- Anton Beletskii added an answer:2Any advice on why KLF2 is not appearing on Western Blots?
We are trying to detect and quantify the presence of KLF2 which should appear as we have memory cells (CCR7+/CD62L+). We're using western blotting for that and we've tried both the traditional "long" method, and we've tried ProteinSimple's Wes Machine. Neither can produce a confirming band.
-We've tried multiple antibodies from different companies
-We've tried checking our cultured cells from D0 through D6 every day to see if KLF2 may have come and went
-Maybe we thought that KLF2 exists in small quantities, so to exclude the noise from extracting all the proteins, we've done a nuclear extraction and ran a western blot on that- No band
- We've tried varying both input protein concentration and antibody concentration
Any suggestions would be great! We've been trying to detect this elusive KLF2 for a while.
We've ran western blots on many proteins, and those work fine. Only KLF2 doesn't work for some reason...
I am not an expert in this particular target, but it seems that proteasome inhibitor Velcade is boosting levels of KLF2.
You can try to boost KLF2 levels by treating your cells with Velcade, or at least treat them with protease inhibitor cocktails (cOmplete) during protein isolation to ensure integrity of KLF2.Following
- Radosław Kaczmarek added an answer:4Can someone recommend a good anti-his tag antibody for western blotting?Most of my proteins are his-tagged (6x). I'm shopping for a robust antibody to exam these protein with western blotting.
We have recently used a monoclonal anti-polihistidine (clone HIS-1) cat# H1029 from Sigma-Aldrich with good results.
It worked well in Western blotting and in flow cytometry.
- Julia Bartels added an answer:4Problems Blotting specific Proteins, any Suggestions?
I am having Problems Blotting specific Proteins on PVDF and Nitrocellulose from the Spore Coat of B. subtilis. The proteins are visible in the Coomassie stain and are loaded as a mixture of all proteins as well as the purified version. It is not a general Blotting issue, as I can detect one of the Proteins on the same Blot. It also does not seem to be a Antibody problem, as I have no problem in detecting them in a Dot Blot. There are a few things I will try in the future, but I wanted to know, of there is some specific advice, or if someone already had such a problem. I will try to equilibrate the Gel in MeOH for the PVDF membrane, I will use a stack of 2 membranes to exclude overtransfer, I will stain with Ponceau to see how the lanes look like (even though I already stained the gel after transfer and it is more or less emptly). Any other suggestions?
Thanks for all the suggestions!
@Kristýna Hrazdilová: The Proteins are all between 15-20 kDa so I use a 15 % SDS-Gel, but have also previously tried 10 % with similiar results for the Blotting.
@ Ianina Pokholenko: So the thing is, that in order to extract the Proteins from the Spore I cook them 2 hours at 95 °C with 0,1 M NaOH, 0,1 M NaCl, 0,1 M DTT and 1 % SDS bevore using them for either the Dot Blot or the Western Blot. I do not fully know, but guess that they are denatured from this treatment. If they are not, they will probably also not be in the SDS-Gel. The epitope we are trying to detect is a his-tag and we are using the penta-his antibody for the detection. But we may try a different antibody.
@Imran Khan: Yes I have stained the Gel after the transfer, and get almost empty Gels with a little bit residual proteins for the higher molecular weight (not the range of my protein though) and sometimes a middle molecular weight band, though not as strong as without transfer and also bigger in size as my proteins. The Proteins I want to detect are between 15-20 kDa and the same size as the one which I can detect. Nevertheless my Transfer Buffer contains 0,1 % SDS. But I have no problem with Overtransfer, as I see nothing on the second Blot if I use two membranes, even from the Protein, which I can detect. I have tried PVDF, Nitrocellulose and Nylon, where Nylon did not work at all, Nitrocellulose worse than PVDF which had the best results. But I might test different blocking reagents (Milk and BSA, or do you have other suggestions?)Following
- Sophie Irwin added an answer:12How does the protein extraction affect the western blot?
I'm currently looking at the phosphorylation of STAT1 and 3. Both the pSTATs are working fine on a western blot, and the STAT3 is fine as well. However the STAT1 is not showing up, to the point that no matter how I've adjusted the protocol, and had a colleague repeat the standard protocol for it, nothing comes up bar the ladder.
My colleague suggested analysing how I was processing the protein from my neutrophils, which is a valid point. It's quite a big protein, around the 85-90kDA mark, so wonder if that is a key factor and the way I'm denaturing and reducing it is not enough to break the bonds and creating the primary structure (so basically making it too big to run down properly) and that the pSTAT1 doesn't have this problem because its phosphorylated.
I've had my supervisor check on other samples that have worked for her, and even she's struggled a bit with it so we think it may be the antibody
thanks for everyones commentsFollowing
- Imran Khan added an answer:4Why does the probing for this antibody look so bad?
lately I have been struggling a lot with getting a good signal for a specific antibody on my western blot. I am particularly worried cause this antibodies works well for other lab mates of mine but we couldn't narrow it down to the mistake i am possibly making.
The protein of interest runs as two isoforms above and below the 188 kDa band in MES running buffer. I attached a picture of one of the several attempts I did and pretty much they all look the same.Note that lane 4th and 8th are EMPTY! That means I really don't know where the smear comes from. And most of my colleagues in the lab seem to think that this doesn't look as protein degradation.
I think it should't be a transfer issue cause if I use antibody for other high molecular weight proteins they work well and i get nice and sharp clean bands.
I tried to load less protein, dilute the antibody in milk versus BSA but nothing solved the problem.
I am trying now to load MORE protein to see if this changes something but after this and remaking again my transfer buffer I wouldn't know what do.
Any advice will be more than welcome!
Thanks a lot
- Amit Kumar MISHRA added an answer:2How do I remove scratch marks background on chemiluminescent western blots?
When I developed the protein signal with ECL on PVDF membrane, it displays many scratch marks background on the membrane. The blot was covered in plastic sheet to prevent drying. These scrath marks are the same one that appear when the membrane white-dried. (picture attached)
This membrane has been stripped two-three times before this time reprobling. The membrane was white-dried and resoak in 100%methanol for a few minute until the membrane turn grey, then washed with TBST before stripping>blocking with milk>reprobing. The primary antibody that was used in this time is high background one. So I washed it about 1hr wiht many change before developping.
On the previous stripping&reprobing with other antibodies, this problem had never occured, even the blot whited-dried and scratch marks appeared during the development.
What is a cause of the problem? How can I solve it? Can I sill use this membrane to probe this antibody?
- Adam Karpf added an answer:4Would it be better to measure NF-kB activity by western blot or Real-Time PCR?
Using anti-phospho IkBa with western blot or IkBa mRNA expression by Real-Time PCR
Activity of a transcription factor is best determined by a promoter reporter assay or by endogenous DNA binding assay by ChIP of a target promoter. In the absence of this, nuclear translocation of NFkB (suggested by Jordan Yaron) is probably the best alternative. Increased protein or mRNA levels are not neccessarily accurate reflections of transcriptional activity.Following
- Parthasarathy Chandrakesan added an answer:2Does someone have experience in detecting ATM, ATR, Chk1/2 via Western Blot?
I want to check the signal of several proteins via Western Blot (ATM, ATR, Chk1, Chk2 and the phosphorylated forms). The problem is: I don't get any signal. I tried different whole cell extracts but unfortunately without any result. The cells I'm working with are VH10tert (immortalized human fibroblasts). Does anybody have any experience with these cells? Is there anything special about preparing cell extracts for those proteins?
I have faced kinda similar problem when i started looking ATM/ATR DNA repair pathway. I used gradient gel, wet transfer for 3 hrs or overnight transfer, probing with primary antibody for 12hrs at cold room and my problem solved. I agree that some time we need to activate the cell or induce the cell for injury for DNA damage response. However, which is not necessary for every cells. One final advice don't use glycerol during probing/blocking.Following
- Aminu Abba Yusuf added an answer:7Is Western blotting a reliable and clinically validated means of detecting BCR-Abl fusion protein in chronic myeloid leukaemia?
In the absence of quantitative real-time PCR as a means of detecting the oncoprotein BCR-Abl, can one reliably use Western blotting (WB) in the diagnosis of CML? Since BCR-abl is a 210kDa protein when the major break point cluster region is involved, and 190kD when the minor region is involved, it is therefore theoretically possible to detect these proteins using WB. My question is, is WB validated for such purpose? In practice, WB can as well be quantitative, and can therefore be used for both diagnosis and measurement of minimal residual disease for BCR-Abl and similar fusion proteins such as PML-RARA, etc. We are contemplating introducing an assay for BCR-abl in our hospital but do not have qPCR facility, so I was wondering if we can reliably use Western since it appears cheaper (at least for a resource-poor setting like ours), and less technically demanding than a qPCR. I would appreciate any reference, please. Thanks.
Thank you everyone! I now understand that Western isn't a great way to detect or quantify these fusion proteins in diagnostic specimens, and that q PCR or flow cytometry remain the best way to go. Your answers have been both great and quite helpful. Thanks Jester,Following
- Surendar Jayagopi added an answer:3What is the procedure to measure insulin stimulated Akt phosphorylation in cerebellar neurons?
I am planning to measure Phospho-Akt/Akt ratio in cerebellar neuronal cultures. Do I have to stimulate my cells with insulin before harvesting cells for western blot. If so, what is the time and concentration of insulin to be treated?
Thank you so much Sebastián and Francesco for your useful inputsFollowing
- José Navarrete-Perea added an answer:5Western blot: why would a primary polyclonal antibody raised in a rabbit, probed with a 2ndary goat anti-rabbit antibody produce weird bands?
I have protein extracted from cells transduced with a lentivirus expressing ZsGreen. The cells go green beautifully, and at least 75-90% are green. I am trying to demonstrate that the zsgreen protein is there via western blot analysis, and have purchased a rabbit polyclonal antibody to Zsgreen from the same company that provided the lentiviral backbone. Running the protein on a WB, this afternoon, to optimise it I ran 4 different primary antibody concentrations (1:1000 (recommended by clontech), 1:10000, 1:50000 and 1:100000), plus a lane with no primary antibody as a negative control. Then washed as per normal in PBS/Tween, and probed with a Goat Anti Rabbit Secondary/HRP that I have used successfully on other rabbit primary antibodies from clontech. I did it at 1:5000, and incubated with my negative control lane, and when i exposed the blot, all my lanes had the same pattern in (like a ladder) including my negative control lane that was only treated with the secondary. So clearly my secondary has bound non-specifically in each lane.
I am lost as to why this may have happened given that my primary is rabbit, and my secondary goat anti-rabbit, and my cells are expressing ZsGreen (or at least they are green and the controls not). I am sure I added the right antibodies too. I used the primary in 5% milk in PBS/Tween, and this works for my other rabbit antibodies from clontech. Help!
Could be related to blocking, usually 5% of skim milk in PBS or 1% BSA is enough. incubate the first antibody in the blocking solution, after that wash the membrane (Tween 20 0.1% in PBS) and incubate the secondary antobody in the washing solution.
- Samuel Coulbourn Flores added an answer:48What could be the reason protein is expressed in an uninduced E. coli culture as well?I am expressing a Leishmania protein of about 35kDa in E. coli. My problem is that I always see the same band in uninduced bacteria control lysate which is quite similar to the induced culture (1mM IPTG) and the protein does not express well (low expression level). I tried different E. coli strains such as M15, BL21, TG1, XL1 blue with pQE41 vector, and got the same. My insert starts with ATG, does start codon may interrupt protein expression? Any explanations or suggestions?
Thanks guys! Never realized social media could be so useful. To me it was all about pictures of kittens.Following
- Michael McAlpine added an answer:3If two proteins run in sds page with different molecular weights, if both show bands with same intensity, does it mean they have same concentration?
If two proteins run in sds page with different molecular weights(having difference of 60 KDa),if both shows bands with same intensity,does it mean they have same concentration?
Coomassie blue will bind to all proteins regardless of size which makes it a good way of measuring the total amount of protein in a sample. However, because of this it is not possible to say that an observed band contains only your protein of interest. For example, if your protein is ~60kD it is possible that the band you are seeing around 60kD is not only the protein you are interested in but also other proteins in your sample which could be of similar size (i.e. 59/60/61 kD). Assuming you loaded samples containing several different proteins it means that you can't say for certain that the intensity/concentrations of the proteins are equivalent.
However, if you did load a sample that only had your two proteins of interest and there were two bands of equal intensity you could theoretically say they have equal concentration. But in practice coomassie dye will detect some proteins better than other due to their biochemistry, and some differences in intensity may not be detected because of the sensitivity of the dye. So even then I would say that you can't for certain say the concentrations are the same.Following
- Dacie R. Bridge added an answer:3Should running buffer get hot during a Western Blot?
I ran a gel yesterday, and for some reason the running buffer was extremely hot after one and a half hours of running the gel. I feel like this shouldn't be allowed to happen but am I right?
I was using the Bio-Rad Mini PROTEAN tetra system tank for my blot. In this system, you have a device that holds two gels in the same system. This allows you to fill the space in-between the two gels and cover the electrode in running buffer. However I was only running one gel, so I filled the whole tank with running buffer. I did this as I like to ensure my wells in my gel are submerged in running buffer before adding anything to them. However, I believe this may be the reason for heating up, I'm assuming because it increased the resistance to much. The gel also took unusually longer to run-although it was a 12% gel (i am investigating an 18kDa protein), after running for 1hr45 minutes at a higher voltage it still had not reached the bottom of the gel. As the tank was getting too hot, I had to put it in an ice tray for the last 30 minutes, and chose to stop after 2 hours of total running to prevent overheating.
Despite my worries, I carried on with the transfer onto a nitrocellulose membrane, treated it with antibodies and still got the protein bands I expected. i was running a test at the time, as the last time I ran a blot with these samples I had excessive binding. I was testing with different antibody concentrations to check whether we needed more diluted antibodies, however my control, using the exact same conditions as last time, worked fine this time. I'm not sure if the heat could have reduced protein content which could lead to my problem with banding being "fixed" for now but just because of this.
Apologies for the long answer, but my question in principle is-is it ok for running buffer to heat up during electrophoresis, and what could be potential effects on a blot if it isn't?
It sounds like you set up the gel apparatus incorrectly. Even if you are only running one gel you still need to have the glass plates on the other side to create a separation between the buffer between the gels and the buffer in the outer chamber for the electrical current to run properly through the system. This can be remedied by placing empty glass plates on the other side.Following
- Boaz Barak added an answer:6Any recommendations for a good antibody against BDNF?
I'm looking for a good antibody anti BDNF to use in Western blot and IHC to characterize BDNF in the mouse brain.
Preferably, an antibody that works good for both applications would be better.
Thanks a lot Sina! This is very informative.Following
- Tiffany Bellomo added an answer:1Does anyone have experience with blue transfer buffer?I've been doing western blot the last two years and I have never had a problem like this. When the transference is finished, the transfer buffer has turned blue. It seems that the sample buffer gives this colour. However, the Whatman paper and the sponge are completely white, without a trace of bromophenol and I am totally sure that the sandwich is placed in the correct position. Can anybody give me the reason for this?
We have had the same problem! We have no idea what to do.Following
- CHANDRA SHEKHAR DASARI added an answer:5Why is untransfected control in my western blot also showing a band?I carried out transfection in HEK293 cells with a plasmid containing myc tagged ORF of a gene. On performing western blot, I used a primary antibody against the myc tag. Problem is that both my untransfected and transfected cell lysates are showing a band. Why is my negative control showing a band?
Aditi, I found the same reason. And I solved it by doing ALP instead of HRP. And repeated with previous samples also. Surprisingly I did not observe extra band in ALP. All the best.Following
- Jinn-Li Wang added an answer:9How do you detect Human papillomavirus (HPV) 16 E6 in western blotting?
In my research, I need to detect HPV 16E6 protein in western blotting. However, I always failed to detect this protein in caski cell line in western blotting. The primary antibody for HPV 16E6 is the antibody of Santa-crus technology. I have tried more than 50 times using different lysis methods, including herpes lysis buffer (NP-40), Tris lysis buffere (N-40) and Triton lysis buffer. No band found in western blotting. Anybody can help me to resolve this problem? Thank you very much.
Thank you very much
I will try again,Following
- Bedair I Dewidar added an answer:10Do you have a good protocol for Western Blot with secreted proteins?I would like to perform Western Blot with proteins which are secreted.
One of them is 45 kDa and the other is 30 kDa.
In one condition, I have BSA added to the cell culture medium, together with N2 and B27 supplements.
In the other condition, cells are grown in 10% FBS.
I think the best method is to concenterate the protein first using a suitable filter. Then, you can add your loading buffer and run western blot like normal. Usually, this method doesnot have internal control (which is a big disadvantage). That is why, for released protein I usually use ELISA method, but it is more expensive.
- Sreeja Karathedath added an answer:20Why protein from SDS PAGE is not transferred onto nitrocellulose membrane?While I'm transferring the protein from SDS gel to Nitrocellulose membrane only marker is getting transferred but not protein. I used 100v for 1hr to transfer. Can we reuse a transfer buffer? Is there any problem in composition of gel buffers I used? I loaded 60microliters in a well (12microliter of Sample Solublizing Buffer and 30microliter of protein (20microg/ml) rest is PBS)
Composition of buffer I used:
Transfer buffer composition- Tris 3.03g; Glycine-14.41g; methanol 200ml - make up to 1000ml
5xRunning gel buffer - Tris-1.5g; Glycine-7.2g; SDS -0.5g - make up to 1000ml
I also need to clarify certain doubts regarding the same question Asiya Parvin addressed.
Which is the ideal voltage and time we can use to transfer 100kDa protein? Should we avoid methanol and methanol from the transfer buffer to get an ideal transfer in this case? Can we reuse the transfer buffer? As the isoelectric point of the protein is 4.7 should we adjust the transfer buffer pH to+2/-2 from this value to prevent any hindrances during transfer .Following
- Jean paul Delgado added an answer:13How can I extract as much protein as possible for western blot analysis on tiny tissue samples?
I need to extract proteins to perform western blot and my samples are kidney from medaka (4cm fish). The size of the kidney is thus very small and I would like to have your advice on how to get as much protein as possible.
Does anyone have experience on that?
Lysating the sample directly in loading buffer. Another option that works well is doing a normal lysis, then precipitating the protein of the sample with TCA, then resuspending the pellet with the volume of loading buffer you will use to load on the gel (20-50ul)
- Saikat Chakraborty added an answer:6Western blot band analysisI must compare the expression of a certain protein X in control mice, sick mice, and mice resistant to the desease. For each type of mice I have 5 protein samples. With each sample (15 in total) I have made western blots so now I have the bands for my X protein for each sample, and also their respective loading control, actin. With these bands I put the image in the program Image J and obtain the intensity of each band. Now I dont know how to proceed. In examples of how to analyze western blot bands they typically use 1 control and 1 experimental band, with which they normalize the experimental bands to the control and the actin band, but I have 5 samples for control, so I dont know how to analyse my data, should I just take a mean of each group of samples?
If I want to know how my protein expression is changing with drug treatment and compare with the Beta actin to target protein ration , what would be the best way to present the data?Following
- Terence P Herbert added an answer:3Persistent White Band at 50kDa in Western Blot with any antibody
I've been facing a problem finding a single white band at 50kDa in my western blot with any of my antibodies (c-fos, fosB, CREB, BNDF), in any animals, and any tissue. I don't know what kind of problem is. I've tried to reduce the exposure time, diluting primary and secondary antibodies, any ideas how to solve this?
- Christopher Malcolm Brown added an answer:8Have you also experienced smeared and fuzzy bands after WB when doing semi dry transfer?
We use the stainfree system from bio-rad. We use 4-15% TGX criterion stainfree gels and have recently observed membranes with fussy band safter transfer, pre-stained standards which have "ran" out as well as samples smeared out. After running the gel we made stainfree pictures of the gel, and the gels looked all fine. The problems we have observed are after the transfer. We use bio-rad turbo transfer, set at 2.5amp 25V 7 min (premade protocol from bio-rad).
It actually sometimes looks as if the gel has melted, and we have run samples 6 - 12 month ago with good results and now the same samples suddenly look bad. We do not prepare membranes for transfer ourselfes, we use readymade turbo transfer packs PVDF membranes also from bio rad, and have done so for 2 years with very good results. Has anyone here experienced anything like this?
Now we use wet transfer, using the same gels, preparing our own membranes (PVDF) and it looks very nice.
We are discussing if it could be the buffer in the ready made transfer pack, the membrane the turbo transfer mashine or perhaps the gels.
The two files reprecent the gel and membrane from the same experiment, stainfree pictures taken with a chemidoc camera ( bio rad ).
Recently we had a very similar issue to what Helle showed. Suddenly, everyone in the lab had issues with bands smearing, and even appearing to slide off of the PVDF. After a huge trouble-shooting blitz over 2 weeks, we eliminated every variable, including gel batch, methanol, transfer buffer, even blot transfer module. We did not expect PVDF to be the problem, but we narrowed it down to a new box of PVDF from BioRad that appeared at the time the problems started. Sure enough, side-by-side comparisons of this PVDF to another box of the same product from a colleague showed clearly that the new lot was the problem. Incidentally, the colleague who gave us the PVDF for our comparison shares the lab and uses all the same equipment and reagents, except for the PVDF. He has had great blots during the same period when we generated 25 smeary, streaked blots that we cannot use. (Obviously, the losses go beyond a roll of PVDF, and include the accompanying reagents, samples and worst of all, student and personnel time).
Rather than take this quality issue seriously, get ahead of it and contain/mitigate it, BioRad technical support has so far refused to acknowledge that it could possibly be the PVDF. Tech support insists it is the module, though the product failure occurred with all of our blot modules (but NOT with our new PVDF). Their logic would mean that all of our 6 modules suddenly began to fail at the same time; coincidentally, this would also be when we opened a new box of PVDF.Following
- Masood Sepehrimanesh added an answer:6Has anybody encountered this symptom on SDS-PAGE in which glycoproteins do not migrate to the resolving gel and remain in stacking gel?
Actually, I am working on Maillard Conjugated Products and in order to determine the molecular weight of glycoproteins formed through Maillard reaction, SDS-PAGE is used. I have been encountering some problems lately on SDSP-PAGE, in which glycoproteins do not migrate to the resolving gel. However, the pure protein and marker resolve clearly in resolving gel.
Here I have attached two images of two different gels. The first and second are 3%-10% and 4%-12%, respectively (these gels ran at 120V). In gel images, right to left, the first line is pure protein, and the 2th, 4th and 6th are glycoproteins.
I appreciate all your advice.
As other friends said: molecular weight of your protein is high and therefore, you have only one options:
Your gel percentage is too low for the molecular weight range of the protein sample after glycosylation. Use a higher percentage acrylamide gel (increase % T in resolving gel).Following
- Gabriel Osvaldo Gallo-Oller added an answer:7What will be the best protocol for transfer of separated high molecular weight proteins from SDS-PAGE gel to nitrocellulose membrane?
I am facing problem in the transfer of high molecular weight proteins to a nitrocellulose membrane from SDS-PAGE gel. Experimental target protein has a molecular mass of ~125kDa, and I tried different concentration of gels. I try both wet and semi-dry procedure but never achieve complete transfer of proteins. What will be the possible solution for this problem?Following
- Ingrid G. Haas added an answer:7What could cause a negative band in WB?
I'm blotting against STAT3 and pSTAT3Y710. My pSTAT3 antibody works great, but I observed a negative band (a white band that is lower intensity than background) at the predicted MW for STAT3 when blotting for total STAT3. What could be the cause of this, and more importantly, how do I fix it?
high amounts of antigen concentrated in a single band will bind high amounts of enzyme-linked antibodies that rapidly uses up the substrate thus causing a "negative" band at this position of the blot. Wash the blot, use fresh substrate and expose for a very short time period. Or repeat the blot using 10x lower amounts of antigen.
Best luck, IngridFollowing
- Cecilia D'Alessio added an answer:2With an ECM1 on a western blot, why is one band much lower than the rest?
I ran a WB looking at expression of ECM1 in several cell lines. The results were as expected, however for one of the cell lines the band was significantly lower (around 60-65 kDa) than for the other cell lines (all of which showed a band at around 85 kDa). I was using a Santa-Cruz ECM1 antibody, N-17. Has anyone else experienced anything similar, or know why this might be? I wondered if either this cell line was expressing a different isoform of the protein (ie ECM1b) or if the protein was less glycosylated?
Glycosylation may depend on the cell line. You could try digesting with PNGaseF to see if N-glycosylation is responsable for the difference that you see.
Another possiblity is that you have more proteolysis in a particular cell line tan in the others, or that in a particular cell extract there is more protein degradation due to less protease inhibitor...Following
- Andrew Hoffman added an answer:1Has anyone come across a substantial unidentified band in the protein isolated human umbilical cord exosomes?
With exosomes isolated from human umbilical cord cells I keep seeing the same high flourescence band in at the same molecular weight 58Kda on a 10% SDS-page gel. This is across different exosome preps which have been lysed in RIPA buffer. I use FBS which has been ultra-centrifuged to remove bovine exosomes but could the unidentified band from contaminants from this FBS?
Agree with Jason. We have performed HPLC on many ultracentrifuged samples of culture medium (FBS exosome depleted) and albumin is a prominent contaminant (~60 KD) necessitating multiple washings, or sucrose gradient, or elimination of contaminant proteins by size exclusion HPLC (although significant loss of exosomes depending on system).Following
- Amanda Fitz added an answer:32Western blotting transfer problemI am regularly using western blotting technique and often I get distorted (ladder and /or protein) bands after the western blot transfer, though there is no band distortion during the gel run.
I read that it could be due to air bubbles and inadequate uptake of buffer by the filter paper. I noticed no trapped air bubbles each time. I normally do 2 - 4 blots at one time and some of them show the distorted bands but not all of them.
I cant figure out what is the exact problem. Could you please suggest possible reasons and how to overcome to these problems?
Have you tried doing one blot at a time. I have noticed that though transfer apparatuses are made to do more than 1 blot they usually do 1 blot well and more than 1 poorly. Whenever I do two one of them always transfers incompletely. but as for your smears- do you set up the sandwich totally submerged? What about the temperature, do you transfer at 4 deg. lowering the temperature usually helps me.
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.