Buffer - Science method

One reason could be uneven heating. Try running the gel at a lower voltage to reduce the amount of heat generated.

Another reason could be that the composition of marker samples and the samples in the adjacent wells differ substantially in ionic strength. Salty samples tend to spread sideways into lanes occupied by low-salt samples.

Make a small 50 - 100 mL glass container of 50% acetic acid by equal volume parts glacial acetic acid and equal parts water

Find a pH meter, calibrate the probe with a pH 7 and pH 4 NIST standard solutions. Dissolve 820.3 milligrams of sodium acetate in about 140 mL of water with a stir bar. Hold the pH meter probe in the solution. Add dropwise 50% acetic acid till pH is 5. If you overshoot, add dropwise sodium hydroxide concentrated stock solution to get the pH up to 5. Then fill with water to final volume 200 mL and adjust the pH with a few drops if it drifts with dilution.

Thank you Mr Adam. I will try it again with reduced buffer concentration.

2-mercaptoethanol (or β-mercaptoethanol) is a liquid with a concentration of 14.2 M. It is very smelly, so use it in a fume hood, if possible.

Sodium dodecyl sulfate is a solid. The dust from SDS can irritant the lungs, so it's a good idea to wear a filter mask when weighing it.

2% in this context means 2 grams/100 ml of solution.

For 100 ml of the solution, you need 2 grams of SDS and 7 ml of β-mercaptoethanol (1 M/14.2 M x 100 ml). To somewhat less than 90 ml of distilled water in a beaker, stirred with a magnetic stirrer, add the β-mercaptoethanol and dissolve the SDS. Bring the volume to 100 ml with distilled water.

To reduce oxidized hemoglobin (Fe3+) to its reduced form (Fe2+), you can use a reducing agent. One commonly used reducing agent for this purpose is sodium dithionite (Na2S2O4). Here's a protocol to reduce Fe3+ hemoglobin to Fe2+:

you can attempt to create a generic PCR buffer with common components and optimize your PCR conditions. Here's a basic recipe for a generic PCR buffer. Generic PCR Buffer:

10x Tris-HCl buffer (pH 8.3 to 9.0)

50 mM KCl

1.5 mM MgCl2 (you may need to optimize this concentration)

You can adjust the pH using concentrated Tris base or HCl. The pH range is broad because different enzymes have varying pH optima. For Taq polymerase, a pH around 8.3 is often used, but some enzymes may work better at a slightly higher pH.

Regarding Mg2+ concentration, you'll need to optimize it. Different polymerases have varying requirements for Mg2+. For a start, you can prepare a range of concentrations (e.g., 1.5 mM, 2.0 mM, 2.5 mM) and test them in your PCR reactions to see which gives you the best results.

The ideal case is to use the same buffer in which the samples are dissolved. If the buffer contains nothing that would interfere in the assay (detergents or organic solvents, for example), it is OK to use distilled water.

See page 6 of this document for a list of interfering substances.

something must have been modified in upstream processing (assuming recombinantly produced mab is the case), leading to scrambled refolding products. Are you able to confirm the protein is holding fully the same PTMs and PMF profile...As Edward Michelini indicated, if all the reagents and even containers, concentrations, and experimental conditions are the same, there might be any change occurred at your protein level.....Since w/o refolding antigen binding capacity, efficacy, and titer cannot be tracked, Mass spec analysis can tell much at this point and may give some clues about your unsuccessful assay reasons.

Dear Himadree Das Just have a look here https://www.aatbio.com/resources/buffer-preparations-and-recipes/hydrochloric-acid-potassium-chloride-buffer-ph-2

Indeed as you already anticipated somewhat it is a bit strange since this combination of just NaCl and HCl does not really give a buffer, but anyway apparently it is still called this way.

Best regards.

Is the elution fraction with 20 - 40 mM Imidazole relatively pure? If so then there should be a problem just elute your protein with 40 mM Imidazole. Maybe other groups studying the same protein are using a different brand of Ni-NTA resin that they bought 6 months ago, whereas yours is five years old. Stripping the Nickel with EDTA and recharging the resin with nickel surely cleans your resin, but prolonged stripping with EDTA causes the Ni-NTA to not work as well as it used to when you bought the resin.

Looks more like a voltage issue during running, sometimes I have observed such band when the buffer in the inner tank leaked out resulting in improper electric field throughout the run. When lanes get overload you would not see such clean sharp bands for the low intensity proteins and it would be a whole merged lane.

This RG thread might answer your question.

Run the gel at constant current (I) about 40 mAmps if one gel or 60 - 80 milliamps if two gels. Constant current results in shorter run times because at constant current your samples and the dye move at the same rate through the gel regardless of resistance of the gel which increases over the run time. So your gel will get hotter at constant current, but your samples will move at the same rate regardless of resistance of the gel. If it gets too hot just lower the current.

I assume you mean it's a stain that is a safer alternative to Ethidium bromide, like SYBR Safe?

You do not need to mix it into your DNA loading buffer or the DNA samples. Just add the SYBR to the melted agarose, swirl well to mix, and pour the gel.

Robert Adolf Brinzer thank you so much! You were really very helpful!

Thanks a lot Didier Fesquet

You could try measuring the pH of your buffers on a different pH meter in someone else's lab, just to double check.

You might also double check your recipes to make sure there wasn't some simple arithmetic error that crept in.

You might see if someone else in your department is also doing blots and borrow some buffers from them to rule out equipment or materials failure.

Dear Shradha Surin

which kind of mutation do you performed?

If the mutation is located outside the histidine tag and the protein is not unfolded by the mutation, theoretically there are no reason why it has to affect the protein binding a Ni-Nta coloumn. It seems that the mutation induce protein unfolding, therefore your protein aggregates (as happen some times when membrane proteins are fused with MBP) and therefore the his-tag are buried inside the aggregates and not able to bind the coloumn.

The overexpression level and solubility in the gel of the WT and mutant protein is comparable?

best regards

Manuele

Referencing the Beer-Lambert Law:

The formula for calculating the total anthocyanin content is based on the Beer-Lambert law, which states that the absorbance of a solution is directly proportional to the concentration of the absorbing substance, the path length of the light, and the molar absorptivity of the substance (REF. 11).

The formula can be derived as follows:

Let C be the concentration of anthocyanins in the solution (mol/L), then according to the Beer-Lambert law, we have:

A=ϵ×l×C

where A is the absorbance at 510 nm, l is the path length of the cuvette (1 cm), and ϵ is the molar absorptivity of cyanidin-3-glucoside (26,900 L/mol$\cdot$cm).

To convert C from mol/L to mg/L, we need to multiply it by the molecular weight of cyanidin-3-glucoside (MW), which is 449.2 g/mol. Therefore, we would have:

C×MW=ϵ×lA​×MW

To account for the dilution factor (DF), which is the ratio of the final volume of the mixed solution to the initial volume of the extract solution, we need to divide both sides by DF. Therefore, we have:

DFC×MW​=ϵ×l×DFA​×MW

Finally, to convert from L to mL, we need to multiply both sides by 1000. Therefore, we have:

DFC×MW​×1000=ϵ×l×DFA​×MW×1000

This then is equivalent to:

Total anthocyanin content (mg/L)=ϵ×l×A×MW×DF×1000

(Devil's always in the details and Appologies: I think I missed keying the X sign (between the l and A) in the original note) 😊

Gel filtration chromatography is not an equilibrium procedure. If the affinity of the proteins in the complex is not high enough at the concentration applied to the column, the proteins will separate as they become diluted while passing through the column. If you start with a much higher protein concentration, you may get some of the complex to remain together through the column.

I haven't tried this but had success with Qiagen's FFPE RNA kit. They have a kit for proteins https://www.qiagen.com/us/products/discovery-and-translational-research/protein-purification/fractionation-and-depletion/qproteome-ffpe-tissue-kit

Thank you for the helpfull tips!

The formula is here if you need to calculate it: https://www.aatbio.com/resources/faq-frequently-asked-questions/What-is-the-absorption-coefficient

Or were you looking for a table somewhere?

There are 3 options for polymer dye buffers:

1. Super bright staining buffer

2. Brilliant stain buffer

3. Brilliant stain buffer plus (more concentrated version, which requires less volume)

One of these buffers should be used when using 2 or more polymer dyes to prevent dye-dye interactions. These buffers don't need to be included in single stain controls. Most compensation issues are due to poor single stains. If you have dim single stains on cells you should consider compensation beads.

I hope this helps. Happy flowing!

Yes. My protein aggregates in the presence of metal.

In general , Why researchers want to remove EDTA from their protein?

The baseline drop in chronoamperometry when using an N2 gas pump for continuous buffer flow in a plastic microfluidics channel could be due to a variety of factors. However, without more information about the specific experimental setup, it is difficult to say for certain. Here are some possible explanations based on the available search results:

Hi there, this is the reason this kit (see below) was invested. It is compatible with almost any buffer.

I hope it helps!

found a brilliant website to makeup buffers at

which gives you the amount of salts to weigh in to achieve a given molarity and pH. Tried it for 1M and 0.1M with sodium tetra-borate-decahydrate /boric acid and got the same problem that the pH shifted down by 0.5pH units when using the higher molarity. Interrestingly, the pH paper stays constant, so who should I trust?

When adjusting the buffer diluted from the pH 8.5 stocksolution (pH 9 to 8.5 using HCl) the pH paper gives a pH of 8.0, thus 0.5 pH units lower than the electrodes. Hope someone has a chance to reproduce this effect or disprove it.

Ok, looking at your progress curves a few things spring to mind. Firstly, V0 is initial rate and should be taken from the first linear part of the curve, before it starts to level off. This is in practice somewhat arbitrary, but if you do this for your experiment here you should get different inital rates.

However, in your case this brings us to another issue, which is that, particularly in the higher substrate tests, your reaction is essentially over before you start measuring, so you will not get accurate initial rate estimates with this data. I would reduce the enzyme concentration or lower the temperature to get curves that look linear for longer (you need to slow down the reaction). Ideally, you highest substrate concebtration curve should look something like your lower concentrations do now.

DTT reduces the Ni-NTA to Nickel solid making a brown precipitate. Did you observe that? Phosphate salts are have low solubility especially at cold temperatures in the cold room so if the sodium phosphate is precipitating, your protein is dissolved in it so it is precipitating with the sodium phosphate. Tris and Hepes are more soluble buffers and are better options and I think Tris is cheaper.

Your protein is least soluble when the pH of the solution equals the isoelectric point of your protein (pI) because the protein is uncharged at the isoelectric point. But for binding to Ni-NTA, the pH of the buffer has to be pH >= 7.8 because at lower pH, the polystidine tag is protonated and so it can't bind to Ni-NTA.

Plug your protein sequence into ProtParam to estimate the isoelectric point:

Try to avoid having the pH of your buffer being near the isoelectric point.

If you need a solution other than dialysis, FASP would be the most convenient approach to perform buffer exchange. Since 8M urea and CHAPS composition is present in your lysis buffer, protein solubility is high and it is not easy to precipitate the targets. Therefore precipitation approaches cannot be recommended unless you perform buffer exchange to get rid of detergent and urea components. By implementing FASP or S-Trap you can perform on membrane cleavage (in solution digest) which occurs at Ambic environment. Alternatively, you may run your sample through PAGE and perform in-gel digestion, this also removes the buffer components. Your lysate buffer is highly denaturant (conveniently). Therefore, if you remove the solubility agents and precipitate them (TCA for instance), you will need similar solubility agents for resuspension, Ambic alone would not be efficient to solubilize proteins.

The concentration of Coomassie Brilliant Blue G-250 (CBB G-250) in Laemmli Buffer for SDS-PAGE can vary depending on the specific protocol and desired staining intensity. However, a commonly used concentration is around 0.025% to 0.05% (w/v). It's advisable to start with a lower concentration and optimize based on your specific needs. Keep in mind that the optimal concentration might also be affected by factors like the protein samples being analyzed and the type of gel system used.

Just to update, I've successfully transferred the small proteins / peptides from 20% acrylamide 1.5mm gel to a PVDF membrane using 250mA for 45 minutes. In case anyone googles this question :)

Roy Cohen Thank you so much!

Hi Melissa Stofberg can you explain more on the incubation of BSA? Thank you!

The potential problem with old dialysis tubing is that it becomes dried out and brittle and has cracks in it. So you could possible lose your sample. But if it looks good, I would test it to be sure it holds liquid and doesn't leak before risking your sample.

This is a case of false postive. The 2° Ab might have bound to an Ag present in sample or any other complemntry thing

To get qualified and specific answers you need to provide a bit more information about your experimental approach. Ismail suggested many different options that may be suitable depending on what your overall experimental setup is. As i read your question you are capturing interactors on a Ni-NTA resin with the bait protein bound and then eluting them with imidazole.

There are a wealth of proteomics sample prep methods nowadays that would get you past the imidazole issues, such as protein aggregation capture (PAC) or SP3, which serve to precipitate your protein onto a bead matrix that can be washed thoroughly before trypsin digestion (if bottom-up proteomics is your method). Alternatively, you may be able to digest your protein directly on the Ni-NTA matrix. A C18 desalting step is usually standard procedure before LC-MS analysis and that should give you nice clean samples overall. Gel-band analysis is another approach that should also work fine (if a bit more laborious).

I would suggest your consult with a proteomics specialist at your institution to figure out the most viable approach for your experiment.

Yes, you can elute DNA or RNA from purification columns using ultrapure water. This is preferable for a lot of downstream applications that are incompatible with EDTA. I almost never use TE buffer.

This product information is for CM Sepharose Fast Flow from CytivaLifeSciences:

Pressure/Flow Specification: 300-600 cm/h, 100 kPa, XK 50/30 column, bed height 15 cm.

This is for CH Sepharose High Performance:

Pressure/Flow Specification Base matrix: 100-200 cm/h, 300 kPa, BioPilot 60/600 column, bed height 30 cm

This is for HiPrep CM FF 16/10 prepacked 20-mol column:

Pressure max. (over the packed bed during operation)1.5 bar [0.15 MPa] (22 psi)

Here is product information from Sigma for CM Sepharose FastFlow, which includes cleaning information, but not pressure limits.

Thanks Pr. Ne be-von-caron for your answer, I will check the manual once again.

Some general information that may help you address your concerns.

1. Protein loading and recovery: The capacity of the Elutrap device to load and recover proteins may depend on factors such as the size and concentration of the proteins, as well as the specific model or size of the device you are using. It is recommended to consult the product manual or contact the manufacturer (GE Healthcare) directly for detailed information on the maximum protein loading capacity and recovery efficiency of the device.

2. Buffer selection: The choice of buffer for protein extraction and purification depends on the specific requirements of your protein and downstream applications. Commonly used buffers include Tris-HCl, phosphate buffers, and various salt solutions. It is important to consider factors such as pH, ionic strength, and compatibility with downstream assays or quantification methods (such as BCA).

3. Compatibility with BCA quantification: BCA (bicinchoninic acid) assay is a widely used method for protein quantification. Most commonly used buffers, such as Tris-HCl and phosphate buffers, are compatible with the BCA assay. However, certain components in the buffer, such as reducing agents or detergents, may interfere with the assay. It is recommended to optimize the BCA assay conditions and perform a compatibility test using your specific buffer and the BCA assay kit you have.

To address specific concerns about the Elutrap device, it is best to consult the product manual, reach out to GE Healthcare's technical support, or contact the manufacturer directly. They will have the most accurate and detailed information about the device, including recommended protocols, buffer compatibility, loading capacity, and recovery efficiency.

Good luck

credit AI tool

There is actually a formula to calculate this using the k factor method. The k-factor is a measure of the sedimentation efficiency, which is inversely proportional to the time required for a particle to reach equilibrium in a centrifugal field. The smaller the k-factor, the faster the pelleting efficiency: https://lab.plygenind.com/compensate-lower-centrifuge-speed-increasing-time

Gel buffer can be reused a few times but eventually nucleases grow and the nature of the buffer changes. It is best to have the same buffer concentration in the gel and the tank and if you are reusing the same running buffer then mix it well before use as it becomes acidic at one end and basic at the other end and results are variable. You may be getting worse results that the others because your bands are smaller so reach the ends of the gel quicker and are more effected by pH and salt effects. You could also try running at half the voltage to avoid heating effects and you may find that the bands look better if they do not run so far and the results will still look good even if run less far

These plates and plaques look much better. It does not look like the water or condensation is affecting anything, but if you are worried then be sure the plates are sufficiently dry before using. You could leave them for a day at room temp or a few hours in the incubator before you use them.

Tris HCl buffer is commonly used in RNA isolation protocols due to its ability to stabilize the pH level, which helps maintain the integrity and stability of RNA during the isolation process.

Tris HCl is a popular choice as it resists pH changes when exposed to acidic or basic conditions, providing a consistent environment for RNA extraction.

The typical concentration of Tris HCl used in RNA isolation is around 10-100 mM, depending on the specific protocol or application.

Hello Yi Sak Kim,

another reason for your tissues problems could be unsufficient fixation. Maybe the tissue is to soft. you can try to solve the problem with those factors which I have mentionend above. Did you perfuse your animals? If yes, leave them for 24-48 h in the fixation solution before you start with your cryoprotction. And you can add 0,25% glutaraldehyde to your 4%PFA- solution to get the tissue firmer.

Hi there,

Instead of using a formula you don't get, calculate the dilution factor as it doesn't require any volume data at first! Demonstration: You have a 1M stock and you want a 50mM solution (0.05M) so the dilution factor is (1M/0.05M)=20. So you need to dilute the stock dilution 20 times. How to dilute 20x? It's simple: take one given volume of the stock and add 19 volumes of diluent! In your case, you need 10mL then these 10mL are representing the 20 volumes of the final dilution which means 1 volume is actually 10/20=0.5mL. So mix 9.5mL of diluent with 0.5mL of the stock and that's it!!!

In a more general context, including any value for stock and final solutions:

Dilution factor DF is the ratio stock/final=DF. The preparation of the dilution consists in mixing (DF-1)volumes of diluent with one volume of the stock.

I see, it is indeed something to take into account, the added ions could be detrimental to the fermentation process... Thank you for your answer

Interesting question:

To prepare an acetate buffer with a pH of 7.98, you need to use a mixture of acetic acid and sodium acetate. Recall the Henderson-Hasselbalch equation ratios: the pH of a buffer is equal to the pKa of the acid plus the log of the ratio of the salt concentration to the acid concentration. Let's do the math: The pKa of acetic acid is 4.76, so you can solve for the ratio of salt to acid:

pH = pKa + log([salt]/[acid])

7.98 = 4.76 + log([salt]/[acid])

log([salt]/[acid]) = 3.22

[salt]/[acid] = 1662

The calculation would indicate that you need 1662 times more sodium acetate than acetic acid in your buffer solution. You can choose any convenient concentration for your buffer, as long as you maintain this ratio. For example - if you want to make a 0.1 M buffer, you need 0.1 M acetic acid and 166.2 M sodium acetate. However, as this is not a practical concentration for sodium acetate, so you might want to use a lower concentration, such as 0.01 M. In this case, you would need 0.01 M acetic acid and 16.62 M sodium acetate.

To make 1 L of this buffer, you would need to dissolve 0.6 g of acetic acid and 1369 g of sodium acetate in water and adjust the volume to 1 L. Alternatively, you can use a buffer calculator to find the exact mass of each component for your desired volume and concentration.

You can also try this, for making an acetate buffer with a pH range of 3.6 to 5.6, but you would need to adjust the pH with NaOH or HCl to reach 7.98.

Evan Kerek and Saddah Ibrahim thank you for your answers

Wolfgang Schechinger Finally, i got the 150 linearity with single point calibration. Thanks for your reply.

Adam B Shapiro My specific operation is as follows: use this to extract NADPH and NADP+ in cells or tissues to make tissue homogenate, then take 50 μl of tissue homogenate and add 100 μl of buffer, incubate for 10 minutes, and then add 10 μl of chromogenic solution. However, it is strange that the reaction seems to be inhibited using this extract.

The components of the buffer are：Tris-Hclbuffer ， Mg2+ ， G6P and G6PDH.

The components of the chromogenic solution are：1-MPMS and WST-8

Thanks. Yes, of course, it is always RNase-free water but my question was whether autoclaving is adequate to inactivate any RNases present?

Hi there,

Use the 10x stock of the buffer specific to the polymerase you intend to use.

According to the product literature from Sigma for a ~0.1 mg/mL (about 0.3 mM) solution of cannabidiol in methanol, there is an absorbance peak at 273 nm with an absorbance of about 0.4. The product page (https://www.sigmaaldrich.com/deepweb/assets/sigmaaldrich/product/documents/266/034/c6395-slbk4474vdat.pdf)

doesn't say what the path length was, but I'll assume it is 1 cm, which is a pretty standard path length for a quartz cuvette. What this means is that your solutions are probably too dilute to detect the 273 nm absorbance on the scale you show. A 10 µM solution would have an absorbance of about 0.013, if it were in methanol.

You diluted the CBD solution from methanol to HEPES, which will have an effect on the absorbance spectrum, and may be a problem for solubility of the compound as well if you try to make a more concentrated solution.

Sirvan Abbasbeigi Thank you sir..

Often it is magnesium phosphate that is the precipitate. Usually you formulate your phosphate buffer to be the right pH based on the mix of mono- and di-basic phosphate that you add and then autoclave. You then add the magnesium when cold from a separate stock solution.

Disulfide bonds should be stable to heating in the absence of reducing agent.

Kits can be very expensive. If you need to do a lot of experiments, it is cheaper (and easy) to prepare the reagents yourself. You only need the enzyme, nitrocefin (https://www.sigmaaldrich.com/US/en/product/mm/484400),

a buffer, and microplates if using a plate reader.

Prepare a concentrated nitrocefin stock solution by dissolving the solid in DMSO. Store the stock solution in small portions in the freezer (-80 if you have one). It will last a long time if stored properly.

For Class A and C enzymes, you can use almost anything for the buffer, such as 0.1 M sodium phosphate buffer at pH 7.0. For some class D (OXA) enzymes, you have to include sodium bicarbonate, at 10-50 mM. For class B metalloenzymes (e.g. NDM) , you should avoid phosphate buffer because you have to add Zn²⁺. I've used HEPES at pH 7.0 with 1 µM ZnSO₄.

If you are using microplates for the assays, you should also include a little bit of a nonionic detergent in the buffer to keep the enzyme from sticking to the plate. I've used 0.005% Triton X-100.

You can buy 96-well clear polystyrene microplates with flat-bottom wells in packages of 25-100 plates. The least expensive ones are not sterile, and not tissue-culture-treated. These work very well. There is a significant cost outlay, but it will save in the end if you need to do many experiments. An example is VWR catalog number 76446-956, 100 plates for about $200.

One method is to prepare 0.2 M solutions of each, then mix them together in a certain ratio to get the desired pH. You can use a pH meter to monitor the pH as you gradually mix one solution into the other while stirring with a magnetic stirrer. Keep in mind that pH is temperature-dependent.

Another method is to use a calculation. For example, here is an online calculator:

You may also be able to find a table that gives the ratios of the two solutions for various pHs, although it is more usual for such tables to give recipes for 0.1 M buffers.

You can repeat the binding experiments with different experimental conditions, with the goal of getting more data points to see if you can fit to 2 different binding sites. It looks like you have enough of a heat change with your current concentrations to try smaller injection volumes (like 1 uL) - if you have the MicroCal PEAQ-ITC or ITC 200 you can do 38 or 39 injections.

If you contact me here I will help - https://www.zimmerpeacocktech.com/contact-support/

Hello,

Yes Robert, I am using the same protein as in the paper.

Thank you, Sunit. I was thinking about that too. Do you also use NBS plates as well?

Thank you for your help!

The problem is not what you loaded into the wells since it spans the entire gel. It looks to me like something in the upper tank buffer is causing this dark band.

The problem with buffered peptone water is that you are likely to get some cell growth over time. So if you are doing your serial dilutions and plating quickly then it may not matter, but if the cells are going to be resuspended for some period of time then there might be time for the bacteria to grow.

is it wrapped in clingfilm?

If it is wet it with methanol, this stops any static occurring between the clingfilm.

Hi there,

Coomassie staining might not be sensitive enough.

You may need to go to WB.

The bands you have might just be antibody HC and LC...

You can directly dissolve it in RPMI. the concentration that I use usually is 50 unit/ml of Benzonase in medium and it works perfectly!

The density of the sample should be greater than the density of the solution in the well, so that the sample sinks when dispensed. Rincse out the wells with the upper chamber buffer before loading the samples, as suggested by Paul Rutland . If that isn't sufficient to solve the problem, make a denser sample buffer by increasing the concentration of glycerol.

You are welcome

What quantities of each reagent do you use?