Science topic

Enzymes - Science topic

Biological molecules that possess catalytic activity. They may occur naturally or be synthetically created. Enzymes are usually proteins, however CATALYTIC RNA and CATALYTIC DNA molecules have also been identified.
Questions related to Enzymes
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I am preparing Enzymatic Antibiotic API by using Penicillin G Acylase Enzyme in aqueous medium but the problem is that after the condensation reaction is complete, the reaction goes backward by hydrolysis of the final product very fast.
It is very difficult to control the reaction. Can anyone suggest how to stop the hydrolysis?
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To add on to Francisco's answer... The equilibrium can be pH dependent and the reverse or forward reaction can be favored at low or high pH. One can drive the reaction in the direction they desire by adjusting pH.
A couple of things to think about..
1. Stopping the reaction once the condensation rxn is complete. I would recommend using small spin filters that work in a micro centrifuge (bench top) to stop the reaction by removing the enzyme, they come in different molecular weight cut-offs (10, 20 , 30Kda) etc...
2. Maybe the hydrolysis is non-enzymatic- removing the enzyme will help you understand if this is enzymatic or non enzymatic hydrolysis.
Hope some of these thoughts help..
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Generally, ATP content of bacterial cells is determined by using disruption of cells, and then ATP is quantified in the extract. This method can determine the ATP in a population. I would like to know about the ATP content of each cell by flow cytometry.
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Normally inhibition possibly can be differentiate into various category but when you go through different books or articles sometimes we get confused in these terminologies
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You are talking about two different enzyme kinetics here. When referring to enzymes obeying Michaelis–Menten kinetics then competitive, non-competitive and uncompetitive can be differentiate by the Lineweaver–Burk plot. Competitive inhibition affects substrate binding site so Km is affected(changes). Non-competitive inhibition does not affect the substrate binding site so Km is constant but Vmax is affected(changes). However in uncompetitve inhibition both the substrate binding site and the other side are affected so both Km and Vmax are affected (changes).
For allosteric enzymes, since they are made up of more than one subunits and have allosteric binding sites that can influence the substrate binding sites, they do not obey Michaelis–Menten kinetics and cannot be differentiated by the Lineweaver–Burk plot.
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Proteinase activity and whole lysate protocol question. Does anyone What about detergent and reducing agents?
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thanks Jose. The issue is to measure cysteine proteinases. So DDT will be needed.
Again many thanks
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Based on some articles I've found Xylanase is a good option, what about protease? What will be the necessary parameters to make the enzyme work in removing the melanin?
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I know where the active site is and thought about the measure fit tool or mergestructs plugin but I only have pdb files no psf and for measure fit I dont want to align my substrate with an existing part of enzyme. I want to place it in an empty space in the active site
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Why you are persisting on using vmd for docking? actually most of docking software are free and straightforward.
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I want to do subcloning of two genes (gene X and gene Y) in pcDNA 3.1+ expression vector. Gene X is having KpnI and NheI restriction site, gene Y having BamHI and NheI restriction sites.
Firstly, I cloned both the genes in 18T-Vector and got positive colony and confirmed by sequencing, then for subcloning I isolated the plasmid and digested it with respective enzymes, but after dual digestion, I am not getting my desired size of my inserts.
Gene X and Y both are approx 1600 bp but after digestion I am getting separate and clear bands at 1000 bp instead of 1600 bp.
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Ms. Ruheena: Like suggested by Tiziani, check to see if both of these enzymes are single cutters on your recombinant plasmid. Make sure they don't cut inside the genes!!! Additional sites for either enzymes may be present in the vector backbone, insert, or may be generated at the site of ligation. These are rare events, but may happen, especially if initially not considered . Also, rarely, some enzymes may start showing the star activity.
Unlike PCR cloning by way of poly T ligation, you may try introducing the specific restriction sites directly by designing forward and reverse PCR primers containing the desired sites at the 5' ends, amplify the genes of interest, and then clone directly the restriction enzymes digested PCR products into the expression vectors. Use error proof PCR enzymes,and then sequence verify the recombinants.
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I want to know whether I can determine the enzymes GPT, GOT (by GOT and GPT KIT) and total protein (by Lowry method) from fish liver and muscle dried at 60 C, or do I just use fresh tissues?
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For any enzymatic assay, the fresh tissue samples should be preferred for getting accurate activity measures. It is difficult to assay the enzyme activities in dried samples. Total protein can be estimated in supernatent of homogenized dried samples with appropriate buffer or NaoH , but the moisture content of the samples before drying is very much important for arriving the exact values.
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It might be oligomeric or multimeric enzyme, and the shape might appear due to co-operative behaviour of different subunits.
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You answered your question correctly. Your enzyme could be allosteric and may have multiple binding sites / multiple subunits. Binding on one site triggers activation on a second site. See Biochemistry text books on allosteric enzymes and their kinetics.
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Please share your experience. Thank you.
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We work on map kinases, PI3K and PP2A in the context of cellular transformation, carcinogenesis and drug resistance.
How to do a enzymatic activity assay with recombinant overexpressed enzyme?
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I want to overexpress an recombinant enzyme in HT22 neurons and measure it's activity (via LC/MS-MS measurement of the enzymatic product). The problem is, that this enzyme is also expressed endogenously in these cells. So when I extract protein/enzyme samples from the cells (in order to incubate with my substrate), I will always have a mixture of recombinant and endogenous enzyme in there which makes it impossible to determine the activity of *just* the recombinant enzyme...right? Or is there a possibility that I don't see?
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I agree with Michael' suggestions above - using a tagged version would allow either partial, specific purification of the recombinant protein and/or quantitation of the relative fraction of the recombinant protein and calculate the relative amount of functional protein from the total specific activity. In either case, the assessment is not perfect since - as Michael stated - the purification may not be quantitative and in the second case, the total expressed protein may not be functionally active. Another approach is to measure the activity of control cells which contain the empty expression vector - this should provide an estimate of hte native activity and subtract this from your recombinant expression cells. This requires care that the control purification steps are treated identically and that your assay is highly specific to your enzyme reaction with minimal interference from background reactions. In most cases, LC-MS-MS assays are highly specific, so background interference is not usually an issue.
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Recently I tested my kinase activity by using phos-tag, and I need a control to confirm the kit is working. Do anybody know where can I by the cheap kinase and the substrate together?
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My experience with phos-tag acrylameide is that it's highly protein specific. Its common for two similarly sized phosphoproteins to have very different rates of migration through the gel. The best negative control for a Phos-tag acrylamide gel is a sample of your substrate that is not phosphorylated. Unphosphorylated substrate should run faster (farther) on the gel than phosphorylated substrate. If you see no retardation of the migration of your substrate protein band upon treatment with kinase than you are either not phosphorylating your substrate or the Phos-tag acrylamide is not sufficiently slowing the migration of your specific protein. Unfortunately, if you do see a phosphorylation-induced retardation in migration of a different substrate-protein that doesn't tell you whether or not your substrate-protein is being phosphorylated. In my opinion, the better positive control would be to use radiolabeled ATP as your phosphodonor and monitor where the radiolabeled band appears in the gel with film or a phosphoimager.
On a practical experimental note you have much more flexibility if you pour your own Phos-tag acrylamide gels than if you use the precast gels. You will likely have much better looking and more quantifyable gels if you spend some time optimizing Phos-tag, cation, acrylamide, and crosslinker concentrations than if you use the stock precast gels.
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During storage of crude extracted enzyme I observed that the number of bands are varying during gel electrophoresis.
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Enzymes survive better in buffers that have a certain pH. I don't know your optimum buffer conditions. Straight water is not advisable. To look at auto catalysis there would be two methods. The enzyme has to be pure. First look at degradation over time with SDS PAGE gel . Second, do an enzyme activity assay to see if your protein is loosing activity. If it is a crude preparation, then any pro teases present can degrade your enzyme. It is best to add a protease inhibitor cocktail mix. You can buy pills, premade by many companies. But the key is to first find conditions where your enzyme is stable. So you will need to empirically test many buffers, etc if the work is not published yet.
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I am needing to create liposomes from 3 phospholipids for use with membrane associated cytochromes. Thus far manufacturers instructions (Avanti polar lipids) has not yielded acceptable results. Can anyone recommend a book, review or protocol paper that highlights in detail various methods associated with liposome formation?
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Not necessarily. However, it is likely safe to also sonify at 25 degrees if you make sure the temperature does not increase further. For saturated diC12PC, Tc is around 4 degrees Celsius, for unsaturated diC12PC it would be less. DiC12PS would be above 20 degrees for a pure diC12PS bilayer, but lower for a mixture.
For Tc of lipids, check out the Handbook of lipid bilayers, 2nd edition by Derek Marsh, which appeared earlier this year.
If you replace the native lipids around enzymes by synthetic lipids, the activity could go down, as some native lipids occupy 'hot spots' on the enzyme to ensure its stability. This could possibly be the case for cytochrome reductase as well. It probably depends on reconstitution conditions.
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What are the common or frequent structural features of the protein which make it work as an enzyme. Like the presence of certain kind of electronic environment or any chemical group or frequently occurring particular amino acid etc.
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Hi Amar, the structural features of a protein that make it an enzyme are as diverse as the substrates of those enzymes (very diverse). Some general rules are that enzymes are often globular instead of elongated like structural proteins, and they frequently have a pocket on their surface with complementary charge to their substrate. Probably the best way to find out if new protein is an enzyme would be to use homology modeling with online tools such as Dali (with its structure) or BLAST (with the amino acid sequence). If the structure or sequence looks like a known enzyme then it probably is as well.
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Low affinity but high catalytic activity?
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There is plenty of such a examples, but I suggest to look for it rather in patents, than articles...
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My protein of interested was cleaved into 2 fragments in the cells. Is there any website can predict what enzyme does it and where it cleaves?
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I don't think there is a website. But you can use different protease inhibitors in your system to narrow things down. If its is a type I or II membrane protein, then things get narrowed down easiLy. If it is cytosolic, it could be a caspase, and there may be consensus type sequences for those. So depending on the sequence of the protein, maybe you can classify what it is, and this will help you determine what type of protease. In terms of a cleavage site, I would look for a consensus amino acid sequence between species, that would be near to where your two fragments are cleaved. One could predict where from the MW of the fragments.
One can buy serine, cysteine, metalloproteinase and other inhibitors from sigma or Roche diagnostics. Also one can do mass spec to find the cleavage site, and this may help you. But I think the protease that does the cleaving can be determined sometimes by what type of protein is being cleaved.
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This protocol is reported in a journal written by Paolucci-Jeanjean et al. (2000).
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Sometimes with SDS, one has to boil the sample to completely denature. I would recommend hIgh concentrations of acid or base for instant quenching. Or add proteinase K to the SDS as Jovencio mentioned.
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Seeking advise on an enzyme analysis method
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Iam also same doubt if you know that kindly help me sir.
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Enzyme-surface interaction
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Hello!
In my  opinion you should freeze the boundary atoms of your system by imposing heavy atomic mass. The interesting reaction in your case will occur within the system. The appropriate base set to describe quantum mechanically the system must be chosen for the rest of the system. I hope it helps you. 
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I want to study on genetics diversity of Diospyros rumphii Bakh., in Sulawesi, Indonesia. I has searching literature related to D.rumphii and I have got limiting information about it, especially about it's genetics information.
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No one enzyme is sufficient to genetic diversity. You will have to utilize all the available information on various Isozymes, and then select 10-14 enzymes which you could perform in your lab. The best book will be Isozymes in Plant Biology by Soltis and Soltis.
If you need some more help feel free to contact me. Even my review paper will be of great help to you.
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Could you please clearly explain the extinction coefficient and its importance in enzymatic assays?
Should we consider it for all type of enzyme assays?
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An extinction coefficient along with a particular wavelength that can be used depends greatly on the enzyme (or actually its substrate(s) and/or product(s)), so no, there is no particular wavelength and extinction coefficient that applies for all enzymes in general. These vary between cases and to find out a suitable method of following your reaction you should look into literature.
Now, if there is a wavelength that can be used to monitor (directly or indirectly) the conversion of substrate to product, knowing the extinction coefficient basically allows one to quantify the conversion i.e. how much product you actually are forming and what you ended up with. After all, the turnover of substrate to product is always in an equilibrium, so if your [S] = 5 mM, in the end [P] (which you are measuring) is actually determined by the equilibrium constant. Bare in mind that when using a marker enzyme with a certain ext. coef., one also needs to be aware of the stoichiometry of the reaction. Also, another way is to measure the disappearance of S. In this case the extinction coefficient allows you to determine the remaining amount of substrate after reaching equilibrium. But, just to answer simply in the context of Michaelis-Menten kinetics, you NEED the extinction coefficient to convert your absorbance units to concentration units if you want to be able to produce Michaelis-Menten parameters.
A typical example: an enzyme oxidizes substrate S to product P and as such reduces NAPD+ to NAPDH in the reaction. As NADP+ converts to NADPH, an increase at 340 nm is observed. The extinction coefficient of NADPH is 6.22 mM-1 cm-1, which can be used to directly quantify the formation and at the end the total product formed during the reaction, assuming the concentration of substrate S is limited in solution and NADP+ is available in excess. This is a very straightforward example as the reaction stoichiometry is simple. Now as you have your concentration series and reaction curves, you use the extinction coefficient together with the Beer-Lambert law (A = ε c l) to work out your conversion rates in quantity over time, like mM/s. Only after this you can construct you MM plot and work out the kinetic parameters.
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I have used French press to disrupt bacterial cells. After ultracentrifugation, the supernatant was used for determination of enzyme activity (PQQ-Glucose dehydrogenase).
Now, I would like to determine the effect of some inhibitory compounds on this enzyme. Should I also purify this enzyme or is the crude extract enough?
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The first question you have to answer is how reliably you can measure the activity of your enzyme in trhe crude extract, regardless of inhibitors. If you can get good numbers without undue interference from other activities in the extract, then you need to think whether there is any reason why other components in the crude extrtact might either destroy your inhibitor or just bind and sequester it making it unavailable to your enzyme. Assuming the answers to those questions are provisionally encouraging, give it a try. If there are already known inhibitors, it would be a good idea first to see if you can demonstrate that predicted inhibition using the crude extract. If not, then just try. It could give you a convenient rough screen to select those compounds that are worth looking at further. However, for good quantitative data for those compounds you eventually decide to concentrate on, I would always want the final experiments to be done with the pure target enzyme
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Drugs either bind to the substrate or cofactor binding site but which one is better to target?
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As Dr. Maria says, each case should be considered carefully.  But, there maybe some ground rules....For e.g., If an enzyme is part of a gene family, then its active site is conserved, and targeting active site of 1 family member may be futile. But, if an enzyme is not part of a family, or if its an isoform with tissue specific expression, then one could design an inhibitor against active site and this should work . Hope this helps!
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Recent advances in modern biotechnology have revolutionized the development of new molecules. Microbes are relatively easier sources than plants and animals, and their enzyme productivity can be improved using genetic modification techniques.
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A team of researchers from Newcastle University have identified an novel enzyme NucB from the microbe thriving on seaweed. This enzyme is a potential weapon against biofilm producing pathogens causing numerous diseases. It has been found effective protection against dental caries and can also help in curing sinusitis. Efforts are being made to incorporate this potential bioproduct into commercial products like toothpaste etc....
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I am currently looking for enzymes that accept halogenated substrates. They are likely to be found in species that live in halogen-rich environments. To scan databases for such enzymes it is very useful to know which species/phylum is associated with halogens.
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You might wish to consider taking a look at some reference books or reviews on extremophiles (or applications). Check the examples below, but you'll likely find better suited sources in their reference list:
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Homology modelling is the type of comparative - template based modelling. What can be the structure that homology modelling estimate - Primary, secondary, tertiary and quaternary.
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Template dependent modelling may also be dependent on the availability of resolved structure that are deposited in the protein data bank. For a quick search of homologous sequences whose structure have been resolved, you can try FUGUE (http://tardis.nibio.go.jp/fugue/prfsearch.html)
The hits that you get here may not necessarily have 100% homology with you sequence but can serve a start template. You can however also model your molecules on multiple templates using tools such as Modeller http://salilab.org/modeller/ to get a more accurate model
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Do you know some enzyme that consequently isomerases and oxidases a substrate? what class would it be? Oxidase?
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Ketol-acid reductoisomerase (EC 1.1.1.81) is another example. In the non-physiological direction there is an oxidation (by NADP+) followed by an isomerisation.
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I determine the enzyme activity in fish liver and muscle by using the diagnostic kit. This mentioned in kit that after calculation the activity is in IU/L, since the kit is designed for serum analysis. I used the same kit for enzyme analysis in tissues, therefore I need to convert the unit IU/L to mg/g. Could anybody help me out?
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You can convert the IU/L of your enzyme into IU/g wet fresh tissues very easily knowing the volume of extract you obtained from 0.2g of tissues, dividing the total units (IU/Lxvolume of extract) by 0.2. You should also measure the protein concentration in your samples. At this point you can also express your results as IU/mg of proteins, dividing IU/L by mg/L of total proteins. The usual protocols for protein determination give you the mg/ml of proteins but you can easily calculate the mg/L. I think that to express the enzyme activity in a crude extract as U/mg of total protein is better than U/g of fresh tissue.
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We have elastase-high purity from EPC company, but the dissolve methods are not clear. Some companies suggest to dissolve it in alkaline first then reconstitute it in the media you want, but I need to dissolve the powder and store it in fridge for further use, so what's the optimal method to dissolve and store this elastase?
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From the specs I read from 3 companies, some store in tris buffer, pH 8 after reconstitution from powder. This seems to work. One method called for adding H20 to the lyophilized powder. I think they stored at 2-8 degrees. As a powder, it is stable for months at 2-8 degrees. I will keep looking.
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I recently purchased a dansyl-fibrin glycoppetide from a US based company. The company has not been helpful at all so now its time to try ResearchGate.
I have tried pretty much everything for dissolving the lyophilized peptide - Nothing seems to work.
Here is what I tried so far - If you can chip in with more suggestions it will be much appreciated:
- 25 mM Na-P pH 7.5 + 150 mM NaCl
- 12.5 mM Na-P pH 7.5 + 75 mM NaCl + 50% DMSO
- 12.5 mM Na-P pH 3.5 + 75 mM NaCl + 50% DMSO
- 25 mM Na-P pH 11.5 O/N on blood turner
- 50% methanol O/N on blood turner
- 50% methanol + 10 min in ultrasonic waterbath
- 50% acetonitril O/N on blood turner
- 100% DMSO O/N on blood turner
- 100% DMF O/N on blood turner
All of the above at Room temp and 50C
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Hi Jonas,
Dansyl chloride is unstable in dimethyl sulfoxide, which should never be used to prepare solutions of the reagent http://en.wikipedia.org/wiki/Dansyl_chloride
Try trifluoroethanol, hexafluoropropanol, deep eutectic solvent from choline chloride+urea in a 1:2 mole ratio. If nothing helps, check decomposition on hotplate - it may be inorganics.
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Directed Evolution is very complex and many techniques were used, and it is not easy to just list them here shortly. Basically you need a mutagenesis technique and afterwards an assay to identify your desired variant. Enzyme production and cloning methods are also involved.
- mutagenesis, random: many techniques: most popular epPCR and uncountable derivates, shuffling and again uncountable derivates
- screening / selection: always a new methods has to be designed, depending on the enzyme activity and task
You can check a lot of very nice papers form the groups of Frances Arnold, Manfred Reetz & Karl E. Jäger, Uwe Bornscheuer and many many more.
Here are some new articles of my old group:
Bornscheuer, U.T. (2013), Protein engineering as a tool for the development of novel bioproduction systems, Adv. Biochem. Eng. / Biotechnol.,
Davids, T., Schmidt, M., Böttcher, D., Bornscheuer, U.T. (2013) Strategies for the discovery and engineering of enzymes for Biocatalysis, Curr. Opin. Chem. Biol.,17, 215-220
Bornscheuer, U.T., Huisman, G., Kazlauskas, R.J., Lutz, S., Moore, J., Robins, K. (2012) Engineering the third wave in biocatalysis, Nature, 485, 185-194
If you need more information, please come back to me, Marlen
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Most of data I read about alpha klotho assays ELISA kits indicate inconsistency and wide variability. I have frozen samples that I wish to check human soluble Klotho levels in them but not sure what is the best assay to use, if any.
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we used eastbiopharm klotho kit and the result was good
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I digested pcr products by MboI enzyme, expected 2 bands one at 296 and another at 72bp. I got respectively in few samples which means they are resistant, the wild type will not show any cut bands. But I got 3 bands in few samples which have two bands also (296bp+72bp). Is this due to what problem? However I am planning to keep again for the problematic samples.Gel picture as follows: 1 well 100bp ladder, 2 nd well is uncut pcr product,3-12 are resistant type, and 13 is wild type. In few wells there are 3 bands which are highly unexpected.
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Did you try adding more enzyme and digesting longer?, since incomplete digestion is the simplest explanation. What DNA are you using as a template for the PCR, could the DNA have a mixture of WT and resistant genotypes?
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Are there specific cocktails of inhibitors used in assaying liver alkaline phosphatase? What is the role of the amino propanol buffer, and can Tris be substituted? Does just running the reaction at alkaline pH exclude the other phosphatases in sera from reacting with pnpp?
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Hi Marsha. I thought that this would be a fairly easy thing to determine, but there are a boatload of papers describing how to do this with various methods:
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I have several amino acid sequences of enzymes, and i would like to extract features of these enzymes. Does anybody know what software or online platform I could use?
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Hi Islam,
I would start with Blast, it should be straightforward to determine the enzymatic class from the answers there.
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Aurora A, B, C are collectively known as pan-Aurora. Why?
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"Pan" is another word for "all", usually associated in medicinal chemistry with non-specific inhibitors. For example, a pan-Aurora inhibitor inhibits Aurora-A, -B and -C with approximately equal potency.
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Need to produce food enzyme using the strains.
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Obviously, viable industrial standard bacterial strains optimized for high density growth and large scale protein production will not be distributed without charge by any commerical institution...
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Doing fluorescence analysis, I've identified a peak related to a NADH-dehydrogenase complex, but I don't know which type of dehydrogenase is. Is there any kit I can use for this? Or any other protocol?
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Do you fractionate as a part of your protocol? If so can you see the protein(s) on a gel? Is there more than one? My immediate inclination is to MS/MS the fraction, identify proteins and look at their functional annotation to see if there is any NADH dependent anzyme present, or BLAST search with the MS/MS sequence to find an annotated homologue from another organism
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In some ligand/enzyme systems I have found very good correlation between the in silico docking of ligands in the active-site of an enzyme (i.e., calculated interaction energy score) and the activity of ligands in an in vitro enzyme assay and a cell based assay.
However, recently I have seen disjunction between those three assays. While the most potent drugs in silico (based on interaction energy score) were still the most potent in vitro and in cell culture, some compounds that have good in silico scores fail completely in an in vitro enzyme assay, but are active in a cell based assay. Obvious issues include the modeling software and force field (I am using a tested software package-Discovery Studio with CFF) and the solubility of the ligands in the in vitro assay.
I was wondering what your experience is in seeing ligands that were modified within a cell, in a cell-based assay, into a more soluble and/or potent drug since that would explain some of my results. Of course the ligands could be hitting targets other than the enzyme I am studying, but the ligands are highly chemically related with only single side-group modifications (i.e., a H3CO- for a HO- at the same position on a ring). Thank you for your feedback.
UPDATE: part of the problem it turns out was that the compounds were not stable to freeze/thaw in 100% DMSO, one freeze/thaw reduced activity, and a second freeze/thaw reduced activity by up to 90%. This affected the in vitro assays, however, the person doing the cell culture assays was not putting the compounds through multiple freeze/thaw cycles. First time I have seen this issue with compounds stored in DMSO.
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i have also encountered similar situation. on further literature survey I found that
1. IC50 or Ki values are more correlating then % inhibition.
2 docking score cutoff - only enhances probability of getting the hits
3. for a given analogue series correlation may not be established may be due to solubility issues. change in binding conformation of proteins, Temparature and ionic strength of the buffer which will influence entropy. (which is often neglected in docking software)
4. very often we neglect invitro assay principal. we should consider any coupled assay involved if so what is the influence of inhibitor on second reaction should be addressed. influence of inhibitor on quenching in Fluorescent based assay, is inhibitor forming chelates with any ions which are critical for assay etc. therefore IC50 value is more reliable then % inhibition
I have listed this as possible reasons. I appreciate if any one encountered with such case shares there view in this regard
Does anyone know the best method to measure lignin peroxidase activity?
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You may refer to one of my publications i.e in Enzyme and Microbial Technology .Comparison of two assay procedures for li p
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How to confirm it is an endonuclease or exonuclease and how to find it out?
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Try NEBCutter V2.0. Its good.
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Restriction enzyme
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You have some options: gel filtration in Sephadex G-25 (fast and efficient), a mixed-bed resin or dialysis. If you choose desalting using Sephadex, GE and other companies (Pierce, Thermo Scientific, etc.) offer several columns ready to use.
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How should I prepare the enzyme substrate solution and when I will use a detection antibody with horseradish peroxidase conjugate?
Thanks
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Product Description
3,3’,5,5’–Tetramethylbenzidine (TMB) is a chromogenic substrate suitable for use in ELISA procedures, which utilize horseradish peroxidase conjugates. This substrate produces a soluble end product that is blue in color and can be read spectrophotometrically at 370 or 655 nm. The reaction maybe stopped with 2 M H2S04, resulting in a yellow solution that is read at 450 nm.
Each tablet contains 1 mg of TMB substrate.
The product is available in packages of 50 or
100 tablets. Custom packaging and bulk purchase information available upon request.
Precautions and Disclaimer
This product is for R&D use only, not for drug, household, or other uses. Please consult the Material Safety Data Sheet for information regarding hazards and safe handling practices.
Preparation Instructions
0.05 M Phosphate-Citrate Buffer – Dissolve one Phosphate-Citrate Buffer Tablet (Product No. P 4809) in 100 ml of deionized water with stirring to yield a 0.05 M phosphate-citrate buffer, pH 5.0.
OR
Add 25.7 ml of 0.2 M dibasic sodium phosphate (Product No. S 0876), 24.3 ml of 0.1 M citric acid (Product No. C 7129), and 50 ml deionized water. Adjust pH to 5.0, if necessary.
TMB Substrate Solution – Dissolve one 3,3’,5,5’–tetra- methylbenzidine tablet in 1 ml of DMSO and add to 9 ml of 0.05 M Phosphate-Citrate Buffer, pH 5.0. Add 2 μl of fresh 30% hydrogen peroxide (Product No. H 1009) per 10 ml of substrate buffer solution, immediately prior to use.
OR
Dissolve one 3,3’,5,5’–tetramethylbenzidine tablet in 1 ml of DMSO and add to 9 ml of 0.05 M phosphate- citrate buffer, pH 5.0, containing 0.03% sodium perborate (capsules, Product No. P 4922).
Stop Solution - Reaction may be stopped by the addition of 50 μl of 2 M H2SO4 per 200 μl of reaction mixture.
Storage/Stability
Store the TMB tablets at 2–8 °C. Protect from heat, light, and moisture. Allow tablets to reach room temperature prior to use.
References
Bos, E., et al., J. Immunoassay, 2, 187 (1981).
PCS,MAM 01/05-1
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I am using IgG sepharose to purify a protein A fusion protein. I'm planning on using Endo H to deglycosylate my protein before apply to IgG sepharose resin. I read about the glycosylation of IgG could be critical, so does the Endo H in sample affect the binding or damage the resin?
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Thanks Jan, I have tried this idea and Endo H did not able to affect the binding under IgG sepharose binding conditions (pH 7.5, 4C). The glycan in IgG Fc region is buried so it may not accessible to Endo H at all.
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If I have the data for an enzyme substrate concentration and both reactions rate with inhibitor and without, how then I calculate the Michaelis constant numerically? Some sources showed that this is an alternate way of calculation. If this is the case can someone specify which method is better for this task.
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Use Y = aX/ X + b where b is Km. this equation relates to v ( initial velocity) = Vmax* S/ S + Km. so X is your substrate concentration and y is your initial velocity. a is your Vmax or value where your hyperbola levels off. Km is your x value where the Vmax is half the rate of the value where the hyperbola levels off.
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Can anyone advise of which tubes to use at higher speed (up to 22000g)? The supplier literature is rather contradictory. Many thanks
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Hi Nikhil, Didi and Gerard
Many thanks for your answers. Gerard, I finally made contact with a specialist at Thermofisher this morning who confirmed that regular high quality falcon tubes should be fine as long as they are fairly full at high speeds. I am about to test this but from your experience it sounds as if this should be fine. I hope the adaptors don't stick!.
Best wishes
Andrew
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This SOD is from Sigma cat#S5395.
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Indeed store aliquots of the solution at -20°C. In a cell culture I don't really know, but I have had some success when adding it fresh every 24h. It wasn't brilliant though.
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I'm trying to characterise mutants of a particular enzyme. I have checked the kinetic parameters (KM, Kcat), the stability (Tm Shift) as well as the mass. I don't seem to find any significant changes in the substrate profile and the KM ranges between 2-12 for the different mutants. What difference in kinetic parameters is deemed as a significant change in activity? And are there other techniques that I can use to characterise these mutants or is it safe to assume that these mutations don't have any effect on the catalytic activity of the enzyme?
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You can do lots of experiments to see if there is difference between native and mutant enzymes. As already suggested you can do pre-steady state kinetics, pH optimum, temperature optimum, irreversible thermal inactivation studies, secondar structure melting temperature using Far-UV CD, Tryptophan intrinsic fluorescence spectral analysis and melting temperature, differential scanning calorimetry and melting temp. I am not aware of your enzyme or substrate but may be it does not show difference with one substrate but may show difference with another substrate. You can try Natural substrates using isothermal calorimetry (iTC) if its difficult to assay by conventional colorimetric assays. May be your mutant is inhibited or activated differently etc.
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Partial purification of Protease Enzyme
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Dear Dipankar,
You need to first find out the molecular size (usually in terms of kDa; kilo-Daltons) before you choose your dialysis bag/tubing. The pore size of the dialysis bag should be smaller (in terms of kDa) than your desired enzyme. Then you can search online for the dialysis bag/tubing you need. I'll be using the dialysis tubing purchased from Sigma-Aldrich, just for your information.
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Cysteine protease
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I’m working with spores of Bacillus subtilis but I have some problems with it. I want to attach an enzyme to the coat layer of spores via covalent or adsorption methods for surface display of the enzyme. I have tried many ways (EDC/glutaraldehyde/different pHs and …) but no protein binds or it might be too low that I can’t detect it. On the other hand when I use fluorescent antibody conjugated with FITC to see if any protein has attached, all the spores, even spores that are not treated with protein, become fluorescent.
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have you tried EDC /NHS mediated linkage of non specific antibody-FITC to spore and compare with non specific anitbody-FITC binding to spore.... is the covalent linkage still inefficient? and do you still see nonspecific Fluorescence without linakge?
if the conclusion is inefficient linkage, i would suggest checking the free reactive carboxyl groups on surface of spore (necessary for efficient EDC/NHS treatment), so playing around the pH for reactive carboxyl group will help in increasing the efficiency of the process.
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I need the details about it due to method, I want to specify enzymes activity in bats during the winter.
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The pH of the digestive tract variies depending on feeding status. During feeding process, meaning a filled stomach and digestive processes going on, I could measure pH 4 till 5 for Pipistrellus. An empty stomach should demonstrate a lower pH (about 2 till 3), but I do not have any detailled information about that. For the intestine, I measured (again Pipistrellus) for the duodenum pH5 till 6, where as jejunum, ileum, and colon/rectum was about pH 6.
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I am looking for a good antibody against a lysosomal enzyme, which works for immuno-EM in human cells (HepG2, HeLa). Could you please advise on any?
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I am not familiar of one but one of my collaborators have done Cathepsins, i.e. Karen McGuire in Ohio (Cleveland). However I have no idea where she is as I am in UK  and the lab she worked was dissolved in USA. Please try to ask Summa Health System in Akron Ohio for her contact
How to perform a Hill's Equation curve fit for an interventional study?
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Classical dose dependent
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What do you mean by an interventional study? What are your variables thare are being graphed?
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I am trying a method to optimize the yield of a PCR reaction and one of the solutions would be to destroy the competing bacterial DNA with this enzyme. If it works, it is worth it to import it. If it doesn't, I wasted at least 90 days. So if you can lend me some to try, I could even agree to pay for it.
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Better to use a 4 base cutter like DpnI or Sau3A. Easily heat inactivated too - can add either directly to PCR reaction and incubate, before adding template
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It is often speculated that cellulases need mechanical energy in order to step from one cellulose chain to another. They get stuck on to the chain and thereby become inhibited from moving on unless mechanical mixing can stop them from being reversibly inhibited by their tight binding by their Cellulose Binding Module (CBM) and the cellulose chain or to solve “traffic jamming” by several cellulolytic enzymes. Does anyone know the necessary mixing intensity, free fall intensity or something like this, in order to answer this question? Is there something about the effect of shearing on the hydrolytic activity of cellulases?
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In our experiments we do find that mixing can increase activity on some insoluble substrates but I think this is simply due to giving the enzyme better access to the substrate not do to sheering. However, if the mixing is not done right substrate can end up on the side of the tube and decrease the rate.
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Some microorganisms can produce enzymes such as lipase, elastase, protease. So how can we detect the activity of these enzymes in case they have been presented in our petri dish?
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Dear colleague,
when you need to test the ability of a certain microbial strain to synthetize lipases, it is necessary to include some triglyceride in the culture medium. One medium which is excellent for that purpose is: 8 g/L trypticase peptone, 4 g/L yeast extract, 3 g/L NaCl, 20 g/L agar and 10 g/L tributyrin. You have to emulsify with an ultraturrax prior to sterilize in an autoclave. The presence of lypolitic activity is detected by the appearance of a transparent halo around the colonies.
This assay is indicated for the detection of esterase activity. If you want to confirm the presence of a "true" lipase you have to use a long chain triglyceride. In this case, you have to use the following medium: 8 g/L trypticase peptone, 4 g/L yeast extract, 3 g/L NaCl, 20 g/L agar, 30 mL/L of olive oil and 2 mg/L rhodamine B. The presence of lipase activity is detected by the appearence of orangish fluorescent halos.
I hope that the protocol can be useful for you.
Best regards
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I've already tried by using a comparison of untreated and treated sodium borohydride. Treatment with NaBH4 reduces shiff base to amine, a step that makes trypsine "jump" the catalytic lysine cut and get, once run in the MALDI-TOF, a full piece comprising the previous + the following (peptides) + PLP. Compare to the untreated tryptic digest spectrum, and you'll get the differential peptide which ends with the catalytic residue.
Very easy to say but actually there are a lot of issues because of the high number of lysine residues (42!), other impurities and eventually fragmentations at source (if so, it's too bad!).
Can anyone give me an advice or suggest another technique (which is not alanine scanning)?
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Don't need to scan: you need to know which PLP enzyme fold your protein has. If the sequence alone doesn't tell you that, run a threading program (fold prediction), which will. That will probably tell you, from the structure-based alignment, which lysine it is, or at least narrow it down to a couple. Mutagenesis will confirm.
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I study the ACACB enzyme and need to see how efficient it is without working with radioactive materials.
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By ESI MS methods you can measure CoA derivatives after acid quench
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Can anyone recommend a useful concentration?
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Please note that mouse and human mast cell chymases are more susceptible to inactivation by the protease inhibitors in fetal calf serum than naturally occurring and recombinant tetra-forming tryptases. Thus, you probably will have to use serum-free conditions if you are going to exposure your cultured cells to recombinant hChymase-1 or one of its mouse orthologs (e.g., mMCP-4). In regard to tryptases, the mast cells in mice and humans express three tryptases that are encoded by the TPSAB1, TPSB2, and TPSG1 genes. The human TPSAB1 and TPSB2 encode nearly identical tetramer-forming tryptases which are resistant to protease inhibitors . Having said that, the Human Genome Consortium has identical hundreds of allelic isoforms of these genes that change the amino acid sequence and thereby enzymatic activity. This means that the "hTryptase-beta" preparations used in the 1990s and 2000s that were derived from pooled lung and skin biopsies from many humans actually contained a large number of functionally distinct enzymes. Thus, please be aware of that not every recombinant "hTryptase-beta" will effect your cells in the same manner. In regard to the transmembrane tryptase/tryptase-gamma/Prss31 which is encoded by the TPSG1 gene, please not that this tryptase is also rapidly inactivated by A1AT and other protease inhibitors in serum. Thus, you will have to culture your cells in serum free medium if you use that recombinant tryptase.
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I want to immobilize ammonium monooxygenase enzyme on conducting polymer.
What is a good procedure?
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It depends on immobilization and storage conditions, from few days up to few years
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The reaction is an oxidative decarboxylation catalyzed by IDH.
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When looking to the thermodynamics of the reaction, it's very improbable ... decarboxylations aren't reversible in general, nor oxidations
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Applied nanotechnology in aqua diets
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Dear Dr. Randeep,
Thanks for your critical suggestions and comments
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I have been expressing a cytoplasmic sulfatase (from Pseudomonas aeruginosa) in E. coli DH5-alpha and have found significant enzyme activity in the culture media (either LB or M9 minimal media with kanamycin) after cells are removed by centrifugation. Cultures are grown at 37 °C and handled at room temperature. There is no known signal peptide coded in the expressed gene and expression is regulated by a T7 promoter and IPTG. Has anyone else observed activity leakage from their E. coli cultures? Or, are there any ideas how this occurs?
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We have cloned a wide range of Gram-positive enzyme genes (cytoplasmic or secreted in the original host) in various E. coli cells and always can detect the activity around the colonies on plate or in the culture supernatant. There is also no difference if the signal peptide is present or not - E. coli is not supposed to actively secrete foreign genes anyway. The majority of the recombinant protein is in the fraction with the cells (after centrifugation), even for those proteins having a Gram-positive leader peptide - so what is the heck? Just isolate the protein from the cells.
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I am working on one protein. I need to screen the inhibitors against it. The enzyme activity was known based on the fluorescent reading of the product. As I increased the concentration of substrate, the corresponding fluorescent reading increased to a certain level, later it was reduced. I had the feeling that there was some feedback inhibition. In this case which concentration of substrate should I use to perform my assay?
As per my knowledge I got to use the Km (Vmax/2) value of the substrate. I wonder is it a right choice to perform my assay. I would like to appreciate if any one could give me a solution.
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I am surprised about many of the answers, well intended but they are only partially addressing your problem (by the way feedback inhibition-a term from Monod- refers to endproduct inhibition i.e., when the product of the last enzyme reaction inhibits the first enzyme reaction of the pathway), The main problem of your question is, that you do not described whether you checked for time and enzyme linearity in the first place. Do you have a one substrate one product reaction or a two substrate reaction (thus Lineweaver Burk will not give you the Km and Vmax because you need to do replots (see Enzymology books: MM-kinetics are only valuable for 1S-product reactions). The observed inhibition at higher substrate concentrations can have many reasons some of them are addressed above by the other researchers.It can be that your assay is not time linear, the substrate interferes,with the fluorescens measurement (so get the product and check for linearity, i.e., making a standard curve for product and fluuorescence) that will tell you whether the substrate may interact with the fluorescence measurement. You can have true substrate inhibition (frequently when at high S conc more than 1 S binds at the catalytic side. you can have at high product conc reverse reaction, or competitive inhibition at the S biding site (read up on this. Doing enzyme kinetics with partial knowledge of what is going on is a waste of time, i.e., taking vmax =v=o.5 as the true Vmax. S0.5 values (half velocity in S-saturation curve as a true Km etc.,etc.
Generally to determine KI values you have to do inhibition studies at various I conc. at Km or the apparent constants (see above) and at saturation (If you only do it at S-saturation and lower I conc and high KI (I 0.5) you will not see an effect if you have competitive inhibition) It depends what the final goal of your studies is how much work you must include. But anyway you have to do your measurements at low and close to saturation of your substrate(s) depending on your S-saturation curve after you insure that time and enzyme linearity is secured andyou are not in the Substrate inhibition range. Although, if your inhibitor is a substrate analogue you will get both inhibition effects. Keep in mind that time linearity depends for an endpoint assays also on your S-conc. you should not have under any circumstances converted more than 20% (most say 10%) of your substrate to reduce your error.
Finally and most importantly before you waste your time and money by measuring things you have to do all over or getting false results, get an intermediate enzyme kinetic book and read it from cover to cover (or take a class!). Don't relay on internet info only some of the descriptions are misleading.
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I am working on the identification/screening of ligninolytic enzyme producing strains. Poly R-478 is used in some cases. Can anyone suggest alternate dye for screening of such strains as the production of this dye is ceased?
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you can use Remazol Brilliant Blue R (RBBR). because ploy R -478 and RBBR both are anthraquinonic dyes.
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I am working on the identification/screening of ligninolytic enzyme producing strains. Guaiacol is used in some cases. Can anyone suggest more dyes or substrates for screening of such strains?
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you can use catechol, veratryl alcohol, ABTS and pyrogallol also for identification of ligninolytic enzyme producing strains.
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A protocol from a Paper says they extend PCR at 60 degrees. Is this possible?
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qiagen long range PCR enzyme works best at 68oC..but theoretically it is very possible that it will work at 60oC ..Only issue is that Tm should be low enough not to overlap with extension temp and there are chances of non specific amplification which can be avoided by adding 5% DMSO.. one more thing at low temp , extension time must also be increased to accomplish amplification..
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Have been using the semi-quan API-ZYM test to see the abundantly expressed enzymes by the microbial consortium in a lab-scale Rc treating wastewater. The enzyme "Naphthol-AS-BI-phosphohydrolase" keeps popping out as one of the abundant enzymes, no matter if the feed is synthetic or real wastewater. I cannot find solid info on what this enzyme does. Any idea about that? PS: the substrate of the enzyme in the API-ZYM slot (no12) is "Naphthol-AS-BI-phosphate"
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Hello I am also in the same situation ultilizando sludge on soils with different sludge treatments that enzyme being of the highest active hope can also help us with more information about it. Cheers
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I'm planning to do same inhibition experiments to test the capability of my compounds to inhibit aspartyl proteases. Can you recommend some detailed protocols? In particular, I'm looking for material about the analysis of data.
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Thank you!Unfortunately, I don't have access to the first article you recommended...
Do racemase enzymes exist in any eukaryotic cell?
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Considering that there are only one form of amino acids (L) and sugars (D) in eukaryotic cells such as yeasts, do racemase enzymes exist in them? If yes, what is their role?
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We have racemases in the eukaryotic cells (Amino Acid Racemases: Functions and Mechanisms - J Biosci Bioeng. 2003;96(2):103-9). See some examples: 1) First eukaryotic proline racemase was identified in Trypanosoma cruzi parasite. The parasite enzyme, TcPRAC, is as a co-factor-independent proline racemase and displays B-cell mitogenic properties when released by T. cruzi upon infection, contributing to parasite escape ( Mol Microbiol. 2005 Oct; 58(1):46-60 and Nat. Med. 2000; 6 (8): 890–7). 2) Serine racemase is a protein representing an additional family of pyridoxal-5' phosphate-dependent enzymes in eukaryotes. The enzyme is enriched in rat brain where it occurs in glial cells that possess high levels of D-serine in vivo. Occurrence of serine racemase in the brain demonstrates the conservation of D-amino acid metabolism in mammals with implications for the regulation of N-methyl-D-aspartate neurotransmission through glia-neuronal interactions (PNAS; November 9, 1999, vol. 96, no. 23, 13409–13414). 3) Fungus Tolypocladium niveum (Tolypocladium inflatum) has an alanine racemase that converts L-Ala to D-Ala for incorporation into cyclosporin by cyclosporin synthetase. Therefore, it is possible that C. carbonum also has an alanine racemase to produce D-Ala (THE JOURNAL OF BIOLOGICAL CHEMISTRY; Vol. 275, No. 7, Issue of February 18, pp. 4906–4911, 2000).
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PALA is a potent competitive inhibitor of Aspartate Transcarbamoylase; it binds to and blocks the active sites of the protein, due to it is a Bisubstrate Analog. I have read some papers, where it is provided by the Developmental Therapeutics Program of the National Cancer Institute (USA). But I don't know how to proceed in order to request any chemical reagent from that organization.
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If you want to contact the DTP at the NCI you can do so at ncidtpinfo@mail.nih.gov.
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I will start using SuperScript™ III to make RT reactions. I have some doubts about the ideal temperature in RT reaction and about the amount of enzyme per reaction. In the protocol, they suggest 50 C and 1 microL of enzyme (but I would like to use less quantity because it is very expensive). Any suggestion about the initial amount of total RNA per RT reaction?
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I use SuperScript III for my RT reactions pretty much every day for the past two years with success. Conditions of the RT rxn depend on the quality of your RNA as well as type of your RNA (how structured it is). My typical experiment is set up as follows: first I set up annealing reactions where I add: 5uM of RT primer with ~5 pmol of RNA of interest in total of 12 ul of reaction.
I was successful with as little as 0.25 pmol of my RNA and I was still able to obtain good yield of cDNA products.
I put that initial annealing reaction in PCR block for the following conditions: 85 C for 1 min, 60 C for 5 min, 35 C for 5 min and finally 50C for 50min.
During that time I prepare my RT mix:
5x RT buffer, 9.14ul,
100mM DTT 2.28 ul,
10mM dNTPs 2.28 ul,
SSIII 1ul
1.3 H2O
(total volume 16ul)
I put that mix in heating block for 3min at 37C before I add it to my annealing rxn (which as you remember I set up at the beginning).
When my annealing rxn is at final seconds of the step 35C I add the warmed up RT mix (8ul of RT mix to 12 ul of annealing rxn.
Finally after the rxn is over, I hydrolyze my RNA by adding 1 ul 4M NaOh and incubation for 5min at 95 C. After that I put my rxn on ice for 5 min and neutralize by adding 2ul of 2N HCl. Then, I precipitate the rxn with EtOH and NaoAC, centrifuge and analyze.
Good luck!
Just by knowing the Michaelis-Menten-constant Km and Vmax of a given enzyme and substrate is it possible to draw the EXACT Michaelis-Menten-Diagram?
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Well, this is what the Michaelis-Menten-Equation is claiming. But if I have two given points on my diagram, i.e. (0 ; 0) & (Km ; 50% of Vmax) and a given Vmax there is more than one possibility to draw the hyperbola, isn't there? Therefore I don't understand how the the Michaelis-Menten-Equation can give you the reaction rate for a specific substrate concentration, because I believe there is more than one possibility.
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By knowing those two parameters the Michaelis-Menten equation v=(Vmax*[S])/(Km+[S]) allows you to calculate the resulting reaction rate v for any given substrate concentration [S]. So there is only one way for the graph to be drawn because it is defined exactly by those two parameters.
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I need clone a fragment in a vector, but with only 1 cut in it. So I have to dephosphorylate but i don't know what enzyme is the better for do that
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Dear Susanna, While I am in agreement with the advice posted above, you should first consider whether or not dephosphorylation is really required - if you have enough vector and insert DNA available and you are attempting to clone into a general purpose vector with blue-white screening, then dephosphorylation is simply not required: you can easily spot a white colony amongst a backgrount of several hundred blues on an XGAL plate, so why add the extra step of dephosphorylation? I would suggest that if it is difficult (or expensive) to generate a large amount of vector and insert DNA for ligation, or if you do not have blue-white screening or a positive selection for the insert, then dephosphorylate the vector. I prefer the standard alkaline phosphatase followed by an EtOH precipitation rather than using temperature-sensitive phosphatases and complex clean-up kits. Regards, Andrew.
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I'm going to work on polyphenol oxidase (PPO) extracted from a fruit peel, and from what I read recently I came out with a series of purification steps involving ammonium sulphate fractionation, dialysis and gel filtration chromatography (Sephadex G-200), done accordingly after crude enzyme extraction (using sodium phosphate buffer).
I'm not sure of which substrates to use for PPO extracted from this fruit peel yet, so I might choose 2 of the best substrates with lowest Km values out of 4 substrates that are going to be tested on this enzyme.
So my main question is: Is my current protocol enough to obtain a partially purified PPO? At least 20 times pjavascript:void(0);urified?
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Hi Allan,
In addition to former answers, all of them helpful, you have a lot of literature describing classical PPO purification of a number of fruits. See for instance:
Isolation, purification and physicochemical characterization of polyphenoloxidases (PPO) from a dwarf variety of banana (Musa cavendishii, L)MAM GALEAZZI, VC SGARBIERI… - Journal of Food …, 1981 - Wiley Online Library
Purification of polyphenoloxidase from coffee fruits P de Fátima Pereira Goulart, J Donizeti Alves… - Food chemistry, 2003 - Elsevier
Polyphenoloxidase from apple, partial purification and some properties
A Janovitz-Klapp, F Richard, J Nicolas - Phytochemistry, 1989 - Elsevier
Strawberry polyphenoloxidase: purification and characterization
P WESCHE‐EBELING… - Journal of Food …, 1990 - Wiley Online Library
And many others. A diferent point is the appropriate substrate to follow the activity. Sometimes, the affinity is not the more determinant parameter. A high Vm and a simple colorimetric methods coulb be great although the Km is not very low.
If you need more details. just say. Cheers.
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I have a crude extract of enzymes produced by Solid State Fermentation, but the microorganism produces a black/ brown dye. I already tried to different methods to remove dye from extracts, like filtration, ultrafiltration and carbon active without good results ( active carbon removes dye but in the filtered we can't found protein neither activity). So, I saw a closely association between dye/total protein/Enzymatic activity, in all experiments, when the color it's removed, the activity and the protein are also removed. I used Spectrophotometric techniques to determinate the amount of enzymatic activity and total protein, and the presence of the dye causes interferences because the amount of absorbance are really higher with great rates of standard error.
I'm sure that the amount of protein (Bradford method) and the Enzymatic activity are not product of the error caused for the dye, because I made some dilutions ( when the dye is diluted ) with linear results of activity and protein.
Any suggestion?
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Dear Angel Josue,
Ever run a native PAGE (or SDS-PAGE) or size-exclusion chromatography (e.g. PD10 /G25 columns from the big G-company) to check, if the dye is covalently attached? If not, theres a good chance to use size-exclusion chromatography to further purify your protein.
If so, you're in trouble...
Ever tried to quantify metals in your sample to check for odd metal binding (iron, manganese or similar)?
Do the spectra indicate for coloured cofactors or is it just "black"? Brown colour can also indicate for Fe-S or other cofactors. Maybe you co-purified a protein neccessary for activity?
Cheers, Martin.
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Protein design is the design of new protein molecules, either from scratch or by making calculated variations on a known structure.
Therefore, Is it possible to design highly active de novo enzymes for any desired reaction?
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If you are really talking about de novo design with no selection or directed evolution component, then the answer is no.
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I am purifying an enzyme secreted from a filamentous fungi, and after "salting out" process, I want to proceed with ion exchange chromatography.
Between the two brand, which can give me the closest to 'pure' end product? I'm thinking to proceed with sephadex G-100 after the ion exchange.
Anyone with experience using the Fractogel before?
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Why not you try this with batch mode ! ( HTPD : high throughput process development)
1 : Clarify your sample with centrifuge or filtration to remove suspended insoluble particles and impurities.
2. Find out approximate pI and molecular weight of your target enzyme by Electrophoresis or by literature.
3. If target compound pI is acidic try Anion Exchange ( strong Q+ and weak DEAE+) , If your target compound pI is basic try Cation Exchange ( Strong SO3- and weak CM-) Keep the pH and conductivity of loading buffer and your crude protein ( conductivity < 5 mS/cm)
4.Screen the proper pH and conductivity with consideration of stability of your protein and buffer pka.
5.there are so many Ion exchanger available in the market. You can screen Anion Exchange resin in batch mode as per availability :
(Ge) Q Sepharose™ FF , Q Sepharose™ HP, Q Sepharose™ XL, DEAE Sepharose™ FF ,ANX Sepharose™ 4FF low sub, ANX Sepharose™ 4FF high sub, SOURCE™ 15Q,,SOURCE™ 30Q, Capto™ Q , Capto™ DEAE, Capto™ Q ImpRes ,
(Merck Millipore) Fractogel® EMD DMAE (M), Fractogel® EMD DEAE (M), Fractogel® EMD DEAE (S), Fractogel® EMD TMAE (M), Fractogel® EMD TMAE (S)
Fractogel® EMD TMAE Hicap (M), Fractogel® EMD TMAE Medcap (M), Eshmuno™ Q
(Avantor)BAKERBOND™ POLYPEI ,
(Biorad)Macro-Prep® DEAE, Macro-Prep® High Q , Macro-Prep® 25 Q, UNOsphere™ Q, Nuvia™ Q,
(Chisso )Cellufine® MAX Q-r , Cellufine® MAX Q-h , Cellufine® Q-500 (m) ,
(Life T) POROS® 50 HQ, POROS® 50 PI, POROS® 50 D, POROS® 20 HQ
(Pall) Q HyperCel™, DEAE Ceramic HyperD® F , Q Ceramic HyperD® 20, Q Ceramic HyperD® F ,HyperCel™ STAR AX
(ProMetic) DEAE PuraBead HF
(Tosoh) Toyopearl® DEAE-650C , Toyopearl® DEAE-650M , Toyopearl® DEAE-650S , Toyopearl® SuperQ-650C , Toyopearl® SuperQ-650M , Toyopearl® SuperQ-650S , Toyopearl® QAE-550C , Toyopearl® GigaCap® Q-650M , Toyopearl® Q-600C AR
( YMC ) YMC - BioPro Q30 , YMC - BioPro Q75
3. Find out stability window of pH , conductivity , temp etc of your protein by analysis.
4.Your purity of target enzyme depend on Host related impurities, process related impurities , product specific impurities. Understand more about your target enzyme and impurities.
5. After confirming from batch mode , you can try bind and elute mode or flowthrough mode.
Go through with attachment for more information or write me more details of your target compound and impurities to ymk108@yahoo.com
Best luck.
Which ssDNA ligase has worked best for you?
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ssDNA circularization without the use of complementary sequences - has anyone tried using enzymes which catalyze intramolecular ligation of unmodified linear ssDNA (or ssRNA)? From a search on the internet, I could find CircLigase & CircLigase II from Epicenter and ThermoPhage ssDNA ligase from Prokazyme. Which one is the most efficient of such enzymes on the market? If possible, please provide a brief description of your procedure as well.
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I am currently not using ligase enzymes
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Specifically I am looking for linkage between data sets contained in databases like SCOP, BRENDA, CSA, EC, KEGG. I can manually search what I need in each, but that is labor intensive and not scalable. Is there any resource that has united these data sets as a utility?
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I think the information you want is in the linked EBI resources. The Uniprot annotation for proteins contains information on active site, domains, and links out to structure databases. There are also family and domain databases at EBI: InterPro for example. Use www.ebi.ac.uk as your launchpoint.
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However both salts have Mg ions.
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Hi Megha,
I think that is because some polymerases can be inhibited by the Cl anion. So, you need to use a different magnesium salt.
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I noticed it already some time ago, that in accordance to suppliers MSDS some enzymes have actually higher activity in other-than-supplied buffer. Why do they not provide the best buffer?
If you go to http://www.clontech.com/takara/US/Products/Molecular_Biology/Restriction_Enzymes/Restriction_Enzymes_g_h/HindIII and click on Documents and load the data sheet, there is a table of relative activities. HindIII should have 100% activity in M buffer (supplied), but 200% in K buffer. So why not use K buffer instead of M?
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Also, may be due to the star activity of restriction enzymes.
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I am working with production of cellulolytic enzymes from fungus ( such as Aspergillus sp) and I want to perform Endoglucanase, Exoglucanase and β-Glucosidase Zymograms. I didn't have problems performing the endoglucanase, I used CMC 0.05% and congo red as revelator, I can see 5 different bands from 25 to 40 KDa. But with the Exoglucanase and β-Glucosidase I revised the methods: for exoglucanase, I can use fluorescent substrates such as 4-Methylumbelliferyl β-D-cellobioside in a Native PAGE to see the specific activity. For β-glucosidase I can use another fluorescent substrate such as 4-Methylumbelliferyl β-D-glucopyranoside or Esculin (Kwon et.al, 1994) to see the band.
The problem is that the fluorescent substrates are expensive, and the delivery times are more than two months.
Any other suggestions about how to perform the zymogram?
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I appreciate the problems caused by expense and by delay in delivery but the very high fluorescence of methylumbelliferone gives great sensitivity for the location of the enzyme bands that is unlkely to be matched by any other substrate. You may have to grit your teeth and bare the cost.
Peter Butterworth
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I'm going to extract polyphenol oxidases (PPOs) from a fruit peel and would like to find out their pH optimum so I need to incubate the purified enzymes into different test tubes, containing different ranges of pHs. Besides sodium phosphate salt, what are the other types of buffering salts that can be used to establish the desired pH ranges?
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Dear Francisco, thank you very much for your answer.
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I have a few sequences described as enzymes with lypolityc activity and I'm extracting the domains characteristics of said activity using Pfam and MEME (I'm following a procedure described by Kumar, et al in 2012, a procedure originally used for phytases). Now, with the domains, I'd like to run them against assembled contigs (or ORF) from metagenomic data. Is there any software that can help me with these analyses in a automated way? And, what should be better to use, DNA from the contigs or open reading frames described by gene identification software?
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Once you have identified a domain of interest with Pfam, you can use the corresponding hidden Markov model (HMM) to search against a sequence database with hmmsearch. A potentially much faster solution would be to use PoSSuMsearch2.
As for searching contigs or open reading frames, I'd use both, so to minimize the possibility that you miss something. If a domain is coded by several exons, you might miss it when searching the contigs (I am assuming that you get transcripts from your gene identification software). You might also miss domains when searching the gene models, because these models might just be wrong, or exons or whole genes might be missing. I don't know the extent of this danger, but better safe than sorry.
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I extract maize kernel for doing enzyme assay but the starch present in the samples gives a higher background OD which makes the assay difficult.
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I think it will be hard to measure amylase activity (I assume this from your question) in a solution with some starch in it. However, you can try to precipitate the protein in the solution first using ammonium sulphate or acetone, and then re-solubilized the precipitate to the initial volume, if you want to measure the activity in the initial volume. A paper by Warner et al. 1991 that measure alpha-amylase activity from maize seed can be found here: http://naldc.nal.usda.gov/download/24404/PDF
If you can give the method that you are using, maybe that will be better for colleagues to understand more about the problem. Hope this help..
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Aspergillus niger produces many enzymes of industrial use and is being used in solid state fermentation process. But it is very difficult to control the spread of its spores during harvest time. What would the best possible way to handle this problem be?
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As working with Aspergillus sp. whether it is niger or fumigatus under Solid state fermentation harvest the crude enzyme extract with filtration with buffer
N terminal sequencing of enzyme vs sequencing gene of that enzyme and translating it to amino acid sequence?
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Which approach is better (inexpensive or easy)?
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It clearly depends on what information you already have, and what you are wanting to do with the information. "Better" does not usually mean "inexpensive or easy" (if this is your definition of "better"). Do you have a particular experiment in mind, or, is this an academic type question? If it is experimental, explain to us what you are trying to do and you should get a more specific/relavant answer.
How to digest RNA using ribonuclease T1 from ammonium sulfate stock?
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I would like to remove RNA from bacterial DNA to evaluate the G+C content of my bacteria. I follow the methods of Tamaoka and Komagata (1984) (see attachment). But I am not sure how to use the ribonucleases. For RNase A, I am thinking to use TE buffer to dissolve my lyphilized powder. But I don't have a RNase T1 powder but a suspension in ammonium sulfate (http://www.sigmaaldrich.com/catalog/product/sigma/r1003?lang=de&region=DE). No information are given in the information sheet about how to use/dissolve it. Has anyone information on how to proceed? Furthermore, there is no information about the volume of the enzyme I should use for the protocol.
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Hi thanks a lot. I conducted my digestions using both enzymes. I checked if there was any RNA on a 1.5% agarose gel. I could not detect any band apart from DNA. But I wonder if this method is sufficient to check for residual RNA. Second, some authors only use RNase A to get rid off RNA (so did some of you), few use a combination with T1. Several times I read that RNase A was not enough to completely remove RNA since the enzyme cuts after C and U producing oligonucleotides which will not be removed by alcohol precipotation. RNase T1 cleaves at G positions yielding smaller oligonucleotides when combined with RNase A that can be removed with the alcohol. So, why is it enough to only use RNase A?
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A useful tool for disulfide bridges reduction, for example in proteins, is DTT. As a product of these reductions (in attachment) we have an "oxidized DTT" (http://www.sigmaaldrich.com/catalog/product/aldrich/d3511). Does anyone have knowledge of its RAMAN spectum or any ideas about it?
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In the database
searching for dithiane, you can find the IR of such a compound.
Anyway, I think that if you are interested in distinguishing the oxidized S-S in protein (reactant) from the oxidized DTT (product), Raman is not the best choice. If you use Raman, probably, a better reducing agent is TCEP tris(2-carboxyethyl)phosphine, (no overlap in the 500-550 cm-1 region).
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I am trying to determine the activity of the enzyme β-Glucosidase, isolated from A. niger and other fungi, produced by solid state fermentation. The problem is that the absorbance is very high (A= >1,5). The first assay I used pNPG 1mM (150µL) dissolved in sodium acetate buffer 0,1M pH=4,8, and 10µL of enzymatic extract. The mix was then incubated at 50ºC for 10 min. After that, the final volume was 1ml with NaOH 100mM. The absorbance read at 412nm. I used a p-nitrophenol calibration curve, 100-1000µM with a correlation of R =0.9998.
The next assay uses almost the same procedure, just a change in the last step. Complete the volume with glycine buffer 0.5M, pH 10.8, and the result is completely different i.e. too low. The problem is that I don't have a "standard reference enzyme" to test these protocols. What do you think? Where could the mistake be?
Note: the extracts have cellulase activity, because before these assays we confirmed the cellulase activity in FPU (filter paper units) and CMC, analyzing the liberation of sugars with DNS method.
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One thing I failed to mention is that color of p-nitrophenol is pH dependent. NaOH makes the pH to high, so the color is also more intense. Gly buffer does not result in too high pH so the color is less intense. As Natividad pointed out using Tris to stop the reaction works well. Make sure your standard curve is made in an identical way with glucose (add all ingredients except p-NPG and enzyme).
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High voltage is applied in enzyme solution.
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Depends on voltage and media, in water it starts to denaturate approx. at +1,2 V vs. Ag/AgCl, below 1 V vs Ag/AgCl it is more or less stable.
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I ran an RT-PCR for monoamine oxidase A in trigeminal ganglion, after migraine induction. However, I could not detect any band for the gene. The primers are correct, all the running conditions are also fine. Is it possible to get a blank gene expression for a constitutively active enzyme?
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Check your target gene for splicing variants. You may get enzyme activity of different isoform than mRNA you target is to generate. The other issue can be SNP in case one is located at 3'-end of your primers.
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My alpha-glucosidase enzyme (from SRL) is degrading too quickly when I make working and stock solution. I store it at -4.0 C. How can I prevent it from degrading? Do I need to store it at -20 or -80?
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The enzyme is need to store at high conc .(50 u/ml) in phosphate buffer at -20C then is need to dilute to 0.5 u/ml in working solution.
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I am working in the production of cellulase from fungus (molds) like Aspergillus niger, but I have problems with the precipitation with ammonium sulfate. I don't see any precipitation in the different fractions, neither after centrifugation at 14,000 rpm for 10 min. To produce the enzymes, I use solid state fermentation and for the extraction sodium phosphate buffer 50mM ph=6. I don't see precipitation yet, even at 70% concentration of ammonium sulfate.
I tried precipitation with acetone with good results, but in some cases the protein loses the activity or became insoluble.
I tried different protocols to precipitation with ammonium sulfate but I don't see good results. The concentration of protein in my crude extracts are about 0.05 mg/ml to 0.5 mg/ml (bradford method).
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Try to concentrate the broth 10 fold by ultrafiltration membrane 30kDa MWCO before Ammonium sulphate pptn..Monitor Protein content and enzyme activity at each step. Hope this helps
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Bacterial lipase is more important for industrial application than fungal lipase?
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Microbial enzymes have more advent than fungal lipases due to its easy cultivation method , less incubation period , easy recovery and ease  of  genetic manipulation  to get high yield enzyme .
Application of lipase in biodisel and other pharma industries for formation of enantiomeres .Application of lipase in treating a cancer are the most probably the recent work on lipases.
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I am performing a reaction in which the [S]>>[E]- I'm obtaining and v vs [E] curve. In Michaelis Menten, one plots 1/[S] vs 1/[v] to get enzyme parameters (Km, etc). Can I plot 1/[E] vs 1/v and get those enzyme parameters? Is there a different set of equations for such a situation (ie. in which [E] is being modulated while [S] remains constant?)
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If you are performing a reaction where [S]>>[E], then you must assume that your enzyme is still following "Zero Order Reaction Kinetics". Basically, what this means is, as you increase your enzyme concentration, your observed/measured rate of reaction will increase proportionally in a linear fashion. You should not really see a "curve", in the sense of a MM hyperbolic isotherm, because essentially, you will always saturate out the total number of substrate binding site because, by definition, [S]>>{E]. What you should see is a straight line following y=mx+c. It may drift off as you reach higher [E], but this might be due to experimental set up, such as limitations of measurement of your reaction (i.e., analysis moving out of the linear range on your detector). Only when you titrate icreasing [S] against a fixed [E] will you observe classical MM curve (assuming your enyme follows "standard MM kinetics".
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I am studying the activities of Laccase (EC 1.10.3.2) enzyme in soil. Using ABTS (2,2'-azino-bis-(3-ethylbenzthiazoline-6-sulfonic acid). I cannot observe change in absorbance at 420nm or near about it (410-420nm) activities. I am using protocol, copy is attached here and can also be accessed online via link www.eeescience.utoledo.edu/faculty/weintraub/Laccase_protocol.doc‎. I tried in many different ways with all possible options. But I cannot make any attempt successful. Should I change the substrate other than ABTS?
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The ABTS is from the best or the best substrate of laccase, it is widely used to this enzyme activity. The substrate is very soluble in water then it is very easy to use with the enzymatic protein and the product is very stable and very easy to oxidase in 1 min if you have good activity. I have work a lot with the substrate for laccase activity, the product is green and highly detectable at 420 nm. I have just to remarks:
1- The concentration that you use is good (0.4 mM) but you can increase until 1 mM if you want to screen the little activity.
2- It is not necessary to add H2O2 is not a peroxydase that you test, the seconde substrate of laccase is the oxygen and you not need H2O2.