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Enzymology - Science topic

Enzymology is the study of the properties and action of enzymes.
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I am looking for an enzyme that can hydrolyse a fucoidan polysaccharide into oligosaccharide units. 
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Did either of you find a commercial fucoidanase enzyme supplier?
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I am working in laccase enzyme production by bacteria. In that i have a problem with quantitative estimation of laccase enzyme. I have confirmed the laccase production with plate assay using guaiacol. But i face the problem with quantitative method of laccase enzyme. So kindly suggest me some tips and methods for the laccase enzyme assay. Thanks in advance:)
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Dear I am published two paper from the phD thesis to my Student in (2020):
1. Doaa Khalid Mezaal, Essam Fadel Al-Jumaili and Ahmed Majeed Al-Shammari (2020). Production and Purification of Laccase Enzyme by Klebsiellapneumoniae K7 . IISTE.
Chemistry and Materials Research . Vol. 10, No.5, PP:17-23.
2. Doaa Khalid Mezaal, Essam Fadel Al-Jumaili and Ahmed Majeed Al-Shammari (2021). Study of Production and Characterization of Laccase Enzyme from Klebsiella pneumoniae K7 Isolate, Medico-legal Update , Jan- March, Vol,21 No.1 pp: 216- 223.
if you don't get these papers i can sent to you by your e-mail.
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Dear all,
I tried docking my protein heterodimer to form a hexamer (trimer of a heterodimer) using symmdock server. The output results has given top 20 models of the hexamer and I opened the top model into PyMOL. I see the the parental template has intact structure while the generated partner structures are all broken in the PyMol (Picture attached). When I opened it in coot all the atoms are present and display very well as atoms. But somehow it is messed up with the cartoon representation in PyMol. Please suggest, how can I fix this issue.
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You need to make sure from the beginning (before docking) that the "receptor" and the "ligand" do not have the same chain label! You can change chain labels using the "alter" command in PyMol ( https://pymolwiki.org/index.php?title=Alter&redirect=no )
alter (ligand and chain A), chain='B'
changes the chain label in object Ligand from A to B
You can force a proper display of your faulty complex by using the "retain_order" setting in PyMOL ( https://pymolwiki.org/index.php/Retain_order )
set retain_order, 1
but the ambiguous chain labels may cause problems with a number of other commands and further evaluation of the complex in other programs!
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I need to characterize two extra cellular enzyme produced by bacteria . For that process the purification of enzyme step is necessary before the characterization process? otherwise can i use the crude enzyme extract (cell free Supernatant) for the characterization process?
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If you have a specific assay for your enzyme's activity, you can perform enzyme kinetics on crude preparations. Also helpful is a specific inhibitor of your enzyme (like ouabain for Na/K-ATPase), the activity you measure in its presence is due to other enzymes and hence subtracted.
Many other measurements can also be performed with crude enzymes, if the method is adapted. For example, size determination by analytical ultracentrifugation requires pure proteins, but methods for crude preparations are available (DOI:10.1016/0003-2697(89)90297-2, DOI:10.1016/S0021-9258(18)64180-8).
Indeed, some purification methods (especially solubilisation of transmembrane enzymes) are so harsh, that part of the purified enzyme may be non-functional or (worse) partially functional. This would make measurements of turnover number impossible.
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Acetate kinases are proposed to exhibit either a "direct in-line" mechanism or a "triple-displacement" mechanism. However, I was unable to find a schematic for any of these mechanisms, even less their respective kinetic treatments to derive rate equations.
Can anyone suggest a reference containing possible rate equations for this case, or at least a clear description of the proposed mechanism so I can try to derive the equation(s) myself?
I appreciate any help you may provide. 😊
Best regards,
Gustavo
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I don't work with acetate kinase, but it looks to me to be a pretty standard 2-substrate, 2 product reaction: acetate + ATP <=> acetyl phosphate + ADP.
The kinetic mechanism may depend on the source of the enzyme, and there may be some complications such as cooperativity or substrate inhibition. Here is a paper on the enzyme from some bacteria in which the kinetics were analyzed as either steady-state ordered sequential or rapid equilibrium random sequential, with rate equations supplied.
Here is another paper on the kinetics of the enzyme from a different bacterial source, in which the kinetic mechanism was concluded to be Random:
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Our primary experiment results indicated that food resource arabinogalactan protein (AGP) could combine with phenolic compounds, such as epigallocatechin gallate (EGCG), thus can mask bitter or astrigent taste and function as a good carrier of these bioactive components in food processing. As a kind of proteoglycan, we are planning to modify its sugar moiety and to test whether the sugar moiety take key function in combination with EGCG. Does anyone have experience in the enzymatic treatment of AGP or proteoglycan based on sugar chains?
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Dear all,
Hope you all doing well. I would like to bring my concern during kinetics measurement of an exonuclease enzyme. I am performing an exonuclease assay, by varying substrate concentration (25 nM to 500 nM) and fixing the enzyme concentration (2 nM) and I could see the cleavage products very clearly till 300 nM in a linear range. After 300 nM substrate the enzyme struggles to cleave the product as it is attaining the supersaturation. Now, I have a concern that should I also change the concentration of enzyme during this kinetic measurement ? And if I change the concentration HOW and WHEN I need to divide the enzyme concentration in the reaction mixture, during the kinetic parameter calculation ? Because I am little bit concern about the calculation for adjusting the enzyme concentration. Please suggest.
Thank you
With kind regards
Prem
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Dear, I would suggest you to take 200 nM of substrate (fixed) and change enzyme concs (0.2 nM to 5.0 nM), with 5-6 data point. Draw a curve b/w enzyme con (x-axis) vs product formation (y-axis). It will guide u to select enz conc where product formation linearly increase. Select that enzyme concentration (stick to that) for all kinetic experiments and mutant characterization. Hope that helps. Regards,
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Dear all,
Attached is the image of expression profile of my protein of interest. Starting from the left after molecular weight marker, are total cell lysate (uninduced), total cell lysate (induced), supernatant (Uninduced) and supernatant (induced). IPTG used for the induction was 1 mM. The expression system used was BL21 DE3. Here, the prominent band is in the right range of expected molecular weight. But I am worried I see almost identical expression in both uninduced and induced. Western blot could answer definitely if it is my protein of interest ( need to be done). But is it possible with normal BL21 DE3 cells ? Please give your insights into it.
Thank you
With kind regards
Prem
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Which promoter and culture media are you using?
Which is the MW of your band? If i remember well the E.coli show a more strong band at about 80KDa, and if you are in this MW range is also possible that your band colocalize with it and is not so simple distinguish the over expression of your protein wth the basal expression of E.coli bands.
If your protein carry an affinity tag (His, GST, MBP) i suggest to you to perform a small purification using 100ul of resin on the non induced and inducer to see it is really a basal induction ot not.
good luck
Manuele
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Hello, I am intending to screen a library of compounds for an in vitro enzymatic assay. The target enzyme follows a Bi-Bi mechanism. The assay development phase is completed with kinetics parameters for both substrates are determined. I am looking into a rational approach to choose the concentration of both substrates and compounds to satisfy the following criteria:
-Inhibition Mechanism-blind design: the mode of inhibition and the substrate of which the inhibitor compete with is unknown.
-Minimize the concentration of the compounds to a reasonable concentration that still able to pick up any possible inhibitory mechanism.
Successful compounds will be used to identify Hits with dose response and IC50 later on.
Is there any recommendations/guideline followed in pharma to deal with this? is there any upper limit for a Hit to be accepted?
Thanks
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In my experience, the usual concentration of test compounds from a generic compound library used in high-throughput screening to search for enzyme inhibitors in an in vitro assay is 10 µM. For fragment libraries, a higher concentration, such as 100 µM may be used because the molecules are smaller and simpler, overall, than the molecules in generic screening libraries.
Going to higher compound concentrations may seem appealing, but the result is not necessarily better because you run into increasing problems with compound insolubility and interference with the assay readout as you increase the compound concentration.
As for the substrate concentrations, there is no "right" answer. There are trade-offs. Since you know the kinetic mechanism and the substrate kinetic parameters, you can calculate the ratio IC50/Ki using the appropriate Cheng-Prusoff relationship for inhibitors competitive with one substrate or the other as you vary the substrate concentrations. This ratio can be used as a measure of the sensitivity of the assay to find inhibitors. (For pure non-competitive inhibitors, the substrate concentrations don't affect the sensitivity.) I suggest you prepare a spreadsheet with columns for [A], [B] and IC50/Ki. Vary A and B and calculate IC50/Ki.
Typically, the lower you make one substrate concentration, the more sensitive the assay is for being inhibited by compounds competitive with that substrate (lower IC50/Ki), but the less sensitive it is for finding inhibitors competitive with the other. You can try to find a pair of substrate concentrations that achieve a balance.
To get around this issue, you could run two separate screens, if you can afford to, with one optimized for finding inhibitors competitive with A and the other optimized for finding inhibitors competitive with B.
You also have to consider the substrate concentrations needed to get a sufficient signal to run the screen successfully. Higher substrate concentrations allow a larger signal to be obtained while still under initial rate conditions, but higher substrate concentrations increase IC50/Ki. Cost and/or substrate availability may also have to be considered.
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Hi researchers…
Is it true that calculate kcat of my enzyme with this formula? Kcat=Vmax/km
I saw this formula in an article: Kcat=Vmax/[E]; but I don’t understand which enzyme concentration?! Total enzyme concentration???
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  • 5 substrate concentrations are way too few to get reasonable estimates for Km and Vmax, 12 are considered the minimum, these should be regularly spaced in the range between 0.1 and 5 Km (better 10 Km, if solubility permits and substrate inhibition does not occur). This, of course, implies that you have an estimate for Km, which can only come from pre-tests.
  • This is true only, if the [S]/v-relationship is indeed hyperbolic, which needs to be checked carefully. If there is cooperativity with several binding sites whose affinities vary by orders of magnitude, the required number of experiments explodes. However, the 12 experiments from above, when carefully executed (standard deviation of points ≤ 2%), should at least demonstrate a departure (runs test or, better but more work, F-test to compare sum of squares of the regression with standard deviation of data points).
  • In addition, one should check that the rate really depends linearly on enzyme concentration.
  • You mix up substrate concentration (mM) and substrate amount (mmol). Use units to check whether your calculations are correct!
  • Your formula for getting kcat from Vmax depends on the enzyme being 100% pure and 100% active, as discussed before. Prima facie evidence for this would be one band only in SDS-PAGE (use several amounts of enzyme, until the band is really (!) heavy) and a specific Vmax in the order of what is published.
  • As a beginner, you may find it easier to use Eisenthal & Cornish-Bowden's "direct linear plot" of your data to determine Vmax and Km than a regression of v against [S]. As a non-parametric method, the direct plot is more forgiving when data have scatter.
  • Do not forget to calculate the error margins for all your values, values without error estimates are next to useless.
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I read this report that 3% (v/v) Ethanol increases recombinant protein expression in Escherichia coli. The SDS PAGE gels seem pretty convincing (1).
However, assuming I get great expression of recombinant protein with 3% (v/v) Ethanol, how am I supposed to decontaminate the biohazardous cell waste from the experiment?
Bleach decontamination isn't a good idea because bleach would react with the ethanol.
Autoclave decontamination isn't a good idea because ethanol is flammable.
Is safety just not a priority?
Source:
1) Chhetri G, Kalita P, Tripathi T. An efficient protocol to enhance recombinant protein expression using ethanol in Escherichia coli. MethodsX. 2015;2:385-391. Published 2015 Oct 8. doi:10.1016/j.mex.2015.09.005
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Hi Adron Ung . The reactions of bleach and ethanol I have seen contain at least 70% ethanol. I'm skeptical that 3% ethanol + bleach could cause significant production of dangerous chemicals.
But you can always keep the 3% ethanol + bleach waste in a fume hood overnight just to be on the safe side.
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How to measure the percentage activity of LDH (0.5 ug/ml) using 2 mM of NADH and 10 mM of pyruvate ( 25 °C, pH 7.00 ) and observing oxidation of NADH at 340 nm? Termination of reaction is necessary while measuring the activity?
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Thanks @M Angeles Zorrila Lopez- Perea
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I am looking for Methods in Enzymology, Volume 22, 1971, Pages 248-252. Specifically, chapter 23: Crystallization as a purification technique.
Alternatively, the following: "A technique for the crystallization of proteins" by William B. Jakoby.
My institution no longer has access to older articles, and I am really interested in the method of crystallization.
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Dear Eugene, please note that there are many more useful articles on the crystallization of proteins available. For example, please have a look at the following relevant reference:
Protein Crystallization for X-ray Crystallography
(see attached pdf file)
Also please see the following interesting articles:
A historical perspective on protein crystallization from 1840 to the present day
A technique for the crystallization of proteins
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I need to calculate Vo in order to than calculate Vmax and Km, I know you can calculate it from absorvance but I don't have that. Is there another way? Can I calculate it using the attached diagram?
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What is needed is r vs cS, not cP vs cS. We can obviously assume that r within 10 min is constant and then r is proportional to cP. But that has to be proved.
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I want to calculate an enzyme's energy of activation, but I don't have the enzyme's molecular weight to calculate the rate constant or the pre-exponential factor A. This is part of an assignement I have for a virtual lab of enzymology, we use this virtual lab https://www.ucl.ac.uk/~ucbcdab/enzass/enzymass.htm by ucl and the process of it is basically you choose one of 5 enzymes which are fictional enzymes from what I can tell, and then you can perform "expirements" to determine the enzyme's behaviour in different pH environements etc.
I've attached screenshots of the diagrams the virtual lab has produced about how the enzyme reacts to different temperatures. I've concluded that at around 50°C it produces the maximum amount of protein so I've calcuted the maximum velocity using that at 5.86μmol/min. I had to calculate this because since I don't know anything about the actual enzyme I have to calculate the energy of activation using this equation: logk = logA - E / 2.303 * R * T (where logk=logVmax), our professor told us that we have to use this to calculate the energy. She also told us to use excel to calculate logk and logA and create a y=ax+b diagram. I honestly am really struggling to come up with how to use excel in order to do that since I'm really unexperienced with it.
Please refer to the photos below if you want to see the data I have.
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If we assume that the cP obtained within 12 min is proportional to the enzyme activity, then the Arrhenius equation can be used, where k is replaced by cP. Another issue is the method of determining the values of ko and E. One can use the graphical method, where lnk is plotted against 1/T, one can use nonlinear methods.
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Dear all,
Greetings !
I am a total newbie in enzymology as I started working on this very field couple of months ago. I am working on a nuclease which degrades oligomers of nucleotide from 3' end. Now, I have a setup for the enzymatic assay where I stop the reaction at different time points using a stopping buffer. The oligo substrate is fluorophore labelled and degradation of which can easily be monitored using phosphor-imager that gives the intensity of the substrate degradation or the product formation. My question here is, what will be my approach to calculate the initial velocity, Km and Vmax of the very enzyme using this assay ? Actually, I am looking for calculating those parameters with substrate change (i.e using the top band from each lane) and not with the degraded products formation as it seems complicated to me. Because each nucleotide degradation forms a product w.r.t time and so several products making it complicated. So my choice will be to see how much substrate degraded with time. I have attached the Urea PAGE profile of the assay, where starting from left, lane 1 to lane 5 is is time t=0 min, 10 min, 15 min, 20 min and 30 min respectively. Please help me in this regard.
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1. In Excel, use the trendline function and check the box for forcing the line to pass through the origin.
You can divide by the volume if you wish in order to convert from moles of product to molarity. Since all the reactions have the same volume, this will not affect the Km.
The ideal Michaelis plot is a rectangular hyperbola, with all the data points falling on the curve. The asymptote is Vmax. In real-life, there is usually some scatter in the data around the curve. In your data consisting of 5 points, there is too much scatter to fit the curve.
2. Regarding the amount of product in an exonuclease reaction, I agree that there can be more moles of product than there were of substrate if you count the individual nucleotides released. If that is how you did the calculation, then it may be OK. It is important that the substrate concentration should not decrease substantially during the reaction, no more than 10%, preferably.
3. The reason I don't agree with plotting "specific activity" is that specific activity is defined as the activity measured under one specific set of conditions, which is preferably at Vmax. If you want to just call it pmol/min/mg, that's fine.
4. One of the crucial aspects of Michaelis-Menten kinetics is that the initial rate of the reaction is measured. On a plot of product versus time, you draw a line that starts at the origin and runs tangent to the start of the progress curve. In other words, the data points at the shorter time intervals lie along a straight line. At later times, the curve may start to flatten as the substrate gets used up, but you would not use those data points to calculate the slope of the line for the initial rate. It doesn't matter how long the measurement lasts as long as you measure the initial rate. Rate is change in product divided by change in time (slope), so the amount of time is accounted for in the calculation. It isn't even necessary for the same amount of time to be used to calculate for slope for every substrate concentration, as long as the initial rate is measured.
I imagine that the rate of reaction with the poorer substrate will be slower than the rate of reaction with the better substrate, making it necessary to use a longer time scale to measure product formation, or to increase the amount of enzyme to speed the reaction up.
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Hi to all I immobilized a peroxidase on a new synthesized MOF. The reviewer of a journal paper asked me to consider the inhibitory activity and specific affinity of my immobilized enzyme in the introduction. How I can answer this comment of the reviewer for a journal paper revision? What does it mean? In my opinion, these items are related to biosensors. I reported the influence of immobilization on km in my article but I have no idea about the inhibitory activity.
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The specific activity of an enzyme preparation is the turnover-number (molecules of substrate that an enzyme molecule handles per second) multiplied with the enzyme concentration.
What your reviewer is in effect asking is
  • whether the enzyme might have been partially damaged during immobilisation
  • whether there is any steric hindrance to substrate binding/product release due to the immobilisation.
Any such effect could invalidate your observed Km. However, I am not sure that this belongs into the introduction. Evidence of full activity should be presented under results, and its significance in the context of the paper discussed in the discussion section. An exception would be IMHO if the immobilisation procedure is well established and proven to result in fully active enzyme, than this would be mentioned in the background part of the introduction, with appropriate references. Of course, you'd still have to very that your preparation is fully active, and this should be mentioned under results.
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I want to know normal value of Superoxide dismutase and glutathione peroxidase and reduced glutathione and malinoaldehyde in tissue of rat.
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Superoxide dismutase, glutathione peroxidase and total antioxidant are antioxidant parameters but malondialdehyde are oxidant markers. The values of these factors are different in the articles. Different laboratory kits, different animal tissues as well as measurement methods can be the cause of this difference. Last but not least is the unit of measurement that should be considered.
I think it would be helpful to search for articles with similar measurement methods to your own work and to compare control groups.
Best regards
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Among various factors, the temperature is one of the most crucial factor for enzyme activity. For most of the enzyme assay, researcher use 37oC or normal room temperature as incubation temperature. If we consider the poikilothermic animals like fish, it's natural enzyme activity depends on habitat temperature and each fish (tropical, temperate, polar or cold water species) has their optimum temperature at which it grows best and we know growth is nothing but a consequence of optimum (good) metabolism.
So, is it right to use the mentioned assay temperature for enzyme activity of fish irrespective of its natural habitat?
Or,
Do the researchers need to modify the assay temperature according to optimum natural habitat temperature of fish?
Please provide your suggestions.
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We may not adopt a single temperature for fish enzymes such as 37ºC in case of human samples. It should be specific the species as fish is not a single species. Again, we need to find optimum temp for each of the enzymes as well. For convenience, we can select a constant temperature for all enzyme assays, which can be selected based on the optimum temperature for growth of that species. In the following publication, you will find an optimum range of temperature for digestive enzymes, however its quite a wide one (45-65ºC) https://scialert.net/fulltext/?doi=jfas.2017.264.272#:~:text=The%20optimum%20temperatures%20for%20both,water%20temperature%20of%20the%20swamp.
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It is very much confusing when some papers use this term "apparent Km of an enzyme even if they use single substrate ? Not at all clear ! Please help me understanding this. Thanks in advance.
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Hi there,
The term apparent Km is used basically when the enzyme is not pure and/or when the composition of the reaction mixture is not fully controlled (ie. containing molecules which are not involved directly in the reaction but which may interfere with it): the presence of contaminants during the enzyme assay may affect the results.
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I apologize if my question is not clear. I have a pure chemistry background and I am very new in a Protein Chemistry and Enzymology science.
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the main difference between pcr and sanger sequencing is that pcr has 2 primers facing towards each other but sequencing has only one primer reading the sequence in one direction only. This means that in early cycles of pcr the amount of dna generated increases exponentially ( doubles every cycle) so after 10 cycles we have 1000 time the amount of template dna that we started with. This allows amplification of tiny amounts of dna. In sequencing we start with a lot of template because with one primer only the amount of amplified original tempate is copied each cycle so after 10 cycles we have 10 times as much of the strand of dna that is being amplified. Also in sequencing the reaction is deliberately terminated early in the propogation step by the addition of dideoxynucleotides which stop elongation at different lengths so giving a snapshot of all shorter sequences making up the full sequence of the product. The enzymes used in both techniques are the same and the cycling parameters are similar.
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Hello fellow scientists,
I wish to determine the Dissociation Constant (KD) of a DNA polymerase binding dsDNA. I won't disclose what the DNA polymerase is because it is unpublished work. I have done some binding assays in Agarose gels, but due to the poor sensitivity of the available dyes I had to visualize the relative binding stoichiometrically, and I could not simply just set the protein or DNA concentration around the expected KD.
Previous work in our lab has determined a KD = 20 nm for our DNA polymerase binding a 33mer locked double stranded DNA hairpin.The purpose of using something so complicated was for kinetics assays.
However, I am using a 13-mer dsDNA construct because my goal is to crystallize the DNA complex and a 33-mer is just way too large! My supervisor has advised that I don't believe that my KD is actually 20 nM for my small dsDNA construct.
I am interested in using Isothermal Titration Calorimetry mainly to calculate the KD of my protein to binding this 13mer dsDNA construct. I would titrate my dsDNA into a fixed concentration of protein. I could guess that the KD is 20 nM, but I actually don't know for sure.
I have heard that when you determine the KD you have to have some estimate of the KD and then scan ligand concentrations above and below the KD, measure the response to get a curve of response vs ligand concentration and the KD is mathematically fit or basically it is just the inflection point of the binding curve.
However that advice doesn't tell me if the KD is say 20 nM, what should fixed concentration of my protein be? (I have appreciable amounts of 100 µM protein because I am a crystallographer so excessive protein isn't an issue.). What is the max and min range that I should scan the ligand concentrations? What if the KD is way worse than we predicted and it is actually 1 µM? What fixed concentration of protein should I use and what min and max concentrations of ligand should I use?
Is there a way that I can measure the KD with a certain fixed concentration of protein, and a huge range of ligand concentrations regardless of if the KD is 20 nM or 1 µM? Is that possible?
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  • For statistically valid results and hyperbolic binding curves, the final ligand concentration should cover the range of 0.1 to at least 5, better 10 times Kd (if solubility permits), you need at least 12 ligand concentrations, which should be equal spaced (doi:10.1006/jtbi.1996.0023).
  • For method with reasonable sensitivity, the constant concentration of the macromolecule should be around Kd for best precision. For insensitive detection methods like ITC, you may need a higher macromolecule concentration for a well-detected signal.
  • If your binding curve is not hyperbolic, you'll need a wider ligand concentration range, in that case it may be worthwhile to space the data points logarithmically (1.0, 1.6, 2.5, 4.0, 6.3, 10, 16,...). From these guidelines it follows that you need to do an initial search study for rough determination of Kd. "Rough" means the order of magnitude.
  • Btw, the use of linearisation (say, Lineweaver-Burk or Scatchard plots) for the determination of Kd is outdated, although these methods still have use for data presentation. Use non-linear regression for determination instead, the Nelder-Mead simplex algorithm (doi:10.1093/comjnl/7.4.308) is more stable than Marquardt-Levenberg (doi: 10.1137/0111030), but requires bootstrapping for determination of error margins.
  • You may also wish to think about the detection method you use, ITC requires relatively large amounts of both macromolecule and ligand. Perhaps, surface plasmon resonance (SPR) would be a better choice. After all, you'll need lots of material for crystallisation later ;-)
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I do not know if it is possible to isolate FADH2. We have obtained NAD and NADH. In contrast we found FAD but not FADH2. Looking around we could not find anyone selling FADH2. Can someone help us?
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Hi. I also want to accurately distinguish the peaks of FAD and FADH2 using mass spectrometry.
Have you solved your problem? I would appreciate it if you reply.
Thank you.
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I am working with cellulases, I have expressed my cellulase enzyme in pET28a expression vector. The bacterial lysate has activity against cellulose substrate but after purification of protein with his-tag using GE 1ml His-trap column, my enzyme does not have activity. I elute my protein with 250mM imidazole, but I have dialysed my sample and still i could not find activity. I need some valuable suggestion.
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During cellulase precipitation with ammonium sulphate, the bacterial culture should be grown in CMC..please suggest the viscosity of the cmc (medium or high viscosity) for precipitation? In my case high viscosity CMC particles remained with the pellet after the precipitation
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I do like Cell, Nature, & Science. I find the discoveries to be amazing. But the experiments can be difficult to understand and difficult to replicate without the state of the art scientific instruments. So I have been frequently reading the volumes of Methods in Molecular Biology and Methods in Enzymology. The reason is because these journals give reproducible experiments with simple explanations.
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Dear all,
Apologies for an off topic. I am curious to know if someone wants to pursue a second PhD in order to switch into a closely related field from his/her first PhD, would that be considered as post PhD experience ? The person may want to get into such situation when he/she wants to get trained into that field as most of lab doesn't consider if that background is lacking. It is the matter of interest of the candidate to move into that closely related field. Please give your inputs and suggestions in this regard. Thank you in advance.
Thanks
Prem
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Well, if it is a sufficiently closely related field, in which the background from the first PhD is at least somewhat useful, I would really try to negotiate for a Postdoc position in the second field, rather than a second PhD. Even if you stay within the same general field, a PostDoc is an opportunity to greatly expand your experience and to learn as many new techniques as possible, while gaining more focused insight into your field of interest.
Of course, this will give you more responsibility for your own training - less formal training, but you need to actively approach collaborators to explain to you how things are done in the new lab, and start acquiring new methods from literature and other sources.
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I’m working with a rather complicated ping pong, bi-bi double displacement reductase enzyme that oxidizes NADH and then reduces a substrate. Interestingly, the enzyme has intrinsic NADH oxidase activity in the absence of the substrate. Of course, upon addition of substrate, the velocity of the reaction increases and NADH is consumed more rapidly. The intrinsic activity introduces a baseline velocity and is complicating our kinetic analyses a bit. Complicating it even further is a noted double substrate inhibition pattern (apparently not uncommon with ping-pong mechanism enzymes). We’ve run a full course of analyses, varying the NADH as well as the substrate and we're now trying to fit the data to an appropriate equation. My question is: How do I take the ‘intrinsic activity’ of the enzyme into account (if I need to at all)? Can anyone recommend an equation and appropriate software that can do this? We currently have SigmaPlot, GraphPad, and Enzfitter (for the double substrate inhibition model). Thanks in advance for any guidance, ideas, and/or discussion.
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قياس فعالية الانزيم في الجزء المراد دراسته ومن ثم اخذ برنامج احصائي هو spss
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Hi dear researchers
We used HRP (Biobasic) in phosphate buffer (pH: 6.5) and after adding TMB, the reaction turning to light blue without adding H2O2. After adding H2O2 to reaction media, the brown color was produced (not expected blue color).
There were no changes after decreasing TMB, HRP and H2O2 concentration.
We expect that after adding TMB to the enzyme without H2O2, the reaction doesn’t start and after adding H2O2, we expect that see blue color.
( I have checked and rechecked all calculations, buffer composition, pH etc. )
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That's alright. You have mentioned virtually everything you did but what happened to the addition of SDS to promote the stabilization of the blue colour? It might be what is missing. Please see the attached document:
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I want to compare the amount of an enzyme's expression (laccase for example) or presence in the tissue as a result of the experimental treatment. Is there a protein similar to an antibody that might bind with the enzyme and can be filtered then quantified by spectroscopy? or perhaps by comparison of PCR bands. I am interested to hear any other thoughts or suggestions as well. Thanks very much.
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Adam B Shapiro thank you very much for your reply and the link.
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Dear All, I am purifying my protein (theoretical PI = 8.9) in a buffer of pH =7.1. till Ni-elution I kept the DTT concentration (1mM). Usually, after elution people used to keep DTT concentration at 5mM. But in my until Ni-elution the protein look absolutely fine. But the moment I add DTT (5mM) thread like aggregate is formed. when I run those aggregate, it shows up as my protein. What could be the possibilities that DTT is making it precipitated. As I know DTT is used to solubilize protein as reducing disulfide bonds. I will appreciate your suggestions.
Thank you
With kind regards
Prem
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Dear Prem
the effect of dtt and other reducing agent is not ever positive, it depends from the role of The cisteines, if the cisteines are involved in The formation of a structural s-s ( as for example happen in Cu,Zn sod1) the addiction of Dtt may induce protein unfolding and aggregation it precipitation. On the contrary if your protein contain just 1 cystene or the cisteines do not have any structural role but for examples are involved in The binding of a metal (as happen in Sco1) Then the addiction of Dtt can play a positive role.
you can find a more detailed explanation of it on my blog:ProteoCool (https://proteocool.blogspot.com/) in the presentation:
ProteoCool n°16
(3 Common mistakes in buffer preparation for protein ) at page 4
good luck
Manuele
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I am trying to plan an pulldown assay to find some prey protein for my bait protein.
I am interested in using the flag tag because it is small and doesn't disturb structure and doesn't require additional steps to tag your protein like would be necessary with biotinylation.
Plus apparently, the bait-prey complex can be eluted with Flag peptide.
So that is very interesting to me.
However, I haven't been able to figure out how strong Flag-Tag binding is to Anti-flag antibody?
Also, does anyone have a simple protocol for Flag-tag pulldown?
Flag Tag = DYKDDDDK
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In my hands, Flag antibody is OK but not the best. I find HA is cleaner. One thing to consider in IPs with Flag is the amount of DTT in the lysis buffer. DTT weakens the interactions. I now don't add DTT in my lysis buffer if doing IPs with Flags.
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I have recently started working on improvement of wheat flour for baking. I have come across a research which involved the usage of xylanase, cellulase, beta-glucanase and carbohydrases for improving baking results. I wanted to experiment the same procedure. Since i don't have any background with enzymology i cannot understand how to calculate the dosage that needs to be added to the wheat. If someone can help me in figuring out how to calculate the dosage. I am attaching the link of the article.
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I think I would express the dosage as units (U=micromoles/minute) of enzyme activity added per gram of flour. Enzyme preparations are usually calibrated in U/mg of protein or U/mg of solid. The units are measured using a standardized assay. If you want to use the SI terminology, then replace U with nanokatals.
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Currently I am working with alpha amylase assay using DNS method but I have a problem with the colour which is not changing even I used high concentration from the phenolic standards (Trolox and Gallic acid) and I don't know what is the problem.
I used different concentrations from enzyme (0.1 - 1 U/ml) with soluble starch 1% (w/v) but the colour of samples very close from the control (orange-red colour). In addition, the absorbance of standards or samples are higher than the control, and the absorbance increases with increasing the concentration of standard whereas it should decrease with increase the concentration of standard or sample. As a blank, I tested starch with DNS, the enzyme with DNS and different concentrations of standard with starch and DNS, but I didn't get changes in the colour.
The protocol that I used is:
100µL of buffer or sample + 100 µL enzyme (procine pancreatic alpha amylase) then mixture incubated for 10 min at 25°C, add 100 µL of starch 1%, then mixture incubated for 10 min at 25°C, after that 200 µL of DNS (1%) was add to terminate the reaction and the mixture was incubated for 5 min at 100 °C, cooled to room temperature and then solution was diluted by 3 ml of H2O. The absorbance was recorded at 540 nm.
Can anyone tell me what the problem is or suggest what I can do to solve this problem.
Thanks in advance
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For measuring the amylase activity with DNS solution (Miller Method):
1. Prepare starch solution (10 mg/ml)
2. Prepare enzyme extract filtered with syringe filter
3. Prepare phosphate buffer solution (o.1(M), pH=7)
4. Add 1(ml) enzyme extract to test tube, then add 1(ml) of starch solution (it can be considered as a main sample)
5. Add 1(ml) enzyme extract to test tube, then add 1(ml) phosphate buffer (it can be considered as a blank or control sample)
6. Incubation of main and control sample at 37˚C for 30(min)
7. Add 2(ml) DNS solution to each test tube.
8. Transfer the tube to the water bath at boiling temperature for 15(min)
9. Keep the test tube at room temperature to cool the sample
10. Transfer the solution to the cuvette
11. Measure the absorption of the main sample at λ=540(nm)
It is necessary to mention that for calculating the enzyme activity the standard curve must be prepared by maltose solution.
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I am using KinTek software to do a kinetic fitting. I have determined kinetics of the formations of several species in the time course of 5min and established a model for the enzyme (just as an example: A = B = C = D). I already know that B is always below the detection limit of the instrument during the reaction. Is there a way to restrict the amount of B to lower than the detection limit (such as 0.01 uM) in the software? I only find restrictions for kinetic constants and the starting concentration for species. If there is not a way, is it appropriate to import a sudo-data set (such as a constant amount of 0.005 uM B at each time point, with a sigma value of 0.005) for fitting?
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I use for fitting and analysis of kinetic data the Copasi software (free download from Copasi.org ). It's very powerful and friendly for users. It allows to put restrictions on almost all parameters. You may try. If you try, please, let me know which software is better for you.
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Hi dear researchers.
I have a mixture of nanoparticles and hemin and buffers, …
It was centrifuged and washed several times.
Is there any way to calculate the residual hemin concentration?
( There are not any biological samples)
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Dear all,
I am trying to check the interaction between two of my proteins of interest. However before trying any biophysical technique such as SPR/ITC/MST etc, I used normal superdex-200 gel filtration chromatography. I used protein A (Receptor; ~40 kDa) and protein B (Ligand; ~31kDa) in purified form and mixed together with 1:2 molar ratio at temp. 25 degree for 1 hour. Now when I performed Superdex-200 with three individual runs (Ligand, Receptor and Complex). each time I get a surprising result wherein the complex has more retention volume than the Receptor it self (attached result brown: complex, cyan: receptor and blue: ligand).
When I run the peak fractions from each run through SDS PAGE, I get the two different protein bands in the complex run (attached result; lane 1-3 is the Ligand and Lane 4-6 Receptor and lane7-9 Complex peak fractions). Please suggest what could be the possibility of such result. The complex formation shows about 1 degree (57 to 58 degree C) rise in melting temperature every time as compare to their individual ones. Buffer used for the protein are same 1X TBST pH=7.6.
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There is not much difference in elution time between the three samples. The small differences may not be significant. Although your expectation is that the complex would elute earlier than the individual proteins, this might not happen for a variety of reasons: (1) No stable complex was formed, (2) the complex formed, but it eluted at about the same time as the individual components because of a more compact shape of the complex compared to the individual components, (3) at least one of the component proteins interacts with the column resin strongly enough that this interaction dominates the elution profile of the complex so that it elutes at the same time as that component.
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Hi dear researchers
Is it possible that observed increase of Km for one substrate and decrease for another one (for enzymes with two substrates such as HRP) after immobilization?
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Dear Soudabeh,
I had immobilized my enzyme and observed a change in the Km value. However, mine was a single substrate specific enzyme. I second the opinion of Tapan Kumar; the pore sizes, the immobilization technique used, etc., could play a role in the availability of the reactive site to your substrate.
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I have a doubt in calculating the activity of immobilized cellulase i've used the formula Activity of cellulase (μmol/ml min) = 1000 w/Mvt; where, w is the amount of glucose produced, M is the molecular weight of glucose, v is the volume of
the sample and t is the reaction time. Here in the formula instead of ml (i.e volume) i have used grams (g) of beads taken, then the unit for cellulase would be (μmol/g min). The way i am calculating the activity is correct?
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Dear Nisha S,
You will initially have to calculate the free enzyme's activity - IU/mL (the quantity that was immobilized). Later, post immobilization, please perform the assay ( I performed the IUPAC Ghose assay). I observed a decrease in enzyme activity after immobilization within calcium alginate beads.
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I am searching the pGex or pET plazid of HDAC6, if he is possibly for somebody and would give for me, would be needed for my work.
I would like to test the interaction
between my labor's new protein and HDAC6. If somebody is willing to provide me with this plasmids please send to me at Dr. Tibor Szénási Institute of Enzymology,
Research Centre for Natural Sciences, Hungarian Academy of Sciences
1117, Budapest, Magyar tudósok körútja 2.
Fedex number: 317 080 212
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I don't have that plasmid, but maybe addgene has: https://www.addgene.org/search/advanced/?q=HDAC6
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Hi dears
How do nanoparticles interact with enzymes after immobilization?
How can bioinformatics tools be used for this?
can I use bioinformatics free softwares with simple system to know interactions?
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In computational settings it is difficult to analyze this kind of interactions. The main issues involve are, when we make molecular models of nanoparticles based on observed geometries in literature based on electron microscopy and other envelop deciphering methods...such as for instance for ZnO nano particles, possible geometries are:
1. Simple Box
2. Sphere
3. Cylinderical
4. Cone
5. Frustrum
6. Square pyramid
7. Tetrahedron
We may take molecular structure of the molecules such as ZnO, gold and silver compounds, whatever has been used to make these nanoparticles... after that we may expand those arrays applying the symetries of the crystal structure, based on the available information, we cut out the shapes and dimension of the nanoparticles... after that it may be docked with the enzyme structures... However, in our experience the nanoparticles surfaces was not observed to be same everywhere in terms of composition... therefore it is difficult to predict from which side it may interact with the active amino acid side chains on the enzymes...People are publishing results on this but with too much speculations and assumptions... which may not represent real information.
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Hi dear researchers
I want to test removal of phenol by my peroxidase enzyme.
Could you help me with a ptotocol?
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There are many research articles exploring alpha glucosidase, salivary and pancreatic alpha amylase inhibition as a therapeutic target for reducing postprandial hyperglycemia (PPHG) in type 2 diabetes. Out of these three, which one is more important/superior and why?
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pancreatic alpha shares higher proportion as conmpared to salivary which makes it better. is this the main reason?
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I want to use in SOD activity.
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Dear Andrew , it was useful to me.
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Hi researchers
I determine optimal pH and temperature of immobilized enzyme activity. I want to know how pre-incubated it at different pH for pH stability factor.
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Hi,
the Question is how to best determine the Enzyme stability at various pH values.
Incubate enough Enzyme at the pH values to be investigated, take sample at various times, measure the remaining activity and then calculate the half-time as a measure of the stability. If you have high Background activity at some pH values it is more problematic.
Incubating the Enzyme at a certain pH and then neutralising it before determining remaining activity is also possible, as mentioned above, just Keep in mind that the Neutralisation process produces salt which can influence the activity and thus making a comparison between samples from different pH values false.
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I'm doing an enzyme assay, but I'm having trouble stopping the reaction so that I can get proper kinetic data. 
I can't heat because the substrates are unstable. 
I tried filtering, but it doesn't seem to stop the reaction completely. 
I can't use acid/TCA precipitation because it will mess up my analytical equipment. 
Are there any other ways of easily, and completely, stopping an enzyme assay?
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Exposing enzyme to low temperature and p H changes maybe helpful.
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I used NOBA, a chromogenic substrate for phospholipases, to measure (for 120 min) the absorbance (at 405nm) of PLA2 in crude venoms from different snake species. Later, I plot the absorbance versus time curve. As far as I know, I would need the exact concentration of the PLA2 present in each venom, but I don't have that information. Is there another way to calculate each PLA2 activity?
Technically, the absorbance versus time gives me already a clue, but I tested several venoms and the more I add the less "beautiful" my graph becomes so that's why I was hoping to calculate the PLA2 activity and plot it as a histogram to make it more visible.
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Hi, first you have to calculate the slope (velocity :∆abs/min) at the linear region. then if you know the extinction coefficient value (epsilon: e) of your product you can apply the Beer Lambert law(Abs= e c l),to convert it to μmol / min.Likewise ,you can also compare activities by unit (U) defined as the amount of enzyme that catalyzes the conversion of 1 micro-mole of substrate per minute,or even the specific activities by determining the proteins content using for example bradford method
Although,we can compare between the activities of your crude extracts ,the extraction method as well as the presence ofendogenous inhibitors can negatively affect the apparent enzyme activities .A purification is highly recommended to avoid this problem.
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Dear all,
When titrating a high affinity ligand I am getting Kd values that are very close to the total protein concentration in the system. My understanding on this matter is that, under such conditions, you cannot ignore ligand depletion and assume that your total ligand concentration is equal to the free concentration. My understanding is that a better way to analyse my data is to either decrease the total protein concentration to a value 5-10X below the Kd and repeat the experiment, but I have two questions: firstly, under these conditions, can I assume that my system has reached equilibrium within my assay time frame and get a reliable Kd? And second, would the Briggs-Haldane quadratic equation be a better way to get a reliable Kd under my current conditions (protein concentration close to Kd) without having to change my experimental set up??
Regards,
Ramon
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You can try to apply the quadratic tight-binding equation to your titration. This can extend the range of Kd that can be measured by the FP experiment, but not indefinitely, since the accuracy of the calculation becomes limited by the scatter in the data. Maximize data quality by averaging a large number of flashes/measurement and multiple replicate samples.
Lowering the fluorescent ligand concentration as much as possible is a good strategy to measure lower Kds. This is obviously limited by the ability detect the fluorescence. With a good plate reader, optimized geometry and filters, and a high-quantum yield dye like TAMRA, you should be able to measure 1 nM, perhaps even less.
To avoid loss of the ligand to surface adsorption, include 0.01% Triton X-100 detergent, or some similar detergent, as a blocking agent, or use non-binding surface plates (expensive).
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I'm designing an assay to test for release of pyrophosphate due to dihydropteroate synthase-like enzyme activity using malachite green reagents. One of my substrates is enzymatically catalyzed and one of the byproducts of this step is pyrophosphate; how can I eliminate the pyrophosphate background, because up until now I have been encountering a giant phosphate wall and the absorbance readings have been way off scale?
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If there is a buffer that has been commonly used by other researchers for this enzyme, you can start with that.
If not, the buffer should be optimized for the particular enzyme you are using. The optimization can include choosing the identity and concentration of buffering compound, the pH, the identity and concentration of salt, the Mg2+ concentration (ATPases require Mg2+ for activity), the reducing agent identity and concentration if needed, the detergent identity and concentration if needed, and a stabilizing excipient identity and concentration if needed.
Prepare concentrated stock solutions of all the ingredients you plan to test, and prepare buffer solutions by mixing them together and diluting with Milli-Q water to get the desired final concentrations. This gives you the flexibility to try many different combinations.
You also need to consider the substrates (one of which is ATP), and their concentrations.
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EC 1.14.14.1 is the accession number of CYP1B1 enzyme. I know that the first number '1' refers to the major class of the enzyme catalysis reaction, I want to understand the meaning of the rest of the numbers in this enzyme accession number.
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The answer to your question has been answered perfectly but it may interest you learn why this clunky system was introduced. From the knowledge of enzyme catalysis it was recognised that the reactions ctalysed could be identifiied s belonging to certain groups e.g. oxidoreductases, transferases, hydrolases etc.and each group was given a number (the first digit in the EC code. Other digits are added to denote the chemical group acted upon, coenzymes and a catalogue nnumber n a list of enzymes that all fall into a particular category. For example, Alkaline phosphatase is 3.1.3.1. The older literaure could be confusing because not all authors used the same name for a given enzyme. Editors of biochemical journals used to insist that the EC number was always quoted in submitted publications dealing enzyme work. This acted as an absolute identifier. The requirement seems to have lapsed somewhat but Suzanne has obviously come across the EC system.
Peter
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What's the DEPC water protocol for removing contamination from racks and other utensils in lab?
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Honestly, I've never heard of this. DEPC-treated water is not for cleaning.
DEPC is the substance used to purify the (already bidistilled) water itself, because DEPC inhibits nucleases; in the final step of preparing DEPC-treated water you autoclave it, and DEPC degrades, producing CO2 and H2O (I'm not sure exactly about its degradation products, but they shouldn't be chemically active). So, DEPC-treated water is simply very pure water without DNases and RNases, but it can't remove contaminations from any surface by itself. For this, you will need detergents, solvents, UV radiation, autoclaving and other cleansing methods depending on contamination type and utensils type.
For example, for cleaning lab surfaces from bacteria and grease we use 70% ethanol solution; for removing DNA traces ethanol is uneffective and you need either UV or chlorine-based cleaner (like diluted Bleach). We usually wash lab glassware manually with detergent, then rinse it with dH2O, seal with aluminium foil and autoclave.
However, nucleases are of the most persistent contaminants, they sometimes don't degrade even after autoclaving, so if you work with RNA you should use single-use plasticware labeled "RNAse free" and RNAse-free water as a solvent (you either buy this water or prepare it yourself using DEPC); you also may use specific products like RNaseZAP to clean lab surfaces. Make sure you work in a clean room, wear fresh gloves and don't exhale air in your sample.
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Dear Altruist,
I am running some QMMM jobs for finding the transition state and later on I want to do the IRC calculation if i can find the transition state.For me it is now a big problem to locate the transition state and to do the frequency calculation.The system has two layers.High layer I mean the quantum part contains the Adenosyl cobalamin cofactor and the low layer contains EAL enzyme.So the high layer contains 180 atoms which is treated by QM and the low layer contains 10000 atoms which is treated by MM.I am not sure is it possible to do the IRC calculation using gaussain for this type of big system.If anyone can give me some suggestions it would be a great help.
Thanks,
Mamun
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Hi,
i in a trouble like you. did any one answer you yet or not?
did you find the best solution or not?
please advise me.
thanks,
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Reactions are carried out in 50 mM Tris–HCl, pH 7.5, 5 mM MgCl2 and 100 mM NaCl using 2 µM cysteine desulfurase (Nfs) in presence or not of 2 µM SufE1. BSA 2 µM can be used as control. In addition to enzymes, pyridoxal 5′-phosphate (10 μM) and DTT (1 mM) are used for all reactions. Reaction is initiated by adding 500 μM L-cysteine for 30 min at 25°C. Then the reaction is quenched by adding 50 μl of 20 mM N,N-dimethyl-p-phenylenediamine dihydrochloride (prepared in 7.2 M HCl). The addition of 50 μl of 30 mM FeCl3 (prepared in 1.2 M HCl), followed by incubation for 20 min leads to formation of methylene blue, which is then measured at 670 nm. Na2S (1–100 μM) is used for calibration?
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You can prepare solutions of sodium sulfide of known concentrations and put them through the same assay in place of your cysteine and desulfurase. You should get some methylene blue formation, and you can measure its absorbance at 670nm. If you make a graph of absorbance vs. concentration of sulfide, you should get a linear plot (note: if your sulfide concentration gets too high, the plot will deviate from linearity due to formation of a dimeric methylene blue species). You can then run your assay for a specified amount of time, quench it by forming methylene blue, measure its absorbance, and back calculate the corresponding sulfide concentration.
One additional issue that you may want to be aware of is that sodium sulfide is VERY hygroscopic. If you want a very accurate sulfide concentration, you may want to consider a less hygroscopic sulfide salt (I believe lithium sulfide fits this description).
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 I know the substrate amount ( 5 different concentrations). Absorbance taken for 0 to 60 minute, rate of 1 min for total 61 readings. Enzyme amount was constant.
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From the linear part of the graph first plot Time vs absorbance for different concentrations of the substrate or product (as per your reaction). Slope of each concentration divided by molar extinction coefficient of substrate or product (as per your reaction) gives enz velocity (Vo) expressed in M/min
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While checking the effect of inhibitors on protease enzyme it was found that iodoacetate instead of inhibiting the protease activity it has enhanced the protease activity. I searched alot but could't find the possible reason for this. Can anyone explain what could be the possible reason for the same. Thank you.
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Yes it can as it can alter or stabilizes the active site residues (probably SNH). Once active site residues of protease stabilizes definitely prtease activity could enhance.
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Kynureninase is an interesting enzyme involved in many diseases such as schizophrenia and viral diseases, is these a synthetic ligand that can be used to compare the activity of certain compounds?
As a collaboration request: Where can I perform the test against this enzyme?
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Dear Sir. Concerning your issue about the synthetic ligands are available for the enzyme Kynureninase. The endocannabinoid system comprises of the endogenous ligands called endocannabinoids, the enzymes which synthesize and degrade them and the receptors, to which these ligands bind. Two types of cannabinoid receptors have been cloned so far, the type 1 and type 2 cannabinoid receptor (CB1 and CB2) from rat cerebral cortex and human promyelocytic leukaemia cells, respectively. They are class A GPCRs and belong to the Gi/o-coupled GPCR superfamily and they couple to Gi/o type inhibitory G-protein, thus their activation inhibits cyclic adenosine monophosphate (cAMP) production and stimulates mitogen-activated protein kinases (MAP). I think the following below links may help you in your analysis:
Thanks
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I want to interpolate the amount of product formed (as concentration or % of conversion) vs reaction time in a biocatalysis process. The fitting equation should have as (y) the amount of product and as (x) the reaction time. I thought to use as the fitting equation the integrated form of the M&M but I am not able to find the correct mathematical form. Or should I use another equation?
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Without any knowledge about the experiment parameters i can only guess which equation might suit your problem.
Having the product [P] as (y) results in some pretty ugly equations most of the time. I'm assuming no inhibitory effects.
[P] = [S]° - Km * W { X }
or alternatively (for the amount of conversion)
[P]/[S]° = 1 - Km/[S]° * W { X }
where [P] is the concentration of product P and [S]° is the initial concentration of the substrate S.
W is the Lambert-W function (or prodlog). You can try
X = [S]°/Km * exp[ ([S]°-vmax*t) / Km ]
as an argument for that function. I hope your program can fit this prodlog-function.
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i have the structure of unbound protein, ligand and protein -ligand complex?
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There is are differences between absolute and relative free energy calculation.
You can calculate the relative binding energy of two ligands using molecular dynamic simulations combined with an MM-GBSA or MM-PBSA approach (search for AMBER MM-PBSA.py). If you need very accurate results, FEP calculations might be the right choice. Alchemical transformations are implemented in NAMD using a dual topology system.
Calculating the absolute binding free energy is very computationally demanding and need a lot of experience. NAMD also has a tutorial for such calculations.
Docking scores are not that accurate (depending on the scoring function) but can be calculated with little effort. Knowing the correct binding pose would be helpful. Examples for free Software are Autodock4, Autodock Vina/QuickVina, UCSF Dock.
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Lignocellulose is a renewable biomass which is widely available in nature. It contains cellulose (glucose), hemicellulose (xylose, arabinose, mannose etc.) and lignin. How one can analyse a perticular lignocellulosic (wheatbran) biomass to quantify the above to detect the proportion of each of them. Is there any specific and simple methods are available to analyse. please let me know.
Biofuels, bioenergy, carbohydrates, redusing sugars, biocatalysis, enzymology, sustainable biofuels, renewable bioenergy, bioethanol, biotechnology, pretreatment
Thank you.
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I am getting Km value of a mutant which is similar to its wild type. How it is possible ? The residue is quite important for substrate binding. Is it possible ? please suggest !
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Hi there,
In the general case Km is definitely not Kd. So unless you know your favorite enzyme substrate couple obeys to the fast steady state equilibrium (kcat has to be much less than k-1), you can't consider Km as Kd. As Km=(kcat+k-1)/k+1=(kcat/k+1)+Kd, if kcat/k+1 is the main contribution to Km then Kd increase due to mutation might not be detectable at the Km level.
You also say that thee mutated aa is essential for binding . What is the mutation you made and are you sure it does abolish the contribution to the binding?
In fact it might be that the contribution of the aa is not that essential for the binding or might be compensated by other contacts between enzyme ans substrate.
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Hi,
I want to characterise by immunohistochemistry fibronectin, collagen I and III on a biologic matrix but the heat unmasking scrap my tissue. There for, I am in the need to a gentle and efficient unmasking method to preserve my tissue and reveal these proteins in it.
Any suggestion would be helpfull.
Thanks
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hi
I agree with Anna Answers.
good luck
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I have data from 2 different substracts (in 0 to 100 mg/mL) for the same enzyme at its optimum pH, temperature and time (all of them had a 20 min reaction).
The fact that this data is not related with time, makes it impossible for me to do kinetics (absence of vo).
Is it possible to compare this data in other way?
I'm a student in enzymology and I would appreciate any kind of help.
Thanks !
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Michaelis-Menten kinetics requires that the initial rates are measured if Km and kcat are to calculated. If you have not ascertained that all the rate measurements are initial rate measurements, then it will limit the types of analysis that you can do. You can still make semi-quantitative statements about the relative rates between the substrates, however (Example: After 20 minutes, a certain quantity of enzyme converted twice as much of substrate A to product compared with substrate B, when both substrates were presented at the same concentration.) I would advise you to convert the concentrations from mg/ml to µM or mM, to correct for differences in molecular weight between the substrates.
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I am performing an enzyme-inhibitor kinetics experiment with an incresing concentration of substrate and inhibitor. A few years ago, we used to follow a protocol which uses Lineweaver Burk equation ( a double reciprocal plot). But, recently, I am coming across a few research articles saying such linearization of non-linear data is obsolete and now we can calculate Ki' and Km and Vmax using other computational tools. The point mentioned in papers was the linearization methods were used when the computer-based calculation was yet to be discovered. So now, due to advancement in computation, L-B (double reciprocal) and Bowden plots are un-necessary. On the contrary, I still find recent research articles having  L-B plots based calculations
Can anyone guide me about this? If these methods are obsolete, what computational; methods or software can I use and how? Please mention some related documentation too in the answer.
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Hi there,
Linearization methods are not obsolete but they present specific disadvantages (for instance LB plot uses inverted axis which make the repartition of data points uneven on the graphics and error bar size is also greatly affected by this inversion). There are a lot of free sites on the web where you may run non linear regressions with your experimental data. This one has the advantage to propose an exhaustive list of non linear fittings:
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Hi,
I have a decrease of 1/3 to 1/2 in the gene expression of gluconeogenesis enzymes (Glucose-6-phosphatase, Fructose-1,6-bisphosphatase, Phosphoenolpyruvate carboxykinase and Pyruvate carboxylase) in mice liver between my test condition and my control condition and I would like to know if it can lead to a reduction of the flux of the pathway or not? Also is one of these enzymes more critical than the others in the pathway?
Thank you.
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In addition to what my colleagues have stated, One may actually claimed inhibition of gluconeogenesis however protein quantitation would brings out the beauty of the work and a better conclusion. Thanks
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Basically I have now two proteins that catalyze the same reaction, but with different affinity for their substrate. The first protein has a lower Km since is enough to visualize the reaction (using a scintillation counter) if I use a concentration around 10-50 nM. When I use the second protein I have to add at least 400-800 nM to be able to visualize the reaction and calculate the different slopes varying the substrate concentration to finally obtain both Km.
As far as I know the concentration of the protein is not relevant to calculate the Km (you only have use a concentration much lower than the substrate of the reaction in a Michaelis Menten-like case). Since my knowledge about enzymology is not very deep, I would like to hear of you that is perfectly possible to compare both Km even if I had to use different protein concentrations to perform the assay.
Thank you very much!
 
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Hi there,
You are right : as long as the enzyme concentration used in the assay is far less than the substrate concentration, your kinetics are OK for velocity determination. And you can perfectly compare the calculated Km values.
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Please colleagues, I need to know how to convert the measure of enzyme activities in plants (Ascorbate peroxidase and Glutathione reductase) from the SI unit of U/L to umol/L. I found that most literature did not report in the SI unit, so I want to convert to make the results comparable to other studies. Kindly assist.
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@Shapiro, thank you for the contribution. I appreciate so much!!!!!!!!!!
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I have optimised an assay with chymotrypsin and N-Succinyl-Ala-Ala-Pro-Phe p-nitroanilide substrate. Flavonoids at concentrations ranging from 0.01-1 mM, were then introduced to observe any affects they may have on enzyme activity. Of those investigated, catechin was found to increase activity. However, upon further reading, all research points to catechins having inhibitory effects on serine proteases, yet my results suggest otherwise. I was wondering if anyone (someone with a better understanding of physiochemistry/enzymology than myself) might have a insight or suggestions as to why this might be the case? Assay was carried out in 0.1M tris buffer pH 8.5. 
Thanks in advance! 
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Negative controls were ran with solvent (20% MeOh) minus flavonoid, the absorbance of the flavonoid wasn't subtracted, noticeably there wasn't any colour at 1 mM, but interestingly a similar increase in activity was also observed at 0.01 mM, a concentration where you'd think the absorbance of the flavonoid would interfere much less (this was observed with other flavonoids such as quercetin, which had a prominent yellow colour). But thank you, this idea will be considered as a contributing factor. I have been postulating 3 possible ideas: 1). flavonoid activate inactive chymotrypsin 2) assist in inducing conformational change during substrate binding. 3) Catechin acts as a nucleophile and facilitates regeneration/removal of products of interaction. This is of course up to interpretation and further input would be very very helpful. All the best!   
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Hello,
I've just started helping on a project requiring the extraction of high molecular weight DNA from Chlorella luteoviridis (green algae) for PacBio sequencing. My coworker has tried using a whole bunch of enzymes which she creates a solution of and incubates the algae/agar plugs in overnight for gentle digestion of the cell wall to occur. Unfortunately, the cell walls of algae are extremely variable between algae and thus her methods are not as successful on this new strain.
Currently she has tried 4% concentrations of cellulase, pectinase, hemicellulase, and chitinase all in one enzymatic mixture. Additionally, a subsequent incubation in lysozyme has been attempted overnight yet we are still getting poor yields of DNA.
Is there any advice on other enzymes/methods of cell wall digestion you can suggest? Thank you!
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Hi,
The attached article will be useful for you. All the best
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Enzymology, Molecular biology, Biophysics
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I'm afraid I don't know about them either, but a few things to think about are -is the protein you are working with purified from the natural source? - is it heterologously synthesised (which could lead to missing modifications, mis-folding, missing partner protein necessary for function)? -is the protein getting too old? -was it stored in optimal conditions?
Also, are you 100% sure you are using the right method to quatify a positive response? You have not provided any experimental details in your question... Are you following a published protocol?
I hope this helps,
-Anna
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Hi, I am interested in measuring nitrogenase activity in free living bacteria. Specifically i am interested in measuring through the 15N2 method which measures the organic nitrogen through mas spectometry. Nevertheless, I would l would like to know why it does not take into account  other forms of nitrogen in the supernatant or intracellular. For example Labelled amonnia.
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 Thanks, yes i assume that the organic nitrogen is the main product but i wonder about the inorganic nitrogen that could also be produced.
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During the optimum pH studies of an enzyme, should we change the assay buffers according to the pH or can we change the pH of the buffer itself
Example: For a given enzyme, the assay mixture contains 0.05 M Tris-HCl at pH 8.6. Now if I want to perform the pH stability or optimum pH, do i need to choose the assay buffer according to the pH i looking for, borate buffer for alkali condition and acetate buffer for acidic condition Or can I simply change the pH of the given 0.05 M Tris-HCl from 4 - 10?
Please suggest me the best or right choice
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Dear Yogi Affable,
You have not stated what pH dependence you want to study. It could be
-       the temperature dependence of he enzyme activity at a fixed pH
-       the pH dependence of the enzyme activity (from which you can also determine the pH dependence of kcat and Km) in the pH range where the enzyme is stable
-       the pH dependence of the enzyme stability by determining the rate of enzyme denaturation as a function of pH
the buffers you use must be used 1 pH unit above and below their pK-values as indicated by Nick Gee and Ozer Nazmi above. A detailed list of suitable biological buffers in the pH range is given in
where you also will find extensive information on the buffers but also that the enzyme activity also depends on the ionic strength I, and reference to important primary papers. It is however not explicitly stated that TRIS is not a suitable buffer for  enzymes that form covalent acyl-enzyme intermediates (hydrolases, transferases). The acylated enzyme can be deacylated by the uncharged TRIS amino group. 
Thus for simple Michaelis Menten kinetics the measured reaction rate v is
V(pH, T, I) = [EH] kcat(pH, T, I) [S]/(Km(pH, T, I) + [S])
where [EH] and [S] are enzyme and substrate concentration.
It makes no sense, and would be very time consuming,  to determine enzyme activities at zero buffer content by extrapolations as Ozer Nazmi suggests.
To study the pH dependence of the enzyme the temperature and the ionic strength must be kept constant in the studied pH range. In the internet you find a free suitable tool to prepare buffers of different pHs with constant ionic strengths.
Google for
Buffer Calculator – BioMol.Net.
A recent open access source on recommendations for  reporting enzyme data in the scientific literature, given in several excellent papers, you will find by googling for
Reporting Enzymology Data – STRENDA Recommendations and Beyond.
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I am trying to re-determine the Km of the enzyme I'm working with. For some reasons, I'm trying to do this using a coupled assay. I am having trouble with that, because the Km that I am finding is more than two times what it should be. I was told that it could be because of a reduced activity of the second enzyme in this coupled assay.
I am having a hard time understanding this statement. Can someone explain to me why a reduced activity of the second enzyme would give me a greater Km of a substrate of the first enzyme?
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I don't think product inhibition can be at play, here, otherwise it would be just worse in the non-coupled assay (where  the product of the first enzyme is not removed at all by the second, so presumably it can accumulate).
Other than that, I subscribe to Adam's comments. Twofold is a sizable difference, but not a huge one, especially if your 'reference' Km was obtained using a different technique, different experimental setups, different enzyme stock and possibly by a different operator. You should repeat your expts side-by-side using the two techniques and striving to match as closely as possible the reaction conditions, before assuming the difference is real. 
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best homogenization buffer for fish liver?
how to prepare?
suitable for most antioxidant enzymes?
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That depends on a couple of questions:
Are you interested in extracellular or intracellular proteins. For extracellular use Na+; for intracellular use K+. The physiological pH inside cells is somewhat lower (6.8) than that of the extracellular fluid (7.4).
If you want to use ion exchange chromatography, make sure that your buffer ion doesn't bind to the resin. I.e., for cation exchange use an anionic buffer (like HEPES), for anion exchange a cationic (e.g., Tris).
Osmolytes (glycerol, sugar, mannitol) may be useful to adjust the osmotic pressure, but not all proteins require them.
You may need some ions (Ca, Mg) or small molecules (substrates) to stabilise your protein. Don't forget protease inhibitors!
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My enzyme is a monomer but sometimes exists as homo-oligomer. The kinetic plot of the enzyme was hyperbolic. The current project on the enzyme involves incubating the enzyme with some compounds at 30oC for 1 h before initiation the reaction. Interestingly, the enzyme exhibited sigmoidal kinetic curve after the 1 h incubation. Although, few articles reported sigmoidal for the enzyme.
My major puzzle is, the later kinetic (with the incubation) study has a two-fold Vmax of the former (without the incubation), yet all reaction conditions, besides the incubation, are the same.
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