Your sample appears to be aggregating suggesting that there are free thiols (proteinyl cysteines) that have not yet oxidized to disulfides, if they are suppose to oxidize.  This is not uncommon.  Your main bands are all getting less intense with accompanying appearance or increase in multiple higher molecular weight bands like a ladder of masses in multiples of 12 kD.  One should always run samples reduced versus non-reduced when analyzing by SDS-PAGE.  Try to use reducing Laemmli buffer or mild reduction with  alkylation with NEM or IAA before loading the samples into the gel wells.  You could also use NuPage gels with reducing agent in the cathode chamber.  It may be possible to reverse this by adding reducing agents such as TCEP followed by an appropriate formulation buffer, maybe acidic pH to protonate the cysteines and slow down the oxidation rate.    

The fuzzines is also a classic misfolding indicator.  Other than the mass/charge ratio, proteins are actually sieving through the gel pores according to their Stokes radius.  When a protein misfolds due to incorrect cysteine pairings to form disulfides then a vast number of different conformations form, each with a respective Stoke's radius.  What make a band have sharp edges is the fact that all the molecules essentially have  one Stoke's radius causing them to try to fit into a single point in the gel.  Because they can't all fit then they end up crowding themselves into a small area still trying to fit into one point.  What do you think happens to the band's edges if the molecules have an assortment of different stoke's radius?  

Your sample appears to be aggregating suggesting that there are free thiols (proteinyl cysteines) that have not yet oxidized to disulfides, if they are suppose to oxidize.  This is not uncommon.  Your main bands are all getting less intense with accompanying appearance or increase in multiple higher molecular weight bands like a ladder of masses in multiples of 12 kD.  One should always run samples reduced versus non-reduced when analyzing by SDS-PAGE.  Try to use reducing Laemmli buffer or mild reduction with  alkylation with NEM or IAA before loading the samples into the gel wells.  You could also use NuPage gels with reducing agent in the cathode chamber.  It may be possible to reverse this by adding reducing agents such as TCEP followed by an appropriate formulation buffer, maybe acidic pH to protonate the cysteines and slow down the oxidation rate.    

The fuzzines is also a classic misfolding indicator.  Other than the mass/charge ratio, proteins are actually sieving through the gel pores according to their Stokes radius.  When a protein misfolds due to incorrect cysteine pairings to form disulfides then a vast number of different conformations form, each with a respective Stoke's radius.  What make a band have sharp edges is the fact that all the molecules essentially have  one Stoke's radius causing them to try to fit into a single point in the gel.  Because they can't all fit then they end up crowding themselves into a small area still trying to fit into one point.  What do you think happens to the band's edges if the molecules have an assortment of different stoke's radius?  

In fact, I think your immunoblots may be working too well. If you look at the bottom of the lower blot you will see a ‘clear’ band all across the blot. This usually happens when a signal is so strong it uses up all the ECL reagent before the membrane is even exposed to film. One way to determine this quickly is to look to see if this band is a faint yellow color on the membrane itself. This corresponds to the deposition of ‘spent’ reagent.

The rest of the image just looks over exposed.

How much time passes between the time you remove the ECL reagent and place the membrane on film?

I would suggest you try loading various, lower, amounts of sample in order to optimize the conditions for this target. You could also probably use higher dilutions of your primary antibody.

Thanks Bruce for your suggestion. I'm using the exact same amount of protein. I can't see the yellow color on the membrane. I usually incubate the membrane with ECL for 3-4 minutes before capturing the image. I might send you one more picture showing just one band as crisp one and others as not clear. 

The second blot also has these clear bands. I still think you should consider  a quick experiment using a dilution curve of your sample. This curve should cover at least 2 orders of magnitude (i.e., up to a 100-fold reduction in protein loading as compared to the blots you have shown here)

Dear Veni,

In addition to the sensible comments made above, the appearance of fuzzy bands could also be the result of a loss of recognition of your target protein by your primary antibody. Have you used the same antibody for the 2 blots ? Was it a fresh dilution or did you re-use the diluted antibody from the first blot ? Has your antibody be stored properly and aliquoted if needed ?

Best,

Damien

I concur with above comments. If you have been using precisely the same protocol, the differences in the two blots may reflect any changes in the protein samples. What is the time differences in the running of the electrophoresis? Were the samples stored at 4 degrees or the samples were frozen and thawed after freezing. These subtle changes may contribute the differences between the two blots.   

To me it looks like free POI might have too much substrate etc...leading to white appearance due to overexposure. This will in turn enhance the rest of the blot, i.e., darken the other bands.

 Dear Veni,

I too feel that this has something to do with your primary antibody dilution, incubation times, etc conditions. Try using a fresh aliquot and perform blocking for not more than 1 hour.

Regards,

Shalini