- Megha Sah added an answer:2How can I effectively release activated small G-Protein from heterologous cell pellet to carry out an activated protein profiling?
I am trying to study the activation profile of a small G-Protein upon treatment with various kinase inhibitors. To assess the activation profile of this G-protein we employ a protein pull down method using appropriate bait protein conjugated to glutathione agarose beads.
The protocol I follow is : Cells are harvested in lysis buffer( 50 mM Tris-HCl, pH 7.5, 100 mM NaCl, 2 m0.5 mlTM MgCl2, 0.1% SDS, 0.5% sodium deoxycholate, 1% Triton X-100, 10% glycerol, and HALT protease inhibitor cocktail), spun down at 10,000 rpm for 10 mins. The resultant lysate is incubated with the bait protein conjugated glutathione beads and the pellet is saved separately for analysis.
The problem is that after certain treatments, the g-protein is translocating to the membrane and is getting stuck in the pellet(as seen by immunoblotting). I have tried sonication and extended incubation time with lysis buffer but it doesn't help in releasing the protein from the membrane(as evidenced by the immunoblot)
What can I do differently to access this membrane bound protein to make it available for a pulldown assay? Any pointers will be greatly appreciated.
Thank you, Cristian. I'll let you know how it works out.Following
- Quang Nguyen added an answer:4How can I compare my infected human sample results from western blot versus ELISA?
I need to compare the results from western blotting and ELISA (human samples infected and uninfected with Schistosoma mansoni). How can I do? I would like references about this. Thanks
The conditions of your coated in ELISA and those loaded in the well in gel for Western blotting may be different. It also depends whether you are doing it in native or denatured western and the reagents used in ELISA, e.g. BSA vs. Superblock as diluent. Geberally, ELISA, I think, would allow you to calculate EC50 if true binding at certain titration of the primary antibody, but it is not very specific as in Western unless you are loading a lysate of sort. ELISA of course is more high throughput and less pain especially compared to native blue Western. You may also consider Luminex and SPR. Hope it helps.Following
- Wolfgang Nellen added an answer:25Is it possible that over-expression of gene leads to self degradation of its transcript?
I have made constitute over-expression construct and transformed into the plant. To check the level over-expression in different transgenic lines, transcript accumulation was checked using RT-PCR but none of the lines showing increase in the transcript level, rather all the lines were behaving similar to wild type and vector control lines. Further to rule out the efficacy of RT-PCR , western blotting was also carried out but same observation was also made at protein level. I know the fact that if transcript is not present then how protein will be formed. But my query is that apart from my construct is not working , is there is any possibility that because of some kind of self regulatory mechanism in plant which preventing the over-expression of the gene?
I guess what Daria means is not a fusion but just GFP or YFP in between your promoter and terminator (best could be to even leave the 5' and 3' UTR of your gene in and just exchange the coding sequence)Following
- Avishekh Gautam added an answer:3How can I get a good signal of Myc associated factor X (MAX) protein in Western blot?
I have been trying to overexpress c-Myc and MAX in HEK 293 cell either alone or by co-transfection. c-Myc is over-expressed with high signal. However MAX has very low signal. I tried to check the constitutive expression in breast cancer and colon cancer cell lines as well. c-myc is detected in some of the cell lines with very good signal. However I am not being able to achieve a clear band of MAX in Western blot. I use Goat anti Max ANTIBODY for Max , and also flag tag antibody while over-expressing in pdream vector, in HEK 293 cells.
I would be grateful for any suggestion. Please help me out. I forgot to mention I am working with human c-Myc and MAX.
Thank you Michal Jeremy for your response. Considering first suggestion, I have confirmed my DNA sequence and its fine.I have used two types of antibodies flag tag and also max specific. The flag tag is fine with other protein so I think that may not be a problem. May be I have to use other more sensitive reagents for detection as you have suggested in point no three.
I use lysis buffer Tx-100. I have been using 2%SDS and beta mercaptoethanol before I boil, then I load the sample. As you have mentioned I shall try to pas through the needles which I have never done.
Furthermore, I use Fugen 6 for transfection, but I have been using media with Penicillin plus streptomycin 1%. So I will rectify when I try next time.
Thank you for your suggestion and time.Following
- Oscar Marcelo Lazo added an answer:3How can I verify a decrease in endocytosis by Western-Blot investigation of a Pathway?
in a recent experiment I could show that upon a certain stimulus the endocytosis of cultivated cells decreases drastically. I did this using fluorescently labeled dextrans, but I would like to show that it really affects cellular metabolism. I thought that it would be a good idea to verify via western blot that a certain marker decreases or increases. However, my biochemical knowledge is too small to come up with a good idea;) So, here is my question: Has anybody a suggestion which marker I could use?
Thanks a lot!
Since you should want to distinguish only the fraction of a cargo that has been endocyted (not the protein in the plasma membrane nor along its synthesis route), I can think in two alternatives:
i) Biotinylation of surface proteins followed by separation of plasma membrane and intracellular vesicles by subcellular fractionation (centrifugation). Then you can calculate a ratio between surface-associated proteins and the intracellular membranes-associated ones. This ratio only can be altered by endocytosis.
ii) If you want to analyse a particular receptor (which endocytosis kinetics is well described, as EGFR or TfnR) you can immunostain surface proteins in live cells and after endocytosis to wash the remaining antibodies from the surface, as has been described in Chen Z-Y et al., 2005 (MBoC, 16:5761-5772). Although it was originally designed for microscopy, I guess the method may be easily adapted for being revealed by westernblot.
I hope you enjoy those experiments. ;)
DON'T FORGET YOUR POSITIVE AND NEGATIVE CONTROLS.Following
- Anna Morales added an answer:4Has anyone had difficulty western blotting mouse DAT?
I'm blotting striatal and cortex samples with a DAT1 antibody from Novus (NB300-254). At this point, I'm concerned I may be denaturing my samples for too long at too high a temperature (15 minutes at 95C). I've read samples can be denatured at lower temperatures such as 37C. My control, Actin, has been chemiluminescing during each attempt but DAT remains elusive. Thank you for your answers.
Madhur, thank you. We are loading 40 ug at this point and we've increased our DAT primary concentration to 1:750. We did these a while ago and still received less than satisfactory results (multiple bands and evidence that suggests DAT barely left the wells). So now we are trying increasing the concentration of SDS to 5% and 7.5%.
Has anyone tried increasing SDS concentrations before?Following
- Goutham Genomics added an answer:16Can anyone suggests a good method to reconcentrate proteins after isolation? Precipitation, Speedvac or Lyophilization?I need to reconcentrate proteins with a very low concentration which are dissolved in RIPA buffer to do a westernblot.
Hello every one as an add on kindly let me know ur experiences with protein concentrating filters when you are working with proteins present in bacterial culture supernatant . which could be the better option.?Following
- Leonardo Bianchi added an answer:5How can I be sure that my protein is on a western blot ?
I've been doing Western blots for months now to show a protein (Cyp17) that weight 56 kDa. I finally found a "specific" antibody but my protein appeared upper (just under le 75 kDa marker). For me it's not a problem because I know for sue from the litterature that this protein is going under some post-translational modifications that could rise its molecular weight but I guess that if i want to publish my blots I better come with something else. That's why i am asking. How do I prove its identity ?
I suppose you could just cut your band, strip the antibody, treat with trypsin and analyse peptides by mass spectrometry.
Attached you will find an article dealing with this, but sure you can obtain satisfactory results with other methods.
If you know what modification (glycosylation?) occurs, you could try a deglycosylation protocol (I used the EDEGLY kit from Sigma and it worked quite well) and run the treated and the control sample in parallel... the two bands should show at different moleculare weights.Following
- Eiji Kinoshita added an answer:43Accuracy of Phos-tag gelsIs anyone working with Phos-tag gels? I have just started out working with them and I understand that with such gels, the protein marker is not an accurate indication of the weight of the protein of interest. However, after blotting for my protein interest, it seems that the darkest bands seen are in the range of 25- 35kDa where in theory, it should have been seen at around 60 - 70kDa. Is this some non specific binding of the primary Ab or is there more to the phos tag gel that I do not know about.
Hi, Kayleigh, I think the Mn(II) procedure is also good as the first your choice.
In genral, the separation in the Zn(II) procedure is better.
Regarding STAT1, we do not have an experience by using the Mn(II) procedure.
If you have any troubles, please contact me.
- Closed account added an answer:4Has anyone an idea how to fix a ponceau staining?
I would like to do a dot-blot with lysates with an additional membrane for comparison of protein content (aim is to do it in a quick and "dirty" approach without proper protein concentration determination beforehand).
Has anyone ever tried fixing the ponceaustaining permanently or an alternative to ponceau that can be fixed?
(also found this now: http://www.bio-rad.com/de-at/product/total-protein-blot-stains), nice.Following
- Sanath Kumar Janaka added an answer:6What are the important considerations in detecting large M.W. proteins using SDS-PAGE western blot?
I am intersted in detecting the p-mTOR protein, which is around 290 kDa. Can anyone give me some tips on how to efficiently blot for proteins in this size-range?
At the moment, I am using 4-20% precast gels using the TGX buffer system (Bio-rad) and loading ~15 ug total protein. Importantly, I'm also doing my transfers with the Trans-turbo system from Bio-Rad.
Additional points to consider during transfer---- If you are using Methanol in the buffer for transfer of your proteins, I would suggest using between 0 to 10% methanol. That increases the transfer efficiency. Standard liquid immersion transfer systems work better for large protein transfers, rather than semi-dry systems.
If you do find a good condition that works for you, please update this forum with your conditions.
- Seungheon Shin added an answer:7Can anyone help me to solve uneven distribution of protein sample lanes?
I have no idea about this problem.
Sometimes my western blot samples are transferred well, but sometimes not.
This is always bothering me.
First image is the western blot result of my research protein, and second image is the western blot result of actin.
After Ponceau S staining, there were no air bubbles observed.
But after semi-dry transfer and antibody incubation, sometimes samples are not transferred equally.
I used 12% of SDS-PAGE gel and ran in 90V, 20min and 120V 1h 30min in 4`C.
Transfer was performed in 15V, 40min in 4`C and I used +methanol transfer buffer with SDS and nitrocellulose membrane.
Please save me from hell.
Thanks a lot guys!Following
- Rahul Das added an answer:9Any advice on the detection of low MW protein by western blot?
I am trying to detect insulin (~5KD) and other small ppetide fragemnts (about 5 KD also) from cell extracts using western blot with 16.5% Tris-Tricine gel supplimened with 6M urea. This method is not working well-(i) Mature insulin band (~5 KD) is wavy (ii) Blotting does not work immediately after transfer, I have to strip the blot to see bands. Can anyone help solving this problem or does anyone have a different protocol for detecting small proteins? Following are the conditions I use-
1. Gel- 30v Stacking, 120 V resolving, room temp.
2. Transfer- Tank transfer, 100V for 40 mins, HyBond PVDF membrane ( Methanol activated).
3, Western- Blocking with milk, Overnight incubation with primary and 2 hrs. with secondary.
I am aware that Elisa works great for detection of Insulin. But our antibody does not distinguish between pro and mature insulin, and I want to detect pro form as well as mature form.
Thanks in advance.
Thanks Marisa! I am using urea gel specifically for insulin due to its low MW. I tired using the Biorad gel too but it did not work. As you predicted, I do not use BME because insulin contains disulfide bonds. Should I try using BME/DTT?Following
- Paul Digard added an answer:1Can Sarkosyl solubilized proteins cause issues during Western Blot?
I have interacted multiple proteins using GST Pulldown Assay (one protein is His-tagged and the other is GST-tagged) and I usually do a Western Blot to confirm each interaction. I have had many positive results (including Western Blot). But I have been having trouble with 2 proteins that have both been solubilized using Sarkosyl. No bands show up in the Western Blot, not even the control. Does anyone know if Sarkosyl solubilization can cause antibody binding issues during Western Blot? If not, what else could be causing binding issues?
I used to routinely include sarkosyl in the initial GST solubilisation step of the bacteria - I forget the exact %, but it was from a published protocol - and it never affected downstream SDS-PAGE and blotting of the material.
- Go J Yoshida added an answer:1What is a solid method to separate nuclear and cytoplasmic proteins from lung tissue?
I've been using commercial kits to separate cytoplasmic and nuclear proteins. I noticed that many advise against using their kits for frozen tissue because the nuclear membrane can break and leak into the cytoplasmic portion. However, all my tissue have to be frozen because I bring them from another facility. Has anyone come across a good method to separate cytoplasmic and nuclear proteins from frozen tissues or account for the slight leakage? They have only been frozen and thawed once before analysis via western blot.
Thank you -- Greg
Given that your available sample is quite small in amount but stored in fresh conditions, I strongly recommend you to use the NE-PER Nuclear and Cytoplasmic Extraction Reagent Kit to obtain both the cytoplasmic fraction protein and cytoplasmic fraction. (strictly speaking, which includes cellular membrane proteins) from thawed tissue.Following
- Ranu Pal added an answer:5How much lysis buffer should I use in order to extract cytosolic proteins for hemisphere homogenates?
I'm preparing to run Western blot analyses on rat tissue and have read some estimates where ~5mg of tissue is used with 300uL of lysis buffer. However, a hemisphere of a 4-month old rat weighs about 0.9g and therefore would require >30mL of buffer which is out of the question.
I have read about different lysis buffers besides RIPA that have been used and have an aliquot from a few fellow researchers (Tris-Triton buffer); again, I'm just not sure how much of this to use for my tissue in order to locate good amounts of my cytosolic proteins of interest (SOD-1 in particular).
I usually use extraction/homogenization buffer (containing higher concentration of sucrose) at 1:9 ratio. After homogenization I usually save aliquots for BCA assay to know the protein concentration. The rest of the aliquots are stored at -80 C. Pls do not forget to add protease and phosphatase inhibitors in the buffer at the last moment. Hope it helps. RanuFollowing
- Sean Patterson added an answer:3How can I do western blot with the very acidic sample?
I have to do immunoblot with gastric fluid which are very acidic.
I've tried without any modificiation first, then...
- the gastric fluid was not mixed with 5x loading buffer,
- some particle was made after mixing with loading buffer,
- and finally, the blot was messy.
(c.f. The volume of gastric fluid is just around 50~300 ul, and the target molecule to be measured is not much to detect.)
I've searched for some specific protocol but couldn't find a good one.
If you have seen any paper/protocol for immunoblot with acidic sample, please let me know.
- Thank you.
An alternative to the above would be a CHCl3/MeOH precipitation of your samples (10 volumes, 1:1). This would both remove the acid and concentrate your samples to help with detection. A warning, though, some proteins are soluble in acidic organic solvents.Following
- Paul Rutland added an answer:7Why do I get more signals on one side of the membrane than the other side for Western blot?
I want to detect AIM2 (39kDa) expression in response to treatment in a cell line. I got something like the attached photo consistently in several repetitive experiments. The ECL signal intensity is much higher on the right that it even stains the ladders but not the ones on the left. It happened to my beta-actin (43 kDa) loading control as well but, curiously, when I switch to a smaller loading control, profilin (15 kDa), this effect is almost completely gone.
Initially I suspected that it was the transfer that went wrong. I tried semi-dry and dry transfer but still got the same problem. Does anyone know what was going wrong?
Beta actin: 1:2000, 1:5000, 1:10,000 still the same
all secondary 1:2000
if rotating the gel in the blotter gives the same result I still feel that the problem can be in the electrophoretic separation.something like the gel temperature rising and some leakage happening as the sample runs into the gel.WHen you take the gel off the e/f rig ,or even while running, do the intensities of the loading dye bands look the same intensity or do they mimic the transfer intensities?.If the latter then you have some form of leak in the system I think.Following
- Ana Maria Calderon added an answer:8Any suggestions with an unusual western blot issue?
We have come across a strange issue with a western blot optimisation.
We are probing for Akt (p). We keep (done multiple times, because I thought it was operator error) getting completely blank membrane, no ladder even! We are using tris glycine gels, magic marker on the gels.
The confusing part is that we have also run the gel, blotted then split the membrane to run two different primaries (same secondary and ECL reagents). The results of the other primary look good, we can see the ladder etc, but the pAkt has nothing, not even ladder. We have even tried overnight low voltage blotting in case we aren’t getting complete protein transfer, although it doesn’t explain why the ladder is missing in the pAkt blot and not the other. We can try increasing the protein loading, but still dosent explain the missing ladder.
This is especially confusing as my technician had this assay working (using Tris acetate) gels a couple of years ago. …and yes we have bought fresh antibody and tried old and new.
You can try your first antibody activity by dot blots or double immunodifussion in agarose or agar.Following
- Elena Guillén added an answer:15Can you help me with unexpected band in western-blot?
Hello and thank you,
I used Allprotect Tissue Reagent to stabilize human kidney samples. After some months I extracted DNA, RNA and protein with the AllPrep DNA/RNA/Protein Mini Kit from Qiagen. Now, I have started to work with the protein extracted with the kit (precipitation) and solubilized with 8M urea. I tested the proteins with a b-tubulin antibody and they work perfectly well, but when I started to do the western blot with other different antibodies, I found an unexpected band at 100 kDa aproximately in all the blots. The only difference between the b-tubulin antibody and the rest of antibodies is that the first one was incubated with 5% milk and the rest with BSA.
Do you know if the protein extraction method can have something to do? I have tried with six antibodies (from rabbit and mouse) and always is the same. Do you think that the BSA can interfere anyway? Tomorrow I am going to test it, but I would know if you have some similar data. Thank you.
Thank you for your advice. I have contacted the manufacturers and I'm waiting for their answer. Anyway I've repeated the same blot I showed you but incubating the antibody with milk and it seemed to work (Signal is OK but a bit weak, but there isn't the 100 kDa band). Now, I want to check the others and add some changes to improve the signal.
- Annemieke Ten Bokum added an answer:2Can anyone help with an antibody/gene to "quantify" macrophages in tissue lysate?I have tissue lysates only and would like to check how the population of macrophages changes over time after an inflammatory stimulus. The problem I am facing, is that all the markers commonly used (e.g. F4/80) change expression during inflammation and would hence inappropriate markers for my purpose. Does anyone know of a protein/gene (I have mRNA as well) that is expressed in macrophages only and does not respond to inflammation?Following
- Brittany Wingham added an answer:10Has anyone had issues with western blot samples seeming to exclude ECL reagent?
I am interested in confirming whether a targeting peptide we have developed is successful in binding its target receptor protein. To do this, I have been running pull down assays to detect expression of the surface receptor.
I have been running two western blots in parallel. In the positive control pull down, I am using AG labelled magnetic beads, incubated for 1hr with a monoclonal specific antibody, followed by 1hr with the cell lysate. This western blot appears fine.
In the second, I am using streptavidin labelled beads, incubated with the targeting peptide for 1hr followed by the cell lysate. This blot came out quite dirty, and was as though in a negative print; the background was dark, and samples appeared white.
I washed off the ECL incase it had over developed and re-imaged - at this point, the samples did not show up, although the ECL and colorimetric ladders still appeared fine.
I just wondered whether anyone else had had this problem, and what the reason could be?
Thanks very much for any advice!
Incase anyone else comes across this issue - it seems that it was a combination of two problems. First, I decreased the amount of protein loaded. (I had been loading high concentrations, as I was working will a streptavidin bead bound pull down of a protein with only low expression to begin with. However, the streptavidin dissociated from the bead kit was leading to the band excluding dyes. Problem one solved. The second issue appeared to be my blocking method. I changed from 1hr at room temp, to 1.5 in the fridge, and this resolved the issue of the remaining membrane appearing too dark.
Thanks for all your input!
- Robert Williams added an answer:7What is wrong with my western blot?
Anyone with good Western Blot trouble shooting skills? I am repeating a western that has ALREADY BEEN DONE successfully. And I have done it twice now and there is ZERO Protein on my blot. I can see the visible bands from the ladder, but thats not developing either. I tried a different developer, nothing. I used ponceau today and there is no protein on the blot.
I use the TGX Turbo transfer system so the program is set exactly the same every time so I didn't over transfer or under transfer.
I am processing the lysates exactly the same...
Does anyone have ANY advice?
I spent a good part of my PhD perfecting my WBs as I had similar problems like yours!
A few tricks I learnt is:
- Make sure there are no air bubbles between the gel and blotting paper
- Coommassie stain your gel afterwards (as said above) to see if the protein is still in the gel
-is there a possibility your protein has/is degraded/ing? Is it running through an SDS PAGE normally? This happened to me and majorly effected my transfer...
- Temperature overload when transferring?
- I know it sounds basic but do you have the blotting paper and gel the correct way so the current forces the protein to migrate towards the blotting paper?
- Maybe try PDVF or nitrocellulose (depending on what you used before)
I hope you sort your problem out! :)Following
- Selva Madre added an answer:8Is there a correlation between dot-blot technique and western-blot?
I have used an antibody (polyclonal rabbit) against an specific type of my protein (an ubiquitinated variant) in a WB. I`ve seen the band, which is alright (molecular weight checked with marker and a different antibody that recognizes the wt-protein and uni-protein). It works really fine and it is detected with the Licor-odyssey system.
But when I did the dot-blot, I can no longer see a difference between my negative control and my positive control. (Negative means that no signal is expected) What I get is the same amount of signal (according to image studio lite) in the positive and in the negative controls which is really above zero.
Thank you I solve this issue using less primary antibody and lower detergent concentration.Following
- Lisa Corcoran added an answer:6Can I use gelatin coating plates for attachment my primary cell?
I am working on primary cell culture (neuron cell) in gelatin coating plates and protein expression. I want to know can I use gelatin coating plates for attachment my primary cell and then check the protein expression of them with western blotting assay? Do gelatin effect on western blotting respond because of the protein structure of gelatin?
Thank you so much in advance.
I agree with the use of 1% gelatin for C2C12 cells. I used Poly D- Lysine for my neuronal cell culture and collagen for HepG2 cancer cell line. I have done ELISA with the HepG2 and found not trouble with the collagen. Good luck.Following
- Muralidhar Katti added an answer:5What is causing the black edges on my membrane and how do you improve band definition for a Western Blot?
I have been doing my first ever western blots and I am trying to replicate the results from a fellow student however my bands are not as defined as hers and I am getting strange black patches around the edge of the membrane. I am following the same protocol as her however my results are not as clear as hers.
The running of the gel seems to go fine as well as the transfer. I stain the membrane with Ponceau after the transfer and it looks good. I also Coomassie the gel and I cannot find any bands on the gel.
During the immunostaining I do not think the membrane has dried out at any stage and I always use a shaker when incubating. My primary concentration is 1/100 and the secondary is 1/10000 (I know that they are compatible).
I'm using 10% precast gels, my protein concentration is 10microgram/well, TBST (0.1% Tween 20) for my washes (I do three washes for 10min each, for each washing step), and my blocking buffer is 10% hydrolysed casein and 1% bovine serum albumin.
I have attached an example of one of the western blot results. The first lane is the ladder (unstained) and the next two lanes are my protein (I am expecting to get multiple bands for the protein).
Do you have any idea of how I can improve my western blot to get a clearer image or what I am doing wrong?
Reasons for not getting precise bands may be chewing of your proteins. Hence include protease inhibitors cocktail while making antigen preparations. Use fresh aliquot of antigen preparation every time. As suggested earlier by Nathan Rout-Pitt, you have to be clean while handling membrane while transfer and developing immune blot reactions.Following
- Sophie Briggs added an answer:18How can I correct unequal loading in Western Blots?
I am currently doing western blots with BTIC cells (Brain tumour initiating cells) which grow as neurospheres. I have done western blots before (with monolayer BTIC's) and had no issues whatsoever. However, I cannot seem to get equal loading on any westerns at the moment. I have changed pretty much everything about my technique bit by bit and still no luck. I was just wondering if anyone has any suggestions as I am at a compete end now and not sure what to try.
So you have a rough idea of my technique, I wash my pellets in PBS, lyse and sonicate, spin down and measure protein using bradford assay (biorad). Then add 40ug protein with ddH20 and 4xSDS loading dye (40uL total), spin breifly, boil for 5 mins at 95 degrees, spin breifly again and load onto my gel.
Any help would be greatly appreciated. Thanks.
Variations are quite substantial, often more than 50% difference.Following
- Ellen R Weiss added an answer:11How can I get phospho-protein signal in Western Blot?
I am trying to find some phospho protein i.e. Aurora, Btk, Bcl2 etc in WB.
Even if I get very strong total protein signal but I failed to see any phospho protein signal!!!!
Is there any special techniques I should adopt to get phospho protein signal in Western Blot?
It will be great relief for me if you people share your experience.
I agree about not using milk for transfer and using a phosphatase inhibitor cocktail. We also sometimes put the inhibitors in the SDS-Laemmli buffer (2X), heat it and solubilize the cells directly in the hot buffer. You have to leave the dyes and the beta-mercapto out of the Laemmli buffer if you want to quantify the protein levels in your samples before running the gel. You add them just before you run.Following
- Susan K Morton added an answer:99+Western blotting - using BSA or milk? And TBS-T or PBS-T?I want to detect phospho-proteins as well as full-proteins. Does anybody have experience with using BSA instead of milk for blocking and using TBS-T instead of PBS-T?
I block with 2.5% skim milk in PBS, apply primary and secondaries in PSBT (0.05% Tween), wash in TBST (0.05%) and have been using Phospho-AKT antibodies. I have had the antibody work fine, but the other day we ran out of PBS and so I instead used TBST (0.05%) to incubate my primary and secondaries. Suddenly the antibody no longer works AND some of the non-specific extra bands have sort of changed their profiles despite the same samples being run. Has anyone found that TBS inhibits their western blots? The TBS is a couple of years old (one of those 5L cubes of 20x), would this likely affect chemistry/concentrations etc?
If its not a transfer issue, its something else! Western blots do my head in!!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.