Science method

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
  • asked a question related to Western Blot
Question
4 answers
Hi everyone,
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?
Thank you.
Relevant answer
Answer
Can you show the source that you should use MW-U for small samples? I dont think that this would be a good advice.
The decision, which statistical model is appropriate for your data cannot be decided without further information. Maybe both, MW-U as well as t-test are not suitable. Both have assumptions, both answer different questions. You have to be clear what you want and which data generating process underlies your data.
  • asked a question related to Western Blot
Question
3 answers
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?
Relevant answer
Answer
Cross contamination of the cell line maybe.
  • asked a question related to Western Blot
Question
4 answers
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!
Nicolle Platt
Relevant answer
Answer
At times, it happens that upon stripping protein bands are not at all visible in the membranes. This is because the stripping buffer may change the pH of the membrane and hence prevent the antibodies from binding to the protein in the membrane. Do not worry, repeat the WB once again with the new antibodies. I hope you will get the results.
  • asked a question related to Western Blot
Question
2 answers
Can you explain to me why BSA works better than Milk as a blocking solution for GTPase proteins?
Relevant answer
Answer
BSA is better for blocking than defatted milk powder, however it is very expensive in comparison 100Euro for 100g vs ~5 Euro for 500g.
You can get non-specific interactions, biotin which is endogenous in milk makes using streptavidin conjugates impossible and it's possible to have poor background.
BSA is not optimised for blocking, it's just available and relatively cheap compared to other proteins and functionally useful for many scientific purposes, e.g. a carrier protein, where you have very little of your valuable protein and need something to "carry" it through processes to avoid loss (like sticking to the side of your tube).
You could use casein equally well.
Binding capacity of defatted milk powder is not a problem either, you just need to have enough.
Supercheap defatted milk powder is unlikely to have been properly defatted and this can cause problems with background. However, don't use NIDO as it has all the fat in it still!!!!
  • asked a question related to Western Blot
Question
1 answer
Any recommendation for TXNIP western blot? Adult mouse cortex samples
Relevant answer
Answer
Rabbit mAbs AEBA-20, RB30652 and 8A11 are supposed to react with mouse TXNIP in WB. They are available from BosterBio and antibodies-online, and no doubt others as well. These suppliers also have pAbs directed at mouse TXNIP.
  • asked a question related to Western Blot
Question
3 answers
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.
Relevant answer
Answer
My postdoc once made a gel comb with one big deep tooth, then poured a custom 2-part gel using the usual formula. By running at low voltage at first, the protein concentrates at the stacking/running interface.
  • asked a question related to Western Blot
Question
3 answers
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
Relevant answer
Answer
One reason could be uneven heating. Try running the gel at a lower voltage to reduce the amount of heat generated.
Another reason could be that the composition of marker samples and the samples in the adjacent wells differ substantially in ionic strength. Salty samples tend to spread sideways into lanes occupied by low-salt samples.
  • asked a question related to Western Blot
Question
3 answers
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.
Relevant answer
Answer
2-mercaptoethanol (or β-mercaptoethanol) is a liquid with a concentration of 14.2 M. It is very smelly, so use it in a fume hood, if possible.
Sodium dodecyl sulfate is a solid. The dust from SDS can irritant the lungs, so it's a good idea to wear a filter mask when weighing it.
2% in this context means 2 grams/100 ml of solution.
For 100 ml of the solution, you need 2 grams of SDS and 7 ml of β-mercaptoethanol (1 M/14.2 M x 100 ml). To somewhat less than 90 ml of distilled water in a beaker, stirred with a magnetic stirrer, add the β-mercaptoethanol and dissolve the SDS. Bring the volume to 100 ml with distilled water.
  • asked a question related to Western Blot
Question
3 answers
Hi all,
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.
Thanks,
Joel
Relevant answer
Answer
Agree with Yannick. However, some antibodies work for both WB and imaging applications. A relatively easy way to test a primary ab for imaging is usually by light (fluorescence) microscopy, using applicable secondary ab (fluorophor conjugated). Also keep in mind that most ab do not bind after fixation with standard glutaraldehyde concentrations used for EM.
  • asked a question related to Western Blot
Question
1 answer
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!
Relevant answer
Answer
I would recommend using well-established stimuli (LPS, TNF-a, IFN-y etc.) in a time-dependent manner. Moreover, you can try cross-checking of your WB antibody reactivity.
Kind regards
AB Bayazid
  • asked a question related to Western Blot
Question
2 answers
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).
Relevant answer
Answer
Dear A.B. Bayazid,
The antibodies should all be reactive to human according to the manufacturer. Is there a varying specifity and resulting variation in intensity?
The lysate in different experiments originiated from different donors, I think that would exclude the possibility of diseases (if you meant that by biomarkers).
  • asked a question related to Western Blot
Question
2 answers
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.
Relevant answer
Answer
When you perform Western Blot detection for tissue lysates, there are signals from endogenous IgGs and nonspecific secondary antibody binding may obscure detection of proteins especially if you are detecting low abundant proteins or proteins of low molecular weight, which may be masked by IgG heavy and light chains generated during the denaturing and reducing steps of tissue sample preparation.
These IgG chains are detected by conventional secondary antibodies that bind to both IgG heavy and light chains. This problem is common while using tissue samples, but it is more pronounced in tissue samples such as thymus or thyroid that are part of the immune system and therefore contain a larger amount of endogenous immunoglobulins.
The solution to this problem is to choose a labeled secondary antibody cross-adsorbed against the species of the experimental tissue, if it is possible. For example, if the tissue you are using is human and the primary antibody is made in mouse, use a labeled anti-mouse IgG that has been adsorbed against human.
In another case, if you are using mouse tissue and this tissue containing immunoglobulins is probed with a primary antibody made in mouse, then the endogenous immunoglobulins should first be blocked with a monovalent Fab fragment of anti-mouse IgG.
Further, there are certain steps you need to follow which are given below.
1) High concentration of antibody causes high background. By diluting the antibody, only higher affinity interactions are sustained. The lower affinity interactions (to remotely similar epitopes) will not last at lower antibody concentrations. So, you reduce the concentration of your primary antibody still further. You may go up to 1:5000 dilution for the primary and for the secondary antibody you may use 1:20,000 dilution. Dilute the secondary antibody in wash buffer only, for example TBST, because diluting the antibody in buffer containing BSA or milk, due to contamination of bovine IgG, sticky immune complexes may form due to cross-reaction.
2) Never block with BSA or BSA-containing solutions because BSA is generally a weak blocker which may result in more non-specific antibody binding. Use non-fat dry milk instead for blocking. Proper blocking conditions may also help to prevent low affinity interactions.
3) You will generally get a higher background if you are using PVDF membrane. Therefore, washing carefully is very important. Wash the membrane 4 times for 5 minutes in TBST with gentle shaking, using a generous amount of buffer.
4) Are you using fresh buffers? If the buffers are too old, you may face such a problem.
5) Another point to note is that too high of an exposure may also lead to this problem. Therefore, it is advisable to check different exposure times to achieve an optimum time.
6) Finally, as mentioned by Denis Komarov use less amount of lysate (10-20ug) to be loaded on the gel unless you are detecting low abundant protein.
Best.
  • asked a question related to Western Blot
Question
3 answers
Hello,
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!
Relevant answer
Answer
Because samples containing detergent tends to stick to the inside of the pipet tip, if you set the pipettor volume equal to the sample volume, you will always expel air at the end of the dispense.
Also, if the samples were heated, some of the water will have evaporated and be located at the top of the tube, lowering the sample volume. Centrifuge the samples after heating and allowing them to cool to room temperature to return the evaporated water to the sample.
Set the pipettor volume to a few microliters less than the sample volume to allow for the small portion of the sample that remains stuck to the pipet tip. Release the sample slowly into the well to allow time for most of the sample to drain off the sides of the pipet tip. Do not push the plunger to the bottom stop - stop pushing out the sample when the last of it has left the pipet tip.
This approach is easier when using a 20-µl pipettor than a 100-µl pipettor because the spring on a 20-µl pipettor plunger is not as strong. Reducing the sample volume from 25 µl to 20 µl will allow you to use a 20-µl pipettor to dispense the samples into the wells, and this will give you greater control over the dispensing.
Finally, if you allow the sample to fall into the well slowly from above, due to its greater density than the buffer (because of the glycerol), instead of placing the pipet tip near the bottom of the well, if a bubble if air is accidentally dispensed, it will not disturb the already-dispensed sample very much and will just float to the top without causing any trouble.
After all the effort that went into making the samples in the first place, spending a little extra time loading them onto the gel carefully is worthwhile.
  • asked a question related to Western Blot
Question
1 answer
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?
Relevant answer
Answer
If your serum concentration and consistent passage of cells, and the conditions (pH, Temperature) are standard, check if the loss of receptor phosphorylation could be due to increased phosphatase activity. You can try adding phosphatase inhibitors to your lysis buffer to preserve phosphorylation. Hope it would help. Otherwise check carefully all the standard parameters. If still the case use another validation method to verify the quality ingredients and reagents.
All the best.
  • asked a question related to Western Blot
Question
4 answers
What are the rules when choosing antibodies
Relevant answer
Answer
Dear Dr. Malgorzata Krok
In Western Blot when you decide to choose the primary antibody make sure that the primary is specific to the protein of interest and should be of a different host species than the sample. Also, the primary antibody should be validated for use in western blotting, the information for which will be made available in the package insert for the antibody which you will purchase.
Another point to remember is that the primary should be specific for either the denatured or native conformation of the protein of interest, which will depend on the type of PAGE (either SDS or native) you perform.
Further, choose a secondary antibody directed against the species of the primary antibody. So, you will need a secondary antibody that is raised in a species different than the host species of the primary antibody. For instance, if your primary antibody is raised in a mouse, you will need an anti-mouse secondary antibody raised in goat, rabbit, etc.
If you would want to perform multiplex Western Blot, use primary antibodies from different host species for each target being probed. Ideally, use a combination of antibodies from two distantly related species such as rat and rabbit, avoiding the combinations like mouse and rat or goat and sheep. This will help in the selection of appropriate secondary antibodies to minimize antibody cross reactivity that could affect your end results.
Regards,
Malcolm Nobre
  • asked a question related to Western Blot
Question
2 answers
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?
Relevant answer
Answer
Since 2-mercaptoethanol/DTT work at RT as well thus you can add them after the samples have been boiled and cooled as well.
  • asked a question related to Western Blot
Question
3 answers
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
Relevant answer
Answer
Western Blotting can be influenced by a variety of factors, and it can sometimes be challenging to pinpoint the exact issue. Here are some key areas to consider:
Antibody Species and Host Origin:
  • Primary Antibody: Always make sure you're using the correct primary antibody for your target protein. Check the datasheet or product information to ensure specificity for your protein of interest. Additionally, the species from which the antibody was generated (e.g., rabbit, mouse, goat) is important to note.
  • Secondary Antibody: The secondary antibody you use must be directed against the species of the primary antibody. For example, if your primary antibody is a rabbit anti-protein X, you should use an anti-rabbit secondary antibody. Also, ensure that the secondary antibody is conjugated to the appropriate enzyme (like horseradish peroxidase or alkaline phosphatase) for chemiluminescence or fluorescence detection.
  • Detection: ECL Reagent: If you're using an ECL (enhanced chemiluminescence) reagent for detection, make sure it's fresh and that you're using the right volumes. Exposure Time: Sometimes, the signal might be too weak if the exposure time is too short, or it could be too strong and get saturated if it's too long. Experiment with different exposure times on the Chemidoc.
  • Controls: Positive Control: Always run a positive control if available. This can help determine if the problem lies with the sample or elsewhere in the protocol.
  • asked a question related to Western Blot
Question
3 answers
Hello,
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)
Relevant answer
Answer
You could try measuring the pH of your buffers on a different pH meter in someone else's lab, just to double check.
You might also double check your recipes to make sure there wasn't some simple arithmetic error that crept in.
You might see if someone else in your department is also doing blots and borrow some buffers from them to rule out equipment or materials failure.
  • asked a question related to Western Blot
Question
5 answers
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?
Relevant answer
Answer
Numerous causes might be to blame for this kind of problem. What I can recommend is that you lengthen your blocking period to at least 2 hours and that you avoid incubating the secondary antibody for longer than 2 hours.
Use freshly prepared buffers and make sure about the proper transfer. Hope you will get a good result next time.
  • asked a question related to Western Blot
Question
3 answers
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.
Relevant answer
Answer
Are you using siRNA? If yes, then you are most likely to face this problem.
You need to consider protein stability which is highly variable. It may so happen that your target protein may be highly stable with a longer half-life. In such a case, you may consider choosing shRNA-mediated gene silencing method over siRNA. While making a choice, you will have to consider the length of the assay as well as the half-life of the target protein.
siRNAs are transiently expressed in cells, while shRNA is stably integrated into the host cell genome. As cells divide, the shRNA is passed on to daughter cells. Using lentiviral vectors for expression of shRNA, provides permanent knockdown without needing to transfect the cells multiple times.
The protein levels subsequently go down over time because the shRNA constantly keep suppressing mRNA.
So, if you are trying to knockdown the expression of protein that has a long half-life, then stable expression of shRNA may be required.
Best.
  • asked a question related to Western Blot
Question
9 answers
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
Relevant answer
Answer
Thank you all for your immediate response. I would like to share the western blot image of serial dilution of my protein. So, I found that due to the vast number protein in the cell lysate, the antibody couldn't detect the protein in the cell lysate. When I load the different dilution of my cell lysate, I could see the expression clearly.
Thank you all for your feedback and explanations.
  • asked a question related to Western Blot
Question
2 answers
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?
Relevant answer
Answer
Sample prep solution for 2D-PAGE experiments called 2D rehydration buffer which includes non-ionic detergents and urea may work. Recipe like;
(8 M urea, 4% CHAPS, 40 mMDTT)...You may also modify RIPA by increasing the SDS, adding a reducing agent, and adding urea as chaotropic agent up to 8M to induce solubilization...You may also investigate novel methods for increased solubilization at inclusion bodies...
here is one of them;
  • asked a question related to Western Blot
Question
3 answers
Hello,
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!
Relevant answer
Answer
it happened to me twice. And it turned out to be the issue with running buffer.
Try a fresh running buffer and keep an eye on the gel while its running that it does not get short of running buffer. As sometimes the buffer on top is a bit foamy and not liquidy.
  • asked a question related to Western Blot
Question
7 answers
Hi everyone
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.
Thanks.
Relevant answer
Answer
Could it be glycosylated? If there is N-linked glycosylation, treating with PNGase F to remove it should result in the protein migrating at the expected mol. wt. for the polypeptide.
  • asked a question related to Western Blot
Question
1 answer
Hi!
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...
Thank you
Relevant answer
Answer
Do you have a working positive control?
  • asked a question related to Western Blot
Question
9 answers
Hello everyone,
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?)
Relevant answer
Answer
Are you sure you're seeing growing bacteria, not precipitate of some kind? TBS alone can hardly support bacterial growth, and you even have it supplemented with NaN3. By the way, people rather use 0.01M or 0.1% NaN3 instead of 0.01%. If you are concerned, you may increase NaN3 concentration.
  • asked a question related to Western Blot
Question
5 answers
Hi researchers,
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,
Mathangi
Relevant answer
Answer
You may try using a more dilute concentration of beta actin antibody. But it may not help. I make the primary and secondary antibody dilutions in 3% BSA in TBST and blocking is done in 5%BSA in TBST. The rest of the protocol seems okay to me.
Besides the above, I would like you to know that although actin is often thought of as a single protein, in mammals, it consists of six different isoforms encoded by separate genes. Each isoform is remarkably similar to every other isoform, with only slight variations in amino acid sequence. So, there is a high possibility that the beta-actin antibody which you are using may not be specific.
You may try the most cited anti-beta-actin antibody clone AC-15, which is used in Western Blot. AC-15 belongs to the subclass IgG1 and was raised in mice against the N-terminal synthetic peptide of beta-actin. The clone specifically targets the beta isoform, with no cross-reactivity against the other 5 isoforms of actin family.
I have attached a link below regarding the anti-beta-actin antibody clone AC-15 for your reference, which may help.
Best.
  • asked a question related to Western Blot
Question
4 answers
Hi,
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!
Relevant answer
Answer
Update for anyone also having this issue:
1. Incorrect lysis buffer-> NP-40 lysis buffer (Pierce recipe) works best for me.
2. The conformation-specific secondary can cross react w/ antibody if the antibody is too high. I lowered the antibody and increased washing volume with PBS-T 0.1%. Evan's comment helped with this.
3. Lab-made antibody was not pulling down POI, switching antibodies solved this.
4. Tris-glycine elution helps reduce antibody elution as Selim mentioned. His protocol is helpful as well.
  • asked a question related to Western Blot
Question
4 answers
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?
Relevant answer
Answer
Just to update, I've successfully transferred the small proteins / peptides from 20% acrylamide 1.5mm gel to a PVDF membrane using 250mA for 45 minutes. In case anyone googles this question :)
  • asked a question related to Western Blot
Question
3 answers
Hello,
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!
Relevant answer
Answer
I think this is because during the transfer, the current is too large to reach the maximum current of the electrophoresis power supply, so the voltage can not be increased. You can try precooling the transfer buffer (containing 20% methanol) in a -20 ° C refrigerator.
  • asked a question related to Western Blot
Question
1 answer
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.
Relevant answer
Answer
I’m getting the same result with beta actin using 0.1mg/ml sample. Please let me know if you figured it out!
  • asked a question related to Western Blot
Question
2 answers
Hi everyone,
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!!
Relevant answer
Answer
Hello Mingqi Li
There are some points you need to consider when performing Western Blot for large molecular size proteins.
1. For large molecular weight proteins I suggest you use Tris-Acetate gel.
2. Large proteins can precipitate out in the presence of methanol. So, you should decrease the methanol percentage (10% or less) in the transfer buffer.
3. Just to ensure that your protein does not precipitate out, you may add SDS to a final concentration of 0.1%. SDS adds uniform negative charge to proteins, making it easier for them to transfer from the gel onto the membrane.
4. For large molecular weight proteins always use PVDF membrane because large proteins can precipitate out in the presence of methanol, and PVDF membrane do not require any methanol in the transfer buffer. So, you have a higher chance of successfully transferring your protein to the blot using PVDF membrane. Also, use PVDF membrane of 0.45um pore size.
5. For large proteins, you should use a wet tank transfer method. As large proteins will transfer out of the gel very slowly, I would suggest you perform overnight transfer at 4°C at 40 mA.
The paper attached below may help you with the protocol. Since you are looking out for one protein, you may use a non-gradient gel.
Good Luck!
  • asked a question related to Western Blot
Question
2 answers
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?
Relevant answer
Answer
As stated above MG132 is a very good proposal. In additon, you could perform immoprecipitation for your protein of interest and in the resulted samples, Wwstern blotting against ubiquitin.
  • asked a question related to Western Blot
Question
3 answers
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.
Relevant answer
Answer
Avoid to heat that much, it may lead to aggregate formation. In general, incubation in Laemmli buffer (containg SDS of course) at 37°C for one hour or so must be enough to give correct results on migration and Western Blot.
  • asked a question related to Western Blot
Question
2 answers
...
Relevant answer
Answer
I believe, you can! Try using somewhere between 0.1-0.5%. I think that will work just fine. Let me know! Bappi Sarkar
  • asked a question related to Western Blot
Question
4 answers
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?
Thanks!
Relevant answer
Answer
If the protein is binding strongly to the resin, you could try eluting the protein using a slightly higher imidazole concentration (~300 mM) at a lower pH (~7), as the affinity of the his tag for the metal will be decreased at a lower pH. Before trying this, you should ensure that the protein is stable at the lower pH first.
  • asked a question related to Western Blot
Question
6 answers
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?
Relevant answer
Answer
The zircon or glass beads of 100-200 micron size in a ratio of 1: 1 of intestinal tissue shall work fine for homogenisation or bead beating. Manual homogenisation in an ice bath will give better results.
  • asked a question related to Western Blot
Question
5 answers
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?
Relevant answer
Answer
Bbumba Patrick I agree that immunoprecipitation would be worthwhile. If the antiserum is specific enough, it should result in an SDS-PAGE profile that is much simpler than the crude extract, making it easier to obtain useful information from the mass spec experiment.
  • asked a question related to Western Blot
Question
3 answers
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?!
Relevant answer
Answer
Saif Wahid thank you so much for sharing your knowledge
  • asked a question related to Western Blot
Question
2 answers
Hi, I am going to use RIPA buffer to do protein extraction. However, I am wondering whether I need to heat the samples (eg. 95°C, 5mins) to get better solubility?
(The downtream work is western blot)
Relevant answer
Answer
Dear Qin,
You don't need to heat your samples before extraction, even more - working on ice is encouraged in order to preserve as much proteins as possible. Make sure not to use to much RIPA, it depends on your samples but 20-50 μL is usually enough for cells, and 50-200 to tissues. In order to get a better yield perform sonication/physical disruption (e.g. with a syringe) after collecting samples
Best of luck!
  • asked a question related to Western Blot
Question
3 answers
Hello everyone,
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.
Relevant answer
Answer
Evan Kerek and Saddah Ibrahim thank you for your answers
  • asked a question related to Western Blot
Question
3 answers
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
Relevant answer
Answer
What do you expect the expression to be endogenously? How much protein are you loading? I had a 25kDa protein that I had to load a ton of protein to detect in WB (40ug/well).
The only way I was able to detect our flag tagged antibody was to also use an overexpression flag tagged plasmid to transfect cells beforehand. Then I used flag tagged beads to pull down the protein, otherwise I could not observe protein bands.
  • asked a question related to Western Blot
Question
3 answers
Hello!
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?
Thank you!
Relevant answer
Answer
It's possible that the cells could have undergone changes in their machinery, which could affect translation, folding, or post-translational modifications of the antibodies. Cell lines can exhibit genetic instability or undergo phenotypic changes over time, especially when cultured for extended periods or subjected to passaging. These changes could potentially impact protein expression and functionality.
Using a relatively low passage number from the initial commercial stock (P.7) should minimize the risk of significant changes, but it doesn't entirely rule out the possibility. Other factors, such as variations in cell culture conditions, media components, or reagents, could also contribute to the observed issues.
To troubleshoot this problem, you may consider these:
1. Verify the integrity of your cell line by comparing it to a freshly obtained sample from the same commercial source or obtaining a new batch altogether.
2. Assess the stability of your cell culture conditions, including media, supplements, and any other factors that could impact cellular behavior.
3. Explore alternative antibody production methods or cell lines to confirm whether the observed changes are specific to the current system.
  • asked a question related to Western Blot
Question
4 answers
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...
Relevant answer
Answer
Not used before, but you can surely give it a try, you might see additional BSA band in your blot, if your protein is at a different MW position, then it can be done. About loading amount - it could be 15 to 30 times lesser than usual western loading amount of a tissue/cellular protein mix.
  • asked a question related to Western Blot
Question
4 answers
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?
Relevant answer
Answer
I am assuming you are using an anti-pIgR antibody. Is it rated to bind the pIgR in a western format? The epitope it detects may not exist in the western blot due to denaturing. Try running the western non-denatured and see if you detect something.
  • asked a question related to Western Blot
Question
3 answers
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!
Relevant answer
Answer
Hi Divya,
I would also reccomend to take care of the steps as Tehreem Maradagi mentioned.
No bands can also arise due to many reasons related to antibody, antigen, or buffer used. If an improper antibody is used, either primary or secondary, the band will not show. In addition, the concentration of the antibody should be appropriate as well; if the concentration is too low, the signal may not be visible.
Another reason for no visible bands is the lowest concentration or absence of the antigen. In this case, antigen from another source can be used to confirm whether the problem lies with the sample or with other elements, such as the antibody. Moreover, prolonged washing can also decrease the signal. Buffers can also contribute to the problem. It should be ensured that buffers like the transfer buffer, TBST, running buffer and ECL are all new and noncontaminated. If the buffers are contaminated with sodium azide, it can inactivate HRP.
It is also important to use a shaker for all incubation, so that there is no uneven agitation during the incubation. Once again, washing is of utmost importance as well to wash the background. This problem can also be caused by antibodies binding to the blocking agents; in this case another blocking agent should be tried. Filtering the blocking agent can also help to remove some contaminants.
Good luck with your experiments.
  • asked a question related to Western Blot
Question
3 answers
Hello all,
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!
Relevant answer
Answer
Hi there,
If your enzyme of interest is produced then there might be an issue with its folding/stability (if not properly folded/matured the protein will exhibit no activity), cell extraction (possibly provoking the loss of enzyme activity) and/or with the assay itself (compatibility with the source of enzyme you use).
  • asked a question related to Western Blot
Question
3 answers
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!!
Relevant answer
Answer
is it wrapped in clingfilm?
If it is wet it with methanol, this stops any static occurring between the clingfilm.
  • asked a question related to Western Blot
Question
1 answer
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?
Relevant answer
Answer
Hi Berkay Alpay, it is not uncommon to observe variations in the molecular weight of protein bands detected by different antibodies, even when targeting the same protein. These variations can arise due to various factors such as antibody specificity, post-translational modifications, or isoform-specific differences. In the case of GAT-1 (SLC6A1), different antibodies may recognize different isoforms or post-translationally modified forms of the protein, resulting in variations in the detected molecular weight. Additionally, differences in sample preparation and experimental conditions can also influence the observed band migration.
To address this issue and reconcile the differences in molecular weight, you can consider the following steps:
  • Verify antibody specificity: Ensure that the antibody you are using (antiSLC6A1 from Elabscience) has been validated for GAT-1 detection in previous studies or references. Reviewing literature or consulting with experts in the field can provide insights into the reliability of the antibody.
  • Positive control: Include a positive control in your experiments, such as a cell lysate or tissue known to express GAT-1 at the expected molecular weight. This will help confirm the antibody's specificity and aid in determining the appropriate molecular weight of the GAT-1 band.
  • Compare with other studies: Look for published research that has used similar experimental conditions and the same or different GAT-1 antibodies. Compare their results to gain a broader understanding of the molecular weight variations observed in different studies.
  • Consider alternative detection methods: If the variations in molecular weight persist, you may explore alternative techniques like mass spectrometry or other forms of protein identification to confirm the identity and molecular weight of the detected band.
  • asked a question related to Western Blot
Question
3 answers
Hello! I am trying to determine the best controls for my western experiment. I was wondering what the best controls are with a small kDa size?
Relevant answer
Answer
While deciding the best loading control for a Western Blot, you should take into consideration the following factors.
1. Make sure that the loading control is expressed ubiquitously at high levels in your samples and that its level isn't affected by experiment conditions. For instance, if you are studying a metabolic process, use of GAPDH as a loading control is not recommended due its role in glycolysis.
2. Ensure that the loading control and your protein of interest are expressed in the same location. For instance, Tubulin, β Actin and GAPDH are whole cell proteins, the locality of PCNA is cell nuclear, Lamin B1 is located in nuclear membrane, and so on and so forth.
3. Choosing a loading control that is close in size, but still large or small enough to be easily distinguished from your target protein is crucial. If the target protein and loading control are of the same size, visualization will be impaired and data interpretation compromised. A size difference of at least 5 kDa between your protein of interest and the loading control will help ensure to distinguish the bands.
Since I do not know your protein of interest (whether it is cytoplasmic, nuclear, cytoskeletal or mitochondrial), I would not be able to give you the best controls with a small kDa size. Nevertheless, you could always refer to Table 1 and Table 2 of the link provided below, which gives you the common loading control proteins with their cellular localization and molecular weight.
Best.
  • asked a question related to Western Blot
Question
8 answers
Hi 👋
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 …
Relevant answer
Answer
Here is a tutorial video on imagej western blot analysis
  • asked a question related to Western Blot
Question
3 answers
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?
Relevant answer
Answer
Are you using the same protein aliquot for the inhibition study? If yes, protein concentration should not be an issue but if you are using different aliquots protein concentration might be an issue. And if you are using samples with different drug concentrations on the same blot then the transfer of protein should not be an issue. But it is not necessary that you will get a linear decrease in protein with increasing concentrations of the drug. The difference of 0.5ng/ml and 1ng/ml is less so you can try the inhibition assay with larger drug concentration differences. You can eventually move to decreased drug concentration differences once you get better results with higher drug concentration differences. Have you tested the results with SDS-PAGE? Are you getting the desired results there? You should optimize the results with SDS-PAGE first then move to western blotting. If you are getting good results on SDS-PAGE and not getting the results on western blot then we can say that there is an issue with the western blot procedure. I hope my answer will be helpful to you. Thank you!
  • asked a question related to Western Blot
Question
3 answers
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
Gel%:4-12
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?
Relevant answer
Answer
Do you get a signal in a dot blot or ELISA like setup? Do you have a positive control?
  • asked a question related to Western Blot
Question
18 answers
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.
Relevant answer
I hope you could find a solution to your problem. I also have the same challenge in detecting the gH2AX! could you please share your standardized protocol?
Thank you
Maryam
  • asked a question related to Western Blot
Question
1 answer
Does GAPDH band on Western blot ever comes around 43kDa?
Please see the attached image.
Thank you
Relevant answer
Answer
There is a streak at your PAGE gel...GAPDH gives a band roughly between 37-40 kDa and this image also indicates a band below 43 kDa...It must be ok, just analyzing inaccurately due to the smile effect you had. You should decrease the loaded total protein amount or (more beneficial) applied voltage to get straight bands, then you can agree the band would be at the correct position. Also, run the gel in a cold environment not let it be heated during the process, this also disturbs the band alignment...
  • asked a question related to Western Blot
Question
5 answers
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?
Relevant answer
Answer
Used RIPA, 40 Hz, 4,800 rpm, for 1-min pulses, three times, with a 30-s rest on ice between pulses, using carbide or zirconia beads...
  • asked a question related to Western Blot
Question
5 answers
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.
Relevant answer
Answer
What I usually do is after activating PVDF membrane with methanol for 2-5 minutes, use Western-Blotting transfer buffer (usually is Tris/Glycine buffer containing 20% methanol) to wash the PVDF membrane for 5 minutes before do the transfer.
  • asked a question related to Western Blot
Question
1 answer
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?
Relevant answer
Answer
Hi Sya,
I hold the view that you are one of proficient scientist, so I am pleased to discuss this issue with you.
I am curious if you have ever investigate the level of cell division marker. The accumulated cell cycle (cell growth) causes cell division and we call it as cell proliferation.
So many factors are concerned in cell cycle, like growth factor, proliferating complexes, or division enzymes. And separase, regulating division of cell, could be one of critical factors.
However, as we know, if the level of separase is downregulated, the imbalnced DNA quantity cause the instability of viable cell. Nevertheless, it is critical, the cell cycle is not affected by absence of separase.
This events could be proved by confirming number of chromosome in a single cell.
However I cannot guess the reason why the level of separase influenced by knock-down cell producing process.
My assumption could be wrong, and not be important in your work procedure, but I offer my opinion cautiously.
I'll give you a reference :
And here are some excerpts from the article.
"failure of chromosomal segregation is not the cause of cell death in separase-deficient cells."
"the presence of nonsegregated chromosomes in separase-deficient cells does not appear to cause additional defects in cell cycle progression events, such as DNA replication or centrosome duplication."
Thank you.
Junhyung.
  • asked a question related to Western Blot
Question
4 answers
Hello all,
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?
Thank you.
Relevant answer
Answer
Daniela Liebsch Thank you for your answer. Yes, the gel is totally fine. I always activate the gel for total protein and everything seems normal. I will try to troubleshoot by the possibilities you mentioned
  • asked a question related to Western Blot
Question
2 answers
Dear all,
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.
Relevant answer
Answer
Exosome marker antibodies are commercially available and can be used for western blot analysis to detect exosome-associated proteins.
Or check for the other markers to confirm.
  • asked a question related to Western Blot
Question
2 answers
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?
Relevant answer
Answer
Dear Kriti,
Did you validate your antibody before starting?
  • asked a question related to Western Blot
Question
3 answers
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 ?
Relevant answer
Answer
Dear @Romain Gioia, this could happen if the first polyAbs are against the peptide from the whole target protein. In WB an endogenous protein may mask this epitope/peptide. The other reasons are already mentioned.
  • asked a question related to Western Blot
Question
4 answers
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?
Relevant answer
Answer
Protein expression tells you, how much protein you have. Protein activity tells you, how active the protein is.
Imagine you have workers supposed to dig a ditch. How many workers you have = protein expression. The more workers, the more work they do per time.
But rather than letting them work with bare hands :), you can give them shovels. This would be some activator, for example some allosteric activator or phosphorylation of *some* enzymes.
Or you can let them work without boots and thus inhibit their activity, because who would like to work without boots, right? This would be analogy of inhibition.
Other aspects that affect enzyme activity are for example temperature and pH.
Yes, to determine SIRT1's activity, you need some activity assay. There is surely something published. You would need to have acetylated protein and peptide and measure either removal of the acetate from the protein/peptide or formation of the O-acetyl-ADP-ribose (product), or decrease of NAD+ in the reaction.
There seem to be several activity kits available:
  • asked a question related to Western Blot
Question
2 answers
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.
Relevant answer
Answer
Thank you ! I used serum starvation medium without glucose for 3-4 hours and then used low glucose/ high glucose medium in the absence of FBS while insulin stimulation time.
  • asked a question related to Western Blot
Question
4 answers
what are the key steps included in this to ensure specificity and reliability
Relevant answer
Answer
You can ensure the specificity and sensitivity of Western Blot results if you follow these measures.
To carry out a successful Western Blot, the quality of the primary antibody is a critical factor. Therefore, validation of the antibody will be important to avoid inaccurate results. Validation would include determining the optimal antibody concentration for the protein of interest. After validating that the antibody specifically recognizes the target protein it is important to determine the linear dynamic range for the target protein using the validated antibody against the samples being investigated. Different antibodies show different linear dynamic ranges, especially at high total protein levels. It is critical that the western blot be carried out within the linear range of the antibody being utilized. This can be easily carried out by doing Western Blots using different sample dilutions.
The specificity of antibodies can readily be determined using positive and negative controls. The best positive controls would be purified proteins or lysates which overexpress the target protein, while the best negative controls would be tissues from knock-out animal tissues or cell lysates.
Another factor to consider are the housekeeping proteins which are excellent normalization references when validated for the tissue or cell being investigated. But recent findings suggest that these housekeeping proteins are substandard loading controls under many conditions. Reasons include the presence of numerous poor-quality antibodies to each housekeeping protein, the relatively high expression of housekeeping proteins in many tissues and cells, and the change in expression of housekeeping proteins in some tissues under certain experimental conditions.
Both the protein of interest and the housekeeping protein need to be within the linear detection range for their expression levels to be accurately determined. Housekeeping proteins are high abundance proteins and therefore are often overloaded, particularly when large amounts of total protein are loaded to detect low abundance target proteins, so that housekeeping protein expression levels cannot be quantified accurately. Another overlooked aspect of housekeeping proteins is that they are highly post-translationally modified, which can also potentially affect quantification depending on the epitope of the antibody utilized.
Total protein quantification by Coomassie blue, Ponceau S, and the Stain-Free method has been shown to have advantages over housekeeping proteins for normalization of Western Blots mainly since this normalization does not depend on expression of a single protein.
There are some more measures which you could follow to ensure that your Western Blot results are reliable. You may have to refer to the article attached below.
Best.
  • asked a question related to Western Blot
Question
1 answer
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.
Relevant answer
Answer
We are using the same system, to give a helpful answer, can you elaborate the type of gel, membrane you would like to use and the blotting system?
  • asked a question related to Western Blot
Question
1 answer
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.
Relevant answer
Answer
1. Could the treatment actually affect expression of these proteins?
2. Maybe try a different quantification method? Maybe there is some error with the quantification?
3. Error introduced by pipetting during sample prep or loading the gel?
4. Problem with transfer? Sometimes the transfer will occur unevenly.
If you do a ponceau stain on the membrane right after transfer you might get an idea of what is happening. That should indicate if the amount of protein being loaded is actually even, and/or if there is an uneven transfer issue.
  • asked a question related to Western Blot
Question
2 answers
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!
Relevant answer
Answer
If we have to exclude problems related primary AB and other technical issues, you would make sure about the source of these bands by knocking down or out the gene and detect again for the presence of these strange bands.
  • asked a question related to Western Blot
Question
8 answers
Could anyone tell me how can we reuse the PVDF membrane in the western blot?
Relevant answer