SDS page and affinity chromatography

[SJfox]

Hi! I'm an undergrad student doing a course in Protein Sciences.

I have a question.. so I've just done a lab.. and it involved purifying a protein in a affinity chromatography column then measuring absorbances of 5 of the first fractions collected. Then, the fractions were run on an SDS page. The fraction according to a standard curve made with the highest concentration of protein of interest in it was fraction 4.

However this had the SMALLEST band on the SDS Page, and Fractions 1-3 had much thicker bands... why could this be? Is this something to do with impurities perhaps in the first 1-3 fractions?!
ANy advice would be much appreciated!

[tgd]

Hello,

There are a couple of questions that need answering.

1 How are you measuring the protein concentration? Is this absorbance at
280 nm, or are you doing a specific protein assay such as the Lowry method?

2. What are you eluting with? Could this compound be absorbing at 280 nm, giving spurious high estimates of the protein content? Alternatively, could the eluting agent be interfering with the specific protein assay such as the Lowry method. As you probably know, a lot of substances interfere with the Lowry assay!

3. What volume are your fractions relative to column size? It is surprising that there is protein in fraction 1. I presume you have washed the column and that the protein of interest sticks to the affinity resin.

4. How are you visualizing the protein on the gel? Is this my Coomassie staining? Is Coomassie known to react with your protein?

5. Are any of the bands that you see the correct (subunit) molecular weight?

A fuller description would make things clearer. Can you say what is the affinity resin and what method you are using to elute?

Good luck

tgd

[SJfox]

Hi! Absorbance is being measured at 595nm. I’m eluting with an elution buffer containing imidazole. The column is Ni2+ agarose resin. The gel is being visualized with Instant Blue solution/stain. I do get strong bands the exact size of my specific protein of interest for all 5 fractions, however it is the thickness of these bands which concerns me! With fraction 4 having the thinnest band. There are some faint bands I can see also of differing sizes for this fraction on the gel.
Why should fraction 1 not contain any protein?

[tgd]

Hello,

I still do not fully understand the procedure, but that is probably just me!

1. I take it you have a crude protein sample containing a His-tagged protein which you apply to a Ni-agarose column (vol of resin?), wash the resin and then elute competitively with excess imidazole. You collect 5 fractions (what volume?). The highest protein concentration is in fraction 4 by protein assay but on the gel fractions 1-3 have the most protein, and the bands in these fractions are much thicker

2. In situtions like this, I recon, 'gels don't lie'. Most of your protein is in fractions 1-3. The fatter bands just indicate more protein, and the fact that the bands become gradually weaker in 4 & 5 indicates that elution is nearing completion. (You are getting a 'peak').

3. If the column were 10 ml and you were collecting, say, 2 ml fractions you would not expect any protein in fraction 1 as there will be a 'dead' volume. This may be roughly calculated as 30 percent of the bed volume. However, if the resin volume and the fraction volume are comparable (say 1 ml in each case) then you might not notice the dead volume effect. (I am presuming the protein STICKS to the resin and that there is no 'dribbling off' during the washing procedure).

4. Why are you measuring at 595 nm? Is this measurement made directly on the fractions (I suspect not), or after a protein assay has been performed which relies on colour development? If it is the latter, have you considered the possibility that the eluting agent (imidazole) might be interfering? The way to check this include a sample of the eluting buffer in your protein assay samples (If, say, you are using 0.2 ml of each fraction for protein assay, take 0.2 ml of the eluting buffer and assay that under otherwise identical conditions). You should also include a buffer control and a water control (but these have probably been done). [What is the principle of the protein assay? Is it the Bradford dye-binding assay, or something like that?]

5. If you are pretty sure that there is nothing in your fractions that will absorb at 280 nm other than protein, one crude way of estimating protein is to measure the absorbance at 280 nm of each fraction where (very roughly) an absorbance of 1 indicates a 1 mg/ml solution. (This method has the great advantage of being non-destructive - you can keep the fraction, and no harm is done).

6. With protein purification there are two great calculations that will be of interest to almost everyone.

(a) What is the yield? What is the total yield (how many milligrams did you get?) and what is the percentage yield (if the original sample contained 100 mg of your protein, and assuming you can calculate this, what percentage you you get back)? 50% or greater would be very nice.

You should be able to give an estimate to the total yield in milligrams (this is the most important one!) You can even do this by looking at the bands on the gels and estimating the protein content (say 10 ug per 'fat' band) and by 'working backwards' estimate the total amount of pure protein you have


So...... How much did you get in milligrams?

(The key question in protein purification. Molecular biologists have it too easy!)


(b) What is the fold purification? This is just the ratios of the specific activities in the pure sample and the crude sample. (You make not have enough information to calculate this)

Perhaps you should construct a purification table? This does not have to be complete (you may not have all the info available) but in my experience it always makes thinks much clearer.


Good luck

tgd.