- Rakesh Kumar added an answer:2Does the anti-TAP antibody detect CBP tagged protein (Calmodulin binding peptide)in Western Blotting?
I am using a fission yeast strain which has a Calmodulin binding peptide(CBP) tagged protein.I want to check whether this CBP tagged protein interacts with my protein of interest by CoIP.
I plan to IP my protein of interest using rabbit polyclonal sera followed by western with anti-TAP antibody to detect whether the CBP tagged protein is associated.
Thanks a lot Dominique!Following
- Christian Lipfert added an answer:8Any advice on a protein purification problem with His tag protein and Ni -NTA column?
We are facing purification problem with a His-tag cloned gene. The lysis buffer is 50mM tris pH- 7 , 100 mM NaCl and 1mM PMSF. The Protein (27 kda dimer) is bound to Ni-NTA Matrix and then washed with increasing conc. of imidazole upto 100 mM and then eluted at 200 mM imidazole. Even after proper washing with 50 mM-100mM Imidazole, the final eluted protein shows a number of bands other than our expressed protein. i m using pre packed HiTrapIMAC column linked with FPLC. Expression system is Rosetta Blue as my protein contain amber stop codon. Post NI-NTA i have done GFC also, But in that case my protein is coming in void volume with all impurities. Also, yield of my protein is very less. for clarification i m attaching gel picture, the marked region is my protein of interest.
Thanks in advance.
Hi Sheenam, have you tried a linear gradient to check when the protein of interest eluates best? Then you can add a flat gradient nearby that point and swap to a step gradient.Following
- Bhukya Bhima added an answer:1Is there a low cost option to isolate human IL 1 Ra at high concentrations?
I performed routine centrifuge separation on 55cc whole blood/5 cc ACD, extracted PRP and PPP and contacted this mix with 2 gms Cellulose Acetate .50 micron beads(Avicel). After incubation, I have been unable to isolate/separate the resultant concentrate (anticipated average 15 cc) via syringe filtration (.22 micron 25 mm cellulose acetate syringe filter) or vacuum filtration (Corning .45 micron vacuum filter tube). Is the only way centrifuge filtration, and if so what is the most reasonably priced but effective approach to take?
As such there is no low cost method available till nowFollowing
- Venkata Vepachedu added an answer:19Why bacterial cells were not sonicating properly after a long duration of sonication?I have used BL21 DE3 for expression. After overnight induction at 16°C, I have harvested the cells and sonicated at 40% amplitude for 30sec on/off for several cycles, but still my cells not lysing properly.
Lysis buffer: 50mM Tris(pH 7.40),150mM NaCl,Triton X-100 1%,Lysozme and Dnase (50 mg/ml stock each),10mM PMSF, 5mM DTT,2% Glycerol.
Can anyone suggest the reason why it is not sonicating properly?
You can confirm cell lysis by two ways:
1. The suspension turns transparent. The turbidity goes down. If you don't add sufficient DNaseI it also turns dense due to the release of genomic DNA.
2. You can observe the sonicated cell suspension under microscope and guess the % of cell lysis.
Another factor that affects cell lysis due to sonication also could be the stage of cell. Stationery phase cells need harsher conditions than log phase cells.Following
- Xia Li added an answer:9Poor affinity of Fc domain for Protein A columnAnyone have experience with poor binding of Fc tagged proteins to protein A columns? We have an IgG1 Fc tagged protein that is not binding at all to a protein A column. I am thinking about trying a protein G column, but are all protein A columns roughly equivalent?
My Fc domain is a mouse IgG2a domain. It really binds weakly to protein A and protein G. I use the new rProtein A fastflow sepharose from GE health, I collects each fraction of every purification step. Every time there are nearly 20% protein which can bind to the beads. I tried many different methods, and I added iodoacetamide and DTT to the condition media to reduce disulfide bond. But still no help to the binding. Do you have any idea about it? Thank you!Following
- Sarang Mahajan added an answer:7Can someone suggest a way to elute biotinylated proteins from avidin agarose beads?
Generally in all the elution protocols, the proteins are eluted in very harsh conditions. E.g Boiling in SDS sample buffer, elution by using Guanidine HCl. This creates a lot of avidin background when run on a gel. I have to submit my sample for LC MS. In order to do that I need to reduce the noise. Can anyone suggest me any method that can reduce the background noise?
I am sharing the protocol that worked perfectly for me
1. Resuspend the samples in 1X SDS buffer.
2. Boil the samples for 5 minutes.
3. Centrifuge the tubes and load the supernatant.
4. Stain the gel with coomassie brilliant blue or Silver stainFollowing
- Astor Mignone added an answer:4Can you help me with my hydrophobic interaction chromatography setup?
I am purifying my protein without any tag. The first step of purification is ion exchange chromatography and the second step is hydrophobic interaction chromatography. I was wondering if I need to buffer exchange my protein before the second step, since the concentration of NaCl into my sample will be 1M at the end of the chromatography. Otherwise, can I leave the NaCl into the sample and only add 1M Ammonium Sulfate salt to the sample before hydrophobic interaction chromatography?
Thank you for your help,
You can try to change the elution salt of the ion exchange. instead using NaCl try ammonium sulfate and then, if neesary, add some more to the eluted material in order to ensure a proper binding to the HICFollowing
- Manyando Simbotwe added an answer:3Why are my Western Blot antibodies binding to some lanes and not others?
Hi Everyone! Thank you for your help in advance.
I am working on purifying a few different proteins from both Ecoli and mammalian cells and am having the hardest time actually detecting them via Western Blot. Initially, I used an anti-V5 antibody from Thermo and it detected every single protein possible, so much so that it highly resembled the coomassie stain. After a bit of trouble shooting I got a refund and moved onto an Anti His Antibody instead (constructs have both tags).
As shown in the coomassie stain below, I am very sure that at least the second protein (boxed in red), if not 3 out of the 4 proteins I transformed, was expressed in all 3 lanes for my Ecoli transformations (the soluble fraction, the pellet, and the elution). However, an anti-His Western Blot seems to only be detecting the pellet lanes. 12 ug of protein was loaded in 13 uL into every well on both the coomassie gel and the western blot gel so I am not sure why the discrepancy between the different lanes is present.
I then tested if the concentration of the elution might have been too high by performing a 5 fold dilution on two proteins: the elution lane of the protein that was boxed in red in the coomassie stain and another purified protein that also had a His Tag as a positive control. Within 4 lanes I diluted the proteins from 12 ug--> 0.096 ug. After developing for 900 seconds, the most I could see was a single band of my original elution at the original concentration (see picture entitled WB troubleshooting).
My questions are:
1. Why would the same exact dilution of this antibody see the same exact sample at the same exact dilution on the troubleshooting blot but not the original blot.
2. Why would the antibody then not detect the second protein on the WB troubleshooting blot?
3. Should I just toss the antibody as trash or am I possibly messing up a part of the procedure?
4. Where there is a will, is there really a way?
Please note that this is using Anti-His unconjugated clone #RM146 from Millipore Lot QVP15. 03209. Cat: 04-1664. 1:5,000 dilution in TBST with 4% milk (found to be the best concentration by another lab) as a primary antibody and ECL anti Rabbit IgG HRP #NA9340V from GE Healthcare, 1:20,000 in TBST, as a secondary. I incubated overnight at 4 C in blocking buffer and 1 hour at room temp for both antibodies with 3 rounds of 10 minute TBST washes in between. I developed the blot using the Thermo ECL High Sensitivity Kit.
Any and all suggestions are needed and appreciated!
Looking at the expression profile of your protein, maybe your protein was over expressed hence, you predominantly detected it in the pellet fraction.
WB is more reliable. Try to look at the raw data, i.e unedited WB and see if you cannot detect any bands in the other fractions.
For Q1, how about reprobing?Following
- Lisa R. Keyes asked a question:NewWhat is the best kit or assay to detect complement factor H function?
I am looking for a kit, or well described assay to determine the functionality of complement factor H. I have seen brief protocols involving incubating different amounts of factor H with Facto I and C3b on ice, then incubating at 37*C, then running on SDS-PAGE. Is there a kit that has the Factor I and C3b available? Or do I need to purchase everything separately? If there is no kit for this, do you know of a well cited protocol anywhere?Following
- Rosa María Martínez-Espinosa added an answer:12After dialysis, why enzyme activity decreases significantly?
Once I have enzyme fractions from the Ni column, I usually run the SDS-PAGE gel, combine the fractions, and do dialysis to get rid of imidazole. However, when I purify mutant enzyme, enzyme loses its activity significantly after dialysis compared to activity before dialysis. Is there a reason for this?
Therefore, I am doing buffer exchange using PD-10 column instead of dialysis, and not sure if this way makes a difference.
Yes, that is. The mutant could be less stable than the native enzyme. This fact should be tested before dialysis performance.Following
- Gopalakrishnan Chandrasekaran added an answer:2Which set of primers is best for verification of gDNA contamination- Exon_Intron Boundary primers or Exon-Exon boundary primers?
Which set of primers is best for verification of gDNA contamination- Exon_Intron Boundary primers or Exon-Exon boundary primers?
- Manjunath Chavadi added an answer:4Which is the best way to detect low Kda proteins ?
I am expressing recombinant protein, which is 7 and 14 Kda.Due to low molecular weight, i have coudnt able to distinguish the expression of the protein on SDS page, i have tried with different percentage of resolving buffer but the results were the same couldnt find much difference, so kindly suggest the solution for the above query
Thank you all .............Following
- Becky Leifer added an answer:7Why is there differential detection of His tag with Tetra-, Penta- or 6xHis tag antibodies?
I have several his-tagged recombinant, purified proteins that are being detected in varying degrees by differentially his-tagged antibodies. I understand that sometimes part of the tag can be hidden, so that a 6xHis-tag antibody may not be able to pick up a tag where only 4His residues are available. What I am unclear on is why (this happened yesterday) a 6xHis-tag antibody would pick up a 6xHis tag well, but a penta-His tag antibody or tetra-His tag antibody used to detect the same protein would give no signal?? All experiments are done under non-denaturing conditions so proteins are in their native state. Thanks!
Update: I wasn't able to track down the epitope that these antibodies are raised against. It may be proprietary information for the companies. I am now trying to understand how the tag can be partially "hidden". Is this due to some ionic interaction between the tag residues and protein residues? It seems to me that the tag itself should not really have tertiary structure, and should be exposed to the solvent for the most part. When looking at protein structure in pdb or pymol, I wonder if I should be looking at the water accessible regions for portions where the tag may not be water accessible, or some other category like a space filling model. Can anyone comment?Following
- Misty Wakler added an answer:8Why is the expression of protein not consistent?
I have cloned gene in pRSETB vector and expressed in BL21(DE3) host. After fresh transformation protein is expressed. However, there are following difficulties:
1) Even only one week old glycerol stock from same colony does not express the protein.
2) Protein is expressed at 20 degrees C but not at 37 degree C.
3) Expression is not consistent.
pRSET A, B, & C Thermo Fisher: Transformation protocol for Top10F' and BL21(DE3) introduction the pRSET vectors are pUC-derived expression. Expressed corectly and contain the N
First screen: Express a protein in BL21 (DE3) from modified pET-vectors with host strain BL21(DE3) Protein expression is induced byFollowing
- Geoff Margison added an answer:10What percentage of glycerol do you use to store proteins at -80 degrees?
I have seen some conflicting advice on this topic. I believe that for storage of proteins at lower temperatures, such as -20 degrees, 50% glycerol is required, but when flash-freezing a protein preparation in ethanol-dry ice for storage at -80 degrees, is that high of a glycerol concentration needed? Any advice on this topic is appreciated. Thank you.
Thanks for all this exchange - especially Amanda and Nicholas. Last week I had an unexpected (= Kurt's experience!) minus 20 protein precipitation problem (expressed from a his-tagging vector I hadn't used previously), so this is all very useful. A fresh prep didnt precipitate in 50% glycerol at minus 20, so I'll test what happens at minus 80! [Note: I don't have regular access to dry ice or liquid nitrogen]Following
- Emmanuel Planel added an answer:3How to purify tau from AD brain?
I'm preparing a tau inoculum from AD brain however I keep getting a contaminated (haem contaminated?) red pellet after my 200k centrifugation. Can anyone suggest a way around this?
It depends what kind of tau you want to isolate from the AD brains. Is it insoluble or soluble tau? The gold standard to isolate insoluble tau from AD patients is described in the seminal paper of Greenberg & Davies in 1990:
We also discussed different methods in a paper from our lab:
But I think that you want to isolate tau from AD brains to inoculate in mice. Then, it is maybe better to follow the protocol from Goedert's lab:
As for the reddish brown color in the pellet, it is hemoglobin, which also precipitates in the insoluble fraction.Following
- Tomáš Hluska added an answer:5Can anyone help with proteasomal β5 subunit isolation ?
I am in a fix... One of the reviewer asked us to prove that disassociated subunit (namely beta 5) of human 20S proteasome is catalytic active. Any suggestion will be helpful. All that comes to my mind is use a mild detergent treated 20S proteasome and further purify (affinity chromatography?) beta5 subunit. please help
I didn't fully read this paper, but I got the impression they claim that b5 supports expression of catalyticaly active b1 and b2, don't they?
If detergents do not work (did you check that it is intact indeed?), how about some salts or maybe 2M urea? Are they covalently bound (disulfide bonds)?Following
- Vicente Bernal added an answer:5How do I overexpress vector PBAD+insert in wildtype E.coli K12 MG1655?
I am working the project to determine the phenotype of my gene in E.coli. Then, I inserted my vector into mutant E.coli K12-MG1655 ( in which the reference gene is knocked out ). I induced with arabinose 0.4% at 37 degree for 2 hours and 30 degree for 4 hours and then took the sample to run SDS-PAGE. The problem is that there is just slight differences between 2 groups: non-induction and induction. I checked the sequence of my vector as well as the existence of my vector in mutant E.coli K12 MG1655.
Have anyone had the experiences in overepression with wildtype bacteria? Could you please giving the instructions how to do thi better?
Thank you very much for your precious time,
Wild type E. coli strains are able to use L-arabinose as carbon source, which is the reason why you might use an ara deficient strain. E. coli LMG194 proposed by Dr. Duez is one option. If you are interested in K12 phenotypes, you could use the E. coli BW25113 strain or the mutants constructed from this strain (which is also ara deficient). The KO collection from Keio University has KO mutants for almost all E. coli genes.
Moreover, if you want to be sure that the protein is expressed, you could perform O/N induction at 20-25ºC with your plasmid in a good producer strain such as E. coli BL21. This strain is deficient in proteases and protein yields are usually much higher than in K12.
Two hours of induction/expression is usually too short to see a clear overexpression of a protein through SDS-PAGE. Try to make a time course experiment to determine optimal induction time for your system.
- Telmo Graça added an answer:14What are some common issues with SDS-PAGE that could be causing some problems our lab is facing?
Our lab has currently run into some problems involving protein migration in our SDS gels, and I was wondering if anyone has experienced something similar, or could provide some insight into what might be causing our issues.
I have attached an image for you to see.
We have suddenly had an issue where when we try to separate protein samples with SDS-PAGE, our higher molecular weight proteins merge into a single line (which is also visible as a translucent "bump" going through the gel). Oddly enough in this image, this line happens a bit lower (right above the dye front). I tried using Coomassie to see if maybe it was our ladder having issues migrating, but the protein stain also showed the protein samples were running exactly how the ladder appears.
The gel will start off running normally, with good separation of our ladder (each band is visible), but then the higher weighted bands will merge back together as a single line, and will slowly overtake every other band. Additionally, these gels have been running quite a bit more slowly than normal. The image attached to this was after about 2-3 hours at 120 volts.
So far, we have remade all of our solutions, have used another labs solutions, tried changing power packs, and have tried different cassettes incase it was faulty wiring. Nothing has been working, and we are all out of ideas.
The time that it takes to run the gel suggests that you have a much higher gel% concentration than you think or, too much resistance in the electric current conductivity (too much salts). Since you are able to load the gel I would look at the separation and not at the stacking part of the gel. I would also look at:
- Loading sample composition: As Nancy said, look at the loading sample buffer and your sample composition. Be sure you don’t have to may salts, Guanidine (precipitated with SDS and affects contiguous gels) or interfering detergents.
- Gel and Buffers percentage: Make sure that when preparing the gel you are diluting your stock solutions and not using them 10X
- Reagents: Finally, I think you should check the expire date of your reagents.
These may seem silly, but I have encountered all of them. Good luckFollowing
- Preeyanan Anwised added an answer:14Any suggestion for di-sulfide rich protein(~6% cysteine of total aa) expression and purification techniques in Escherichia coli to try further for?Hi,I am working on a protein of M.W. ~34 kDa (289 amino-acids), having 18 cysteines. First I was trying to purify it using 8M urea and then refolding by dialysis; tried with 6M Gn-HCl also. I used to get pure protein all the time, but not in it's native form as the 15N-1H HSQC showed only the random coil chemical-shift(not well dispersed peaks) every time. But, the protein is functional as concluded from western-blot test .
Then I tried to co-express it with Chaperone proteins(GroEl-GroES), where I was not able to purify the desired protein from the chaperones using neither gel-filtration nor additional ATP-MgCl2 wash through the purification steps.
Can I get some suggestion who has worked with or, handling this kind of di-sulfide rich Protein expression and purification in E.coli to get well-dispersed peaks of the native protein in 15N-1H HSQC spectra? Any further suggestion on the methods I have mentioned above also to try for? Thanks.
I have trouble as the same Somanth, I tried to express hemoglobin from Crocodile both in E.coli regardless of strain and express them in fusion protein system. However, I don't get the functional active our protein from these system. Later, I move on to eukaryotic expression system as yeast (Pichia pastoris). But I can not get the active hemoglobin in right conformation. Because my protein of beta chain hemoglobin has a Cys-free residue in its polypeptide chain. However, I'm not sure. Any further suggestion on my trouble, I have mentioned above also to try for? ThanksFollowing
- Valesca Anschau added an answer:3What is the best Glutathione S-transferase (GST) column "non agarose matrix" for protein purification ?
I'm trying to purify a GST tagged protein and I need a GST column to purify my control protein.
Glutathione S-transferase (GST) is a 211 amino acid protein (26kDa) whose DNA sequence is frequently integrated into expression vectors for production of recombinant proteins.
Glutathione is a tripeptide that is the specific substrate for glutathione S-transferase (GST). When reduced glutathione (GSH) is immobilized through its sulfhydryl group to a solid support, such as crosslinked beaded agarose, it can be used to capture pure GST or GST-tagged proteins via the enzyme-substrate binding reaction.
Binding is most effective in near-neutral buffers (physiologic conditions) such as Tris-buffered saline (TBS) pH 7.5. Because binding depends on preserving the essential structure and enzymatic function of GST, protein denaturants are not compatible.
After washing an affinity column to remove non-bound sample components, the purified GST-fusion protein can be dissociated and recovered (eluted) from a glutathione column by addition of excess reduced glutathione. The free glutathione competitively displaces the immobilized glutathione binding interaction with the GST, allowing the fusion protein to emerge from the affinity column.
This affinity system commonly yields greater than 90% pure GST-tagged recombinant protein from crude bacterial or mammalian cell lysate samples. Glutathione-based affinity purification of GST-tagged fusion proteins is easily done at either small, medium or large scales to produce microgram, milligram or gram quantities.]
More information here:
I hope its helpsFollowing
- Amanda Solem added an answer:8Can I induce protein from BL21 E.coli, pET32-b vector without adding IPTG?
I was thinking that whether I can induce protein from BL21 E.coli, pET32-b vector without adding IPTG? Since I was told that IPTG might be toxic to the cell, so I'm thinking of changing the temperature to induce the protein. Is it workable?
1. Double check to make sure your BL21 cells are DE3 - meaning they make T7 RNA polymerase. (I know it sounds obvious, but people often have both BL21 and BL21(DE3) in the lab so they can have a no expression control)
2. BL21s are standardly used with IPTG (in fact I have done little optimization of IPTG concentrations and all of the concentrations I tried worked just fine for BL21 and Rosetta lines). I can't imagine that the IPTG itself would be a problem unless you use an amount much higher than normal recommendations. As some others suggested, were you advised not to use IPTG because your protein is toxic to the cell? Perhaps it would be good to speak again with the person who gave you this advice.
3. Do you have pLysS in your expression strain? i.e is it BL21(DE3) or BL21(DE3,pLysS)? pLysS encodes lysozyme, which is useful for both lysing cells when you want to purify your protein AND inhibiting the T7 lac promoter. If you do NOT have pLysS, you might check to see how much background/leaky expression of your protein you have without IPTG. Depending on your particular circumstance, you may or may not actually need to induce I suppose. This may be what you observe with pRSET.
4. If the problem is really that your protein is toxic (as someone else suggested), then it can help to have pLysS present to repress leaky expression of your protein so your cells survive nicely until a particular point when you cross your fingers, add IPTG and let the cells suffer for a short induction until you harvest all your protein.Following
- Giuseppe Ossolengo added an answer:4Are there tricks to improve the efficiency of on-column digestion by TEV protease?
We are trying to remove a His-tagged fusion tag from the target protein by on-column TEV protease digestion during Ni-NTA purification, but the efficiency was quite low compared to the digestion during dialysis, which requires a second Ni-NTA purification step to remove the His-tagged fusion tag. Are there tricks that can help improve the on-column digestion efficiency?
if the imidazole should be the problem of the on-column cut, it should be enough to perform the last column wash in a buffer without imidazole, like 50 mM Tris-HCl (pH 8.0), 0.5 mM EDTA and 1-2mM DTT (that's indicated for TEV), with or without NaCl depending on the stability of your protein in low or high salt.Following
- Jaime Guillén added an answer:8Why do I get an unexpected MW overexpressed protein in E.coli strain?
I am expressing a protein having a MW of 32kDa by using pET21d+ vector and Rosetta gami strain.
I have tested several temperatures between 16 and 20°C and IPTG concentration between 0.3 and 1mM for the expression and all these experiments gave me always the same result. A protein is expressed in high quantity but it has not the expected molecular weight because SDS-PAGE and MS analysis give a MW of 40kDa, 8kDa more than the expected 32kDa. NO BAND appears at the desired MW.
I have observed also other interesting phenomena:
- the protein (40kDa), which has a 6xHisTag, is not well retained on the Ni-affinity column and, mostly, it goes in the washing (washing buffer contain tris 20mM pH 8, NaCl 150mM and it does not contain imidazole);
- The fractions collected before and after the imidazole gradient seem do not contain HisTag (test with antibody against HisTag);
- I found that transformation in rosettagami is very difficult and not always successful (perhaps due to the need of three antibiotics: tetracycline, kanamicine, and ampicillin)
The first thing that one is led to think is a problem with the sequence, but one of the ORF in 5'->3' direction on my plasmide is in agreement with the expected protein and 6xHisTag. A problem with a unrecognized stop codon is also possible and could be able to extend my protein sequence, but the stop codon is TGA, that is the suggested one for E. Coli strains. In addition, it is after HisTag as it has to be. However, also by considering a wrong recognizing of the stop codon is hard to think that 8kDa are added to the original protein.
By considering these phenomena, I am thinking that the protein that I would express is toxic for the used strain (it is not a pLysS or a pLysE) and that the band at 40kDa is an other protein expressed by the strain in this particular condition.
Is this plausible or can other explanation be possible?
What do you suggest to perform the expression in these cases?
Thanks for your suggestions.
I suggest you identify the protein by MS (peptide analysis) if the 40 KDa protein not is your protein you are wasting your time.Following
- Nick S Berrow added an answer:9How should I choose a protein affinity tag?
Hey guys, I need to express a small protein domain(70-80 AA), I use GST tag to express it and then cut the tag. but I find there are some truncation of protein. Does anyone has idea about this?
Is your protein truncated if you look at it by MS (SDS-PAGE probably not obvious with only 8k protein depending on the truncation) before cleavage from the GST?
If it is OK prior to cleavage from fusion then what is the source of your protease for fusion cleavage? This could be another source of (unwanted) proteases. Add protease inhibitors throughout lysis etc. as Marcia suggests if you do not already include them, remove from elution buffer for fusion cleavage.Following
- Daniel Joseph Slade added an answer:3Can someone please suggest a commercial antibody that detects an outer membrane protein from E. coli or multiple bacterial species?
We are currently expressing an outer membrane protein in E. coli and we are fractionating the soluble and membrane fractions, but I would like to have an antibody for an outer and even inner membrane proteins from E. coli to verify where what we are seeing during expression and fractionation. I've looked around extensively, but I can't seem to find many options. Thanks in advance!
Thanks Russell and Adam. Both are great responses and it's always nice to have an extra set of eyes. Cheers!Following
- Anna Chen added an answer:8How can I extract protein from Skeletal Muscle Tissue?
I am a PhD candidate at the University of Chicago. I am trying to conduct a western blot on some skeletal muscle tissues isolated from mice. I am running into problems with the sample prep step. Normally the tissues are harvested as a whole tissue, immediately frozen with liquid nitrogen, and stored in -80oC. When I want to do a western I will remove the tissue, suspend it in RIPA buffer that contains protease and phosphatase inhibitors, mince the tissue with scissors and homogenize before I spin the samples down (15 min at 12K in 4 oC) and use the supernatant. According to my Bradford assay I have protein (10-40 ug/uL normally) but when I do a western my internal control ( I've tried GAPDH and Beta Actin) are very very low and my target protein does not exist at all. Does anyone have suggestions on how to better extract proteins? Thank you in advance!
I'm happy to report with a combination of changes I got a WB that worked! Here's what I did. I removed the TA muscle from -80 oC freezer ( they were stored there after being snap frozen upon dissection). I added 10x vol of the buffer that Jose kindly provided above. I homogenized it with a new metal homogenizer ( I was using one with plastic blades before) for 3 passes of 10 secs. I then left it agitating on the vortex overnight and then spun it down for 10 min at 13k in 4 oC. I loaded 50 uL on the gel and ran the blot and finally got protein expression. Thank you all so much for the advice!Following
- Gertrudis Rojas added an answer:8How can I compare ELISA and BIAcore results?
Higher affinities result in higher ELISA signals, of course. But now Im working with several mutated ligands and their receptor. The wt ligand has an affinity to the receptor around 20nM. Several of my mutated variants have a severely decreased affinity (100-fold less), but in some of them the change is due to a decrease in association and in others to an increase in dissociation. I measure the affinities immobilizing the receptor on a chip and injecting the ligands with a BIAcore 2000 machine. When I do the ELISA coating with the receptor, adding the ligands (2h) and detecting with an anti-ligand conjugate (1h incubation ), those ligands with decreased association have no residual binding, while ligands with the same overall affinity but due to increased dissociation rate show positive ELISA signals at teh same concentration, although clearly lower than the wt ligand. Is the higher influence of association decrease in ELISA signals a common finding or not?
Second, two of my ligands are always giving me not only the same overall affinity (100-fold reduced), but also similar kinetic parameters (greatly increased dissociation). But in ELISA they consistently show a difference. The first ligand needs to be 10-20 times more concentrated than the second to obtain the same signal. The second one is very similar to wt in ELISA, just slightly lower. Is that possible? Should I check the quality of my kinetics measurements or you envision other factors (besides affinity and on- and off-rate) that could explain the difference in ELISA. I was thinking in a different proportion of active molecules in the different samples, but this is difficult to test.
Thanks for your answer. I actually tested all of them on BSA-coated plates, without evidences of non-specific background. But right now I think I found the explanation, which as it happens was rather simple. The affinities were not correctly calculated (the problem was in the evaluation, not in the experiment itself).There is actually a 7-fold difference between the two molecules (due to a kon change).Following
- Dennis Flierman added an answer:3How can I use 2 capture antibodies to detect cardiac Troponi I, in sandwich ELISA?
Just wondering how I can stick the 2 antibodies to the wall of the plate? as I have them in separate aliquots.
Should I incubate the first Ab O/N then next day incubate the other for another night before blocking?
Or incubate both together?
I tried to google similar protocols, got nothing.
Hi Zahran, you need to explain in more detail. Why do you want to coat with two antibodies? If you really want to coat with two antibodies, then it really depends on what you want to do with the assay. The two antibodies could have varying affinities, and mixing them 1:1 could therefore bias binding of one epitope. Most likely, you are looking to bind one Ab and detect with the second, as this is the common use of ELISA.Following