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Hi
The question is: Determining hexokinase activity in rat brain where 10 µl of a 10% (100 g/L) brain homogenate (containing hexokinase) is mixed with 1990 µL of reaction medium (containing the substrate glucose and the coenzymes ATP and NADP+ and the auxiliary enzyme glucose-6-phosphate dehydrogenase). The reaction mixture is placed in a cuvette (light path = 1 cm) in a spectrophotometer at 37°C, where the hexokinase reaction can be seen as an increase in absorbance at 340 nm. The reaction proceeds at a constant rate during the measurement period, and during 10 min a total increase in the light absorption at 340 nm of 0.10 is recorded. The absorption coefficient of NADPH = 6300 x M-1 x cm-1
I need to find the enzyme activity of hexokinase in µmol/min/g. I don't understand how I need to interpret the first line (10 µL of a 10 % (100 g/L) brain homogenate) - and how should I use these numbers?
Thanks in advance!
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10 µL of a 100 g/L (= 100 µg/µL) homogenate contains 1000 µg = 1 mg = 0.001 g.
That will be the number in the denominator when you divide by the number of g to calculate µmol/min/g.
The change in absorbance of 0.1 unit is converted to the concentration of NADPH formed using the extinction coefficient of 6300 M-1cm-1 and the Beer-Lambert Law: absorbance = extinction coefficient x pathlength x concentration.
An absorbance change of 0.1 for NADPH with an extinction coefficient of 6300 M-1cm-1 in a 1-cm pathlength cuvette corresponds to a NADPH concentration of 0.1/(6300 x 1) = 1.587 x 10-5 M = 15.87 µM.
This amount of NADPH was formed during 10 minutes, so the rate of NADPH formation was 1.587 µM/min. This occurred on a reaction volume of 2 mL, so it can also be expressed as 1.587 µmoles/(L-min) x 0.002 L = 0.00317 µmoles/min.
This happened when you used 0.001 g of material, so the specific activity was (0.00317 µmoles/min)/(0.001 g) = 3.17 µmoles/min/g.
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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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I found the dns method to measure the amylase activity, but I cannot get the dns reagent. Therefore, could you suggest me an alternative method or alternative reagent to dns reagent?
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You can Use Nelson- Somogy Method.
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Sometimes the enzyme may not be activated or may not function properly unless the it is co-expressed with other protein for post-translational modification or assembly into a functional catalytic complex.
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Does a better docking score or stronger binding energy between a mutant enzyme with its substrate always translate into an improved enzymatic activity?
How should RMSD, RMSF, radius of gyration, solvent-accessible surface area (SASA), etc. from molecular dynamics simulation be used to guide enzyme engineering with the aim of improved product synthesis?
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You may want to Google "simulation of enzymatic catalysis"
Some interesting hits:
Two Sides of Quantum-Based Modeling of Enzyme-Catalyzed Reactions: Mechanistic and Electronic Structure Aspects of the Hydrolysis by Glutamate Carboxypeptidase
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Hi, Iinserted gene of Reverse transcriptase into pPLc245 plazmid. If I'm right, this plazmid doesn't code cI857 represor. For expression od reverse transcriptase from this plasmid I used E. coli DH10B. RT was expressed after increase of cultivation temperature to 37 °C. Before this thermo induction I cultivated E. coli DH10B with pPLc245 at 28 °C. But at this temperature RT was not expressed. Why is this possible? Is E. coli DH10B coding cI represor or pPLc245 could contain gene for this represor?
Thank you for all responses.
Bohuš
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If your plasmid really is pPLc245 (and not some derivative of it) then it should not carry the lambda repressor gene and you should get constitutive expression at any temperature. So I can't really explain your result unless RT does not fold properly to have activity at RT or 28.
I'm actually a bit surprised this worked at all for you though, generally plasmids with superstrong promoters like lambda pL are not very stable under conditions where there is no repressor.
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Hi, every body
I am investigating the enzyme profile of a white-rot basidiomycete fungus. In the enzyme assays, we detect enzymatic activity by color change. However, my fungus produces pigments that do not precipitate with centrifugation, so it causes errors in the results. Does anyone have a solution? Thanks.
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Dear Khajezade,
Greetings and have a nice time.
You can remove the pigment easily by using acetone or chloroform. Any one of them is useful to dissolve the pigment well. You can try any one.
With regards.
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I would like to ask you, when I measured the enzyme activity of papain by the national standard method, but the sample tube and the blank tube became zero and negative values when subtracted from each other. The blank tube was added with TCA first, which means that it was caused by the casein itself and showed a light blue color.
How can I adjust the OD value of the diluted papain, which is also light blue, to 0? Is there a problem there?
There is no air bubbles in the measurement, and the instrument operation and personnel operation have been excluded.
Translated with www.DeepL.com/Translator (free version)
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Hello, I agree with Engelbert Buxbaum, apparently there should be no problems following that methodology, I would also suggest that you review some aspects that may be affecting the analysis, such as:
-preparation of reagents and buffers (verify pH, ionic strength, etc.)
repeat the procedures in the same way for the blank and the sample, eg the incubation time, the same centrifugation time to remove the pellet
check the detection range of your spectrophotometer and the wavelength selected for analysis.
You can also do a quick test and use the reaction blank without enzyme (control -) as the autozero and use the reaction medium plus the enzyme plus the TCA as the blank to inactivate it before the incubation process, after this time centrifuge and then that it would be your positive control to correct any color that could be caused by amino acid residues and then read your sample
I hope I have helped you in some way
regards!!!
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I am about to calculate alpha-amylase activity for some fungal samples. I followed the protocol of Sigma-Aldrich (https://www.sigmaaldrich.com/DE/en/technical-documents/protocol/protein-biology/enzyme-activity-assays/enzymatic-assay-of-a-amylase) and incubated my samples in 0.2 % starch solution for 3 minutes.
According to the protocol, the equation for the U/ml value is:
U/ml = (mg of maltose released * dilution factor) / (ml of enzyme applied)
I think I also have to consider the incubation time of 3 minutes, and divide my result from the equation by 3?
Thanks in advance for help.
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Yes, at least an International Unit should be used.
Amylases hidrolyze glycosidic bonds. When a general substrate is used, as starch or glycogen, the hydrolysis will be done for ALL enzymes capable to hydrolyze alpha 1-4 and alpha 1-6 bonds between glucoses. Then every broken bond will produce a new reducing sugar. So, if you measured reducing sugars, to calculate the amylase activity, you will be able to know how many glycosidic bonds were hydrolyzed, by time unit (min). 1 unit of amylase is the enzyme required to release 1 micromol of reducing sugar (from starch or glycogen substrate) by minute.
A maltose standard curve will be necesary to get the molar extinction coefficient, to convert absorbance (ie. 550 nm by DNS method) to equivalent moles of maltose, as reducing sugar.
Hector
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I have been failing to express a very large trypanosome (160kDa) protein using various expression systems. The protein is a chimera of 3 enzyme activities and I can express truncated forms containing at least one of these enzyme activities. So, I was wondering if I can express and purify these enzyme 'domains' individually and somehow get them to associate, in order to perform enzymatic assays assessing their potentially interactive (regulatory) roles.
For instance can an inducible di-cre recombinase-type system be used? Ideally any recombination domains (or tags) that I may fuse to my recombinant construct would be as small as possible.
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Look, unless your project is to design methods for expressing protein domains, I'd forget your idea of making it in pieces and hoping they will somehow combine properly. It is 1) unlikely to work and 2) very time-consuming. And 3) you'd need to have the entire protein as a comparison anyway!
Try finding a better way to express the native protein. Have you tried putting it in yeast? E. coli strains that are optimized for large proteins? Under an inducible promotor? Lots of good protocols already exist.
Good luck!
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I have diluted my chrysin in DMSO, however when run an enzymatic assay in phosphate buffer pH 7.4 (50 mM) and pH 7.6(10mM) it formed precipitation? do you have any suggestion to stop it but still using the same buffer? Thank you
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The buffer will have be different or the precipitation will happen again ; ) Some have used cyclodextrins to encapsulate compounds with limited solubility, like chrysin, but you'll need controls to be sure its still accessible to your enzyme. Some nonionic surfactants may help, often bellow their CMC, but controls will need to be done to be sure the assay is still functional. Enzymes that catalyze membrane bound or insoluble substrates are a real challenge, good luck!
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The Vigna radiata plants were subjected to Cd stress ranging from 50-200 ppm. Why does the enzymatic activity (SOD, APX, and CAT) increase under increasing metal stress despite the decrease in protein content?
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And what about TAS??
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Hi, I'd like to ask whether it is necessary to precipitate MMLV Reverse transcriptase before affinity chromatography purification (Äkta)? My colleague must do this step with his Taq DNA polymerase. He use (NH4)2SO4 or Na2SO4 + PEG. Without this precipitation is polymerase inactive.
Thank you for your responses.
Bohuš
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the so called Pluthero method to purify Taq is quite old (1993) and while simple it can result in low yields and DNA contaminants (see "A simple and efficient method for extraction of Taq DNA polymerase" 2015)
I don't think the ammonium sulphate precipitation is required for activity (please correct me if I' wrong) but it may be worth a try on MMLV RT, as they are very different proteins. Yields are sometimes not very important and Am sulph precipitation can be a very useful method to isolate/purify and concentrate protein preps. I might just use a modern MMLV RT prep like this https://www.protocols.io/view/recombinant-protein-expression-of-mmlv-rt-h-yxmvmxmw9l3p/v1?version_warning=no
iff your constructs are similar, good luck!
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How to know whether a particular protein is an auto cleaving protein or not (i.e. Self cleaving) ?
Also suppose a particular protein is degrading immediately so How to know whether this is due to Auto cleaving nature of the protein or due to some other proteins (i.e proteases) in the supernatant.
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If you want to know if the protein is cleaved during its biosynthesis in a cell or following secretion from the cell (either autolytic or by some other protease), you could follow its mass over time following induction of the gene. This could be done using 35S-methionine labeling, immunoprecipitation and SDS-PAGE or by Western blot, both requiring a specific antiserum or antibody.
I think it is difficult to establish whether the cleavage is autolytic or not when the protein is expressed in its native setting. It might help to express it heterologously, so that whatever activating enzyme would normally be present would instead be absent. Then if the protein is expressed in a soluble form but is not cleaved, you would have evidence that cleavage is not autolytic.
Another approach would be to overexpress the protein in bacteria such that it formed inclusion bodies (which often happens but is not predictable). Then, the protein would be unlikely to be cleaved because it would have precipitated before folding. Refolding the protein from inclusion bodies would be necessary to render it in a soluble form. Because proteins in inclusion bodies can be purified before refolding, you could find out whether the pure protein cleaves itself once it refolds by SDS-PAGE or protein mass spectrometry or N-terminal sequencing.
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Hi, has anybody experience with addition of ammonium ferric citrate to expression media? I decided to ide this because of its perfect solubility in water.
Thanks for your responses!
Bohus
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When I want to express proteins with Fe-S clusters and purify cluster loaded protein (holo-protein) I add 0.2mM (final concentration) to my E.coli culture. I induce with lactose and also add L-cysteine as S source. As I grow my cells in LB, I don't think the extra nitrogen I am adding is doing much. I don't think by adding ammonium ferric citrate will impair E.coli growth or protein production, for sure it will not harm the induction experiment under normal conditions. If you could specify a bit more your objective, we might give you more insight.
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I am trying to assess the activity of PFK enzyme in mESC lines. Recently in the metabolomics analysis, I discovered that although the cells show an increased uptake of glucose 6-P and fructose 6-P, there is a major drop in the levels of downstream metabolites starting from fructose-1,6-bisP all the way up to pyruvate. (except glyceraldehyde-3-P, 1,3-bisPglycerate and 2-phosphoglycerate)
There is also an increase in the levels of glucosamine-6-P, mannose, mannose-6-P and of 6-P gluconate and ribose 5-P. These levels could be increased as G 6-P and F 6-P could be shunted into the pentoseP pathways.
Therefore, I am planning to check if there is a blockage between the F 6-P and fructose 1,6-P by detecting the activity of the PFKinase enzyme.
So far the kits I've found to do this, are based on the calorimetric assay where PFK activity is determined by a coupled enzyme assay, in which fructose-6-phosphate and ATP is converted to fructose1,6-diphosphate and ADP by PFK. The ADP is converted by the enzyme mix to AMP and NADH. The resulting NADH reduces a colorless probe resulting in a colorimetric (450 nm) product proportional to the PFK activity present. One unit of PFK is the amount of enzyme that will generate 1.0 mmole of NADH per minute at pH 7.4 at 37 °C.
I was wondering if there is an in cell method, more precise for cell extracts.. I am not sure how sensitive or specific this calorimetric assay would be.. Any comments/ suggestions would be much appreciated.
Many thanks.
Best,
Pooja.
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Did you solve your problem?
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Can I say for a given same enzyme, more number of substrate A is changing into the product than substrate B per second ? Please make this this this clear to me. I know Kcat/Km characterized high efficiency. However, I am getting into this situation somehow. Is it possible ? How I would justify or rationalize this in the paper for publication ?
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As described in Cornish-Bowden Fundamentals of Enzyme Kinetics, if the substrates are both mixed together in the reaction with the enzyme, the ratio of the velocity of A to the velocity of B is (kcat/Km*[A])/(kcat/Km*[B]), so if the samples are mixed and the substrate concentrations are equal, A will be converted faster at all concentration, thus the name specificity constant for kcat/Km. However, if the reactions are separate and the concentrations are near saturating, the kcat will determine which rate is faster.
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I am a student from a relatively developing country and the commercial enzyme is expensive for us. My graduation thesis is desperately in need of an available standard curve for my AchE assay so that I can calculate my results using it. Please is there anyone who currently doing the AchE assay who can share it with me or where can I find it, are there papers or textbooks that I can find it in?
Your help would be very much appreciated.
Thank you
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A standard curve must be prepared under the identical conditions as the assay. It is not likely that one that you did not prepare yourself will be accurate.
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I can only start with raw feather from chicken/duck, but not feather powder.
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I'm having the same problem! Any update?
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I'm trying to run an assay for Caspase 3/7 activity to visualize apoptotic cells in flash frozen, unfixed mouse skin sections. The assay involves a non-fluorescent reagent that is processed into green fluorescence by endogenous, active Caspase 3 or 7 in apoptotic cells.
After thawing the slides for 3 minutes and washing once in PBS to remove OCT, I added the reagent to the unfixed tissue (I've tried 1 hour at room temp, or 30 mins at 37C). After the incubation, I post-fixed the sections with 4% paraformaldehyde, added DAPI mounting medium then visualized. 100% of the cells had green fluorescence, and the nuclei appeared enlarged so it looks like the cells are bursting or autolysing during the incubation. I'm wondering if I should fix the tissue immediately after thawing to stop the damage to the cells.
What would be the best method for post-fixing flash frozen mouse skin so that endogenous enzymatic activity is maintained? Does anyone have experience specifically with using the CellEvent Caspase 3/7 assay from Invitrogen in frozen tissue sections?
PS. I have also tried a TUNEL assay which generated no signal.
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I would suggest you to carry out ELISA or WB analysis of Caspases 3/7 for the detection of apoptosis in flash frogen tissue. If you fix the tissue, the enzyme activity will be lost, hence you can homogenize the tissue and take the sup to anayse capase-3/7 activity using ELISA and level by WB.
As you mentioned in your content, you can go ahead with the TUNEL Assay that is very sensitive ,(For example R & D Systems USA) and you can analyse DNA fragmentation in a single cell of you sectioned tissue.
Best
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Most people say that imidazole doesnt affect the majority of downstream applications for purified proteins but it is a chelator, so you would think that it might chelate the Mg2+ in solution and inhibit Mg2+ dependent reactions. Anyone seen any papers that discuss what concentration Imidazole will chelate magnesium or manganese ions in solution?
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Not only ligand concentration but pH is also an important factor of chelation .
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Hi everyone, I have been trying to calculate the Km value of a recombinant HCV NS3/4A protease using a FRET substrate, the cleavage of which can be detected at 500 nm using excitation wavelength of 355 nm. I have a RFU vs. time graph of different concentration of the substrate (attached). Next, I am calculating the initial rate which is the slope of the linear initial part of the progress curve, with the ultimate goal to plot the 1/[S] vs. 1/[V] graph and calculate the Km from the Lineweaver-Burk equation. However, if I calculate the slope from this graph for the time range between 0-10 minutes which is the initial linear part, the intercept for the 1/[S] vs. 1/[V] graph is negative but Km value can not be negative. I was wondering if anyone can guide me on how to calculate the initial velocity correctly from the graph I have? Thank you so much!
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follow Dr Adrian and Dr Shapiro's advice. The curve does not appear to reach Vmax (Fig.1 ).
Equally spaced substrate concentration- 0, 0.05, 0.2, 0.8, 3, 6, 10, 20 and 30 uM should do the trick as the Km is approx 4.7 uM, see
Do triplicate to get narrower confidence intervals, (Fig. 2) simulation
Get more data at the climbing stage (<4 min), maybe every 30 sec, as this is the region you calculate initial velocity (Fig. 3).
Use nonlinear regression software- GraphPad for e.g. as it has an inbuilt MM curve.
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Hey guys,
I'm using (for me new cells) E. coli Arctic express for protein expression (reverse transcriptase). I tried expression at 20°C and 10°C. At 20°C solubility of my proteine was 50% and at 10°C it was about 57%. I'd like to know some your experiences and tips how to use these cells (the best media for them, optimal temperature for night culture or growth up to induction and after induction, how to eliminate chaperonins after expression, how long should lasts expression, some supplements to media, concentration of antibiotics in night culture, concentration of IPTG ... )?
Thank you for all advices!
Bohus
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I've not used Arctic Express directly but have some experience of overnight protein expression systems.
We would grow the starter cultures up to whatever the required OD600 density was (this took some optimisation for each construct so no fixed answer on this point), then turned off the heat and propped the incubator lid open to the ambient temperature (somewhere in the teens). This was using Autoinduction medium containing glucose, glycerol and lactose (there are various recipes for this online), so it only starts induction by metabolising the lactose when really high OD after the glucose is exhausted and derepresses the promoter.
Key things we found improved expression levels were adequate aeration - using 3 litre plastic baffled flasks containing at most 1L of culture would give a very aerated culture, and selecting "good" colonies - I engineered a cleavable fusion GFP reporter upstream of the reading frame so you could identify high expression clones by putting the plate on a blue light box, as well as being able to see where the protein was during purification stages. Sometimes it was necessary to go for colonies that didn't express the highest fluorescence to maximise solubility.
Although it costs more, carbenacillin beats ampicillin hands-down for the same plasmid - it breaks down more slowly and is more stable in acid pH that can arise as cultures are prolonged and denser.
Elimination of chaperonins after culture - there is a protocol to wash some of these off your protein on column by addition of ions and ATP - it is a number of years since I've done this so I can't remember the precise details. Again though, this is published data and can be found by searching for this.
The final answer is 'try try try'. Every protein is different, so asking questions here and doing literature searches, then trialling your findings in your own laboratory is often the only way to optimise the system in your hands - what works for one person may not be completely applicable in another laboratory.
Very best of luck!
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Hello everyone,
I'm working in my doctoral thesis in the purification of a lipolytic enzyme from an halophile archea. Currently, I'm trying to purificate the recombinant enzyme using Haloferax volcanii as an expression system. The problem is that suddenly when filtering my crude extract through a 0.45 um nitrocellulose membrane I lose 90 % of the enzymatic activity, when that did not happen before and at most I lost 20 % of the activity.
Has the same thing happened to someone else? Or do you have any ideas that could help me? Thanks!
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Yep, nitrocellulose is a poor choice
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We are used the protocol for urease extration adaptated (to C. neoformans) from Amin et al.,2013:
Briefly, the broth cultures were subjected to centrifugation (5,000 × g, 4 °C) and the recovered mass was washed twice using phosphate-buffered saline (pH 7.4) and then stored at -80 °C. Subsequently, was thawed to ambient (room) temperature, followed by mixing with 3 mL of distilled water and protease inhibitors (TLCK, Tosyl-L-lysyl-chloromethane hydrochloride) and sonication for 60 s. After centrifugation (15,000 × g, 4 °C), the supernatant was desalted by eluting through SephadexG-25 column. The resultant crude urease solution was mixed with an equal volume of glycerol and then preserved under refrigerator (4 °C) for further uses.
Okay, we ran the suggested protocol, but after 2 days, enzyme activity was no longer observed.
Then, we tried to exclude the TLCK from the protocol, again the activity was observed on the day of extraction, but after it lost the activity.
We also tried to activate urease with sodium bicarbonate and Ni solution, without success.
In another procedure, we repeated the protocol and lyophilized the solution resulting in a white solid. So we tested it on this day, we prepared a urease solution (10 mg / mL) and the activity was good, but after a few days, again the activity was lost.
We always work with refrigeration, and we are careful to keep the solution on ice when we handle it.
Please any suggestions?
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Something I would have done differently is to have used a pH-buffered solution during the procedure to maintain the pH at the optimal pH of the enzyme.
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Hi, I did heterologous expression of reverse transcriptase in different strains of E. coli at 3 temperatures, 37 °C, 28 °C and 20 °C. As I expected, BL21 provided the best solubility at 20 °C and the worst at 37 °C. On the other hand strain MC4100 had the best solubility at 37°C and the worst at 20 °C. Has anybody similar experience with E. coli MC4100?
Thanks for responses!
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What expression system are you using? The typical pET based vectors used in BL21(DE3) strains do not work in MC4100 as it lacks the T7 RNA Pol gene required.
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As I am characterizing an 3' exonuclease and calculated the apparent Km and Kcat for two different substrates. What I observed is little bizarre. A substrate (S1) which looks like (qualitatively) degraded slowly by the enzyme has Km lower (~60 nM) and lower Kcat (5.1 sec-1) while other substrate (S2) apparently degraded faster, has higher Km (~175 nM) and but slightly higher Kcat (~8.2 sec-1). Now I have difficulty in understanding why is this happening. One more thing I observed is that S1 gets inhibited after 150 nM substrate concentration. While S2 continue to maintain it's saturation plateau even at 8000 nM. I am little perplexed how to rationalize this result. Any help will be appreciated much in this regard. Need an explanation about preferential degradation of the substrate by the enzyme.
Thank you in advance.
Prem
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I've edited this answer, hopefully a little clearer now, esp. with an example uploaded.
I think the issue is perhaps that you are not taking into account the fact that S1 displays substrate inhibition. Because S2 does not show this effect you are not comparing like with like. Effectively the enzyme is operating in the presence of an inhibitor with S1, whereas with S2 it is not.
However, even ignoring this effect, the Kcat/Km ratio (or Vmax/Km ratio) does not actually predict which is the better substrate in a real assay situation. You need to consider [S] too.
To see the effect of varying S you would need to plug numbers into the equation v = Kcat [E] S/(Km + S). I think you will find that at low S, S1 has the edge and at higher concentrations S2 has a higher rate. This is actually too simplistic for S1 and Adam B Shapiro has given a more complex equation which is relevant to the situation with an inhibitor in the system.
I have now uploaded data for two hypothetical substrates S1 and S2, neither of which show substrate inhibition, using the standard equation v = Vmax[S]/(Km + S). The tabulated data show that there are conditions in which you can get more product with a substrate with a LOWER Vmax/Km ratio. In fact, you can get almost any rates you like by varying S, thus Vmax/Km (or Kcat/Km) ratios alone do not tell you which is the better substrate.
Hopefully, you can now see some possible explanations why your S2 substrate was better than you expected.
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Hi, I'd like to try expression of reverse transcriptase with low temperature, 15 °C, due to solubility. Should I use some special plasmids for cold expression? I ordered cells for this purpose - E.coli Arctic express. Is it enough or is it better to combine these cells with plasmids for cold expression? Thank you all!
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you can try the pcold vecotrs of takara
i used in some year ago for a protein fusion that was degraded with expression at 20-25-30°C in pet vectors and performing the expression at 17°C, I obtained good results.
you can see an example of it at minute 6' 10'' of the following video
present in my blog: ProteoCool
Sincerely in other cases, with more stable proteins they provided lower expression than vectors based on T7 promoter therefore i suggest to you to test it in parallel with T7 and not replace it.
good luck
Manuele
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Hi guys,
Is there anybody who adds betaine (trimethylglycine) to media for protein expression? I read that betaine is able to increase solubility of proteins. If you use it for this purpose which form is the best? Is there any proven concentration?
Thank you for all answers!
Bohus
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Dear Bohuš Kubala please have a look at the following original research article in which the addition of osmolytes such as betaine has been studied in detail:
Native folding of aggregation-prone recombinant proteins in Escherichia coli by osmolytes, plasmid- or benzyl alcohol–overexpressed molecular chaperones
Fortunately this paper is freely available as public full text. I hope it helps.
Good luck with your research!
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Hi, I'm trying to produce reverse transcriptase. I need to increase its solubility. I would be grateful for some experiences and advices. Thanks so much.
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1. Try adjusting the temperature as you have, even trying at 30 could generate some soluble protein,
2. Adjust the amount of IPTG that you use to induce the expressions
3. Adjust the time you carry out the expression for, if you get soluble proteins at 20 C then leave over night to generate more protein.
4. If there is any information out about you protein of interest, does it require a chaperone protein to assist its folding, if so this will need to be co-expressed with the protein,
5. Check, double check, and get someone else to check that in you sequenced clone your gene of interest is in frame with the His Tag, you will be surprised how often the cloning is done incorrectly.
6.a Change the tag or place it at the c- terminus in the construct, it may be that your tag is affecting the folding of the protein, but less likely here as you have tried to express fragments and they are still stubborn. I would try to express the protein with no tag to see if it is soluble in the cells you are using.
6.b If I have an insoluble protein then MBP (maltose binding protein) is one of the best bets, a bulky soluble protein that can be easily cleaved off in many cases.
7.Change the cells you are using to express, as it is a viral protein, the codon bias may be restricting the accurate synthesis of your protein, therefore it may be worth trying Codonplus BL21, or other derivatives such as Rosetta gami strains
8. Another option is to purify the inclusion bodies of you insoluble proteins, and refold them, this can be done either by refolding on a his-trap or sequential dialysis in decreasing urea concentrations
I hope that offers you some avenues to try out!
9. For the non specific bands contaminating your sample, that is normal - always try to extend the washing step on the purification column to remove non specific binders, the other option there is to identify the concentration of imidazole that your protein of interest elutes at, then create a gradient and collect very small fractions to minimize contaminations. APS cuts before this step also reduce the amount of protein you load onto a column, overloading can generate many non specific interactions!
Good luck!!
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Hello!
I am looking for examples of industrial processes which are catalyzed by enzymes (either natural or recombinant). Particularly, I am interested in processes that end up with enzyme removal or inactivation (thermal or chemical), and I am trying to find examples from different areas - the food industry, textiles, pharmaceutics, fuel, etc.
Could you help, please?
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Also, Enzymes are used in industrial processes, such as baking, brewing, detergents, fermented products, pharmaceuticals, textiles, leather processing. Here are a range of processes showing how enzymes are used. See the RG link:
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First of all, I apologize for my poor English.
I would like to measure the enzymatic activity of COMT, referring to the following reference, but is it theoretically possible to substitute norepinephrine bitartrate hydrate?
Thank you very much.
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It is just a salt form of NE. I haven't done this assay but Googling NE + bitartrate + COMT + assay produces results that show this substrate form has been used by others.
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Anyone know how to measure catalase activity in brain homogenates in terms of μmol/min/g of brain. Reaction was stopped by addition of dichromate and absorbance was measures after 1 min at 570 nm. Controls are without H2O2 for each samples.
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There are several tests available to measure the catalase activity in brain homogenates. Hope you are trying to analyze enzyme activity using colorimetric assay kit. We have used the sensitive kit purchased from BioVision, Inc., CA that is very sensitive reproducible. If you are using the colorimetric assay kit, follow our paper published in Brain Research Bulletin 95 (2013) 54– 60. For more detail, see our research paper. Follow the kit mannual for calculating the catalase activity. All the best.
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Can you plzz provide some references for effect of Mn fertilization on soil enzymatic activities
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Please have a look at some enclosed PDFs...
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Hi!
A special primer has to be used with only 19 bp poly dT (Tm 44.9) instead of 30 bp in original study (Tm 55.2). Will this induce failure of mRNA capture? Will this cause RNA detachment in Reverse transcription using Superscript II (@42 ℃)? If this is a problem, how about start reverse transcription @ 37℃ or 40℃ for 20min, then rise it to 42℃? Will superscript II be completely block at lower temperature?
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19T-V is better than 19T in targeting the beginning of the polyA site, yes. I believe 19T-VN just gives a slightly tighter clamp.
As to the reaction conditions, yes: heat your RNA + oligodT to 65 for 5 mins, crash cool on ice, then add the buffer/dNTPs/enzyme, mix well, spin down briefly, leave at RT for 10mins, then over to 42 for 25mins.
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I have Laccase from trametes versicolor with 0.5U/mg activity.
I need enzymatic activity of 50U/g in solution. What should be the amount of enzyme I add to obtain enzymatic activity of the above said?
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If you have a solution with 0.5 U/mg is the same of 500 U/g, so if you wish 50 U/g you must dilute 10 times that solution in the appropriate solvent or activity buffer.
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I am working with erythroid progenitor cells in culture and want to assess the activity of an intracellular (cytoplasmic) enzyme. I have a protocol from the literature for the activity assay, but I can't seem to find any detailed info on the lysis buffer and methods used prior to the assay. For westerns I lyse with RIPA on ice and then sonicate, but I know I can't use RIPA for this, so my two questions are:
1. What lysis buffer recipe do you recommend to extract cytoplasmic proteins for an activity assay?
2. What lysis method do you recommend in conjunction with this buffer? Details would be appreciated!
I know there are many lysis buffer options and I'm just not sure how to figure out which one is both easy to make and will work for this purpose, and if one lysis method (freeze/thaw vs sonication, etc.) is more/less suitable for retaining enzyme function for downstream testing?
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You may also have a look for this option;
Thermo M-PER™ Mammalian Protein Extraction Reagent for RIPA like non-denaturing lysis buffer...
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A full-term female infant failed to gain weight and showed metabolic acidosis in the neonatal period. A physical examination at 6 months showed failure to thrive, hypotonia, small muscle mass, severe head lag, and a persistent acidosis (pH 7.0 to 7.2). Blood lactate, pyruvate, and alanine were greatly elevated. Treatment with thiamine did not alleviate the lactic acidosis. Which of the following enzymes is most likely deficient in this patient?
a) Alanine amino transferase
b) Phosphoenolpyruvate carboxy kinase
c) Pyruvate carboxylase
d) Pyruvate dehydrogenase
e) Pyruvate kinase
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Pyruvate carboxylase
Rationale: A biotin-dependent enzyme belonging to the ligase family that catalyzes the addition of CARBON DIOXIDE to pyruvate. It occurs in both plants and animals. Deficiency of this enzyme causes severe psychomotor retardation and ACIDOSIS, LACTIC in infants.
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Hi every one,
I am extracting enzymatic extract from plant (50 mM Sodium phosphate buffer (pH 7.8), 0.1 mM EDTA, 0.1 % (v/v) Triton; 1 mM PMSF). I mesure the protein content and enzymatic activity fom this extract.
So I want and I need to know if the enzymes extract can be conserved for further enzymatic activity measurement (SOD, CAT, APX, ...).
I really need your help, thank you very much
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I have been successfully storing at -80 °C, making a separate working solution for other immediate assays, to avoid multiple freeze-thaw.
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If I isolate/enrich lysosomes from mammalian cells through either differential centrifugation or using commercial kits, what is the best method of storage to preserve enzymatic activity? Is there a specific buffer that would be the most appropriate? Organelles should be intact at this point so would slow freezing as with cells be more appropriate than snap freezing with liquid nitrogen?
NB: structural integrity of lysosomes is less important than enzymatic activity.
Thank you,
Sam
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Thank you for your suggestion Maurice Ekpenyong . Although I’m hoping to look at enzyme activity rather than any cytochemistry. Reading the paper, it sounds like DMSO and snap freezing with LN2 similar to Freezing live cells would be enough. I may have to run a comparison.
best wishes.
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I'm studying digestive enzyme supplements and a large proportion of them contain a wide variety of proteases (Serratiopeptidase, DPPIV-peptidase, pepsin and others) as well as some other, mainly carbohydrate-digesting, enzymes. These are all often contained within the same non-enterically-coated capsule so liquid can leach inside once the capsule is consumed. Would it be expected that the proteases start digesting each other as well as the other enzymes, and would this then reduce the effectiveness of the supplement?
Thank you for any answers
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Hi. Yes . It breakd down proteins into amino acids. lipase enzymes break down lipids (fats and oils) into fatty acids and glycerol.
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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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For some reason, we need to have sodium chloride (NaCl, 0.05-0.1M) in our system. Will this affect the reverse transcriptase and block the SMART reverse transcript?
Thanks
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1. Km much, much lower than the prevailing substrate concentration ([S] >> Km)?
2. Km around the prevailing substrate concentration ([S] ≅Km)?
3. Km much, much higher than the prevailing substrate concentration ([S] <<Km)?
Please give examples too. Thank you!
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Hi there,
I think the best example to illustrate your question is the hexokinase/glucokinase which are in charge of the first step of glycolysis. Hexokinase exhibits a low Km compared to glucose concentration because it has the role to incorporate glucose into glycolysis in order to produce ATP at the end. As the cell needs ATP at any time hexokinase exhibit a low Km compared to glucose concentration in cell. Then its activity is more or less constant and near maximal capacity whatever the situation. Glucokinase has the same activity as hexokinase but only at the hepatic level (whereas hexokinase acts in any cell). Its role is to sense glucose level in order to regulate insulin/glucagon balance therefore glucokinase exhibits a higher Km so that glucose concentration variation will induce also a variation in glucokinase activity which will initiate a cellular response in order to optimize glucose metabolism. So basically the same reaction is used in 2 different ways by 2 different enzymes belonging to the same enzyme family: their properties are in accordance with their respective roles in the cell.
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I was wondering if the methods of enzyme immobilisation reduce enzymatic activity. I am aware that everything dependes on enzyme kinetics and the type of immobilisation, ¿but do the enzymes immobilised via covalent or ionic bonding experience a reduction in their catalytic activity?
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  • The equilibrium constants that describe binding of it substrate to the enzyme and dissociation from (k1 and k2 and hence Km) it will change, simply because the substrate can no longer approach the enzyme from all angles. One way to look at this is that the effective substrate concentration seen by the enzyme will be lower than the molar concentration.
  • The electrical charges of the protein and the counterions (some of them on the matrix) will create an electrical double layer which charged substrates have difficulty crossing.
  • Some enzyme reactions produce or consume protons, creating a pH gradient between the surface and the bulk medium.
  • Binding of the enzyme to the matrix may change its conformation and/or its flexibility, which would affect kcat and hence Vmax.
  • The velocity of the enzyme inside a matrix will increase with [S] only until diffusion of substrate and product through the pores of the matrix become rate limiting, this lowers apparent Vmax. Increasing [E] beyond an optimal value will increase enzyme cost w/o the benefit of increased reaction rates. This effect depends on the Sherwood number Sh of the system.
For all these reasons (and may be some more) the measured enzymatic activity will be lower by a stationary efficiency factor ηs = turnover of immobilised enzyme / turnover by free enzyme ≤ 1, or by the operational efficiency ηo = τf / τi ≤ 1 (where τ is the relaxation constant, the time for [S] to drop to 1/e ≈ 37% of the starting value). Note that the proper way to determine τf is after destruction of the matrix by ultrasound.
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I want to study the activity of a mannosidase against a substrate using a 96-well plate.
Following the protocol, the reaction lasts for 1h and I will end up with a absorvance value at 450nm.
How can I convert that to enzymatic activity (umol/min/mg)?
I know I have to create a standard curve but, should I create it using known concentrations of the product formed (mannose) or would be okay to do it using known concentrations of the enzyme?
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Knowing you are wanting to measure the Enzymatic Activity of a Mannosidase, you should be using a p-Nitrophenyl Mannopyranoside as a substrate, where the product you measure in the reaction is the P-Nitrophenol, which has a specific Extinction Coefficient. You can then simply measure the Activity of your mannosidase in the appropriate buffer system and using the appropriate Stop reagent, to obtain a Delta A405 of a test sample verus a Blank Sample and with Volume of reaction known, dilution of the Mannosidase known, Time of reaction known, and the Extinction Coefficient, obtain your desired umol/min/mg. See example in Attachment.
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I have a batch of fish liver samples than I would like to preserve for internal controls during EROD assays. Would lyophilizing the fish liver samples prolong the length of time that I can keep them stored -80C without affecting EROD activity?
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Juan C Pérez-Casanova - I would not recommend freeze drying the basic liver samples - that's a complex tissue to freeze dry and preserve the enzyme. Solid tissues, even small samples, generally take days to freeze dry, and then the challenge is to reduce the residual moister to <2%. Freeze drying can preserve enzymes well, when properly prepared, most PCR reagents and enzymes are freeze dryed, for example. I suggest that you extract your enzyme, aiquot in buffer and freeze dry the aliquots, then store in the freezer. Do watch out though, even freezing can affect activity as there can be a pH shift in buffer, when you freeze.
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I'm preparing enzyme-responsive polymeric nanosystems and according to the manufacturer's information, the enzyme is in
-» "50 mM sodium acetate buffer, 1 mM EDTA, pH 5.0.";
-»"≥10 units/mg protein" and
-» unit definition "One unit is defined as the amount of enzyme that will hydrolyze 1.0 µmol of compound x per min at 40°C, using 100 mM Na+/K+ pH 6.0, with 1.33 mM EDTA and 2 mM DTT as the activation buffer."
The flask has 50 micrograms of enzyme.
Concentration: 0.451 mg/mL.
416.0 U/mgP
I wanted to prepare 0.5 UN/ mL solution.
Thanks very much
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@Marco: you are definitly right! I made a slight calculation error... ;)
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I would like to know how one can decide when to double the volume of total reaction and that of the individual components of an enzymatic reaction such as restriction digestion, PCR, in vitro transcription etc. rather than keeping the same reaction volume while increasing the concentration of the individual components?
For instance, if I want to double the units of T7 RNA polymerase than usual (100 U to 200 U) for in vitro transcription, as I am increasing the template concentration (from 1ug to 10ug), should I keep the total reaction volume constant (50 ul) or should I increase it too (100 ul)?
The example I stated is more of an experimental specific. It would be really helpful to get both, a general reasoning fit for all the enzymatic reactions, and also for a specific experimental setup such as the above.
Thanks in advance !!
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This depends on what you are doing as many components in reactions when not at the correct concentrations can affect the outcome
For example a restriction enzyme (RE) digest has both buffer and enzyme components that are required to be at certain concentrations. Too much RE in the reaction can lead to problems such as star activity (NEB HF enzymes overcome this to a certain extent). Buffers contain salts that at the wrong concentration will also affect the reaction. As a general rule we dilute RE 10 fold per reaction i.e. 1 uL RE per 10uL reaction to keep the glycerol concentration at 5% or less.
As for IVT changing the ratio of components may compromise the reaction as the conditions are no longer ideal. This may well have to be tested empirically. As you increase the amount of template by 10 fold you also need to increase the concentrations of the other components to ensure these are not rate limiting and hence the volume of the reaction.
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Hello. I wanted to relate N mineralization with enzymatic activity in the soil after wheat residue incorporation. Urease is the most widely used enzyme in these kind of studies. However in my experiment, the chemical fertilizer treatment is not ureic. I see studies where they do not specify the type of fertilizer. Is urease suitable to assess N release from plant residues? I have read that nucleic acid can break down into uric acid, but not urea.
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Yes ,you have correct read the Nucleic acid break into uric acid. This is well known in the plant kingdom, where soil bacteria can help breaking down uric acid or its products introduced into the environment by animal excreta or fertilizers. The principal problem in getting rid of uric acid is its poor solubility in water. the total Earth’s DNA base pairs is estimated to weigh 50 billion tons, which translates into a huge abundance of uric acid in our planet .But , Uric acid in the environment is a rich source of nitrogen (33% of its weight) to plant life, hence its important role in the universal food chain. Most animals get rid of uric acid, thereby replenishing the ecosystem with a precious nutritional ingredient. The principal problem in getting rid of uric acid is its poor solubility in water.Animals that cannot afford the energy requirement for intestinal excretion of uric acid prefer to convert it to a more soluble compound, being blessed by a functional uricase enzyme . Mammals other than primates, and carnivorous dipteras, can thus metabolize uric acid into allantoin, which goes with urine. Amphibians and telecosts can take allantoin further down the road to urea, which is even more soluble and readily excreted in urine. Nucleic acids are the main information-carrying molecules of the cell, and, by directing the process of protein synthesis, they determine the inherited characteristics of every living thing. The two main classes of nucleic acids are deoxyribonucleic acid (DNA) and ribonucleic acid (RNA).
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I have Started Working on Diabetic Rats and I need to assess the activity of below mentioned Enzymatic activity
Hexokinase
Aldolase
Phosphoglucoisomerase
Glucose-6-phosphatase
Fructose-1, 6-diphosphatase
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A good place to look for protocols to assay basic metabolic enzymes is at Worthington Biochemical's web site.
Here is an example of a protocol for hexokinase.
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I'm new to enzymes and I would like to receive all possible help for this. I have a product that initially contains 57.9g of starch, which I would like to hydrolyze with alpha amylase, that has an activity of 1,11,793 U/G. How much grams of this enzyme should I add to the product so that all of the starch (57.9g) breaks down?
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hey guys,
i want to know what enzymes are there in draft beer, which produced by beer yeast?
as far as we know there are: Protease A, acetyl coenzyme A and Sucrose Invertase, and what else? anyone have a list of them?
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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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I’ve read that soil samples for soil enzyme assays should be kept cool and at or near field moisture levels.  But I cannot find information on how long these samples are viable for under these conditions.  Is a month or 2 in the fridge too long to wait to perform the assay?  Also, is freezing the samples until they are all ready to be analyzed acceptable, or would this compromise the sample?  Reason being I plan on having samples collected from different sites over a 2-3 month time period and need to coordinate sample collection, storage, and shipment.  And this leads to my next question…
I’ve also read that comparing soil enzyme assay data from different sites and times is not recommended due to differences in site soil conditions (pH, temp, moisture, etc.) as well as changing conditions over time (rain events, etc.).  Is there a way to overcome this, maybe by controlling moisture content?  I am essentially trying to compare a treatment effect (+ vs -) on soils under 2 plant types, grown in 3 different climates, and over time following treatment.  Any advice here would be appreciated.
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Hi Michael,
Fresh samples are always best analysed. However, experience shows that freezing is less harmful than drying samples. Although freezing also gives different results than storing at 4°C. However, the effect of the storage method itself is different depending on the soil type.
According to Dadenko et al. the greatest changes occur during the 12 weeks of sample storage. After that they are less pronounced.
So to be able to compare, samples should be stored under the same conditions and for the same amount of time before analysis. Or they should be analysed freshly just after collection.
See:
and a similar question:
And as far as the changing conditions of the soil environment are concerned, the influence of factors other than fertilisation etc. on enzyme activity should always be taken into account when analysing the results. That is, compare it with the parameters of each soil and, of course, include moisture and temperature in the statistical analysis. Ideally, the controls should be the same at each site without any additional factors, i.e.: without fertilisation and without plant; with fertilisation and with plant; without fertilisation and with plant. Then you would get the result of the activity of this soil under these climatic conditions and then you can compare the different soils with each other.
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The experiment has total of 16 days and its collect sample in 0 hours, 24 hours and 48 hours (7 days) and in 14 days the same. Doing the analyses in the spectrophotometer, I checked that in 7 days did not show any enzymatic activity and in 14 days it present a good enzymatic activity of 144 U/l in 48 hours.
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Hello Miyu
Laccase is an inducible enzyme, I see your medium too rich in C source and nutrients, this could cause that the fungi repress the laccase enzyme, because is not required to obtain carbon. Remember that laccase is used to degrade lignin and access carbohydrates to the lignocellulose substrates.
I could be possible that, when sucrose decreases, the fungi start to produce laccase. I recommend you decrease the sucrose or use another carbon source, a lignocellulose substrate could work.
Good luck!
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I have an experiment with two levels of two different enzyme and a control treatment (without enzyme) in a completely randomized design. Therefore, a 2 x 2 + 1 factorial design. Can I modeling like a nested design or exist a different model for it?
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Dear all researchers
In the following manuscript, there was a factorial design including thirteen treatment groups: no added α-tocopherol in the feed (0 dose) or four different doses (50, 75, 100 and 150 mg/kg of diet) of three sources of α-tocopherol (RRR-α-tocopherol, RRR-α-tocopheryl acetate or all-rac-α-tocopheryl acetate). As you can see, there is just a control group (0 dose), and the most important highlight in paper is to find the effect of source, dose, time and interactions.
What we did to resolve the problem was to randomize control group between three sources. Therefore, for the analysis, we could have a control per each source. There are two important points that you need to be aware of: 1) you need to have enough replicates to distribute between treatments (in our case, we had 12 replicates, and after randomization of control, we still had four replicates per each treatment, and 2) after randomization, you have incomplete design for the analysis (due to different level of replicates in control compare to others). After passing the peer review in nature scientific reports, I think it can be an alternative approach in this kind of study design. For more details, please have a look on the following link.
Best regards
Saman
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GDP is the product of GTPase reaction. In the biochemical setup containing enzyme GTP and GDP, GDP acts as an inhibitor as the enzymatic activity was found to decreased. Km increases and vmax and kcat decreases.
The kinetic parameters of the above reaction are same as in mixed inhibition. Some of the previous reports stated that Product inhibition is a part of mixed inhibition where product of the reaction act as an inhibitor.
What can be a major difference between product and mixed inhibition?
If the above reaction is not a mixed inhibition then what else it could be?
Kindly help me in this please.
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Dear Engelbert Buxbaum, The question was directed to the author of the question, and not to you. For me the question was not clear and therefore I asked for a further explanation. However, it just so happens that I know perfectly well how to describe an enzymatic reaction with the accompanying mixed inhibition and product inhibition - and more. I would like to add that the times when graphical methods were used are gone. Best regards,
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I have 4 conditions to be investigated:
  • Enzyme concentration (3 or more levels)
  • Substrate concentration (3 or more levels)
  • Solvent: taurocholate (6 mM), taurocholate (12 mM), Triton X-100 (2%), Triton X-100 (5%)
  • Incubation time (3 or more levels)
1. What's method to optimize those variables before I can conduct the cholesterol esterase inhibition assay? (not the one-factor-at-a-time one)
2. or because my final goal is the inhibition assay, can I optimize the cholesterol esterase inhibition assay? (I do not know what type of inhibition of my extracts)
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Fractional factorial design is a method that is used to find the optimal set of conditions when there are multiple variables, without testing every possible combination:
However, in this case, I don't think that is the right approach.
If you just wanted to find the combination of conditions that gives the largest signal in the assay, you would use the highest enzyme concentration, the highest substrate concentration, and the longest incubation time, then test the 3 solvent (detergent) conditions. But that is not necessarily going to be the optimal result for the inhibition assay.
To properly set up an enzyme assay for inhibition screening, you need to measure the kinetics of the enzyme reaction, specifically the Michaelis constant (Km) for the substrate. To do that, you have to measure the initial rate of the reaction at a range of substrate concentrations.
Since the detergent type and concentration are a consideration, and might affect the Km, you should determine the best detergent condition early on, even before the Km measurement. To do this, you just have to identify a single enzyme and substrate concentration that gives a small but robustly detectable signal in one of the detergent conditions, then see which detergent condition gives the largest signal using that enzyme and substrate concentration. Then measure the substrate Km using that detergent condition.
If you have an assay with a continuous readout, this is a simple task. You just have to set up reactions at various substrate concentrations and monitor them over time, then measure the initial rate at each substrate concentration. You may have to experiment with different enzyme concentrations in order to find the right range to allow you to measure the initial rate in a reasonable amount of time (10-60 minutes, let's say). If the assay is an endpoint assay, meaning you measure the product concentration after stopping the reaction, it's more work because you have to run a separate reaction for every time point in order to measure the initial rates. It is essential to measure initial rates when determining the Km.
Once you have identified a suitable enzyme concentration and measured the substrate Km, you have to decide on the substrate concentration to use in the assay. This choice may depend on what sort of inhibitors you are interested in finding.
If you care only about competitive inhibitors, then you should try to keep the substrate concentration as low as feasible, while still running the reaction under initial rate conditions. This will maximize the sensitivity of the assay to competitive inhibition, allowing you to identify relatively weak competitive inhibitors.
In contrast, uncompetitive inhibitors have the greatest potency when the enzyme is fully saturated with substrate. If those are the ones you want to find, then you should use a substrate concentration at least 5 times the Km, if feasible.
Finally, for noncompetitive inhibitors the substrate concentration doesn't matter. Use whatever is convenient, but make sure to run the reaction under initial rate conditions.
If you don't know what kind of inhibitors you are looking for, set the substrate concentration close to the Km, to balance the sensitivity to competitive and uncompetitive inhibitors.
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Any significant progress in the design or innovation of solid catalyst for biomass hydrolysis and sugar fermentation into alcohols in a biorefinery? If yes, I would like to have some recent updates regarding that subject. Thank you all
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Thank you for your answer,
Pedro Nakasu
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I'm measuring fluorescence intensity of the product in an enzyme reaction in 37 celsius degree. I set a kinetic measurement in 1 hour. The fluorescence intensity of the BLANK ( contains only buffer and substrate ) decreases by time and its graph is not a straight base line. I'm thinking if other samples I'm measuring has the same change as the Blank solution, so should I normalise the data by dividing the blank's fluorescence intensity ?
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Hi Phuong,
I agree with Andrew and Adam's answers and will add a couple other hypotheses worth considering. In general, you are looking for the reason your blank signal has unexpectedly decreased.
1. photobleaching - This will be an exponential decay with time, though it can look linear if the rate is slow relative to your time window. As Adam mentioned, you want to decrease the amount of light reaching your sample. Narrowing the excitation slit will do this. If you want to maintain the same signal level, match this with a wider emission slit. If you don't have control over slits, you can put a neutral-density filter between the excitation source and the sample.
2. instrument drift - I used to work for an instrument manufacturer and drift is just a difficult thing to minimize. Warm up your instrument for longer than the manufacturer recommends -- I suggest at least 1 hour. You may need to do a measurement to completely initialize the instrument.
3. material translocation - you have enzymes in solution, and proteins to some degree like to be at interfaces. Your instrument is examining the middle of the cuvette volume, but with time your proteins will migrate from the bulk solution to the air-water interface or cuvette-water interface to some extent. I have seen papers showing this on the 1 hour timescale. If you can add a stir-bar to your cuvette, this could help maintain solution homogeneity. If there are long times between measurements, maybe take the cuvette out and mix it by inversion between measurements.
If you suspect hypotheses 1 or 2, and cannot totally eliminate the baseline artifact, then subtract the blank. If you think you have material translocation and cannot mix the solution... good luck :)
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I need the standard curve for POX but I didn't find it anywhere in the papers. You help will be highly appreciated.
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Yes, you need to prepare this yourself as the type of buffer matters. You can select different methods for assay of peroxidase by using an appropriate colorimeteric chromogen.
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Based on this paper
,TEV protease can cleave between the Gln and several amino-acids (besides Gly/Ser) with acceptable efficiency in its recognition site.
Therefore, it's practically possible to purify many proteins (without an extra residue at the N-terminal end), by using affinity chromatography.
I was wondering if anyone could share their experience/knowledge using TEV protease to cleave between Gln and Met?
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Sorry, I have no experience with the specific case of Q/M cleavage, but remember that most proteins whose second residue is a small sidechain aminoacid actually have their N-terminal methionine removed by MAP while still being elongated in the ribosome.
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I am trying to measure GUS expression, and was wondering if anyone knew what concentration hydrogen peroxide inhibits the GUS protein???
Thanks ! Any help is appreciated!
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Very much interesting question but haven't worked with it.
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Generally, the 0.25M sucrose solution is the most common organic solution which is used for tissue homogenate preparation. There are many specific buffer solutions which are used for tissue homogenate preparation for specific enzyme assay. Please provide some references about the use of 0.25M sucrose solution and its advantages as well as disadvantages associated with its use over other buffers. Thank you.
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Krishna Pada Singha sir In majority cases sucrose is used in tissue homogenate preparation as it acts as a cushion, and give you better separation of cell fractionation, less contamination between these fraction , also it maintains osmotic strength of the buffer while having low ionic strength. Furthermore, for low ionic strength, sucrose is fine in case of higher ionic strength Mannitol is recommended. Mannitol elevates blood plasma osmolality, resulting in enhanced flow of water from tissues
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I have been trying to test PPO activity in vegetable juice. I extracted the enzymes using a buffer solution containing potassium phosphate buffer, NaCl, Triton X-100 and PVPP. I tested for PPO activity using catechol as substrate and measured the change in absorbance using uv-vis spectrophotometer. However, after the addition of catechol, the reaction mixture turned turbid, which prevented me from obtaining the initial velocity gradient. Has anyone faced this problem? Any help is appreciated.
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Dylan, you address your question to a large range of different professionals who all have their own jargon. Thus, dont use abreviations to avoid miss-interpretation. e.g. PPO has different menaning to say Polymer Chemist, Phyto Chemist, Bio Chemists etc.
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I used Kakade method with by adding 0 to 2 millilitre (i.e., 0, 0.5, 1.0, 1.5 and 2 ml) of extract into duplicate sets of test tubes and each tube extract and adjusted to 2ml with distilled water. After that, 2 ml of trypsin solution [4 mg trypsin (Sigma Chemical) was dissolved in 200 ml 0.001 M HCl] was added to each test tube and kept in water bath at 37°C. To each tube, 5ml BAPNA solution. 
For the calculation, I first took the average of the absorbance values of each tube and divided by 0.01. Then calculated the differences between each tube and divided by the volume of extract to TIU/ml. Then I plotted the TIU/ml against volume of extract and extrapolated to zero. But I dont know how to calculate TIU/g using the extrapolated value. What dilution factor should I use? Im confused. Can someone please help me? 
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many thanks for your usefull explanation regarding to my question.
To determine the trypsin inhibitor of soy I use following reagents:
Trypsin: Trypsin from bovine pancreas ≥ 7,500 BAEE units/mg (Sigma)
Substrate: BAPA N-α-benzoyl-DL-arginine-p-nitroanilide hydrochloride
- Is the mentioned Trypsin from bovine the correct choice or preferable?
- Which kind of Trypsin standard you are used in your work to determine the TI (mg/g)? Could you please give me Information (e.g. manufacture, BAEE- Value, Kind of Trypsin)?
Best regards
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Recently I added catechol to distilled water and incubated it overnight in a shaking incubator. By morning the solution was all brown. What may be the reason? Does the brown coloring signify production of melanin from oxidation of benzoquinone which may be due to non-enzymatic oxidation of catechol to benzoquinone ? 
Please note: in this above experiment there was no enzyme added like catechol oxidase.
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To test auto oxidation you can use argon or nitrogen, keep the system overnight in a closed Slenck (So the is no oxygen on the system) and then evaluate the color at the same time next day
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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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for enzyme assay i am using crude enzyme source instead of pure protein. is there chances that i would be able to get maximum result or enzymatic activity.
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To elaborate on a point made by Engelbert Buxbaum, having available a potent and specific inhibitor of your enzyme of interest in a crude extract is very useful for at least 2 reasons. (1) You can use it to find out whether there is any background activity in the assay not due to your enzyme and, if there is, to measure and subtract it. (2) You can use it to measure the concentration of the enzyme of interest in the crude extract, allowing you to measure kcat (see my first comment for caveats), by doing an active site titration.
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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 have stored the tissues in -80 degree. I need to check the activity of mitochondrial enzymes and Calcium accumulation. I want to clear, how long we can store the tissues, whether mitochondria is stable in stored tissues. 
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Yes, it is possible to isolate mitochondria from frozen tissue. Tissues can be stored for more than 7 years in -80 degree, but isolated mitochondria can be stored for 2 years only. Yes, mitochondria is stable in stored (-80 degree) tissues. 
Important points to be considered to check mitochondrial enzyme activity are
(1) Post mortem interval (PMI) of tissue (more the PMI the more chances to see a reduction in the activity)
(2) how many times the tissue has undergone freeze-thaw. Generally, we check the activity of mitochondria at different freeze-thaws, we find more activity at 3rd freeze-thaw. If it is more than that we see reduction in mitochondrial enzyme activity.
For more information on mitochondrial enzyme assays refer to the attachment and also check other papers from the same team.
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Hi! I've been studying a particular enzyme that in in vitro experiments exhibits two optimal pH's. All other parameters are the same (incubation time, substrate, temperature, salt concentration, substrate and enzyme concentration) except for the pH.
I'm wondering what the possible mechanistic/molecular physical explanation for this is? I've been looking all over the internet but it's hard to find any discussion on multiple optimal pH's. If anyone could point me in the direction of some literature I'd appreciate it.
Thanks in advance!
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Hello, the question is a rather new phenomenon described in some publications.
See attached info that may be useful for you.
I think the multiple optimal pH values can be a result of both, which can cause the conformational changes in both enzyme and substrate depends on the pH value of the reaction mixture.
Goog luck!
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I have been searching the literature about this and I have not found an answer. If the atom that accepts a hydride is made more electrophilic, can this speed up the rate of hydride transfer?
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It will certainly increase the speed of hydride transfer, but it depends mainly on how easily the hydride migrates from the donor molecule (basicity or nucleophilicity of hydride).