Buffer - Science method

I want to prepare 0.05 M bicarbonate carbonate, pH 9.6 as coating buffer for ELISA.

Could anyone help me with right recipe, procedure, and or reference in the preparation?

Thank you in advance.

I am purifying 1-deoxy-D-xylulose 5-phosphate synthase (DXS) enzyme in E. coli BL21 (DE3) cells. I used one-step nickel chromatography to purify it, but I analysed the sample in the SEC column, and it showed an aggregation peak that eluted in the void column. I followed the protocol of a student who have already graduated. I also analysed her sample, it also contained the aggregation peak but was much smaller than that of mine.

Now, I am confusing what this aggregation is, where it occurs, and how to prevent it.

Media: M9/H2O

Lysis buffer: 20 mM Tris, 250 mM NaCl, 2 mM MgCl2, 0.5 mM TPP, 5mM DTT, pH 8.0 containing Complete protease inhibitors, 250U/mL Benzonase. The cells were lysed by sonication (20s on, 40s off, 70%, 5min) after incubating on ice for 30min, then centrifuged at 19,000g, 40min at 4 °C, the supernatant was filtered and applied to Histrap HP 5ml column.

IMAC Equilibration buffer: 20 mM Tris, 250 mM NaCl, 2 mM MgCl2, 0.5 mM TPP, 5mM DTT pH 8.0. 60mM imidazole as the washing buffer and 200mM imidazole as the elution buffer.

DXS is a dimer, the WM of the monomer is ~68kD.

I analysed the fraction without any treatment after nickel purification, it also showed two peaks as shown in Figure A. I have collected these two peaks, the A260/A280 of the first peak (aggregation) is ~1.0 and that of the second one is ~0.7. Did this mean my protein associated with the nucleus? The SDS-PAGE results are in Figure B, and the Native-gel results are in Figure C. In the native gel, the aggregation couldn't run into it.

I have changed the sonication to cell disruption, but it didn't work. All I could think of was to improve the lysis step, and I was not quite sure if the expression step could affect the aggregation. The Bensonaze I used is >90%, and I'd like to use a higher-purity one. Will this help? Did anyone meet this before, how to improve it? Many thanks for your suggestions.

I am performing western blot and recently i have been obtaining faint bands for the samples i had already run and had got darker bands. I wish to determine the concentration of protein in those samples, but they are now gel-ready (loaded in laemmlli buffer). can anyone please suggest a way.

Hello, I am currently trying to optimise a bacterial expression system using BL2 1DE3 bacteria and IPTG. In order to just have a quick look, whether I can see any protein overexpression on a Coomassie stain, I was wondering: can I lyse my bacterial pellets in 4x Laemmli buffer (+5% beta mercaptoethanol) for a whole cell lysate and then directly run this on a gel?

Problem 1: In my research, I encountered an issue with my wet transblot procedure: despite using 8-10ug of RNA sample and applying a voltage of 10 v for 2 hours in TBE transfer buffer, I observed incomplete transfer of the top band from the UREA gel (8M) to the nylon membrane upon examination.

Problem 2: for northern blot, I conducted prehybridization at temperatures ranging from 55°C, 60 °C, and 65°C, followed by membrane washing with SSC buffer and blocking with blocking solution. Subsequently, I proceeded wash the membrane in wash buffer and soak in detection buffer and applied CDP-Star on top of the membrane. The entire membrane exhibited fluorescence (is it normal?), later the resulting X-ray film exposure did not reveal the desired bands. Background noise quite bad. I would appreciate any professional guidance or suggestions to address this discrepancy."

Dear researchers

How to prepare buffer solution with pH of 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13 using NaH2PO4 and Na2HPO4?

Should we use NaOH and HCl?

I'm a beginner in ITC, does anyone have an idea why when measuring the heat of dilution and in measuring the interaction of HSA with the drug or even with the buffer itself, the curve goes up? HSA measurement at HEPES pH 7.4 and pH 6 give similar results. Different buffer concentrations and different ionic strength do not solve the problem...

how to prepare 0.02 M sodium phosphate buffer (containing 6 mM NaCl, pH 6.9)

I am running SDS-Page western blot using 10% acrylamide gels. However, my samples are not migrating more than 55 kDa. The bands are not defined. I am using 4x Laemmli buffer with LDS from Biorad. The cell lysates are human whole brain lysates. I am wondering if the LDS has something to do with this? I tried to boil the samples at 95 degrees for 5 min; heat at 70 degrees for 10 min, all did not work.

We received a limited amount of an antiserum antibody as a gift from another lab and are trying to avoid having to purify it. Our initial titration blot with blocking in 5% milk successfully detected the target protein, but encountered significant non-specific binding likely from high albumin-IgG in the serum.

Would using BSA for blocking decrease the non-specific binding, or could it exacerbate the issue due to additional albumin from the BSA? We have never performed westerns with unpurified antiserum antibodies before so any help or tips would be appreciated!

Edit: This is NOT a phospho-specific antibody

After treating my membrane with the primary antibody, the destaining buffer was accidentally used instead of tbst buffer. Immediately after use (about 3 seconds later) it was replaced with tbst buffer (10 minutes, 4 times wash), will it affect the antibody signal? The composition of the destaining buffer is acetic acid, methanol, and 3dw.

I have problem for removing polysccharides in the dna extraction of bacteria. when I want to collect supernatant after adding extraction buffer, suppernatant isn't sufficient and DNA concentration is law and isn't sharp. But RNA qualifiction and quantification is better than DNA. Can anyone help me how can I improve the quantication of DNA?

I have animal feed sample. I am using 1 g feed and 5 ml buffer. After protein extraction. I diluted my sample 20 and 40 times. but the concentration and Percentage of protein varies in both dilutions but sample is same. For example, for 20 times dilution, i got 12% and for 40 times dilution, i got 14 %. I need same values for every dilution of same sample.

For separation of Human from bacterial DNA

I made a buffer solution dissolving 79 g of NaH2PO4.2H2O in 1000 mL DI water. Then tried to adjust the solution pH to 6.0 using HCl or NaOH. Initial pH was below 6, so I added NaOH into the solution. It started increasing the pH gradually, and solution pH came to 5.3. Then I added more NaOH (drop by drop and stirred thoroughly for a good amount of time) but pH was constant at 5.3 for quite some time. Then suddenly it jumped to 7.02. After that, I added HCl (drop by drop and stirred thoroughly) to decrease the pH, but nothing happened. pH was constant at 7.02 for a long time, then suddenly dropped to 5.3. Tried several times but pH jumps between 5.3 to 7.02 and vice versa. I can't seem to find any pH level between these two values. What should I do?

I tried glycine buffer but the yeast cells acidify the buffer so the pH goes down to 7-6 overnight. I need something that will stay around 9 and that isn't toxic to the cells.

Hello, everybody.

I am looking to add a DNAse treatment step to my bacterial lysis (gram-negative) procedure for crude enzyme extraction because my lysate always ends up being too viscous. We only have Zymo Research DNAse I in the lab and its information sheet says not to "avoid phosphate buffer and calcium chelators". However, I am using a chemical lysis procedure using Promega Cell Culture Lysis Reagent (following their bacterial lysis protocol) since we do not have a sonicator available. According to their information sheet, CCLR has 25mM Tris-phosphate (pH 7.8) and 2mM 1,2-diaminocyclohexane-N,N,N´,N´-tetraacetic acid. Does anyone know if adding DNAse I to my lysate will still work?

I am not sure of the composition of Zymo's DNA digestion buffer, but I was thinking of supplementing divalent ions to the lysate to counter the 2mM chelating agent in the lysis buffer. However, I am not too sure how to circumvent the phosphate problem. I am not sure how phosphate affects DNAse activity exactly. On the other hand, Thermofisher has a protocol for removing DNA from protein extracts (extracted using lysis reagent in phosphate buffer) using DNAse I. Will the phosphate component of my buffer significantly affect the activity of the DNAse?

Any insight on this matter will be greatly appreciated. Tips on how to solve the viscosity problem altogether are also greatly appreciated.

Thanks

I have synthesized a sensor, which has sulphonic acid grp (-SO3H) and Boronic acid as well in it. And has a Molar mass of around 899 and also has an amide bond in it.. I am using Reverse phase HPLC to purify using ACN and Water(0.1%) as mobile phase. As I separate the compounds of one peak, the next very day it is splitting into two peaks in the chromatogram. Is TFA creating a problem? I could not figure it out. Should I need to use a buffer as an additive instead of TFA? Which buffer and how much is good for use? Could you please suggest?

My western blot had a problem with the protein band. I just checked it and observed my loading buffer is expired. does anyone explain it is due to expired loading buffer or any other problem i have with it?

Hello, I want to do EMSA with native PAGE to check protein-dna interactions.

The PIs of my proteins are between 8.1-8.5. I know that the pH of my buffer must be higher, so that the net charge is negative and the protein goes "downwards" to the anode. But do I have to adjust the pH (e.g. let's say 9.5) of everything? So separating gel, running buffer and loading dye? Or is the gel enough? I cannot find anything about running buffer and loading dye.

I my group we only did discontinous native gels so far, but in all recipes the pH of the stacking gel is around 6.8. Then my protein would run out of the gel, wouldn't it? Can I also change the pH of the stacking gel without changing the purpose of the stacking gel? I also found continuous native gels on the internet. Does that really work without getting a big smear?

Dear Research Community,

I am encountering a significant hurdle in my research involving enzyme inhibition testing. The inhibitor I am investigating exhibits solubility exclusively in DMSO, rendering it insoluble in aqueous environments such as the 100mM phosphate buffer I am utilizing for enzyme kinetics studies. Upon attempting to incorporate the DMSO-dissolved inhibitor into the reaction mix, it precipitates out, leading to haze formation in the solution and hindering accurate data collection.

I am seeking insights and suggestions on how to effectively address this challenge. Specifically, I am interested in methodologies , or alternative solvents that could facilitate the integration of the inhibitor into the reaction mix without inducing precipitation. Additionally, any advice on modifying experimental conditions or buffer compositions to mitigate this issue would be greatly appreciated.

Thank you in advance for your expertise and assistance.

I regularly have low DNA concentrations on the vetigastropod tissues I extract. I have tried fresh tissue and older/museum tissues. We use the Thermo Scientific GeneJET Genomic DNA Purification Kit.

Heating the elution buffer and using less buffer to elute does help, but concentrations are still quite low. Downstream applications are PCR and sequencing.

Super desperate and would appreciate any suggestions on how to increase the yield from extractions!!

We are interested in detecting aggrecan (250 KDa) by WB. No luck so far, are there any recommendations, such as running buffer, sample loading buffer? Thank you in advance!

I was wondering if the routinely used NativePAGE sample buffer with a pH of 6.8 causes protein precipitation if my protein has a pI of 7? Could I increase the buffer pH without affecting the electrophoresis? Thanks a lot for your help.

i am trying to stain different proteins of interest in human paraffinized section.

my signal should be the vessel only, but regardless of the antibody I get circle shaped spots , I tried antigen retrieval with trypsin, and I tried different dilution of antibodies (1:100-1:1000) and blocking in 5% and 10% donkey serum

why I have those spots? how can I reduce them? I have the same issue at different wave lengths (regardless of secondary antibodies tag)

this is my protocol:

•Thickness of sections : 10µ

•Deparaffinization by heat 55degrees for 20 min then (xylene - xylene: ethanol - ethanol) 10min each*twice

•Hydration (ethanol 95% - 70% - 50% - tab water) 5min each*twice

•Antigen retrieval : citrate buffer 10min microwave then leave in buffer to cool

•Peroxide treatment: 3%H2O2 10min at room temp

•Blocking: 5% Donkey serum + 0.3% Triton-PBS 1 hr RT

•Antibodies:

•Primary in 1%BSA: VWF (1:200)

•Secondary 1:500 of Rabbit 594

Hi,

im looking at PAL activity in blueberry samples. I use L-Phenylalanine as a substrate and absorbance at 290nm. The absorbance values of my blanks are higher than 1, even my milli pore water is above 1. I diluted buffers, substrate... even remade everything. i am out of options.

my protocol is as follow:

acetone powdered tissue extracted with borate buffer pH 8.8 and PVPP.

75ul sample extract

150ul (l-phenylalanine 30uM and borate buffer 30mM)

after 30mins rxn stopped with 4N HCl.

absorbance at 290nm

my latest results are very close to each other (0.8-0.999) my blanks are above 1.

any ideas to what might be the problem?

I purified a protein containing His tag via Ni-NTA chromatography wherein the protein was eluted in elution buffer containing 500mM imidazole, 50 mM Tris, 500mM Nacl. Since my next step of purification was Ion exchange chromatography, I dialyzed the protein in dialysis buffer containing 50mM Tris and 5mM beta mercapthoethanol. However, in the third round of dialysis, I observed extreme precipitation of the protein.

Kindly let me know what can be done to reduce the precipitation.

According to Bradford assay, the concentrations are pretty low. Apparently I added way too much extraction buffer to my lysates. Is there a cheap and reliable way to concentrate my protein extracts?

Dear all,

I am looking for guidance relating to a formula to calculate mass of reagents based on the desired pH and molarity. Is there a formula for this?

I am looking to create, and justify with calculations, a phosphate buffer. I am intending to create phosphate buffers 0.1 M pH 6.0 and 0.2 M pH 6.8.

Thank you in advance for any guidance :)

Kieran

Cannot separate between 10~25kD, always have a line around 25kD...

(Resolving gel buffer: 30% Acrylamide/Bis, 1.5M Tris-Cl, pH 8.8, 10% SDS, 10% APS, TEMED)

We prepared alginate beads 2% in CaCl2 200 mM, after stirring for 1h the beads were washed with water, and incubated with the substrate in the presence of 100 mM tartrate buffer pH 5.5 after 1h incubation we noticed huge weight loss of the beads

I am working with leishmania threonine synthase (LdTS) enzyme of aspartic acid pathway which is ~75kDa protein. I purified the protein and confirmed by western blot too. The condition used for induction is -

0.1mM IPTG concentration at 20°C for 12hours.

For purification I am using Tris buffer and PBS buffer both with Imidazole concentration as by Ni²⁺ NTA chromatography-

The problem which I am facing is that I am non specific bands along with my band of interest. How get rid of this? As well as how I can improve my yield? Currently my yield is 4.146mg/L with Tris and 3.886mg/L with PBS buffer.

I have attached the gel images and blot image below.

As an alternative to looking at the conservation of cysteine residues amongst homologs of a protein of interest, is anyone aware of a server that can predict whether disulphide bond formation within a protein is likely to be required for correct folding/oligomerisation? That way one could add a reducing agent to the buffer to reduce the chances of unwanted aggregates forming. Essentially it might be useful to reduce the need for an optimisation step i.e. detection of aggregation after a gel filtration run.

What are the differences between 50 mM glycine-hydrochloric acid buffer (pH 3) and 100 mM glycine-hydrochloric acid buffer (pH 3), especially regarding osmotic pressure? Why do they have the same pH value but different glycine concentrations.If I need to use it to wash cell , which concentration of glycine buffer is closer to isotonic, thereby ensuring cell survival?

I am trying to purify a his tagged protein using IMAC. The homologs of this protein purify very well. However, this protein precipitates on elution.

Buffer composition- 20mM Tris-HCl pH7.4, 500mM NaCl, 0.5M Imidazole-HCl pH7.4

Since the homologs are soluble I am not expecting any drastic changes in buffer compositions to be required - this might be a bit naive.

One theory I have is that the protein is eluting too concentrated - causing precipitation. Might it be worth eluting the protein into buffer to dilute the fractions and reduce the likelihood of precipitation?

This protein has a signal sequence which was cleaved during cloning. It might be that I have taken away some of the surface charge that makes the protein soluble. Something I will also check.

Any advice is much appreciated!

I have been trying to subclone a gene into the pEGFPC1 vector, and chose BspEI and SalI as my restriction sites. As a control, I tried to perform a single digestion (2hrs, 37 degrees) of the empty vector separately using the two enzymes (BspEI and SalI HF) in NEB Buffer 3.1 (both enzymes show 100% activity as per NEB). However, only BspEI worked, and SalIHF didn't. Could anyone point out why SalI HF was not able to digest the vector in NEB Buffer 3.1?

PS:

i use 1.5% agarose gel and 5ul ethidium bromide for 300ml TAE buffer

I am preparing the NAP Buffer described in Camacho-Sánchez et al., 2013. The protocol calls for dissolution of 700 grams of Ammonium Sulfate in 1L of water. This step is described as "taking hours" and should be done at "low to moderate heat", but I've had the buffer on a hotplate stirrer for more than 16 hours at 55°C and it still doesn't dissolve completely. Has anyone else had this problem? What temperature range do you use? How long does it usually take you?

Thanks!

Hi, This is E. Park. I'm trying to purify a protein which was produced in E. coli as inclusion bodies. I already solubilized the inclusion bodies using a 6M guanidine-HCl buffer. Here is my question: I want to store this solubilized inclusion body for 12 hours at 4 degrees Celsius. Should I store this at a lower temperature (freeze) or is it okay to proceed with this method?

I am new to CO-IP.

I am following this protocol for CO-IP

0,25ml 1M Tris-Cl

0,15ml 5M NaCl

0,5ml 10% NP40

0,25ml 10% Nadeoxycholate

MQ 3,85ml

1/2 pill EDTA free

250ul 1M Tris-Cl

500ul 5M NaCl

50ul 10% NP40

25ul 10% Nadeoxycholate

MQ 4.2ml

1/2 pill EDTA free

250ul 1M Tris-Cl

50ul 10% NP40

25ul 10% Nadeoxycholate

MQ 4.75ml

1/2 pill EDTA free

1- Resuspend in medium

2- count cells

3- Wash once with cold PBS

4- Add 1ml cold lysisbuffer 1 with EDTA free protease inhbitor.

5- transfer to an eppendorf tube

6- Add 5ul benzonase

7- Incubate 30min @ 4Co in a 50ml tube on a roller bank

8- Pass 3-5x through a blue needle

9- Spin 10 min at 10000g at 4Co

10- Transfer supernatant to a new tube

11- Save 150ul for total lysate

12- Add 850ul to 50ul beads agarose A and add 10ul Ab

13- Spin 16 rpm o/n @ 4oC

1- Short spin 20"

2- remove supernatant

3- Wash 1x with lysisbuffer 1

4- Short spin 20"

5- remove supernatant

6- Wash 1x with lysisbuffer 2

7- Short spin 20"

8- remove supernatant

9- Wash 1x with lysisbuffer 3

10- Short spin 20"

11- Remove all wash.

12- Add 150ul elution buffer (0.1M glycine Ph2)

13- Incubate 10' @ RT with rotation (do not exceed 10')

14- Short spin 20"

15- Transfer to a new tube

16- Add 15ul neutralization buffer (Tris HCL ph8.1)

19- Add loading buffer and Boil samples @95oC for 5'

20- Short spin and load on a 4-20% gel

the question is Why I m seeing this massive band along the membrane and how can I optimize the protocol.

thank you in advance

I am been working for a couple of months without success on setting up an assay based on GTPases loading with bodipy gdp and then measure the exchange to GTP in presence of various GEFs.

Reading on literature, the idea is that once the Bodipy GDP is loaded onto the GTPase, there is a significant increase in fluorescence (compared to Bodipy GDP alone) , which decreases when adding GEFs, which exchange it to GTP and thus releasing the Bodipy GDP.

I have been stuck on the first step, because after incubating my GTPase with Bodipy GDP I saw that there was no difference in fluorescence compared to bodipy gdp alone.

Among different protocols that didnt work, here is one:

I store my GTPases in a simple Tris based buffer, tried to buffer exchange them into HEPES buffer that the paper uses but nothing. There are other assays that instead of Hepes use Tris, I have tried those too but nothing.

i think my GTPases cannot load the GDP for some reason and i dont understand why.

If anybody had been through this assay and would like to share any tip in protein storage handling or the assay i would be grateful.

thank you!

hello ，I want to use exogenous ligands to activate receptors on the surface of the U87 cell line. How can I remove endogenous ligands before the experiment to ensure a low baseline receptor activation level? I tried treating the cells with a glycine buffer at pH 2.7 (30 seconds * 3 times), but after treatment, my cells died. How can I optimize my experimental conditions? thanks for your kindly help.

I want to prepare reduced form of NADPH. I am using 100mM of Tris (pH8.0) to make the buffer but the absorbance is showing to be 1.6 at 340nm which should supposed to be 0.6.

can someone please help me to resolve this?

Hi everyone,

Recently, I bought a new cell line named Tenocytes from a company.

I followed the manufacturer's instructions and used their medium and coating buffer.

However, I observed that the cell was not attached to the bottom, as shown in the pictures I attached below.

As you can observe here, I saw all cells are still alive. However, they do not attach to the bottom.

I would greatly appreciate your suggestions or any advice for my experiment.

Best regards,

I am working on quantifying alkaloids in a plant extract. I want to prepare a phosphate buffer solution with sodium phosphate dibasic dihydrate with citric acid at pH 4.7. However, sodium phosphate did not dissolve in water. Kindly request a method to dissolve it properly.

Thank you!!

Exploring the release rate of paclitaxel from a mesoporous silica nanoparticle, prior studies have employed various release mediums, such as PBS buffer, or PBS supplemented with Tween or SDS at varying ratios. I am intrigued by how paclitaxel, being highly hydrophobic, can dissolve in PBS (pH 7.4) without the aid of a surfactant. Can you elucidate the rationale behind selecting the optimal release medium, considering the notion that it should mimic the cellular environment?

Hi, I'm doing experiments reconstituting membrane proteins to liposome.

And have a few questions.

1. If I use buffer while hydration of lipid, buffer can encapsulated into liposome. Then, detergent treatment for membrane protein reconstitution (I usually use 0.75% OG, n-octyl-beta-D-glucoside) results in leakage of buffer from liposome?

2. Will buffer leak from liposome during or after detergent removal by dialysis?

I digested my cas9 vector by adding 5ug of vector, 3ul of NEB 2.1 buffer, 3ul of BbsI enzyme and completed to 30ul with ultra-pure H2O.

I incubated overnight at 37C and added 1 ul of CIAP incubated for 10 min at 37C.

I ran my digested vector on an agarose gel and there is no visible band.

Although I can see a band for the intact cas9 vector, I don't for the digested one.

I had already made this digestion and I could see a band before, now it disappeared and I tried to make some new digested vector and there is no visible band as well.

I don't understand what is happening, if anyone has any idea.

Further, how does one calculate the pH of such a buffer solution? Using a pH meter is impossible due to the high viscosity of the buffer solution... Any chemists here?

I am blocking some microwells with PBS + BSA 1.0 % and then sensitizing them with LPS. Please let me know what range of absorbance values I should expect for a microwell without sensitizing with LPS, but with the blocking buffer used.

Hello everyone,

I use several pcDNA3.1 expression vectors to transfect cells.

The vectors were prepared by midi-prep a year ago and diluted in TE buffer.

Now that I run new experiments, I decided to measure plasmid concentrations again, prior to transfection.

All their concentration have droped by 2 to 3-fold.

260/280 ratio are still good (over 1.8), but strangely 260/230 ratio have risen (from 2 to 2.3~2.5).

Given the good 260/280 ratio, the presence of EDTA in the buffer and the -20°C storage, I'm pretty sure it is not degradation.

It could be adsorption of DNA on eppendorf tube wall but given the 100~500ng/µL range of concentration, I don't think any tube surface could sequester this much vector quantity.

Anyway I heated my vector for 15min to 60°C and votexed it without increasing the measured concentration ?

The only thing I see would be freeze/thaw cycle maybe ? (I did 10 to 20 such cycles...)

Should I add glycerol to my TE so that freezing and ice crystals don't shear my vector ?

Or just aliquot my vector?

Where did my vectors go guys ???? ^^

Thanks for the help you can provide,

Philippe.

Typical neutralization buffer used for Protein A/G affinity separations is 1 M Tris/HCl pH 9.0

Any alternate buffer for 1 M Tris/HCl ?

I am doing protein estimation from fish tissue by Bradford's method but after adding the reagent cbbg- 50mg, ethanol-50ml, ortho-phosphoric acid- 100ml in 1L soln precipitating clot is shown even in the standard BSA solutiin. Each test tube contain. 0.1 ml sample+0.9 ml phosphate buffer+5ml bf reagent.

I have been running a series of ITC to look at the binding affinity of Fluoxetine Hydrochloride (Prozac) (0.5mM) with Human Serum Albumin (0.02mM). Prozac is my ligand and HSA is in my sample cell. I am using a 0.1M Tris base buffer at pH 7.4. I have been adding runs at different concentrations of ethanol (4%, 8%, 12%, 16%, in both the cell and syringe) to see if alcohol will affect binding, and the magnitude of the peaks significantly increased in the negative. Then I ran a "water into water" using buffer at 12% EtOH in both the syringe and the cell and I still was seeing large peaks. I am interpreting these as the EtOH and my buffer were reacting. But I don't understand how that could be if both the sample cell and titrant were at the same concentration of ethanol. Does anyone have any idea what could be happening? I am an undergraduate Biology major and Chemistry has not always been my strongest subject.

Hi! I used the NEB PNGase-F denaturing protocol to denature some lysates and ran them on Western. I usually load 20µg of protein on western; so I did that with my un-denatured lysate control as well as my lysates, using the same amount of the lysate/denatured lysate in the sample prep

2µL lysate/denatured lysate

2µL DTT

3µL lammeli buffer sds (6x)

8µL MilliQ water

Samples were then briefly vortexed, spun, and boiled at 95C for 5 minutes before being run on a gel and probed for the protein. The denatured protein samples show the correct band I want, but when using GAPDH as my loading control, it is evident that the same amount of protein was not loaded. I cannot find anything on the internet about using different amounts of protein for loading after this...

I also made a silly mistake after and thought I could BCA my denatured samples to get the exact concentration of them, but all the glycerol interfered with the BCA and gave super high protein results.

I want to check the mTORC activity and for that I am using rat kidney. I used cell lytic buffer from Sigma Aldrich to extract the proteins from rat kidney (50mg sample homogenized in 200ul cell lytic buffer with Halt protease and phosphatase inhibitor cocktail followed by centrifugation at 13000 g for 15 minutes), took the protein concentration by BCA assay, and then ran SDS-PAGE at 80V for 1.5 hours. I used BioRad's 10X Tris/Glycine/SDS electrophoresis buffer. I then did the transfer to PVDF at 0.18A for 1.5 hours. I used 10X Tris/Glycine as a transfer buffer. I incubated the blot with S6 primary antibody at 4 degrees overnight, washed with TBST 3 times and incubated with secondary antibody for 2 hours at 4 degrees followed by TBST wash 3 times.

When I developed the membrane, I see the bands are not aligned. I see it smeared. Can anyone help me point out what could be the issue? Shall I lower the voltage while running the gel? or the problem could be something else?

I've tried to purify recombinant protein from E.coli and I got a good single band in small-scale while several unspecific bands presented in LARGE-SCALE. I tend to Re-Purify the targeted protein (which still in Elution buffer that contain 250 mM imidazole) using Ni-NTA agarose gel.

Hello everyone,

I need a citrate buffer with an optimal pH of 4.8 for the enzymes I'm working with. I'm using citric acid monohydrate (molecular weight: 210.14 g/mol) and adjusting the pH with NaOH. I'm preparing it as a 10x concentration and diluting it to 1x in the final volume (15-200 µl).

However, I've come across recipes for citrate buffer that use both sodium citrate and citric acid.

My question is whether the buffer I'm making will be strong enough to maintain a pH of 4.8 when I dilute it to 1x in my sample. Is the recipe with sodium citrate and citric acid a better option for buffering at pH 4.8?

I'm currently using a Qiagen Kit for DNA extraction, I saw white precipitate in the P1 buffer, I was wondering if it's normal and will it effect my DNA yield ?

Thank you in andvance

Hi Everybody. I'm senior in a University in Vietnam. This is fisrt time I do SDS-PAGE, so I want to ask about the correction of my loading buffer recipe for SDS-PAGE as a following file.

In addition, If 0.5M Tris-HCl pH 6.8 isn't available, Can I replace it by 1M Tris-HCl pH 6.8 or any other concentrations of Tris-HCl pH 6.8.

Also, how should I mix these components? (In order or out of order)

(Samples I am using in my research is piglet intestines)

The detailed Western Blot procedure

The membrane is first rinsed briefly with 20ml of TBS, followed by a 30-minute incubation, shaking with 10ml of Blocking Buffer. During this incubation, the volume of primary antibody needed for a 1:5000 dilution into 10ml of blocking buffer is calculated, and 10ml of blocking buffer is prepared for the antibody. Subsequently, the Blocking Buffer is removed, and the membrane is incubated for 45 minutes, shaking with 10ml of primary antibody in Blocking Buffer (Rabbit anti-ADH diluted 1:5000). After removing the antibody, the membrane is washed three times for 5 minutes each with 10ml of Blocking Buffer. Meanwhile, a 1:5000 dilution of the secondary antibody in 10ml of blocking buffer is prepared. The membrane is then incubated for 45 minutes, shaking with 10ml of secondary antibody in Blocking Buffer (Alkaline Phosphatase conjugated Goat anti Rabbit diluted 1:5000). Following the antibody removal, the membrane undergoes three 5-minute washes with 10ml of TBS/T. Finally, 5ml of BCIP/NBT liquid substrate (Sigma-B1911) is added, and incubation continues until color develops, with the reaction being stopped by rinsing with distilled water.

I was told I might have washed it with a different TBS (10mM one instead of 5mM)

What could the reason for such a different bands on the well.

I'd like to prepare 2.5% glutaraldehyde/2% paraformaldehyde in 0.1 M sodium cacodylic acid. We can buy this pre-made but we only need to use low volume at a time so buying pre-made buffers (minimum order of 100 mL) is not cost-effective. So I plan to prepare this buffer myself and just store by freezing. Can I do this? Will this affect the property of the buffer? Any tips for storing this long term are highly appreciated.

Due to the high cost of Magnetic IP Kits, our group has decided to make the buffers ourselves. We intend to use SDS Lysis buffer for cell lysis, which should work just fine. However I struggle to find clear information on which wash buffer to use. There are lots of suggestions includung plain PBS, PBS-T, Lysis buffer or a self-made wash buffer (Tris, EDTA, EGTA, Triton x100, NaCl and protease inhibitors) Does anybody have experience or suggestions for which wash buffer to use?

Thank you in advance

I have ordered the SOD colorimetric kit (invitrogen) and a GSH/GSSG ratio assay (abcam, ab205811) and was hoping to use the same brain tissue for both kits. In the methods provided the by the manufactures, the GSH/GSSG assay requires PBS/0.5% NP-40 and the SOD assay just requires PBS. Does anyone have experience with these kits and can advise? Could NP-40 be left out if cells are throughly homogenised with a ultrasonic homogeniser since this is just a detergent?

Really appreciate any advice.

I do ELISA many times, and my result is not the same. I realize that when I harvested the supernatant, the volume each well was different. For example I stimulate cells with 200 ul media, when I harvest it, the volume in each well is different and less than 200 ul. So my concern is, should I make all the supernatant at the same volume (200 ul) by adding media, then dilute it with blocking buffer as I want? Or just dilute with blocking buffer directly without making the same volume first?

Thank you very much 🙏

Hello! I am not sure whether or not my protein samples are reduced or not. Based on the information below, can someone please let me know if they are reduced?

__ uL total cell lysate (exact amount calculated from BCA assay)

1 uL Thermo Fischer 10X Sample Reducing Agent

2.5 uL Thermo Fischer 4X LDS Sample Buffer

1X RIPA buffer to dilute sample to 10 uL

I am trying to optimize my control titration for ITC so that I can ensure my ligand to protein traces are reliable.

My small molecule is stored in DMSO. I have been very careful to avoid buffer mismatch. In short I make a 2x stock solution of my compound in 20% DMSO diluted with protein dialysis buffer, and a 2x stock solution of 20% DMSO (without compound) diluted with protein dialysis buffer. The 2x compound stock is added to an equal volume of protein dialysis buffer (species in syringe) and the 2x stock 20% DMSO in protein dialysis buffer without compound is added to an equal volume of protein dialysis buffer (for control) or 2x protein in dialysis buffer (species in sample cell). This results in a final 10% DMSO and 1x ligand/protein solutions or for my control 1x ligand and buffer (for control experiment). This is also all done with locked pipettes.

Upon titrating compound into buffer I am seeing a significant heat transfer but it is not constant or high with a slight linear decrease, which I know is indicative of buffer mismatch.

Is it possible that this compound into buffer just produces a large heat release intrinsically?

I have attached the raw data below.

Hello,

I have recently switched from doing ligand into protein for ITC because my ligand is in DMSO and has low solubility at 10% DMSO; therefore, now I am doing protein into ligand -- the heat transfers are much better and seem to correlate to the protein concentration loaded in the syringe which is good to see.

However, at the end of my titration regardless of the concentration of ligand or protein added there is an increase in heat transfers. I am also seeing this in my protein injections into buffer alone.

Any thoughts on why this might be happening?

I have tested enzyme activity (b-galactosidase) with cation ions in different buffer and the results differ dramatically. For example, while I used MES buffer calcium acts as an activator but when I dissolved my enzyme in phosphate buffer calcium acts as an inhibitor.

is there any explanation besides I done something wrong?

Kindly ask for help and thank you for any answer.

An unopened Sigma-Aldrich (P4557) phenol solution bottle was shaken (prior to the addition of the Equilibration Buffer) and a gel-like layer formed at the bottom of the bottle. The upper phase is still liquid. The bottle was shaken briefly after the phenol solution was taken out of +4 C. What should be done? Should it be heated in order for it to return to liquid?

I am trying to create a workflow for flow cytometry experiment sample prep. I intend to use LIVE/DEAD™ Fixable Aqua Stain and an antibody cocktail for extracellular antigens.

I intend to use the viability dye first. The protocol I found does not seem to mind the protein content in buffers, as the cell washing is performed with PBS (5-10% fetal cell serum(I imagine they meant "calf")), and the cells are suspended in the same buffer before viability staining. I read that the presence of proteins in the buffers might contribute to a higher background, is that correct? How should I adjust the solution (for washing and resuspension) if that is correct?

Additionally, I intend to perform antibody titration for the antibodies I'm using. There are a few questions here too. We have no prior experience with these antibodies in the lab. Questions:

1. Is it important to do titration for every antibody in the cocktail?

2. Do I keep the same general sample prep with the antibody cocktail, only swapping the antibody cocktail with a single antibody that's in different concentrations? Do I also apply the viability dye as well?

3. The antibody cocktail I am to use includes dyes such as SB and BV etc, which require a special staining buffer or blocking buffer. Do I need these special staining buffers or blocking buffers for titration as well? Is it necessary since titration typically only uses one dye at a time (together with viability dye?)?

I know the information above might not be detailed enough but could you share your personal experience related to these scenarios? The protocol that I base my protocol on comes from this page

Thanks a lot for your input!

If someone has done some comparative experiment I would be really grateful for results and impressions. I need to detect low abundant microRNAs from plants. I bet that there is no difference among them but who knows :)

Hello,

I want to use a commercial preparation of recombinant PNGase, but I do not have the buffer. What cofactors or buffer conditions does PNGase F require? I will add the enzyme to cell lysate and then run western.

Thanks!

I have tried to achieve a separation of the nucleus and cytoplasm by using RLN buffer+ NP40 (0.2%), and then proceeding with GeneMATRIX universal RNA purification kit (the same used for total RNA extraction) and followed cell culture RNA purification protocol. I have managed a separation with this protocol in HeLa cells, but SH-SY5Y cells seem to be very sensitive. I have added the protocols and would appreciate any suggestions.

I was running PCR 2 lines of fax1 potential mutation (with positive and negative controls).

However in the results I have not been able to detect any DNA bands in the gel electrophoresis with one exception.

I had 2 amplification mixes - one for the fax1 gene and one for the tfax1 mutation (insertion of tDNA).

I used 1.5% agarose gel in 1*TAE buffer with 2ul GelRed per 100ml gel.

and inserted 20ul sample in each of the gel pockets.

Where could the problem be?

(i apologies for any unclarities and mistakes in my question)

Thank you in advance!