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9

There is nothing like a general polymerization time, as this is a chemical reaction which depends on many factors. The most important are: APS: Concentration and and the age of the chemical are important. APS decomposes over time when it is not stored completely dry (which it is not on most benches) and also with freezing thawing cycles in storage. APS ...


9

The stacking gel concentrates proteins loaded into the sample wells so that they are resolved as a unified "line" once they enter the stacking gel. The reason for the lower pH is that this "lower ionic strength implies higher electrical resistance and consequently a higher electric field, provoking the faster movement of the proteins and of every other ...


6

The other major difference between the two is the amount of acrylamide in the upper (stacking) gel - it's generally around 4%, while the lower (resolving) gel can vary from 6 or 8% to 20%, depending on the size of the protein(s) you're looking for. When you load your samples in the wells at the top of the gel, then start the current, not all of the sample ...


6

Others have already explained the effect of different factors on polymerization time. This answer is mostly about: If I increase the time then would it affect the band pattern? It may. Some tips: Make a little extra mix so that you can leave some in the beaker to know if polymerization has happened. Always layer the poured resolving gel with water so ...


5

You are correct that molecule mobility depends on the mass-to-charge ratio, and this means that different sized molecules with the same $\frac{m}{q}$ will have the same acceleration. However, the velocity of a molecule moving through a gel matrix depends on the point at which the force exerted by the electric field is in equilibrium with the frictional ...


5

Put simply, the answer is that you could seek to detect viral proteins, but because these proteins would be very minor components of your sample, you would have to use an immunoblotting technique to detect a specific viral protein. While immunoblotting is quite a sensitive technique, I think its fair to say that it can be technically demanding. In contrast a ...


4

If your talking about Sodium Dodecyl Sulfate(a detergent) it has a two main functions: (1) It disrupts all non-covalents bonds in proteins i.e hydrogen bonds from the main chain, or hydrogen bonds between residues which are dependent on protein shape. (2) It coats the proteins with a negative charge which is useful for laboratory techniques such as gel ...


4

The polymerization time is strongly dependent on APS and TEMED concentrations. I do not think that the increased time for polymerization would somehow influence the division of the proteins in negative way. Actually the time varies from 20 min to 1 h, but I personally leave the gel to polymerize longer. Unpolymerized Acrylamide could build adducts with some ...


3

Do the percentage values refer to the percentage of acrylamide in the gel? Yes. The 8% gel is 8 g acrylamide per 100 mL. The “4-12%” gel is a gradient gel, which are useful for separating proteins over a large range of sizes. Read more about the uses and formulations of gradient gels here.


3

The problem here is concentration. Coomassie Blue stain can only detect protein band greater than or near to 50ng, in this case, the concentration of your protein is too low for detection. If you want to stick with Coomassie stain, you can try colloidal Coomasie stain instead because it has a much lower detection limit than Coomasie blue (between 10ng to ...


3

The reason why the sample buffer is more concentrated (typically 2x or 5x depending on your protein concentration) is its dilution when you mix it with the sample. You mix 4 parts of your sample and 1 part of 5x sample buffer, so that the final concentration of this buffer is 1x. If you use 1x sample buffer together with the same amount of sample, the final ...


3

The protocol for SDS-PAGE uses a solution of sodium dodecyl sulfate (SDS, also known as lauryl sulfate) to solubilize and linearize folded protein molecules and give them a negative charge that is approximately proportional to the length (and more or less to the mass) of the protein, depending on its sequence. Some sequences may bind the SDS more readily, ...


3

I'm going to treat this as a partial homework question but provide some guidance as to how you can potentially address your question and have solid theory to back it up. Chymotrypsin preferentially cleaves peptide amide bonds where the carboxyl side of the amide bond (the P1 position) is a large hydrophobic amino acid (tyrosine, tryptophan, and phenylalanine)...


3

APS and TEMED concentrations matter a lot. Even with that standardized, your polyacrylamide on the shelf is losing reactive groups as we speak. Who know how long it sat in the warehouse? It all adds up to the fact that the minimum required time is practically impossible to predict. On the other hand, once polymerized, a gel is stable for days without ...


2

An SDS-PAGE standard curve of Rf vs LogMW is linear because the standard you're given is engineered to behave that way. An expanded SDS-PAGE curve would show that the actual graph is sigmoidal: Structues that are too large can't enter the matrix, and too small they pass right through. Your standard has taken into account the % composing the gel, and the size ...


2

According to PDB (protein data bank — a repository for protein structures) the chromophore must be protected from interaction with water molecules to fluoresce. The chromophore is found right in the middle of the [protein], totally shielded from the surrounding environment. This shielding is essential for the fluorescence. The jostling water molecules ...


2

Usually you would want to keep the amount same, not the concentration. However, if you still want the concentration to be the same then you can add suitable amounts of PBS or your lysis buffer. For e.g. if you have two samples with 1mg/ml and 4mg/ml concentrations and you want to load 20μg of total protein, then you can take 20μl of first sample and add ...


1

You should de-stain your gel longer. I can see at least one thick band at the top, but without de-staining longer you are probably missing other fainter bands. Also, please explain what you mean by "proper bands". The top one looks fine to me. There might be others but we cannot see them.


1

If this is accepted or not depends on your institution. For publications, it is usually accepted to use adjustions of color, tonal values and contrast which are applied to the whole image and which do not alter the statement of the image. These changes are not ok if for example the contrast it raised so much that weak bands disappear. Not accepted are ...


1

As far as I know, it has an effect. If you denature your protein before running the gel, you do a normal SDS-PAGE and seperate the proteins by their size. Since you use SDS, the charge of the protein doesn't have any influence on your gel. If you don't heat your proteins before, you do a so called native PAGE. Here the size of your protein doesn't matter ...


1

It's a guess, as I haven't run a algae sample on SDS-PAGE, but your migration problems might be due the algal polysaccharides that are present in your sample. Check if your protein purification protocol is specific for algal proteins and see if there is any step dealing with the algal polysaccharides.


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