Western Blot - Science method

I am having an issue with the western blot transfer from last week, I see imprints of cassette on my membrane.

I am making a 14% gel and I run transfer for 2hr at 100V, after the transfer I see imprints of cassettes on the membrane.

I thought it could be because of the cassette was too tight or because of the temperature. So I keep in check for the cassette and temperature (1hr 45min) and repeated the western I see the same problem.

I used fresh buffers both the times.

Any suggestions how can I improve?

Hello! I am currently running a western blot using human AD tissue samples. I prepared these samples over a year and a half ago. I am having trouble standardizing these proteins with Beta Actin. I have ruled out other parameters that could be causing the problem. Therefore, I am wondering if my tissue samples are no longer reliable. In addition to this, I noticed that when i thaw my samples, there is a good amount of precipitation. I just vortex and spin down to mix the samples thoroughly.

For my results, I keep seeing bands stuck in the wells and multiple faded bands. Of course when I first ran these proteins, the beta actin signal was clean and neat.

If someone could please elaborate on what could be causing this. In addition, I would appreciate if you could provide how long samples last and if there is a way to troubleshoot this.

Thank you!

P.S. Samples have been stored in -20 C fridge.

Dear Community, I recently performed a Western Blot to test whether the mitochondria marker ATP5A1 is in the experimental cells or not. However, the results did not show up at the expected MW . I would like to ask what happens in this case and my sample 1 even didn't have band with beta-actin marker? Would you like to give me some advice ?Thank you so much .

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.

I'm now doing western blotting for several proteins.

but transfer is not perfectly happened.

bands are looks like smeared or smudged.

And it seems to the lower MW bands are more affected.

what did I do wrong?

I use towbin buffer without SDS / PVDF 0.2um pore / biorad wet blot tank transfer system / during transfer, use stirring and ice block with 400mA (about 200~100V)

Also, when I performed with 20V overnight same thing happened.

I attached picture of my transferred membrane

when I stained membrane with Ponceau S solution, other protein bands looks same as marker band.

I would like to know if anyone tried the western blot with the Instant Blue stained SDS-PAGE gels. Thanks.

Hello, I have isolated extracellular vesicles from a cell culture using the Exosome isolation kit. However, the western blot results did not show up because the concentration was only 0.2 ug/ul. I believe a concentration of 1 ug/ul is required for a successful western blot. Could you please suggest a method to increase the concentration?

I am researching beauvericin, one of the toxin proteins commonly produced by the entomopathogenic fungus Beauveria bassiana. In my research, I plan to use the Western blot method to confirm the presence of beauvericin toxin protein in the fungal filtrate. But I'm having trouble knowing what antibodies to use to detect the presence of beauvericin.

Can someone share their knowledge or experience?

thankyou

Hello Everyone.

I have samples that significantly vary in protein concentrations. can I load equal volumes of samples that have different protein concentrations on basis that later on I will normalize the target protein to the housekeeping protein?

Please I would be grateful if you advise.

Thank you

In my study utilizing anti-Flag antibody to detect a Flag-tagged protein in HepG2 cell line via Western blotting, I have observed multiple bands beyond the expected molecular weight. I seek guidance regarding the potential origin of these additional bands. Could they represent dimerization/oligomerization events manifesting as higher bands, or degradation products yielding lower bands?

How can I substantiate these observations, particularly considering the presumed specificity conferred by the Flag tag?

The truth is that I have had many, many problems to be able to see a specific band of this protein. My problems have ranged from bad transfers, to unknowns that I have not been able to answer. The times I have a good transfer (see image of ponceau s #1), I still fail to see defined bands once developed (see failed attemps). I have tried everything, increasing the concentration of the primary antibody, secondary, using more West Dura, cooling the transfer buffers to avoid burning (see ponceau s #2) and still nothing. Does anyone have any idea what else I could try? Thank you all very much in advance.

Hi!

I´m currently trying to isolate Mitochondria from HCM cells. For my western blots I need an additional antibody against a mitochondrial protein which is bigger or smaller than 14/17 kDa.

I was thinking of a protein which is only present in mitochondria or in the matrix. Maybe transmembrane proteins (TOM or TIM) will work too, but I don't know if the membranes are still intact in my samples. Maybe proteins from the ß-oxidation? Does someone know which antibodies can be used to detect mitochondrial proteins in western blots? If possible only proteins/Abs that are located/synthesized in the mitochondria (matrix). Thanks a lot!

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've done several experiments using western blotting to detect extracellular vesicle markers TSG101 and CD9. However, I consistently observe bands that are larger than the expected size. Notably, my lysate shows the correct band size, but the samples consistently display larger band sizes despite the use of a reducing agent. I've used 1% SDS lysis buffer along with protease inhibitor for sample preparation, and I also attempted using RIPA buffer. Interestingly, the bands only appeared when SDS was used, but they differed in size compared to the expected bands. Any help would be greatly appreciated!

Thanks!

Greetings to all, I have attempted to detect pERK using western blotting with HEK 293T cells, but I am unable to do so. I overexpressed a gene that caused pERK to be overproduced, and I was expecting to see and detect pERK, but that did not occur. In addition to making sure the cell lysis procedure is working as best it can, I have replaced all of the buffers and meticulously completed everything else. Could someone please let me know if they have encountered this issue and how they resolved it? Many thanks in advance!

The lanes contain the same sample and concentration (15ug). The image contains GluN2A, PSD95, and Beta Actin (Top, middle, bottom respectively). Due to this being the same sample every lane should theoretically be the same but the only one close to being the same is Beta Actin. GluN2A and PSD95 seem to be following the same pattern.

Hello everyone, I'm a master student and in my project I'm studying RFP-based constructs (RFP + few amino acid-long tags). I'm now performing a Western blot in SDS-PAGE of my cell-line lysates.

My western blots (as the photo shows) have a problem in the Ponceau staining as the marker is fairly visible and correctly transferred, however my sample lanes show little to no protein (I've loaded 50 ug), especially in the second half of the blot. The same problem is present also with other types of samples (like whole cell lysates) and seen in some of my collegues' blots.

I've tried make a new batch of sample buffer (+ B-mercaptoethanol), new acrillamide (gel is usually 10%) and new transfer protocol (10 minutes, 25V).

Another weird thing I've seen is that incubating an antibody (any kind really, I've tried B-actin), the signal is confined in the area of the blot that corresponds to where the stacking gel meets the running gel.

I thank whoever offers suggestions.

Hello. TetraHis antibodies recognise 4xHis-tag in the protein. What if I have a long His-tag (e.g. 10xHis-tag) and I perform ELISA or Western Blot based on native PAGE? Is there a chance that two TetraHis antibodies will sometimes bind to the long His tag and I'll get a false signal increase? Or is the potential distance between the antibodies too small and therefore after the first molecule has bound, the second molecule won't be able to approach the His-Tag, and it will be always 1 binding per 1 protein?

I started to think about it because the manufacturers of precoated well plates indicate the amount of histidines in the tag of the target protein. Is it possible that they imply potential overbinding?

Thanks.

I tried several times using the Lipofectamine 2000 reagent, extracted the proteins but couldn't detect on Western blot. My plasmids were constructed with PcDNA 3.1+ Please does anyone have any suggestions? Thank you.

Hi, I am an MS student who is studying idiopathic Pulmonary fibrosis(IPF)

My P.I asked to analyze cytokine IL-6 or TNF-a through Western blot despite Many researchers using ELISA for it because of tight budget, can you recommend good IL-6 antibody products for W.B? (It would be better if I could use it at ELISA)

Also, mRNA expression from RT-PCR method is fine too.

Thank you for reading my question.

i am working with membrane protein and there is a his tag 4 amino acid above the c terminus (seq HHHHHHDLEY). anti his antibody does not recognise the his tag in western blotting. i was wondering if it is necessary for his tag to be on N or C terminus for western blotting and purification

Dear Community,

I recently performed a Western Blot on phospho-Erk1/2, phospho-STAT5, phospho-p70-S6K and phospho-STAT3. But the results were very strange. They all showed multiple extra bands and the bands on the correct height were not really making sense (in fact things I blotted before dozen of times with the same result now showed the complete opposite and strange superintense bands occured). I worked with all ingredients before for months without any problems (no extra bands, repeatable results, same protein amount). The only thing I changed was to store both prepared Mastermix (Lysate + Lysis Buffer for Dilution + 1/4 Laemmli 4x) and lysates at -80°C before blotting instead of -20°C. Can this explain my problems?

And if it is the reason can I prepare fresh mastermixes from the frozen lysates stored at -80°C or are they also damaged?

Thank you very much!

Hello everyone. I am in the middle of performing western blotting experiments measuring expression of p-S6(Ser235/236) protein in total cell lysate.

Originally, I was quantifying my data using ImageJ, and normalizing protein expression compared to the housekeeping gene on the same blot. (Ex: HSP60)

However, I was notified by my PI that she read a paper where the researchers compared expression of p-S6 to S6 protein on the same blot. It sounds like the researchers simply stripped the blot and re-probed with unphosphorylated protein since they are both the same size.

Which approach is the best way to normalize expression of phosphorylated protein on a single blot? I am considered that stripping and re-probing the same blot will cause loss of protein & unwanted retention of the phosphorylated protein.

Although I can detect the protein (above 200), leaving it for longer in the transfer (plus 2h), I think I am losing my endogenous protein, as its detection is not getting good. How can I solve it?

I have used CSII to phosphorylate HP1a, in vitro. Now, to test this phosphorylation, I have options such as using an anti-serine phospho antibody or mass spectrometry.

Has anyone ever tested an anti-serine phospho antibody western blot that worked for them?

Any recommendations and catalog numbers would be helpful.

Hello, I'm having a bit of trouble doing the Western blot experiment.

Even though I load the same sample, the band appears inconsistently (in all the trials).

(I usually store the samples at -20°C and boil them at 95°C for 5 min before loading.)

This sample is harvested from bacteria-infected mammalian cells and I extracted only the mammalian cell protein by RIPA-buffer lysis method.

I wonder if anyone has an experience like this and how to solve this problem.

Thank you so much in advance!

I am doing crispr knockout in T cells. After transfection, I am trying to validate knockout by western blot and sequencing. with western blot, I can see truncated protein in knockout cells whereas in scrambled cells, band is at the expected position of the target protein.

With agarose gel, I can see 3 bands for knockout protein and scrambled is one band. My question is how do I sequence three bands. Shall i cut individual bands extract and purify DNA and then send for sequncing ?

what should be my approach? I am attaching pic of gel for reference.

pls help

I am trying to run a western blot on A549 cells with beta-actin as the loading control and alpha-SMA as the protein of interest. However, both beta-actin and alpha-SMA have similar kDas. How would this work?

I am also unsure how loading controls in western blots work. How would my WB process change?

**Note. This is what my lab professor told me to do.**

I'm working on senescence, and I'm trying to observe senescence markers. I have performed western blots for p21 but it seems that the bands are at a much higher molecular weight than expected. The expected molecular weight is aprox. 18KDa but I see bands around 50KDa. Has it ever happened to anyone? why does it happen? (I am using a monoclonal antibody 1:2000 from proteintech 67362-1-Ig). thanks :)

I am doing crispr knockout in T cells. After transfection, I am trying to validate knockout by western blot and sequencing. with western blot, I can see truncated protein in knockout cells whereas in scrambled cells, band is at the expected position of the target protein.

With agarose gel, I can see 3 bands for knockout protein and scrambled is one band. My question is how do I sequence three bands. Shall i cut individual bands extract and purify DNA and then send for sequncing ?

what should be my approach? I am attaching pic of gel for reference

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?

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)

Hi everyone! I have encountered a problem with my western blots that I would be grateful if someone could help me with.

After I had been using the same protocol with good results for over a year, my blots suddenly started coming out with thin, dotty bands (see images). I also add an image of how it used to look before. Does anyone have any idea of what could be wrong?

All our buffers and gels are the same (from BioRad and Invitrogen).

Currently I am trying abcam protocol for it.i checked the cell lysis by trypan blue stain which is more than 90 %. but I am facing following issues

1. unable to remove beta actin in mitochondrial fraction.

2. I am able to see mitochondria pellet (cell-1-1.5 X 10^7 cells) but the protein yield is very low 30 ug. and band of interest-SDHA is also very less on western blot compared to whole cell lysate when same amount of total protein (30 ug)

The old antibody has been discontinued and the currently offered are detecting a band at 62 kDa, which does not reflect the correct molecular weight (47 kDa).

We have recently aquired plasmids with the following configuration:

EF1A>{gene of interest}:P2A:Bsd

When used for transfection (293FT cells and SH-SY5Y cells), these cells expressed the desired protein after 48 hrs (verified several times by western blot and viewing of GFP which is encoded in some of our plasmids). When the selection antibiotic is added, most cells survive, which is expected. However, after a few days, the cells no longer produce the desired protein (verified many times by western blot and viewing GFP).

To be sure, we always use a negative control for the antibiotics, cells which were not transfected, and they all died quickly (36 hrs at most).

Oddly enough, when used for lentiviral infection, there is no issue, and the cells continue expressing the protein even after a few weeks of antibiotic selection.

We have not run into this problem with other vectors acquired from other sources.

Thanks in advance

Hi all, need a little help with this western blot please.

Protein band seems pretty spread out and this isn't the first time it's happened, thinking something must be wrong with the gel or protein degradation?

Any help appreciated!!

Hello everybody, I'm currently performing Western blot and I am a beginners for Western blot, so I want to ask that Can I replace blotting paper by any other papers?

So we have used different membranes and increased the protein concentration to 80mcg during the protein estimation and even though it's been always showing, we have not had any luck getting their expression lately for some reason.

I am doing western blots with an RNA binding protein and believe due to the vertical streaks, that I have RNA contamination. I have tried boiling the samples for longer as well. Another protein I have probed for does not have the vertical streaks and is not an RNA binding protein. Could someone recommend a protocol they have used or know of to get rid of RNA contamination in protein preps for western blots?

Does anyone have experience using this blocking buffer (made with Salmon serum) for immunohistochemistry of mammalian cells or tissues?

Hello

I am working on c1q and lair2 and how their interaction affects lupus (autoimmune disease) .

we used western blot , yet we can't see lair 2 ,

Hi, I´m doing Western Blot and normalizing with GAPDH.

I usually perform it with mouse skeletal muscle samples. I am using GAPDH (D16H11) XP Rabbit mAb #5174 from CellSignalling with an antirabbit secondary Ab.

The problem is that I use to see 2 bands at 37 and 45 kDa when it is indicated to give only 1 band at 37 kDa. More often is the upper band more intense and homogeneous between samples but it sometimes happens with the lower band.

Does anyone know what could be happening? Thanks

Hello,

What would be the best procedure right after extracting brain samples from mice, if we plan to analyze the samples later ?

Would it be best to just directly flash freeze the brain tissue in isopentane before storing at -80°, or to homogenize and add some RNase or protease inhibitors before the freezing ?

Additionally, if the samples are sent to external collaborators for the analyses, what is the recommended temperature for shipping, is -20° cold enough ?

Many thanks,

Benjamin Vidal

Problem

After Transferring the membrane picture with Ponceau S Staining looks like the membrane is burning and has poor transfer. Can someone help with this?

I also noticed something weird on the gel after the transfer, it seems there are some blue spots with Coomassie Blue staining.

12% gel

I load 30 ug protein in each lane with six samples and two ladders separately.

Wet method: 19-hour/constant 30V at working fridge (4 degrees) with ice bucket

PVDF membrane active with the methanol > 1 mins

The transfer buffer is fresh with 25 mM Tris, 192 mM glycine, and 20%methaol but make a 10* stock solution of transfer buffer, which is 250mM Tris 1920mP Glycine, then add 700ml ddh20 and 200ml Methanol to make 1L transfer buffer.

At the protocol I'm using now there is an added note indicating that it is possible to freeze the protein samples mixed with the laemmli to temporarily store them at -20ºC and denature them before performing the western blot. However, one of my coworkers says she tried it once and it didn't work out. ¿Can anyone please tell me if they have done this before and how it turned out? I have a large number of samples and this could save me some time. Tips and suggestions are appreciated.

Hello¸

Im PhD Student in UQTR ( Canada), I have a crucial experience for my project thesis : I have a WB (western blot) membrane that I need to store for a long time for incubation with other antibodies (Precisely Ubiquitin) . I read that I can put it in a bag at -20C.

I want just to confirm if I did it correctly (see picture) or there are other ways.

Thanks in advance

Ayoub

Dear users,

I'm now struggling for almost three months to make a pSTAT3-ELISA (https://www.rndsystems.com/products/human-mouse-phospho-stat3-y705-duoset-ic-elisa_dyc4607b-2) work but was unable to succeed. I completely followed their protocol without the smallest deviation but got no result. I then tried different Lysis Buffers (RIPA with SDS, RIPA with Triton-X-100, Lysis Buffer #6 of RnD) but nothing changed. I doubled the antibody amount with no effect and tried different ug amounts of the samples (1-2,5-5-10-25-50-100ug) but nothing changed. It's always the same result since the beginning: The standard curves are perfect but no signal from the samples. When I use other techniques with the same samples (Western Blot, Dot Blot) I get very good results for pSTAT3. I even performed a Western Blot with the Standards which indicated that my samples should be in the standard range. What can I do to make this ELISA work? Thank you very much for your answers.

Hi! I am a first year PhD student and I have been trying to do western blotting using #Jess #proteinsimple in my lab. My samples show a beta actin crisp band while i dont see any thing for the target protein of interest. Antibodies used - #HNE, #cytokines like tnf-alpha, #autophagymarkers like #P62 and #LC3

Any suggestions on how to optimize?? Any leads are really appreciated..

Thanks

Ritishka

I am unsure of the cause. It is reproducible across multiple blots and cells lines with both the phospho-specific antibody and the total mTOR antibody. Thanks for the help!

Hi all. I performed CRISPR knockout by lentiviral transduction. Cells were grown in puromycin for a period of time, protein was extracted and cells were also frozen down. A western blot showed almost complete knockdown of my target protein. After a couple of months I cultured a vial of these cells and I am now detecting abundant expression of the target protein via western blot. I have no clue how that has happened. I have also tried culturing cells in puromycin again, thinking that perhaps the wild type cells took over the population after thawing, but even in puromycin these cells are growing well. Has anyone had a similar experience or know why this is happening?

I have conducted several western blots to detect the expression of Zonula Occludens-1 (ZO-1), NCX-1, and PMCA1. However, I have observed that the expression levels of these proteins are very low, resulting in faint or even non-existent bands, despite adjusting the dilution factor of the antibodies (up to 1/200) and extending the transfer time overnight. Upon repeating the western blot experiments, the results for these proteins remain nearly negligible.

Could this lack of detection be attributed to limitations inherent in the cell line model I am utilizing, specifically Caco-2 cells? Alternatively, is it possible that these proteins are better detected using techniques such as immunohistochemistry or other methods? Interestingly, these proteins have been reported in some studies utilizing Caco-2 cells. Upon reaching out to these researchers, it was discovered that both parties are employing similar techniques and protocols.

What could potentially account for this variation in results despite consistent methodologies being employed?

I am doing western blots for extracellular vesicle protein and I don't have a good loading control. I want to quantify my western blots to compare the amount of CD81 in EVs collected from two different locations. What is a good way to do that?

Thank you!

I am treating tet on mice with doxycline (76ppm) for 2 weeks and i dont get the overexpression according to Western Blot. In rtPCR i see differences, but in protein level these differences are not reflected. What might be the issue?

Hello, 

I am using a c-myc-tagged plasmid for coIP experiments, and i always get two specific bands on my western blot when I immunoblot with the c-Myc antibody (ab32). I dont understand what can be the second band i see on my western blot. Can anyone Help plz 

Thanks 

Pam

I have utilized pET26b(+) NS2B-NS3 DENV2, and transfected it in E.coli. After transfection the selected cells were characterized based on the kanamycin resistance present in the vector. The selected cells were incubated with 1 millimolar of IPTG at 28 C for 4 hrs induction. While the results seem positive for the experiment, multiple bands occur underneath the protein of interest. the selection was done based on the histidine tag using penta-his antibody. Please help me with this concern. I am using TMB for the development of the blot.

Hi everyone, I transfected HEK cells with a protein that is intracellularly expressed through flow. Then I isolated the protein using RIPA buffer protocol, however, I accident added IP buffer instead of RIPA and now I do the know if it’s worth going through the whole western blot process.

For more context, I only know the protein shows intracellular expression through flow but don’t know exactly where it’s expressed. I looked up and ThermoFisher said IP is less harsh than RIPA so I don’t know if it collects everything or not.

Any advice is appreciated!

I'm wondering if there are any cases in which a nonsense mutation don't trigger non-sense mediated decay. If so, I would like to know if it would be possible to detect truncated protein in a Western blot (if the antibody binds to a region that is present in the truncated product).

Thank you very much.

In our experience, the antibodies #9196L and #9192 from Cell Signaling have not been effective.

Hi

I am working on lair2 and it's interaction with C1q to understand who that affect lupus.

We used cell line to produce lair2 . than we transfected it with c1q.

and now we are doing western blot , to detect the two proteins, the first time we incubate them with c1q antibody . and we see the two proteins . after that we used the same membrane to detect lair2 , we wash the membrane and incubate it with lair 2 antibody and we see the protein .

we used 2 negative controls , in the first detection we saw two faint bands in them . in the second one we didn't see them.

I don't know what's the problem , and how to analyze it?

Hi

I am working on polyacrylamide gel for western blot. I don't know why I get small wells .

when I prepare the gel it sound good , yet when I load the samples it is small.

Possible solution to increase the yield of the protein?

I am performing western blotting using tissue lysates prepared from white adipose tissues isolated from mice. The targets are membrane proteins. The protein quantification was carried out using BCA. I have attached the protein conc results.

Hi there to all of you.

Can I extract proteins with TRIZol? If so, what would be the most effective way to measure proteins following TRIZol extraction? I intend to detect proteins using western blot after using the TRIzol reagent from Ambion.

The issue is that, in order to avoid having to redo the experiment, I now need to obtain the mRNA for qRT-PCR and the protein for Western blot from the same sample. Viral proteins within infected cell lines are the proteins I'm looking for.

Hi,

I recently got in an argument with a new coworker on how to dilute protein samples for WB.

I normally dilute the Laemmli based on the total volume that I want to load in the wells, Example: if I have 5uL of protein sample I will add 2.5uL of LAE( 1/4 of 10) and add 2.5 of H2O, and the 10uL resulted are loaded in the well.

He assert to dilute the Laemmli based on the sample volume.

Example; for 5ul of protein sample He will add 1.67uL of LAE (1/3 of protein sample) and then or load 6.67 in the well or take to a certain volume with H2O.

His argument is that on the LAEMMLI protocol is written "dilute 1:4 with sample" but I dont catch the reasoning for it to be like this, because like this, more concentrated protein sample will have less LAEMMLI than a less concentrated one but the ug of protein is the same.

Hello researchers,

I'm currently working on a project involving LC3B antibody (Cell Signaling & SIGMA) for western blot analysis. Unfortunately, I've been consistently facing issues with dark background and black patches on the membrane. Despite trying various troubleshooting methods, the problem persists. Has anyone encountered a similar issue and found a solution? Your insights and suggestions would be greatly appreciated.

Thank you!"

I looked at the data sheet provided with the antibody, but I have found nothing regarding the recommended dilution range

After collecting the lysates, BCA assay immediately comes next to determine the quantity of the lysates to be used for western blot. However, the scaffolds used are plant protein-based, which hypothetically contributes to the total protein concentration of the lysate considering the mechanical and chemical degradation during lysis procedure. Thus, even when loaded with equal protein concentrations per sample, after western blotting, the cultured meat samples show low expression levels. How can i establish a fair comparison of protein expression given the situation?

I am working with SH-SY5Y cells and differentiating them into neurons using retinoic acid. I need to extract protein from them in order to conduct a western blot, however, I am having trouble getting enough protein. I am using 6 well plates and seeding 250,000 cells per well, with 3 wells per condition and harvest/differentiate at 80-90% confluency. Once they are differentiated I place the plate on ice, wash with 300ul of PBS per well and then add 300ul of RIPA buffer and protease inhibitor solution. I leave this on there for a few mins and shake the plate before using a cell scraper to lyse the cells (scraping about 30 times). I then transfer all 3 wells worth of cells to a 1.5ml eppendorf and centrifuge at 14,000g for 15mins, I do not set a temperature, but by the end of it, the centrifuge cools to 4 degrees celcius. I then transfer the supernatant to another eppendorf and perform a BCA assay. My issue is I am getting very low/undetectable levels of protein and don't know what I can do to increase this. If anyone has any suggestions that would be greatly appreciated, thankyou in advance!

In a Western Blot (WB) experiment, does the concentration of SDS-PAGE gel affect the position of protein bands? In the image, the same protein samples were used for all five lanes. The left side (lane 1, lane 2, lane 3) used a 10% gel, while the right side used a 12.5% gel, with all other conditions being consistent.

Thanks for your kindly help!

Hi, I am new to this platform. I am currently preparing for Western Blot experiments using Young (7-10 mo) and Aging (20+ mo) mice and the protein of interest will be tight junction proteins (ex. occluden, claudin-5, ZO-1).

I have looked through a series of commonly used housekeeping genes (e.g. β-actin, α-tubulin, β-tubulin and GAPDH) but found that all of them will be affected by aging. I have also looked through stain-free and total protein normalization (by Ponceau S or Coomassie staining), which seems to give a more promising result than the housekeeping genes. But since they are relatively new approaches, I would like to seek opinions here about a good way for the loading control of age-related Western Blot.

Thank you very much!

Hello,

For many phosphorylation studies of MAP kinases, cells are serum-starved overnight prior to stimulation. Serum-free media such as DMEM 1X is supplemented with 0.5% bovine serum albumin. Is there any drawback to including NEAA in the serum-free media? Is NEAA known to cause phosphorylation of MAP kinases?

Thanks!