I like Michael's answer. Many enzymes suffer denaturation upon dilution. Often glycerol or PEG can be used to substitute for protein to simulate protein-protein interaction. Sometimes BSA is used. From what Thomas has told us, I suspect his enzyme is a bit fragile. Its activity depends on integrity of disulfide bonds and perhaps other stabilizers.

Some years ago I did some experiments with Triose Phosphate Isomerase which demonstrated that addition of glycerol increased activity several fold over the standard conditions used for activity assay.

Reason 1 is good. Reason 2 is confusing. How much BME do you use? I guess at least in mM, while the enzyme is likely in uM or lower. BME can not be depleted in a stoichiometric manner. There might be some other reasons, such as a loss (precipitation or inactivation due to your concentration methods) of an inhibitor. The best way is to further purify your enzyme and see.

I guess I am totally confused. If you concentrated your partially purified protein 13-50 times then the enzyme activity should have increased by a similar margin. The fact that it only increased 2-3 fold is the surprising part. Am I missing something?

Dear Tomas,

In general increasing the total amount of units during a purification step is due to a remove of a inhibitor, whichever it is. So, my question is: Could be possible that during concentration some small inhibitor is filtered away?

As you say increase of concentration could favor increase in multimerization. So, the question is did you use the same enzyme concentration in the enzymatic assays? If there is a multimerization effect the activity is not linear with protein concentration in the assay. In this case the increase in total units is simple due a difference in the amount of protein used in the reaction.

The effect of bME sounds a little complicated. Usually the concentration of bME is enough high to keep all the free Cys reduced. However if you add more it will break up disulfide bonds. Which concentration of bME did you use?

Do you try with DTT? DTT is usually more stable than bME.

Gastón

so we are talking specific activity? Units/total mg?

I forgot to say that there is a easy way to check if an inhibitor is removed during concentration. Just take an aliquot from the filtered solution (usually discarded during concentration) and add to an enzymatic assay. Compare against an assay without filtered solution.

Duane,

I understand that we are talking about total units (number of units in all the volume). In this particular case and assuming that no loss of protein during concentration the specific activity (Unit/mg protein) should be also increase.

I'm with Duane. It sounds more like you are losing activity.

Do you actually know that your enzyme is a multimer? This first explanation seems quite plausible. It is a very general observation that enzyme stability goes down with dilution, and although nobody ever bothers to find out why, it seems very liekely that beyond a certain point one starts to get significant formation of monomers - if you are working with a tetramer or a hexamer the equilbrium will be hugely concentration dependent. Usually over the concentration ranges we work with the physiological oligomer predominates, but maybe not always - i.e. the dissociation equilbrium could be more prominent with your enzyme or with the buffers you have chosen to work with (e.g. Tris will stabilise hydrophobic interaction far less than phosphate). The second hypothesis seems much less clear and convincing, but anyway why not replave bME with DTT - less likely to lead to complications (and not so smelly!).

Gastón,

assuming no loss of protein and no loss of enzyme the specific activity would not increase but remain constant. The enzyme is a protein so the units of enzyme would increase in direct porportion to the total protein.

What Paul says is correct, very dilute protein solutions have a tendency to denature spontaneously. Therefore we need to have some sort of stabilizers when the real thing is at a very low concentration.

Other than indifferent proteins, even carbohydrates or lipids can stalilize proteins. Purified proteins are usually less stable than their native cousins.

Yes, I assume that you are talking of specific activity of the protein. As you purify the stuff, irrevalent proteins are removed and your specific activity increases. When it reaches a maximum (it cannot be purified further, i.e., you cannot increase the specific activity any more) you stop because you have reached your goal.

Total acitivity, on the other hand, usually decreases because some native protein is lost during purification (we are not 100% able to recover the protein). Therefore we often use the total activity as a measure of recovery of the protein (assuming that the specific activity does not change).

You case is therefore strange.

If you concentrate a protein 10-30 times, most likely you have also lost 50-80% of the original stuff. They have just gone down the drain. But you see an increase in total acitivty that must be due to #1 point you have suggested. The concentration of ME is too high to explain the increase the total activity.

Thomas, you will have to measure the specific activity of your enzyme preps. Until then it is difficult to tell whether you are gaining or losing activity.

I totally agree with Steingrimur . Minimally to have a useful discussion we need you to provide us with the specific activity of your enzyme preps.

Additionally, It would be informative to know what procedure was used for concentration and whether you followed the concentration step with a dialysis step. If you performed concentration with high salt and failed to return the enzyme to more optimal concentrations for your activity assay, your results will not reflect the optimal activity of your enzyme.

Your enzyme is a multimer, however is the catalytic site present as a result of association of all the polypeptide monomers or is the catalytic site on each monomer and able to function independent of the other monomers?

.

enzyme activity depends on pH and ion concentration. By modifying the ionic concentration and/or the pH you might be stabilizing the enzyme. A more stable enzyme has better activity.

As stated above by Peter - your specific activity must increase as you are purifying your enzyme preparation. It is simply a matter of assaying your protein concentration as you go on through the different steps of purification.

Now, once you reach the highest specific activity of your enzyme; it is only then that you can tackle the stuctural aspect of your enzyme preparation by using appropriate analytical methods such as UV/VIS, CD etc..

Best of luck!

Also, many enzymes are much more active when they have other proteins around. They bind to other proteins and use them as a kind of "leverage-maker" for their enzymatic activity. You protein is a multimer, and it probably likes having other proteins around. Several restriction endonucleases show this kind of increased activity when the protein concentration is increased. The enzymes of the urea cycle, Krebs cycle, and urea cycle show "channeling" in which the enzymes that catalyze the reactions within a pathway are much more active in combination with other enzymes of the pathway than they are alone. If your enzyme is a member of a pathway, then it might be showing channeling.

You could also try using TCEP instead of BME (if this isn't even worse for the protein) to test wether it is the reducing agent or not.

Does the effect reverse, when you dilute your sample?

Does the state of the protein actually change? So, did you check by cross-linking and SDS-PAGE?

Duane,

What you say is correct. Assuming no protein loss, the specific activity shouldn't be modified. That is also true for the total units, it shouldn't be changed after concentration. As I understand the Tomas's problem is exactly this: the number of total units was increased after the protein was concentrated.

Do you understand the same?

Thanks for all the answers.

I was talking about TOTAL activity which increased 2-3 times. I didn't measure protein amount in the diluted samples, so I don't know the change of specific activity. However, as I can compare it with the total activity before dilution, it basically returned to original value (let's say before chromatography I had 2 nkat, after chromatography I had 1 nkat and after concentration I had 2 nkat again).

For that reason I don't think it's denaturation because than it had to be reversible.

I don't think some sort of inhibitor would be involved, because the enzyme was purified through 2 chromatographic columns, so theoretically it should be gone before if it was so loose to get lost after concentration.

If your enzyme has a very low affinity for the substrate (high Km), then the enzyme concentration can influence the acivity you are seeing.

Tomas,

Did you use the same concentration de protein in the enzymatic assays before and after the concentration?

I like Michael's answer. Many enzymes suffer denaturation upon dilution. Often glycerol or PEG can be used to substitute for protein to simulate protein-protein interaction. Sometimes BSA is used. From what Thomas has told us, I suspect his enzyme is a bit fragile. Its activity depends on integrity of disulfide bonds and perhaps other stabilizers.

Some years ago I did some experiments with Triose Phosphate Isomerase which demonstrated that addition of glycerol increased activity several fold over the standard conditions used for activity assay.

no, I use standardly 5 ul of my enzyme and then recalculate to get the activity in 1 ml

maybe I'll try the addition of PEG/glycerol/BSA. What final concentration range should I try? If it was true, what does it say about my enzyme besides it's crapy? :)

Hi Tomas (sorry about the lack of accent marks),

If you have access to Methods in Enzymology this might be a good start:

Deutscher MP.

Methods Enzymol. 2009;463:121-7. doi: 10.1016/S0076-6879(09)63010-X.

Maintaining protein stability.

I would stay away from BSA. It defeats your attempts to purify your enzyme. Sugars

also are used. The smaller PEGs, however, have been shown to cause subtle unfolding

of proteins, so one needs to beware and keep the molecular weight up. There

are lots of papers on maintaining solubility and activity of isolated proteins.

Tomas,

If you are concentrating your protein using a amicon or a other comparable concentrators, any protein impurities with a lower molecular mass than the filter cutt-off (eg., 30,000kDa) will be removed during concentration, so your enzymes specific activity can increase, because you are measuring total activity/mg total protein.

Good Luck,

Balajee

You do not mention, How do you measure your protein? also mention the details of measuring the normal versus concentrated?

I think of two possibilities, which might cause this problem.

1) Your enzyme is active only in multimeric form or there is positive cooperativity. As increase in enzyme concentration with purification may cause increased multimerization.

2) your enzyme has lower km value, thus change in enzyme concentration in your assay may reflect on the Vmax and hence total activity.

I hope this will be of some help to you.

Nilesh

@Mary D'Souza if you mean total protein, I use Bradford reagent from Bio-Rad, for my proteinI measure it's activity. I mix all reagents with buffer add the same volumetric amount of enzyme (so the amount of proteins differ).

I would not base any conclusions on results where you concentrate"13-50" time and total activity goes up "2-3" fold. Your variance is way too large to make any claims. Also, after bME has been sitting in solution long enough to deplete oxygen, then it can form mixed disulfides with your protein cysteine residues... adding more bME will almost immediately reverse the situation, but after a while the mixed disulfides will appear again. This is the biggest drawback to bME (besides, it smells bad:-). Use DTT, where this problem does not happen because the formation of the 6-membered cyclic disulfide ring within itself is so much more stable than a mixed disulfide with another sulfhydryl. Also you say you use the standard 5 uL amount for your assay. This is the absolute wrong way to do enzyme assays. You always adjust the volume of enzyme you need to get a rate that gives you a reasonable signal to noise ratio, for a assay time length where no more than 10% ( 5% if you are being very quantitative) of your substrate has been consumed and the rate is linear with time, and, not too fast so that you miss the beginning of the reaction before taking measurements. Hope this helps.

I'm pretty sure my error is less than 100%. And the samples were just after chromatography, so basically in fresh buffer. DTT doesn't work with my protein for some reason.

Due to the increase in the concentration of enzyme, total activity increases.

I think your hypotheses, Tomas are valid, if the specific activity goes up with concentration. However, it could also mean as others have suggested that you have gotten rid of impurities. To test your hypothesis for the multimeric enzyme, you can just redilute your protein and assay again. And as Dean has pointed out, you would need to improve your accuracy with the experiments and do things quantitatively to find out what is going on. If you want to determine if BME is activating your enzyme, which is possible, then do the experiment without concentrating. Just add BME to your enzyme and see if it improves the activity. If your activity has disproportionally

Increased due to the increase in concentration, and you quantify this appropriately, ie maybe the activity goes up 10 fold but your concentration only increases 2 fold, then you may want to do additional experiments. But all the experiments you may want to try should be quite straight forward. What are your units for total activity? This would help clarify things. Is it total units, specific activity (units/ mg protein)?

Hi Tomas,

Many experts have already given lot of valuable suggestion and opinions.

As you mentioned,that your enzyme is a multimeric protein. Then yes, there is a possibility that increase enzyme concentration,leads to increase in multimerization of a protein and in turn your total activity increases by folds (co-operativity effect). In my opinion,this could be possible, as your protein therotically although known to be as mutimer,during purification,it might get dissociated to trimer/dimer/also in monomer form. However, during assay condition or in presence of all the essential pre-requiste substrate and cofactors,it might reunite into a multimeric form,thus leads to increase in total activity. You may compare taking UV spectra of protein preparation used for the assay each time with the protein during purification stages (Gel Filteration Chromatogram). If this is true, your purified multimeric protein might show different adjacent peaks corresponding to dimer/monomer.

2. Make sure that there is no mutation occured in the gene encoding your protein,because sometime a single point mutation (a critical residue) in the gene do affect the enzyme activity. Thus, the total effect you observing could be due to part of inactive/less active enzyme.

3. As mentioned by @Steingrimur Stefansson Sir and Nilesh Aghera, Your enzyme might have a low affinity towards its substrate (higher Km), thus it requires a higher enzyme concentration to attain same or higher Vmax in a given assay time.

Hope,you may find all the suggestions/opinion useful.

Good Luck for your endevours!

Best regards,

Irshad Baig