detailed answer!

Right now our lab is in the process of updating of some instruments and expendable materials that are necessary for Western-blotting. Now we use both nitrocellulose and PVDF membranes (we had only nitrocellulose earlier), and unfortunately I'm not sure from which kind of membrane I obtained these pictures. 0.5% milk wasn't a misprint. I know that typically 5% milk is used, and I block with 5% milk if I do this by classic blocking protocol (just incubate membrane in milk for an hour or more). But recently we bought "SNAP i.d. Protein Detection System" instrument that saves our time: it uses vacuum pump to "impress" molecules into membrane surface and makes it possible to block membrane over 30 seconds and to stain it with one antibody over 10 minutes. And the "SNAP" user manual states explicitly that milk must be 0.5% to be suitable for this protocol. (It seems weird to me, but I do what instruction says). The secondary antibody dilution is 1:1000 (in the same 0.5% milk). And I've never heard about the fact that increasing pH of PBS-Tween increases antibody affinity, perhaps I should read more about this (we currently use tableted PBS, pH 7.4, and I believe that 0.1% tween doesn't significantly change its pH).

I suppose that you're right about the degradation of our marker: we store it at -20°C, freeze and thaw repeatedly, and at least once it was heated to 95°C to denaturate proteins. So you say (if I completely understand) that this smear of degraded marker proteins covers the membrane between the bars, thus preventing it from blocking, and it causes unspecific staining where antibodies bind to membrane? But in this case this area of membrane is covered anyway - not by milk proteins but by smear of degraded marker proteins - and isn't free for antibodies to bind. And if it's not antibody-membrane but antibody-protein interaction, why would my antibodies unspecifically bind only to smear but not to bars? I'm still very confused about this.

Hi Ekaterina, I see your points. I didn't pay attention at the SNAP i.d. Protein Detection System, which I didn't know, and actually your blots show it can work very well, at least with some Abs! I'll have to start thinking at implementing it in our lab...

Coming back to your WB, I would just keep in mind some general principles out of your experiment, which I think overall it's perfectly fine and reliable. First, nitrocellulose always gives higher background problems than PVDF, but it has other advantages, you don't need to activate it in Methanol, it can dry without problem also during a blotting, and preserves proteins on its surface much longer time. Anyway for quick usage of the blots, PVDF is probably better, mostly if you have to use 0.5% milk. Secondly, I know that not too many people know the trick, but pH 8 really increases binding efficiency of antibodies to their antigen (whenever we have a drop in pH in our buffer, often due to older dry milk powder which naturally oxidizes in the air increasing the lactic acid content, we have a marked loss of signal)... although as always in biology there might be exceptions. And you are right, a small amount of Tween or any detergent does not alter the pH of a buffer, provided the buffer is strong enough, but PBS is. PBS is fine for WB, but if you should get spotty blots, TBS is better (PBS is based on phosphate which in the presence of divalent ions precipitates and with some weak antibodies can give problems... just to keep in mind). You have learned everything about protein MW markers and how to store them! Finally, the source of non-specific, "background"  signals in your marker lane: this is always a complicated issue, but in general when you get a smear it means the peptides derived from proteins that were degraded are aggregating, that's why you cover such a long MW range (aggregates always range from very small to very large). When polypeptides aggregate, they easily trap or non-specifically interact with other proteins (think at the effect of the prion proteins in Prion Disease, or of the A-beta peptides and hyperphosphorylated Tau in Alzheimer's Disease, or of Huntingtin aggregates in HD, that all trap other vital factors in cells or the extracellular matrix), namely, in your case, your primary and/or secondary Abs, hence the signal on the whole lane, except where the marker proteins are intact. It is true that you are in SDS loading buffer, but aggregates are typically SDS resistant, normally even boiling resistant and often are actually produced by boiling, because it increases denaturation.

I hope you have a clearer picture. Cheers.

Thank you, these peptide aggregates was indeed the part that I didn't understand. I suppose I should just open a new aliquote of marker and keep it on +4.

(But even if the marker is degraded and has smears it doesn't affect the position of the main marker bars, does it?)

I don't think it does, to answer your question in the parenthesis... If you know the apparent MW of your protein from previous WBs you should be able to verify it (normally you do the reverse, of course! use the marker to be sure of your WB... but in this case your specific signal is very clean, so one should think it's indeed specific to your protein...). Anyway to be sure you can run it next to a new marker aliquot. You  might observe a small slowing down of the run...