Western Blot - Science method
Identification of proteins or peptides that have been electrophoretically separated by blot transferring from the electrophoresis gel to strips of nitrocellulose paper, followed by labeling with antibody probes.
Questions related to Western Blot
I am currently studying differences in the expression of some proteins after treating cells with Western blot. My conditions are Control vs Treated, and I have repeated the Western blot 3 or 4 times. How do I perform the statistical analysis of the band density quantification? Initially, I thought that performing a Mann-Whitney U test was more appropriate since the number of repetitions is low, but I have read that it is common to use t-test. Which one is preferred and why?
I have transfected cells with vector containing my protein of interest which has a Flag tag. The expression is inducible. When I run western blot, I see flag in my transfected induced cells as well as non induced cell control. I even added lysate of non transfected cells as additional control but I see a flag band in there too. I have tried multiple concentrations of Flag antibody, asked my lab mate to prepare a control sample using her cells but I still see the flag band.
What could be the possible reasons?
Hello fellow researchers,
I'm having a puzzling problem in my Western blot experiment, can somebody help me? I conducted an experiment using prefrontal cortex samples, following a WB protocol that has previously yielded successful results (We took quite some time to standardize each step). However, this time around, I'm facing a situation where I'm not able to detect any bands, despite thoroughly checking various aspects of my protocol.
Here are some key details:
- My samples were homogeneized in RIPA buffer + proteases inhibitors as usual, and are relatively fresh, I homogenezeid last month, and I am realizing western blot with those samples since that.
- I run my electrophoresis in BioRad system, at 150V, 400mA, 2h, room temperature (10% acrylamide gel)
- I transfered to nitrocelulose membranes in semy dry transfer 30V, 1h, 164mA, room temperature
- I performed a Ponceau staining and confirmed that the samples were transferred correctly to the membrane (image is attached)
- I used three different antibodies in those membranes in the first time (I cut the membranes in three different sizes), it didn't work and I thought that it could be a old antibody solution problem. So I stripped the membranes and I incubated with new antibodies solutions (I got three new and sealed antibodies, including the secondaries) and none of them resulted in detectable bands.
- I was very careful to see that I incubated the correct primary antibodies, with their respective secondary antibodies
- Blocking (BSA 5%) and washing steps (3x with TBS-T) have been successful in previous experiments with those antibodies of the same brand.
- The protein quantity in the samples appears adequate, as good bands were visible in the Ponceau staining.
- I'm using high-quality and well-maintained Super-ECL reagent.
I'm completely stumped by this situation, especially because even the internal control protein, beta-actin, is not being detected. If anyone has faced a similar issue or has suggestions on what else I can investigate, please share your insights. Any assistance or guidance would be greatly appreciated.
Thank you for your attention and help!
Can you explain to me why BSA works better than Milk as a blocking solution for GTPase proteins?
I recently performed protein extraction with RIPA buffer of some cells I collected. The intended purpose was to use it to load in an SDS PAGE gel for subsequent Western Blot. I made the mistake of lysing the cells in too much RIPA buffer and although there is protein, the sample is too diluted. I want to load 20 micrograms of total protein per lane, and I have roughly 400 microlitres of a protein solution at 0.13 microgram/microlitre. Since there is no gel that supports having 150 microlitres of sample loaded into a lane, I am looking for a way to concentrate what I have rather than just discarding it.
My gel running is going well but as the maker running down towards the end of the run, it got spread out towards the outside of its own lane, not so sure how could it can happen, even I tried to load the empty lane with 1x loading buffer
Hello, I hope analytical chemistry people or biochemistry major fellows could help me. Kindly advise on how to prepare the following denaturation buffer: [2% SDS, 1 M β-mercaptoethanol (β-ME)]?
The buffer is expected to be used with Endoglycosidase H enzyme, extracted from Streptomyces plicatus, CAS 52769-51-4 | 324717.
I am currently planning an experiment that involves viewing E. coli cells tagged with gold-conjugated secondary antibodies using a scanning electron microscope, and I am running into the issue of cost for primary antibodies. I might have the option of using primary antibodies previously purchased for Western blots, but I am unsure if these antibodies can also be used for SEM imaging. I do not yet know enough about the chemistry and reactivity of antibodies to answer this question, thus I find myself here!
On a related note, if anyone has any recommendations of good websites to purchase primary antibodies for E. coli that work with SEM, I would love some! I have found a few websites, but each of them only has 2 or 3 antibodies for this purpose.
Hi all, I need help. I have been trying to assess p-syk (65E4) in THP1 cell extracts unsuccessfully via western-blotting. Anyone has any tricks I should try? So far I have been using PVDF membranes and loading up to 55ug (tried 15, 30ug unsuccessfully). Any tips are greatly appreciated!
Hope you´re all doing well!
I´m currently in a bit of a situation. I have done Western Blot on primary human monocytes . I´ve tested the usability and approx. parameters used beforehand on THP-1 cell Lysate.
In case of the THP-1 cells, every antibody worked and produced a usable signal. In case of the primary human monocyte lysate it didn´t work. The proteins should be expressed in the lysate and Tubulin reacted as well, just the lanes with the other antibodies didn´t react. There are no bands at all.
I´ve used a RIPA Buffer for lysis, the monocytes were isolated via beads, for the transfer I´ve used a nitrocellulose and for imaging an ECL kit.
Does anyone have a good guess what went wrong/ How i could improve? My current only guess is that either the antibodies may have degraded over freeze/thaw cycles (stored according to the manufacturer) or that the proteins in the lysate did degrade/denaturate over time (produced single use quantities and started the blot max. 3 days later, stored at -80°C).
I performed a western blot using different tissues from the mouse(brain,lung,liver,spleen,small intestine, colon, kidney, testis). However, I observed a significant amount of background on the membrane. Why is there more background in the tissues during the western blot, but none in the cell line? What are the differences between them, and how can I reduce the background in tissue samples in the western blot?
For protein isolation, I used RIPA buffer containing a 1X protease inhibitor. I homogenized the sample using a rotor-stator homogenizer and then centrifuged it at maximum speed at 4°C for 30 minutes.
For western blot sample preperation, I diluted samples 50 ug in 16 ul RIPA buffer containing 1X protease inhibitor. I added loading dye with 0.2M DTT then I boiled it at 85 °C for 5 min.
I performed 2 hours blocking step and then incubated overnight with the primary antibody ( I prepared with blocking buffer)
I used primary ab 1:1000 for anti-A protein(from rabbit) and 1:6000 for anti-actin(from mouse) For secondary ab, I used 1:15000 anti-rabbit and 1:15000 anti mouse.
To reduce the background, I checked the antibodies by performing three separate western blots. In the first western blot, I omitted the primary antibody to test the specificity of the secondary antibody. In the second western blot, I used a lower concentration of the primary antibody: 1:2000 for anti-A protein (from rabbit) and 1:6000 for anti-actin (from mouse).In the third experiment, I used tubulin, but I didn't observe any difference.
I am using the costar gel loading tips round 4853 and an eppendorf pipette (100 µl) to load my gels. The volume of my samples is 25 µl. Before loading, I usually set my eppendorf pipette to 25 µl and use the gel loading tips as mentioned above. But everytime I get air bubbles in my tips that result non-optimal western blots.
Do you have any tips/ideas how to avoid air bubbles in the tips?
What I've already tried:
I resuspend the tip to remove the air bubbles but in most cases the resuspending process results in more bubbles.
I pull the pipette slowly to avoid capture any air, but still isn't working.
I readjusted the volume of the pipette to 23,5 µl to set a lower volume to avoid sucking excess air, but then I lose sample since a small amount is still in the tube. So that doesn't work either.
I am thankful for any help and tips!
I'm researching platelet-derived growth factor signaling, and recently ran into a big problem. Specifically, I am determining whether a particular peptide can activate the PDGF receptor. My initial results using western blot and immunofluorescent microscopy were very promising; compared to negative controls, I saw significant receptor phosphorylation after treatment with recombinant PDGF-BB, a peptide previously shown to activate signaling, or my experimental peptide.
However, my results are suddenly no longer reproducible. I now see no change in receptor phosphorylation or downstream pathway activation between any of the treatment/control groups. I have been troubleshooting for months, and have tried different media, different cell types, fresh reagents, different harvesting methods, and different timepoints. For reference, most of my experiments have been conducted at a PDGF-BB concentration of 100 ng/mL, and peptide concentrations of 1 ug/mL. Cells utilized include THP-1 macrophages differentiated using PMA, and HeLa cells. Timepoints have ranged from 30 minutes post treatment to 24 hours post treatment.
Based on literature, even if my experimental peptide does not activate the PDGF receptor, the PDGF-BB treatment should be a reliable positive control. Has anyone experienced a similar issue in which cells no longer respond to positive controls? Does anyone have any suggestions for conditions to test?
Hi everyone, during sample prep, we usually boil the samples with 2-mercaptoethanol in Laemli buffer. However by mistake, some of the samples were boiled with Laemli that had SDS but no 2-mercaptoethanol. Then I found out and add the 2-mercaptoethanol to the samples immediately after they cool down. Can we still detect the proteins with the antibodies like usual?
Hi everyone! I am new in lab and I have been having problems with Western Blot, I use a Chemidoc and when I reveal I see nothing, after reincubate, or incubating with a new antibody, the signal is lost, or, is very very low, when I dye with red Ponceau, I see a lot of protein because I put 40 ug per lane, I don't have idea about what happened, someone could help me, I will be eternally grateful
I have been doing western-blot experiments for 8 months or something (Invitrogen B1000 Mini Blot Module). I used to get good results until 2 weeks ago. Then something happened, I could not transfer my proteins to the membrane. Here is what was changed 2 weeks ago:
-I ran out of 10x running and transfer buffer so I prepared 10X running and transfer buffer according to CSH protocols.
-Before I tried these buffers, one of my friends wanted the western-blot system and she couldn't get the proteins in the membrane. I discovered she used a very old (8-month-old) 1X transfer buffer in the system.
- After her failed experiment, I ran my experiment with my new buffers. I faced with the same problem.
First I encountered with E2 error (biorad power supply), I remade the buffer, but this time ponceau staining was negative although the ladder was okay. What do you think is the problem? Do you think the old buffer (which my friend used) caused any damage to the system?
Here are the last images I got after many trials and I re-remade the all buffers (including gel tris buffers)
I am trying to develop my western blot using an antibody targeting a 17kda protein. It is binding non-specifically and not giving any band(like in the photo). What does it mean is happening? How should i trouble shoot this?
I did a knockdown, after checking the level of protein expression with Western blot, I got some percentage let's say like 60 to 70 % of knockdown, but when i tried checking with qPCR to check the transcription level it was as if there was a complete knockout, what could have been the reasons, I am confused can some help me with an explanation? You help will be highly appreciated.
Hi. I ran the western blot of my immunoprecipitated sample. I don't know why my protein band appears like a smear?
what could be the possible reason? I loaded 3% of input that is cell lysate and bound protein after IP.
I have attached the blot image. Please suggest me the how to solve this
I want to isolate total proteins from mouse brain tissues to run western blots & determine levels of nuclear-rich proteins such as p16INK4a and other cytoplasmic proteins. I used RIPA (50mM Tris (pH 7.4), 150mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 10mM NaF, 1mM EDTA) supplemented with protease & phosphatase inhibitor to homogenize, followed by sonication, rest on ice for 30 min, followed by centrifugation at various g force & duration. According to my Western blot results, there are bands of p16INK4a but only in the pellets, and not the supernatant, when I centrifuge either at low speed (1000g x 10min) or high speed (16000g x 20 min). How can I keep p16 proteins in the supernatant?
I am facing an issue with my polyacrylamide gel for western blot. This is 12% gel and I have used precast gel and made it with the recipe provided by BIO-RAD. I have been using the same method, recipe and voltage for last 2 years. It used to run smoothly but all of the sudden this problem appeared. Can you suggest what could be the issue?
Thanks in advance!
Its a small protein with 5.7kD weight. After doing weight adjustments considering acidic amino acids and also 3XFLAG tag, it should be 11.6 kD.
But on 16% tricine gel, it appears between 15kD and 25kD (almost 20kD).
What could be the reason?
I have checked it in cell extract and also after FLAG purification.
Does anyone have a recommendation for a good phospho-parkin (Ser65) antibody to use for western blot? The cell signaling one didn't really work for me...
I am encountering bacterial growth in my diluted western primary antibodies (in TBS, without any milk/bsa, with 0.01% NaZ). We keep the antibodies in +4 C since we use them frequently (Our incubations are also o/n at +4 C). Almost every 2-3 weeks I observe the contamination. I filter the antibodies with 0.4um filter every 1-2 months.
I am wondering why there is that much of bacterial growth even with NaZ. Also, is there a better way of decontaminating antibodies? Can I keep the antibodies in -20 C (how many times I can freeze/thaw them?)
I have been running a western blot setup optimisation with beta actin as a loading control protein. I am getting non-specific bands at the higher MW range in addition to my target beta-actin band.
These were my key steps:
1. Gel was made at 16.5% (as I was probing a smaller MW protein at 5kDa as well) and samples were run at 120V for 90 mins.
2. Transfer of gel was done at 10mA overnight in the cold room (4 degrees celsius).
3. Transfer done as per bio rad protocol, and following remove of PVDF membrane, blocking was done in 5% BSA in TBS-T for 1 hour and r.t., following which anti-beta actin was added in a 1:10000 ratio (antibody from protein tech, mouse monoclonal). But primary antibody was in 1% BSA in TBS-T.
4. Pri incubation was overnight in cold room as well.
5. Next day, after 3-5 washes in TBS-T (each was 5 minutes), secondary incubation with 1:20000 of anti-mouse antibody (diluted in 1% BSA in TBS-T as well.
Is it something to do with antibody dilution? I have read article that used even more dilute concentrations for beta actin (1:100,000 for eg.), and I am not sure that the higher concentration could have resulted in the extra bands. Or the fact that I incubated my antibodies in 1% BSA, as opposed to maintaining the BSA % used for blocking.
Kindly refer to the attached image for the blot.
If someone has experience in this aspect, it would be great to hear from you!
Thanks and Regards,
I am trying to demonstrate a protein-protein interaction through co-IP. To IP, I use a mouse IgG isotype control and a lab-generated mouse antibody to protein X. When I do WB, I use rabbit-generated antibody from Cell Signaling as well as their mouse-anti- rabbit conformation specific secondary (L27A9). In theory, I should not see IgG bands due to using antibodies generated in different hosts and the conformation specific secondary. However, it appears there are IgG bands both in the isotype control and the sample. Shown here, 500ug total protein was used in the IP with 10ug ms antibody. Antigen was allowed to bind beads for 1hr RT while rotating. I eluted in 40 uL total 1x LDS buffer with 5% 2-ME boiled >95C for 10 minutes. My proteins of interest are protein x (~25 kDA with the antibody we use to detect) and protein y (forms various complexes or in free form, ~65 kDA, 55 kDA, 17 kDA free, or 8 kDA free). The input band should be 55 kDA as this is the most prevalent form. My labmates use the same Ms isotype control, ms protein x primary, and secondary with no issues. I have run my protocol by them and they do not see a reason explaining the background.
Lanes L-R: protein x KO cells (IP with protein x), protein x KO cells (IP with IgG), Protein X input (20ug), protein x over-expressing (IP with protein x), protein x over-expressing (IP with IgG), protein x over-expressing input (20ug).
Does anyone have any idea as to why there appear to be heavy and light chain bands here? What can I do to reduce background? Any ideas or advice would be helpful. Thank you in advance!
Hi everyone, I've tried to transfer initially using a 20% methanol transfer buffer using 250mA max current for 70 mins, all I got was transfer of mid-high range proteins, I suppose all proteins below 20 kDa escaped the membrane from the other side. Anyone has any experience with blotting 5kDa +- peptides and can share tips?
We have a fisherbrand powerbank (cat. FB300Q) that we use for our western blots. During the transfer, we noticed that it would not reach 90 volts and only get up to 77 volts. We thought it might have been the electrodes, so we switched out the cassette and lid, but unfortunately, that didn't resolve the issue. We also thought it might not be cold enough, so we put a fresh ice pack in the chamber, but that didn't work as well. Also, in the past, we used to put our transfers in the cold room (large walk-in 4C) and sometimes the power banks would switch from volts to amps.
Has anyone else ever experienced this during a transfer? Could this problem be mechanical, or something else?
Thanks for all your help!
I recently did western blotting with Jess. My actin bands have been variable between sampes i.e. the actin band width and size for each sample was different. What do you think is the probable reason? I loaded 3 microliter of the protein or cell lysate. im sure my pipetting is not bad.
I met a problem. I cannot see any signal of brca2. I just made a 4% SDS gel, and did transmembrane for 3 h under 300 mA, but there was no signal when I did ECL exposure.
Because brca2 is such a big protein, 384 kDa, it is a challenge to do WB. However, I didn't find any special protocol for this WB. So does anyone have any experience in BRCA2 WB? or in big molecular weight protein WB?
Please share your methods, THX!!
I'm interested if a protein is being degraded. My idea is to western blot my samples, add an antibody for ubiquitin, image, wash off the antibodies, add an antibody for the protein of interest, image, then subtract the images from each other to show only the ubiquitination of the protein of interest.
My concern is that due to survivorship bias, this will not show what I want it to show. I want to know if the protein is being degraded, but what this would show in reality is the proteins that have been ubiquitinated, but have not been degraded. Should I be concerned about this? Are there any better methods for this purpose?
I am working with a protein located in the mitochondrial inner membrane, and I would like to know in which conditions should I perform the denaturation step... maybe the "standard" denaturation at 95ºC-100ºC for 5 minutes does not work for that kind of proteins.
Thank you very much.
I am trying to using Ni-NTA beads to do small-scale protein sample purifications (from cell-free reactions); however, majority of proteins still bind with sticky bead solutions after running western blot. May someone do similar process? and may provide suggestions on this issue?
There are too many information about this topic presented below:
2.4 mm Metal (Steel) Beads are recommended for hard tissues, skin, muscle, bone, hair
2.8 mm Ceramic Beads are recommended for hard tissues, heart, muscle, skin, tendon, tail, whole organs
1.4 mm Ceramic Beads are recommended for soft tissues, brain, liver, kidney, spleen
I am using Tissue Lyser with Ripa Lyzis Buffer to obtain protein samples from certain tissues for western blotting. I wanna enhance the recovery of total protein from the samples. Which type and size of beads should i use for homogenizing the large intestine tissue of rat?
I used some polyclonal western blot antibody, and that datasheet size is 45-50kDa
I also get the band in expected size range, but there's multiple other band(at least 3 bands in other size), and when i found the references about that protein, there's various size (e.g. 35kDa, 63kDa, 75kDa)
In this case, how can I confirm that 45-50kDa band is really my target protein?
Can I use the protein sequencing (service that supplied by company using SDS-PAGE band, N-terminal partial amino acid sequencing) for check that 45-50kDa band is identical with my target protein amino acid sequence?
Is this protein sequencing can apply for target band validation?
I have been treated PC12 cells with 100ng/ml NGF for 10 days, and lysed them using RIPA buffer for western blot. I got no bands for primed cells, but for wild type cells. Does anyone know why?!
Does anyone have suggestions on how to achieve complete or almost complete stripping for the western blot membrane? My protein of interest has a molecular weight of 110kda and my loading control appears around 120kda. We have never achieved complete stripping and end up getting two bands which sometimes become very problematic while quantifying. Since the two molecular weight is very close we don't cut the membrane. We also tried to get a different molecular size loading control since our primary antibody is in-house but that didn't work. Therefore I was wondering whether anyone has any suggestions regarding a good stripping buffer. Thank you.
Hello everyone, I'm currently trying to detect the expression of my target protein through Western blotting. The protein I'm studying has a size of 13 kDa and is tagged with a Flag tag. I've been using a 15% polyacrylamide gel for the separation. However, I'm encountering any signal and expression in my results. For the Western blot, I performed an overnight wet transformation. I have been using 5% skim milk as a blocking agent, and the primary antibody I'm using is anti-mouse. Thank you in advance
For the last 2 years, I have been producing antibodies in 293 Freestyle cells for my PhD project, and tested them against cancer cells, by verfying, first of all, binding to such cells. However, since the beggining of this year, the produced antibodies stopped binding cells. In charaterization by western blot, besides the band I would normaly expect, I see now an extra band with increased molecular weight. I tested the whole procedure, from production to purification, and I don't think the problem is in there. So my question is: is there any chance that the cells would change something in their machinery that would impact translation, folding ou post-translational modification on the antibodies, ou production of any other product? I have been using a relatively low passagem from the initial commercial stock (P.7). Would it be plausible? Any other tips?
My advisor suggested that I use the recombinant protein from the elisa kit as a positive control but I can't find any reference to know how much protein to put in each well...
I used to try Ab1 dilution 1:1000 TNT-5% skim milk because it was my first time to detect PIGR, I got nothing but empty film even I used to applied an advance ECL. 2nd, I used to make dilution Ab1 1:500 TNT-5% BSA, and I got nothing too.. till I tried dilution 1: 100 and still nothing there film. I got fast green detection and there is a band I expected as PIGR.. as well I do succeed in DMBT1 detection using 1:100 Ab with TNT-5% BSA. but it doesnt work on PIGR detection. Any suggestions please?
RIPA buffer was used for protein extraction from rat liver samples. After calculating the protein content of the homogenate through Bradford assay (BSA), 5-30 ug of protein was loaded to 12% SDS-PAGE. Then the protein bands were transferred through the wet-transfer method followed by the blocking buffer [1 % BSA (Himedia) in 1X PBS for 1 hrs], followed by washing thrice for a period of 10 min, later adding primary antibody (overnight) and again same washing step followed by secondary antibody (1.5 hrs). The antibody we ran for was β-actin with anti-mice specificity. Since the protein ladder transferred quite well on the blot, we arent speculating on the transfer issue and also ponceau proves the same (visible after stain). In addition, we did a dot blot, checking the affinity of the antibody. Bands are also visible with CBB stain on the gel. Still we arent getting it on the blot. Please suggest a solution to this. Thank you!
I am over-expressing acid sphingomyelinase enzyme in HeLa cells. Theenzyme shows expression on western blot but I do not have any activity in in vitro assays. I have checked the activity through Mass Spec as well as radioactivity. Please suggest where could I be going wrong? Thanks!
The background of my Western Blots is very uneven. I use premade NUPAGE gels and PVDF membrane. For blocking I using intercept blocking buffer for 1 hour at room temperature. Primary antibody is incubated overnight and secondary for 1 hour at room temperature. I keep the membrane in a 50 ml tube and it is on a rollerbank during all the incubations to keep it from drying out. Does anyone know what causes the background on the blots and how I can fix it?
Thanks in advance!!
I have conducted western blot experiments with rat brain tissue, where I have tried to measure the GAT-1 concentration. I am using the antiSLC6A1 antibody from Elabscience (Catalogue Number: E-AB-66423). Similar to the example in their website, I have detected GAT-1 blots closer to 55 kDa. However, when I look at other antibodies for GAT-1, I see that the blots are closer to 70 kDa (which is a lot closer to the mentioned molecular weight of GAT-1 in antibody manuals). Has anyone got similar results and somehow worked their way around?
i use image j to analyze my band of western blot previously i select band then analyze it by plotting curve and calculating area under curve .. recently when i analyze band this appear with no any curve draw .. what may be the reason ?? Iam new user for this application …
I am doing westernblots. Thee independent experiments. In each experiment i get different result. For example, if i see a decrease in my target protein at 10ng/ml then in another repetition, it is decreasing at 5ng/ml.
I have optimised by dilutions. I am confused what could be the reason behind it? Is it the health of the cells?
I'm working on the expression of gH2AX protein in primary cells using the western blot. unfortunately, with normal ECL I couldn't detect any band! and with sensitive ECL I just see false positives. even between lanes!!! my method is:
gH2AX : 17 kDa
Loading: 20ug protein
Transferring: 1h 25v
blocking buffer NFDM 5% (1.5h/RT)
Primary Ab: O/N 4C
• Rabbit monoclonal Anti gH2AX, cell-signaling/9718s 1:1000
• Secondary Ab: 1h/RT
Anti-Rabbit IgG, Cell signaling, 7074, 1:5000 1h/RT
does anyone have a similar experience?
Hello. My colleague and I have been attempting to perform a western blot for gamma-H2AX in mouse brain tissue samples. Repeatedly, no protein is detected. We have tried 12% and 15% gels, and have transferred at 0.3 amps for 1 hr, 45 mins, and 30 mins with semi-dry transfer, as well as at 0.2 amps for 30 mins. We generally use milk for blotting but have tried BSA. Preparation involved sonication in RIPA, addition of loading buffer, and heating at 95 degrees for 15 mins. We have loaded 30 ug with nothing detected. Actin and GAPDH controls show up fine. If anyone has any special tricks or advice for getting detectable bands, I would appreciate the assistance.
Currently, we use surgical scissors to cut the tissue (previously frozen at -80 degrees) while in a homogenization buffer. Then we place a metal bead in the tube and employ bead beating homogenization at 40-50 osc/min for 1 minute 6 times. We then centrifuge at 10,000g for 15 minutes and collect the super. Is this excessive? Coomassie blue stain shows protein transfer to membrane and bio rad protein assay shows high protein concentration; however, there are inconsistent results from probing. Is it possible that the target protein was denatured in the homogenization process?
I used to activate my PVDF with methanol for 5 minutes, then I did transfer as usual. But these days twice I got the same results, which is there is a block white marks in between after I finished my fast-green staining. I do believe those are not bubbles formation.
Some student suggest me to wash the PVDF first with water then continue transfer and so on.
I confuse, what's wrong with my technical transfer. I have never facing this situation before.
Is it possible that a KD cells showed reduced cell proliferation by WST but during FACS analysis it did not show any change of cell cycle? apoptosis checking by cleaved caspase 3 western blot also no significant change. So what happened?
I am used to using the TurboBlot for Western Blot and had no issues with exception of transferring high MW proteins. That's why I wanted to try the Wet transfer method. However, everytime I try it, the lower bands appear so badly on the membrane (the images are total protein blots). What could be the cause?
I am running the system (Biorad Criterion Blotter) with a constant voltage of 100V in a cold room (the problem is not overheating). The buffer I use is the recommended by Biorad (I do it myself). While I am running, I put an ice pack inside together with a magnetic stir. Could it be the magnetic stir motion causing this?
I am working on identification of exosome related protein markers through western blot antibody probing. In my samples, there is an impurity seen at 66KDa which is obtained during the sample acquisition process. When I am screening for the exosomal marker proteins, the impurity is also binding with the antibody and another unwanted constant band is also seen at35KDa during blot imaging. Here I have used Biorad Precision dual color ladder as a protein standard
I am unable to sort this problem. Help me If any one could.
I have used western blot to check SIRT1 and H3 acetylation expression. I saw change in protein expression of H3 acetylation but no change was seen in the SIRT1 expression. I got same intensity bands across all the samples. Why is that?
I did a western blot with 25ug of protein lysate. As control, i used the recombinant protein and cell who not express my protein. The first antibody (rabbit polyclonal) recognize only the recombinant form. A second antibody (rabbit polyclonal) recognize the recombinant and the endogenous forms. Why does the first antibody not bind the endogenous form ?
I am finding it hard to understand the difference between protein expression and what is protein activity. Like if my protein is getting expressed in western blot how can I relate it with its activity? I used ELISA kit that tells me about the concentration of SIRT1 in samples. But I wanted to know whether the activity of SIRT1 is being inhibited or activated. How can I know that ? Do I have to use some other kit?
Isolated primary hepatocytes and treated with insulin (1, 10 and 100nM ) for 15 minutes. I tried pAKT western blot but I didn't see the band, I would like to know the working protocol, please share the your thoughts.
Hello, I have to transfer a PAA 15% of 1.5mm onto a PVDF membrane in the Biorad Trans-blot turbo transfer system. The protocol says for a mini gel to use 1.3A up to 25V for 10 min. I read about the bad transfer results someone had. Any suggestions to improve the transfer??? My bands of interest are below 25kDa.
The protein extracts are frozen at -80 degrees, and we have tried to quantify by the Lowry method both before freezing and just after thawing and before loading the gel. Even if we quantify the total protein and load the same amount of protein on the gel, the loading control band always appears uneven, we have tested b-actin and vinculin.
Hi, I have run a couple of Western blots for tyrosine hydroxylase in mammalian SH-SY5Y cells and have noticed 2 distinct bands rather than just one, as is shown in most of the literature. Some other papers do show 2 bands, but don't really address it. I thought it could be a phosphorylated form of the protein, but the jump in size looks a little big for that?
Interestingly, the higher band only appears in the second and third lanes where the lysate loaded was from cells I have put through a differentiation protocol; the first lane is from undifferentiated cells.
Does anybody have any ideas about how to interpret this/have experience in blotting for TH and could offer any advice? Thank you!