I recently finished an experiment where I analyzed the rate of ATP hydrolysis of Heat shock protein 104 in three species of fungi. They have shown to all have different rates of ATPase activity. How would I explain these findings based on structure/ATP affinity? The experiment was done in vitro.
The species under consideration: Candida albicans, Saccharomyces cerevisiae, Schizosaccharomyces pombe
As nobody has answered this good question, I'll have a go.
Firstly, let me state that I have little-or-no knowledge of heat-shock proteins. What follows are some general observations and thoughts.
It would not be unusual for the same enzyme from different species to have different kinetic properties. For example, yeast and horse liver alcohol dehydrogenase (EC 1.1.1.1) vary quite dramatically in this regard (see below).
It is to be expected that the same protein from different species will have different amino acid sequences.
Even the sequence of cytochrome C (~104 amino acids), one of the most conserved proteins, differs between species. There are 44 amino-acid differences between human and yeast cyctochrome C, for example (although there is no difference in amino acid sequence between the human and chimpanzee proteins) [see Creighton, 1993, p116, quoted below].
This fact alone could explain the kinetic variation, but it might be very difficult (and require a lot of work) to pinpoint what differences in amino acid sequence (if any) account for the observed differences in kinetic constants.
In the case of hsp104, a quick alignment (out of curiosity) of the amino acid sequence of the Saccharomyces cerevisiae and Candida albicans enzymes showed quite a number of amino acid differences between these obviously homologous proteins.
I used the TCoffee sequence alignment algorithm (a new version of ClustalW) at the xPASy server. (I received an alignment score of 98).
For the record, the accession numbers used are the following:
hsp140 from Saccharomyces cerevisiae, 908 amino acids, Accession Number AAA50477
hsp140 from Candida albicans, 899 amino acids, Accession Number AAK60625
Of course, the observed kinetic differences may not be due to amino acid sequence differences at all, or may not be only due to such differences. Post-translational modification (such as phosphorylation) and differences in quaternary structure are other possibilities.
I notice from this reference (Parsell et al., 1994) that hsp104 from Saccharomyces cerevisiae forms oligomers in the presence of ATP, and this might be very important in any explanation of kinetic differences. (My whole knowledge of hsp140 does not extend beyond this excellent paper).
Yeast and horse liver alcohol dehydrogenase (ADH) are homologues and catalyze an identical reaction. Both also contain zinc.
However, the yeast enzyme is a tetramer, whereas horse liver ADH is dimeric.
Another difference worth pointing out is that in vivo yeast ADH functions as an aldehyde reductase (making ethanol), whereas liver ADH functions as an ethanol dehydrogenase (in alcohol elimination). [I am aware there are other isozymes of yeast ADH, which may have different functions].
As an ethanol dehydrogenase, the yeast enzyme has a kcat value of 455 s-1 (pH 7.05, 25oC; Dickinson & Monger, 1973) whereas the horse liver enzyme has a kcat value (for ethanol dehydrogenation) of only 1.67 s-1 (pH 6.0, 25oC; Dalziel, 1962, 1963).
In the reverse direction, the kcat for acetaldehyde reduction is 3850 s-1 for the yeast enzyme (pH 7.05, 25oC, Dickinson & Monger, 1973), whereas it is only 125 s-1 for the liver oxidoreductase (pH 6.0, 25oC; Dalziel, 1962, 1963).
How can we explain these kinetic differences from an analysis of structure? In my view this is a very tough question. It may be due to some or all or none of the differences I have highlighted.
Perhaps the question needs to be rephrased as follows: Is there any fundamental difference, at any level, in the catalytic mechanism, or in the form of the enzyme in solution, or in the amino acid sequence, or in the tertiary or quaternary structure, that can reasonably account for the observed kinetic variation?
Are the differences worth explaining? Perhaps one needs to go no further than to record the individual variation under rigorously-defined reproducible conditions?
Before such questions may be answered, it is important to establish the nature of the observed kinetic differences. What follows are merely some guidelines, most of which you are probably aware of.
Is the velocity versus enzyme plot linear in all cases at both high and low substrate concentrations? That is, in kinetic jargon, is the ν vs [Eo] linear? As the enzyme is known to form oligomers in the presence of nucleotides (see above), this might be an important control. Does doubling the enzyme concentration exactly double the rate, and does halving the enzyme concentration exactly half the rate (at both high and low substrate concentrations)?
An example of an enzyme where the ν vs [Eo] is often not linear is phosphofructokinase.
I notice you are using a coupled assay with two coupling enzymes. What effect does doubling the amount of one and/or both of these enzymes have on the observed rate? It should have none, otherwise the assay is not valid.
The substate is ATP. Almost certainly, the 'true' substrate for the enzyme is MgATP2- (correct me if I am wrong about heat-shock proteins). How much Mg++ do you have there?
In determining kinetic parameters for ATP-utilizing enzymes one needs to be aware of ionic equilibria during experimental design. Failure to do so may give rise to spurious kinetic effects.
A solution containing ATP and Mg++ will contain many ions, probably only one of which is the substrate for the enzyme, and whose proportion will vary with concentration. It is essential that this is taken into account. The problem and posssible solutions are explained very well by Cornish-Bowden (2003, pp 86 - 89) and by Storer & Cornish-Bowden (1976).
One experimental design is to keep the concentration of MgCl2 in constant excess over total ATP concentration (CB recommends 5mM). This is an important one.
Are Michaelis-Menten kinetics obeyed in all cases? Is there any evidence for substrate inhibition, or substrate activation?
Are the double-reciprocal plots (Hanes plot, Lineweaver-Burk plot, Eadie-Hofstee plot) linear in all cases?. Non-linearity may be an indicator of kinetic complexity, or of poor experimental design.
If you plot the kinetic data for all three enzymes on a single double-reciprocal plot what sort of a pattern do you get? Competitive? (no differences in kcat, but differences in Michaelis constants).
In comparing enzyme kinetic parameters the Michaelis constant has the big advantage that it is independent of enzyme concentration. Thus any differences are likely to be 'real' and not due to errors in, say, protein concentration.
But what about Vmax? If, say, you determine that the maximum velocites differ by a factor of 1.4, can you be certain that you are not inadvertently adding slighly more enzyme in the higher case, and that the catalytic constants are in fact identical?
As I said above, these are just personal thoughts. Most you probably already aware of.
