- Brittany Wingham added an answer:Is anyone using Abcam Anti-CD40 ab13545 for Western blotting?
I have just begun western blots looking at CD40 in immune cell pull downs. The antibody daasheet states the band will be apparent at 41kDa, although the predicted size is 34kDa. I am not too concerned about my blot, as I have just done a ponceau stain which showed a clear band around 35kDa (primary overnight to image tomorrow).
I just wondered whether anyone else had a similar experience with this antibody in particular?
Thanks very much!
Hi Gaetana, thanks very much for your reply. I have had a second repeat over the weekend which showed similar results, so I may purchase the Santa Cruz antibody for my next run. When i've ran westerns on CD40 before with different treatments, the band was also at around 42kDa so to 35 just seemed too low. Thanks again for your help.
- Caroline Loos added an answer:Can anyone help with protein aggregation - Western blot issue?
I'm looking at a protein of approximately 140-150 kDa which is larger than we have ever looked at in our lab so I needed to adjust some steps in our Western blot protocol.
From suggestions I found online I used 6% gels and started transferring overnight (16h) at 20V instead of 1h @ 300mA like we do for other smaller proteins. The transfer seems successful and I'm getting bands but I'm getting these huge bands all the way on the top of my running gel at about 260kDa. I see them even after staining the membrane with fast green (which we do as a loading control for total protein) so its not an antibody issue. I have never had this before with any of my other proteins, therefor I think it must be caused by the different methods used.
Does anyone know if under these new conditions (6% gels + transfer overnight) it might be more likely to get protein aggregation in the top of the gel for some reason?
The muscle samples I used are the same as I have been using before with all my other proteins for the past 2 years. With 8 and 12% gels I have no issues. Therefore I do not think its the sample but rather the method of separation or transfer since those are the only differences between my current method and the ones we have been using in the lab for a long time.
I did not boil my sample because I thought that might help but I still see aggregation in the top of the gel.Following
- Lael L Cheung added an answer:What is western blot actually detecting? How do I mark standards during wester blot imaging?
I am running urine samples on a western blot in attempt to detect IgM (urine samples with known IgM levels). Today we detected numerous high molecular weight bands. I was wondering if someone could share methods to confirm that the western blot is actually detecting IgM or if it's staining more non-specific (i.e. other proteins, protein fragments, or background). Also, we are imaging on a Imaging Quant machine using Imaging Quant 4000 technology. Is there a way to mark the standards so that they appear on the same image as the samples (i.e. write on them with sharpie before imaging, or superimpose the standard image with the sample image)?
I appreciate your time and thank you in advance!
• Per your question about overlaying visible MW standards with target signals in a Western Blot:
Internet searching for Western Blot Pen finds several vendors offering pens with ECL-active ink. After tracing over the MW standard, treatment with detection substrate will cause the standards to appear by ECL. Depending on the fineness of the pen tip, you may also pre-annotate your blot with sample identifiers and other helpful information for convenient digitization as part of the blot image. Prices range from $186 to $49, so choose one that best fits your needs and budget.
Per Dr. Frausto's suggestion, there are MW standards that will automatically appear by ECL. If such products interest you, it would be prudent to review their technical documentation. Aside from price, you may want to consider the mechanism by which these standards appear by ECL. Some of them use standards that competitively bind to target-detection antibodies. Because there will be a lot more standard than target, most of your detection antibodies may sequestered the standard, thereby reducing the attainable ECL signal from your target.
The imager that you are using has been discontinued. If the user manual is missing from your facility, then perform an internet search for PDF copies archived by the original vendor or uploaded by other customers. Note: If your imager is too old, then it may be incapable of overlaying signals from different detection modalities.
• Per Dr. Butnev's explanation above:
Depending on the primary antibody that you used, the multiple bands that you saw in the Western Blot may be due to the pentameric nature of IgM. In its intact state, IgM has five copies of mu-type immunoglobulins linked together by a J-chain polypeptide and multiple disulfide bonds. When subjected to denaturing SDS-PAGE, the IgM (intact MW >900 kDa) will fragment into the following components: immunoglobulin heavy chain mu-type (MW ~65 kDa, 10 copies), immunoglobulin light chain (MW ~25 kDa, 10 copies), and J-chain polypeptide (MW ~15 kDa, 1 copy).
A thorough confirmation of IgM presence by Western Blot would include loading controls of all immunoglobulins that could pass through damaged glomeruli into the urine: IgA, IgD, IgE, IgG, and IgM. For the cleanest signal, use an antibody specific for the IgM mu-type heavy chain. Multiple vendors sell this type of antibody, including HRP-conjugated versions, so choose a vendor that you trust. Verify that the antibody does not cross-react with blocking buffer proteins that coat the rest of the blotting membrane. Otherwise, using an antibody that is not raised against a definitive component of IgM (mu-chain) may lead to multiple bands being detected and thereby complicate subsequent analysis. The J-chain is present in both IgM and IgA, so it is not as definitive for IgM as the mu-chain.
Please excuse the lack of brevity. Hope this information helps.Following
- Peter J Vollbrecht added an answer:Can I Measure Glutamate level by western blot?
I found Antibody against Glutamate is available...but is it possible to perform WB for a single amino acid?
Saikat, I think the best way to measure glutamate would be using microdialysis techniques in vivo. I am not experienced in cell culture techniques.Following
- Heidi Kaastrup Müller added an answer:Do you have any suggestions regarding Odyssey infrared immunoblotting?Blocking buffers (Licor, Rockland) are expensive (up to 200€ per 500 mL). Is there any less costly alternative out there?
I am using the standard 0.2 µm Nitrocellulose membrane (Cat# 170-4159) from Biorad. After the transfer I incubate the blot in TBS for 5 min before adding the blocking solution. For the past 6 months I have used the Odyssey Blocking Buffer diluted 1:3 in TBS and get really nice results for both low and high abundance proteins, including phospho proteins.
I have tested different concentrations of Tween-20 during incubation with the primary antibody and see no difference between 0.05%, 0.1%, 0.2% and 0.4%. Now, I routinely use 0.1% Tween-20 during both primary and secondary incubations. I also have 0.01% SDS present during the secondary incubation. After the last wash with TBS+0.1% Tween-20, I wash the blot in TBS before scanning the image. Also, my TBS solutions are prepared from Trizma Base (pH adjusted to 7.6 with conc. HCl) and not as a mixture of Tris-HCl and Tris-Base. This also appears to give better results.
All the best,
- Eva Mayr added an answer:How can I get rid of slurred bands in protein sds-page/western blot?
Want to immunoblot my 3T3-L1 lysates by sds-page/western blot.
I use a 10% seperation gel (self made), load about 50µg Protein and run it 1h at 50V and then turn up to 100V for an other hour. Via wetblot the proteins are then transfered to a PVDF Membrane, blocked by 5% BSA in TBST and stained with primary AB overnight, then 1 hour secondary AB.
Now the PROBLEM: As you can see in my figures i get acceptable bands for the bigger proteins (60kDa), but the 37kDa Proteins always generate slurred bands. Also the small proteins of my prestained size standard make slurred bands.
How get rid of this slurred bands?
Thanks for your advices! I did the experiment again, with less protein (20µg) and stopped the electrophoresis earlier (the bromphenol was about 0.5cm above the end of the gel. I got clear bands and also my molecular weight standard was sharp. Thank you very much!Following
- Maria Vitale added an answer:How can I get the data from this western blot picture?
As you can see in the picture, I have run two samples on SDS-Page, and signals have been detected by PARP antibody. Control has no band, while the treated sample is giving band, indicating that the PARP has been cleaved. Now I want to get this data for my paper, but due to black background, It is difficult for me. Can someone help me in this regard. How can I use photoshop for getting the desired bands?
Looking at your blots definitely you need to repeat the assay. The dark spots diffused in several point are usually indicatives of inadeguate blocking step of the membrane .Are you blocking the blots in milk, bsa, or other buffer? For how long? A less concentrations of proteins could help in a less smearing and a more clear bands.
A little less exposures time can help also to look better at the bands.
But it could also be possible that some degradation is in place ( high intensity of lower bands ). So it is important to know exactly other details of your experiments.
Definitely not modify the figures of real results.
- Amar Kumar Singh added an answer:What could be the reason if half of the protein ladder in western blot is transfereed on to the membrane and half (the heavy ones) is not transferred?
Hi I transferred my protiens from gel to the blot but the ladder was not found completely transferred can anybody suggest me the reason?or could it be because of voltage setting?
Transfer of larger protein is little difficult. It depend on time of transfer, gel concentration, methanol percentage. Increase in time and decrease in gel concentration would help to transfer protein better. The question to be responded would need more detail to give better response. Details of range of proteins not transferred, transfer image, time, voltage, buffer mix component etc. RegardsFollowing
- Maya Miller added an answer:How can I confirm the presence or absence of a protein after drug treatment using Western blot?
As per my research, drives me to inhibit a particular gene which codes for a protein involves in multiple functions.
After treating the cells with my designed drug against that protein, i have found the expression of that particular mRNA got reduced by Quantitative PCR.
But i want to confirm it at the protein level.
Since the target gene is novel, there is no commercial antibody available to it right now?
Is it possible to find the protein level by any other staining or non-antibody mediated method like fusion of GFP to the target gene etc.,?
since raising an antibody against that protein cost (in lakhs) which i couldnt afford it right now.
kindly help me in this regard!!!!
Following Adam's and Tichaona's advice. I think you can do both. If you know the size of the protein you can identified it through western blot and actually take the band for analysis at LC\MS\MS method. You can quantify it's amount and to be sure that this is your target protein you can sequence it (through LC\MS-MS- just your band).
Regarding Erik's note... You have to do the experiment in different concentrations of the drug. In this way you will have a decreased amount of the protein relative to your control.Following
- Jennifer Pérez-Boza added an answer:What are the best Exosomal Markers for Western Blot?
I work on exosomes isolated (via ultracentrifugation) from MDA-231 breast cancer cells.
When I run westerns to confirm isolation I generally use Alix and Flotillin - these are antibodies I borrowed from another researcher in my department. I get some mixed results - the Alix signal is general weak if present at all. I also realized the Flotillin ab I have is for Flotillin-2 and not Flotillin-1, the more classical exosomal marker (at least in papers I have read) - are both of these proteins supposed to be present in/on isolated exosomes?
In terms of recommended proteins to blot for I have seen -
Are some of these markers more enriched in breast cancer lines/exosomes than others? I cannot purchase all the antibodies at once and am hoping to prioritize by what is most likely to be successful.
I'm also curious about what kinds of protein yields other people are getting from ultra-centrifugation isolation. I generally get only around 3ug of protein total - this is modest considering the starting material is 100ml of conditioned media. The low protein yield may relate to the inconsistencies I am experiencing with Alix and Flot-2.
In Summary - What markers have people had success with when performing western blots on MDA-231 (or other breast/epithelial cancers) derived exosomes? and how much exosomal protein was loaded to detect consistent banding?
CD63 works very well, though it would also be interesting to add a specific surface marker of your cell type so that you can confirm their origin. Also, if you want to buy any antibody, and have where to choose, cell signaling is a brand that usually gives good results...Following
- Sandipan Chowdhury added an answer:Is it possible to know the stoichiometry of two proteins interaction at physiological level?
I tried to know the stoichiometry of two proteins using size exclusion chromatography but i found both form aggregation and eluting in void volume, so could not be able to study. Now i am using whole cell lysate and run on size exclusion chromatography (Superdex 200HR 10/300 GL column) then collecting fractions of 100 ul and checking out them on western blotting using protein specific antibodies but not getting band on desired position according to my positive control.
Is there any way to calculate the stoichiometry of these proteins? please suggest me...
The answer to your question depends largely on 2 factors: (1) affinity of binding (2) how much of the individual proteins you are able to obtain (preferably purified protein). Some of the techniques available are as follows:
(1) Isothermal titration Calorimetry - for this you would need ~20uM of purified protein and the affinity of the proteins needs to be strong for the calorimeter to be able to measure the heat of binding accurately. However, complex stoichiometries (especially if the stoichiometry is at least partially conc. dependent can significantly contaminate results).
(2) Analytical ultracentrifugation - this is actually a pretty good technique if your protein pairs are capable of forming multiple stoichiometries - you will be able to obtain Kdiss of each stoichiometric combination. But you need significant amounts of purified protein for this.
(3) Glutaraldehyde cross-linking - cross-link your purified protein partner with glutaraldehyde (which is non-specific as it relies on exposed lysines) and run a gel (or do a western) to identify higher molecular weight bands. Compare this result with Blue-native PAGE (non-denaturing gels) analysis of uncrosslinked protein mix. While this might be the simplest/easiest thing to try - the results can be quite notorious because often several molecular weight bands will be seen - that said it is quite certainly the first step
(4) Mass-spec is certainly an option - but once again concentration dependent multiple stoichiometry will make results hard to interpret
(5) Dynamic light scattering - using scattering of light to measure molecular radii of different species which exists in solution and thereby relating it to molecular weight and size. Requires significant quantities of purified protein
(6) FRET - tag the two proteins with GFP/YFP - purifiy them and measure FRET. This will need small quantities of protein. But if there are multiple stoichiometries then interpreting FRET ratio could be tortuous (FRET signal will change with stoichiometry of association as well as distance between the FRET pairs). Also GFP/YFP might itself affect the stoichiometry of association, especially if the pair forms a moderate to weak affinity complex.
(7) SIMPUL - this is probably the best technique but it requires a TIRF microscope. Have the two proteins tagged with say Strep and Myc tags. Add fluorophore tethered primary Ab to the protein mix - lay the protein complex+Ab mix on a cover slip and perform single molecule photobleaching experiment. Very tiny amounts of protein will be required - starting sample need not be very pure (actually could simply be clarified lysed cells) - but instrumentation and software needed are not the most commonly available ones.
Hope this (along with other answers listed above) helps you think about what will be the best option for you.
- Richard Christison added an answer:Why would I not be getting recombinant protein from mammalian cells?
I have transfected CHO and HEK293 with constructs for 3 different proteins. I get resistant colonies (compare to mock transfected) which grow up after 3 weeks or so. The proteins should be being secreted but I cannot detect them in the supernatant by western blot or elisa (i have also checked cell lysates). The constructs have been checked and are all in frame with no mutations etc. Could it be that the proteins are aggregating/being degraded?
Checking the mRNA will tell you something, you should at least get a better idea of where to look for you problem.
Unless there is some compelling reason for production of a stable line it would be a good idea to at least look at the other expression systems as some systems suit some proteins better than others.
I am also a fan of checking a pool of transfected cells after a couple of days when setting up stable lines - it can save a lot of work catching a problem early.
- Fuad Bahram added an answer:Any suggestions for myc protein westernblot?
There are two sepcific and strong bands when I use anti-myc(santa cruz 9E10) to analysis the whole cell lysates of transfected flag-myc 293t cell.But I could not detect any specific bands when I analysis the whole cell lyasates of HELA, HCT116,U87,MEF cell.The process of lysing is as below
1,wash the plates twice with cold PBS
2.add 200μl RIPA buffer into plates and scratched the cell into a tube
3.incubate the tube on ice for 20min
4.sonic 5s and break 2s for 12 times
5.centrifuge 13500 rpm for 15min,4℃
6. using supernants for 10% SDS-page immunoblot
Are there any problems?
does anybody konw how to successfuly detect endogenous myc protein?
Should I pay attention to some details specially?
We have been working with Myc in more than 20 years. Myc could be difficult to detect in many cell lines due to the level of the endogenic protein. In case you want to immunoprecipitate Myc then running WB you could Ip with N-262 and for WB use C-33 but biotinylated to reduce the detection of Ig bands. and as secondary add Streptoavidin. Check many of our publications.
- Viktor Y Butnev added an answer:Most of protein is still stuck in the origin of resolving gel after transfer, how can I improve the resolution of the bands?
Stacking gel is 5% and resolving gel is 7.5% and my protein is 260kDa. I loaded 30ug protein in a volume of 50uL. I did a wet tank transfer 120V for 1 hour with cooling. I don't know why most of the protein is still stuck in the interface between stacking and resolving gel. Please help. I have attached a picture in ppt if it helps.
The proteins about 250K are often stuck between the stacking and resolution gels because they are bound to some other membrane (hydrophobic) proteins. I would suggest to try the commercial gradient gels with low concentrations of acrylamide. I would also recommend to use Laemmli SDS-sample buffer (2% SDS and 5% b-mercaptoethanol) for the extraction of the membrane-bound proteins from the cells. After that you will need to make a high speed centrifugation at least at 10,000 g (because a lot of insolubile proteins and DNA will remain in the pellet). Then, you will need to boil your sample for 3 min at 100*C and load it on your gel. Since not all of your protein will be extracted from the cell membrane as a monomer you might want to use a higher concentration of your 1st and 2nd antibodies for your western blots. Hope this might help.Following
- Emilia Manole added an answer:Can we co-incubate the primary and secondary antibodies at the same time for Western Blotting ?
Generally, we incubate the blocked membrane with the primary antibody, wash away the unbound primary antibody, then incubate with the secondary antibody which binds to the primary antibody, and detect the protein of interest on the membrane. Since the primary antibody binds its epitope with its variable domain, whereas; the secondary antibody binds with its constant domain, there should be no reason that mixing these two antibodies together will interfere with the binding. Can we incubate the blocked membrane with both primary and secondary antibodies at the same time for Western Blotting ? If it works, it will save time.
More of this, the time of incubation is different for the 1st and the 2nd antibodies. Sometimes it is better to incubate the membrane with the 1st antibody over night at 4 dgrs for a proper result. If you put the secondary antibody a lot of time with the membrane, or more concentrated, it would be possible to bind unspecific and to obtain many unspecific bands or background. And it is right that the TBST washes between the antibodies have a great importance in obtaining a good and clear result of WB. So the protocol was established very well and for certain reasons in the way to put the 1st and the 2nd antibodies separately.Following
- Liu Jiyun added an answer:Do you have any advice on my protein 1st Dimension Run Problem?
There is problem with the running of 1st dimension of proteins. The sample is isolated from leaf and midrib of banana plant. We have optimized the protocol and getting good results but there is problem with the running of 1st dimension of leaf now. It does not run properly. Usually, 1st dimension run from negative to positive, in these cases it start running from negative to positive but after 1:30 hour, it also starts to run from positive to negative side as well. This problem persist until the run is over and we got the strips in this form(picture attached). Firstly i thought it might be some handling problem or with the samples, so I changes the sample and run with the midrib replicate. Problem persists. Now, i do not know what is the problem with them. New Rehydration buffer and 1M DTT stock were prepared but issue was not solved. Room temperature was 22C.
First I thought that it might be due the improper washing of the focusing tray, but we cleaned it well and still the problem persist. Can this be due to the Protean IEF Cell Bio Rad? It got some problem and now running the strip in this manner?
Hope to hear positive.
I agree with Manuel Sebastián- The phenolic extraction method is more suitable for the banana plant protein extration compared with other methods as it can clear polysacharides effectively. The detailed protocol can be seen in the article: http://www.nature.com/nprot/journal/v1/n2/abs/nprot.2006.102.htmlFollowing
- Wolfgang Schechinger added an answer:What could have gone wrong with my Western Blot?
my Western Blot looks like this (see attachment).
The samples loaded onto the gel are cell lysates, I blotted for 4 hrs at 4°C and 300mA (transfer buffer: Tris Glycine with 20% MetOH), which I've done before and so far it has always worked fine.
Does anyone have an idea what might have happened?
Elizabeth, have you prepared all the samples the same way, i.e. are they comparable? Looks to me there was problem with the gel rather than with the blot - different salt concentrations in your samples on the left and on the right (where you suspect bad transfer)?Following
- Fernanda R C Giachini added an answer:How to store a membrane in Western Blot?I am a beginner in western blot.
Sometimes I need to store the PVDF membrane for later re-probing.
I have asked my colleagues, some said I should dry the membrane and store it in 4 degree
but I was told by others that the membrane should not be dried in any process,
some said the proteins may degrade, some said it will result in higher background signals.
So most of my colleagues store the membrane in TBST in 4 degree.
But when I search in the internet, I found that some people did dry the membrane in the storage process.
There are a few questions I want to know:
1. The rationale of drying the membrane in storage,
2. The consequence of drying the membrane, why the membrane should be kept wet?
3. How long can a membrane be kept, without affecting the result?
4. Which method can store the membrane for a longer time?
Does dry-air storge work for nitrocellulose membranes?Following
- Kyohei Miyazaki added an answer:Does anybody know a good antibody to claudin-5 for western blot?
Hi, I use HUVEC for tight junction research. I struggled to get bands of claudin-5 for westernblott(WB) using some antibodies. I could obtained good results from immunofluorescent stain using same antibodies. Anti claudin-5 antibodies I tried are rabbit anti claudin-5 polyclonal antibody (IBL, Takasaki, Japan), mouse anti-claudin-5 monoclonal antibody (invitrogen, catlog No. 35-2500) , and mouse anti-claudin-5 monoclonal antibody (Santa Cruz, sc-374221) . WB protocol is followed; blocked by 5% or 10% non fat dry milk / 0.1% Tween20 / TBS at room temperature for 1 hour, followed by incubated with 1:500 diluted primaly antibody in 1% or 5% non fat dry milk / 0.1% Tween20/ PBS at 4℃ overnight, and HRP-conjugated anti mouse or anti rabbit IgG as secondaly antibody diluted 1:10000 in 1% or 5% non fat dry milk / 0.1% Tween20/ PBS at room temperature for 1 hour. I could not get significant bands of claudin-5 at 23kDa. Protein applied to SDS-PAGE was about 30 ug/well. Does anyone let me know good anti claudin-5 antibody for WB? Thank you.
Dear Attila Lehotzky
Thank you for your suggestion. claudin-5 is expressed in HUVEC, but very low as you pointed out. Further more, antibodies I used had many non-specific bands, and these bands signals were stlonger than that of claudin-5, so the band signal of claudin-5 was hard to detect. I'll modified with secondary Ab dilution and incubation time.Following
- Trang Mai added an answer:Can you show the picture of your worst Western blot, and explain the reasons for this disaster please?I start this topic in order to collect the gallery of the photos of Western-blots (WB) as ugly as possible. The detailed description of the problem and the technical solutions were applied to resolve it are strongly suggested, however ugly WBs without the clue what is wrong are welcome to discuss it here. The overall idea of this post is to help the young researchers to resolve the problem with simple and cheap tricks. On the photo below you can see the lightning in the middle, dust in the upper left corner, small air bubble in ECL solution in the upper right corner. The background is not fully grey, means ECL was not properly distributed, only band visible is not specific binding of the antibody to the slightly degraded marker band (right).
Sometimes I used the 12% acrylamide SDS gel but it couldn't separate my proteins below 28 kda. All the proteins below 28 kda (including the ladder, the lower red one is 28 kda) were in the same line with the dye. However when I prepared the gel several days before and stored it in the complete running buffer 4°C, the gel worked fine. It could be the pH problem? Both the gel and the buffer contained 0.1% SDS. Thanks for your comments !!Following
- Madhur Agrawal added an answer:Why was Sirt1 detected at 75 kDa but not 110 kDa?
Hi, I'm studying Sirt1 on mice muscle. The antibody has not a specific epytope to recognize, so it should detect the band at 110 kDa. The problem is that I have only a single band at 75 kDa in wild type and mutagenized mice. I know that the 75 kDa is an inactive fragment, but I expect to detect the active form in wild type. Can someone help me?
I would suggest you to contact Abcam once and in the mean while, try cell signalling antibody, it works well for me for 110 KD band.Following
- Bulut Hamali added an answer:Does anyone have any idea why I only have low weight size in western blot?
I have been trying this for more than 2 months, but it is always the same result. I am doing Fasciola hepatica whole cell lysate like below:
Adult parasites were homogenized in 10 mM Tris–HCl, pH
7.2 containing 150 mM NaCl, 0.5% Triton-X100, 1 mM
EDTA, 1 mM PMSF and 1 tablet complete Lysis protease inhibitor. To remove cellular debris, thehomogenates were solubilized for 30 min at 37C, followed
by centrifugation at 12,000g for 15 min at 4C.
When I do Ponceau S staining I do not see any higher bands than 40kDa. I load about 15-20 ug per well, but it did not improve until now.
And Thanks a lot Wolf. I actually did always semi dry transfer but I will think about to try wet blot as well. Thanks a lot for suggestion.Following
- Wu Anguo added an answer:What is the best percentage of gel for a 14 kDa protein using western blot and the best voltage for transferring the protein from gel to membrane?Running a western blot gel using a 14 kDa protein.
12% gel, I usually use 12% gel to analyse 14-16 kDa protein such as LC3Following
- Stephan P Tenbaum added an answer:DNA dot blotting and Licor Odyssey infrared detectionantibody, detected using the Li-cor Odyssey infrared system. We have used the instrument previously only
Quantify DNA and put equal amounts in 20-50 ul H2O or TE. (for tiny amounts you may choose Qubit quantification.)
Dot/Slot blot straight to the NYLON membrane (Amersham Hybond N+, GE Gealthcare) – I use a vacuum blot (Biorad). Do not heat the sample before loading. (If you boil you may get DNA denaturation resulting in unknown amount of ssDNA stretches, since I use dsDNA antibody as loading control heating would potentially alter the result.
I tried UV crosslinking, of the membrane and baking worked better. I boiled samples before blotting and got worse signals. I blotted with SCC-based buffers and they worked worse in my hands then straight blotting in water.
Fix by baking at 80ºC for 2h
Block directly with 5% milk in TBS-Tween for 2h
In the following I combine HRP/ECL detection in combination with fluorescent secondary antibodies so I mix mouse dsDNA antibody and rabbit 5hmC primary antibodies for incubation (o/n 4ºC). So I get the loading ctrl of the same spot.
The same way, I incubate together a mix of secondary antibodies (anti-rabbit-HRP (1:5000/1:10000) and anti-mouse-Alexa flour 680 (1:5000) for 1-2h. Then, I first (!!!) do fluorescent detection in a Licor Odyssey or camera-based system to detect dsDNA signal strait with the washed wet membrane and then I incubate the blot with standard ECL. (The fluorescent signal is easily quantified (linear signal) – ECL is more sensitive as signal is accumulative on the x-ray film and may be exposed longer at will. ECL solutions may screw up fluorescent secondary signal detection if you do the other way round.
I use a mouse anti dsDNA antibody (ABCAM HYB331-01, ab27156) as loading control (1:10000) and 5hmC antibody (1:1000) Active Motive. Cat. Nº 39769. I also tried a mouse Methyl-cytosine antibody once and it worked fine.
I hope this helps, good luck!
My very best,
- Vanessa Mendes added an answer:What are the causes of a bad Western blot result?
I did the Ponceau on the last run, and it came out looking like these two images, attached.
What could cause this, besides bad gel?
I'm having the same problem. How did you solve it? Today I tried more sonicating and DNAse I, but it didn't seem to improve.
- Fernanda Mügge added an answer:Does anybody know of ATF6 (Activating transcription factor 6) antibodies that have worked in INS-1 cells for western and immunostaining?
I am planning to look at the expression of ATF6 in INS-1 cells under high glucose or ER stress. There are number of antibodies out there but I want to know if anyone in researchgate has personally used any of these and has worked without much trouble for western and immunostaining. Thanks.
were any of you successful?
I work with MEFs and use a SCBT antibody. For Western Blot it is VERY difficult to get a good result for ATF6. I've tested at least 4 extraction protocols and the last one, which leads to a very concentrated cell extract is the best so far, but I get a LOT of innespecific bands;
Today I am using this same Ab for immunofluorescence and don't know if it will work...Following
- Narendra Singh added an answer:What is the best protocol to make ECL (western blotting) mannualy at lab?
cancer biology researchers, those do western blotting frequently.
Thanks dear Wassim.Following
- Amy Schutz Geschwender added an answer:Does anybody have experience with blocking reagents compatible for the LI-COR Odyssey?I am using the LI-COR Odyssey system for Western Blot detection. I have high background levels
Yes, every lot of PVDF membrane from LI-COR is QC-tested for low background fluorescence.Following
- Denis Dermadi Bebek added an answer:Are the protein samples extracted for 2-D gel using urea and CHAPS buffer valid for direct 1D western blotting?
I need to confirm some results of 2D immunoblotting by carrying out the western blot (1 D) using same samples that are prepared for 2D using rehydration buffer contains 8M urea, 4% CAPS, 40 mM trisCl 6.8 and DTT. as i know that boiling urea will cause carbamylation. i wonder whether i can use the sample directly in my WB without boiling.
when i perform the 2 D gel of my proteins in the rehydration samples the second dimension works perfectly so i assume loading the samples directly will also work.
is the equilibration step after IEF contribute to the resolution of the 2nd dimension?
incubation at room temperature with SDS and b-mercaptoethanol for 15-20 min helps a lot. I run my samples that were extracted in urea.Following
- Norbert Kartner added an answer:Best way to resuspend Acetone-precipitated protein for Western Blot?I'm using QIAGEN kit to extract RNA and to better use these samples, I decided to extract protein as well, which QIAGEN recommends to do so by adding acetone to the flow through.
Judging by the size of the pellet I've been able to collect great amounts of protein, however, when I try to resuspended in RIPA buffer or PBS alone, the pellet barely dissolves.
Does anyone have a suggestion in how to resuspend the pellet or what buffer to use?
Try PRMM method, doi: 10.1128/AEM.70.1.610-612.2004
Haven't done an acetone precip. in over a decade, but I seem to recall that barely-dried pellets dissolve in a minimal amount of just plain distilled water. Use a flame-sealed Pasteur pipet as a stirrer to suspend/dissolve the pellet. Any ionic strength changes solution time from minutes to hours. You can add buffers etc. after dissolution. BTW, if you use urea, do NOT heat your sample above 37 deg. Urea and heat will transcarbamylate your protein (particularly messy for 2D gels). Methanol precipitation has to be cold, not room temp. if you want anything approaching quantitative recovery for dilute mixed proteins.Following
About Western Blot
Western blot is a widely accepted analytical technique used to detect specific proteins in the given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native proteins by 3-D structure or denatured proteins by the length of the polypeptide. The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are stained with antibodies specific to the target protein.