Enzyme Activity - Science topic

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.

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.

You can Use Nelson- Somogy Method.

May help you

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

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.

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.

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!!!

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

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!

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!

And what about TAS??

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!

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.

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.

Did you solve your problem?

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.

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.

I'm having the same problem! Any update?

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

Not only ligand concentration but pH is also an important factor of chelation .

Antara Chakravarty

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.

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!

Yep, nitrocellulose is a poor choice

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.

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.

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.

Dear Bohuš Kubala

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

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:

Fortunately this paper is freely available as public full text. I hope it helps.

Good luck with your research!

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!!

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:

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.

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.

Please have a look at some enclosed PDFs...

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.

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.

You may also have a look for this option;

Thermo M-PER™ Mammalian Protein Extraction Reagent for RIPA like non-denaturing lysis buffer...

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.

I have been successfully storing at -80 °C, making a separate working solution for other immediate assays, to avoid multiple freeze-thaw.

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.

Thanks @M Angeles Zorrila Lopez- Perea

Check https://pubmed.ncbi.nlm.nih.gov/22410228/

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.

For all these reasons (and may be some more) the measured enzymatic activity will be lower by a stationary efficiency factor η_s = turnover of immobilised enzyme / turnover by free enzyme ≤ 1, or by the operational efficiency η_o = τ_f / τ_i ≤ 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 τ_f is after destruction of the matrix by ultrasound.

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.

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.

@Marco: you are definitly right! I made a slight calculation error... ;)

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.

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).

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.

Check https://patents.google.com/patent/AU2016220533B2/en

Also check https://scienceforecastoa.com/Articles/JNFSF-V1-E1-1002.pdf

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.

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.

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!

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.

https://www.researchgate.net/publication/348891690_Absorption_of_a-tocopheryl_acetate_is_limited_in_mink_kits_Mustela_vison_during_weaning

Best regards

Saman

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,

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.

Thank you for your answer,

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 :)

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.

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.

Very much interesting question but haven't worked with it.

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

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.

Hi Kai-Lin Pan

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

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

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.

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 k_cat (see my first comment for caveats), by doing an active site titration.

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.

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.

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!

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).