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 184.108.40.206) vary quite dramatically in this regard
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
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
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
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
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
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
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.