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Protein Purification Techniques - Science method

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Questions related to Protein Purification Techniques
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I am looking for a protocol to purify proteins from RIPA. I need to do iTRAQ afterwards, so excess urea is not a good idea. Please let me know if you have any suggestions. Thanks.
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Are you looking to buffer exchange your proteins out of RIPA buffer just to get rid of the buffer? Zebra spin desalting columns should do the trick for a low volume, simple buffer change up with minimal sample dilution. Not sure if you will get rid of every single trace of detergent though...
Maybe TCA precipitation?
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5 answers
I have isolated and identified more than 40 hypothetical proteins from E. coli by using MALDI-TOF and LC-MS/MS. Hypothetical proteins are cloned, over expressed and two proteins are characterized by binding study and crystal study.
Most of the proteins are not forming crystals. It is very hard to make deletion mutant of all hypothetical genes and make characterization of hypothetical genes. I have isolated and purified most of the hypothetical proteins by using chromatography. I would appreciate if anyone could give suggestions and advice regarding the protein identification by using any functional studies.
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I think that is very important a refined bioinformatic study of hypothetical proteins for planning a very detailed set of wet-laboratory experiments
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My elution fraction looks very dirty on SDS-PAGE. When I dig through literature some suggestions are
1. Increase salt concentration to reduce non-specific binding
2. decrease detergent concentration
The buffer I use for lyses (sonication) and washing the beads is
50mM Tris pH 7.5
150mM NaCl
1% triton x-100
10mM Beta-Mercaptoethanol (freshly added)
Any suggestions to decrease non-specific binding?
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Hi Kai
Thanks a lot for your suggestions. I am purifying another batch today. I just used what ever i had. I tried to restrict my sonication as well. I did 2 and half mins of 30% duty cycle (olden days sonicator large probe) 3 times with 5 mins interval. I am using for lysis 1xPBS pH 7.4 with 0.5% triton x-100 + 25 μg/ml PMSF + 2 μg/ml aprotinin + 1 μg/ml leupeptin. I will avoid triton in the next steps and see what happens..
Thanks once again.
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43 answers
I am trying to express a protein which I have 6XHis-tagged. I am purifying the protein in denaturing conditions. After elution, electrophoresis and coomassie blue staining I don't see any band in SDS-PAGE. In the eluted sample, a protein concentration of 5-6 microgram/ml was found. Is this concentration detectable?
One more thing, after washing with a washing buffer, the protein conc. of proteins in the flow-thorough was found to be 10-11 ug/ml. Does it mean that binding of the His-tagged protein to the Ni-NTA column is not efficient?
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Sudhir, The manuals always use some standard protein which expresses well. So, before even you worry about the volume of culture, you need to make sure the protein is expressing enough to see by coomassie staining.
I think you might be using ITC or SPR or protein-protein interaction, if so you need good amount of protein. Again, you are working on very big protein. So, I would think you would benefited if you workout the expression of the protein. Daniel has made really good points. for example " A good rule of thumb is that if you can't see the band appear by Coomassie, then you will not have enough material to warrant purification."
1. work on the expression- you can lower the temperature, change the expression strains, changing tag (his to GST or MBP), changing vectors, several other factors could be varied to get this done. If you are getting protein which could be detected only by western blot or silver stain then you could do only pull-downs are the best options.
2. You could even work with domains instead of 238 kDa protein. I dont know if its necessary to use full length protein for interaction studies. Small domains expresses well.
3. once you are sure that you could see the protein induction, then check the solubility by taking small volume and binding with beads. Chose denaturing protocol only if its not soluble.
4. If its soluble, then any manual can guide you through the rest.
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I am about to purify a protein complex and have a difficult problem.
Let's say protein A, B and C with 100kDa, 80kDa and 25kDa are the components. I know that they form inhomogeneous complexes with a range of something like 2xA 3xB and 8-12xC.
So the effective difference between each complex is as low as 25kDa and the average mass around 500 kDa which means their chromatography peaks overlap on any of our size exclusion columns (e.g. Superdex 200).
My first intuition was that we need a much longer column and we have only max. 600mm columns.
How long should such a column be, where do you get them and what would be the best matrix? I think I can manage to get a 2 meter column hooked up to the FPLC, but does it even make sense to try?
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You can plug 2 superdex columns in tandem, or even three or four, this is no problem to increase column length, but I doubt this will be enough to separate such complexes (500 vs 525 is 5% MW difference, i've never heard of such resolution of SEC).
I think, all the techniques of separation based on the (apparent) molecular mass / hydrodynamic radius or other physical caracteristic of your complexes will fail.
If you can get your protein C highly charged compared to the others (choosing appropriate pH), you could try to IEX, but this is also unlikely to work and could lead to complex dissociation.
How do you get your complex ? Is it in vitro reconstituted ? If you could manage to get the protein C in a limitant amount, you could increase the homogeneity of the sample shifting to the lower content of C in your complexes.
Good luck !
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I have cloned a gene in a plasmid which is without a tag and I am going to use the baculovirus expression system for its expression and purification. Since there is no tag present with the gene and there are no antibodies available for this protein, how do I go about its purification?
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Clear the lysate by centrifugation.
Ammonium sulphate precipitation. Identify the cut with your protein.
Gel filtration of this to narrow down to proteins of similar size
anion and/or cation exchange
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8 answers
I have used the methanol/chloroform purification method to purify proteins many times with good results. Last week, however, this method purified proteins pellets (from total protein isolated from a mouse cell line) that I have not been able to dissolve it in anything, including 8M urea, 2% SDS and other similarly harsh conditions. Do you have advise how to dissolve them? What factors might contribute to the formation of such insoluble protein pellets?
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I was using methanol/chloroform method few times. You should watch not to overdry the pellet.
But the most effective way of dissolving the pellet was sonication. Try it, I am quite sure it will help. I was dissolving the pellet in 1X sample buffer for SDS-PAGE.
@Toufic el arnaout: proteins do not remain functional, they are denaturated.
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7 answers
I have expressed my protein in E. coli. and I want to purify it. I performed the ammonium sulfate precipitation with 15%, 25%....75% in the indicial samples and ran both the supernatant and precipitates on SDS page. I noticed that my protein starts to precipitate even at 15% (at this conc half of the protein was in pellet and half was in supernatant). at 35% all the recombinant protein was precipitated and was present in gel. Now I want to do purification of that protein by Hydrophobic interaction chromatography using P-sepharose. I am wondering:
1. What should the concentration of ammonium sulfate in the sample be? (Normally it is recommended to be 1 to 1.5 M but at this my protein precipitates).
2. What should the conc of ammounium sulfate be for column equilibration?
3. What is the appropriate ammonium sulfate conc for wash buffer and for elusion buffer?
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I think that you have answered the question yourself you cannot use HIC if you use ammonium sulfate. You need not use ammoniun sulfate for HIC you could use any high molarity salt such as NaCl. However, I suspect that you will have issues with precipitation with any high concentration salt.
Are you using an existing protocol? What protein are you purifying? knowing these this would help.
I have used HIC and it is fantastic for some proteins but for most applications the resolution is not very good. If your ammonium sulfate fraction is pure enough you may get away with SEC I use superdex 200 pg. Good luck
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The proteins I am working on are 75.9 kDa with pI around 10.2 and 67.3 kDa with pI around 9.5 respectively and i´m am trying to find the perfect buffers (PH, composition, etc..) to use in purification for both. Can anyone help please?
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Here's a set of opinions from a LOT of experience, not all good:
Tris should be avoided. It is very temperature sensitive, an amine, and is a good general base catalyst.
For any buffer, you need to maintain control of ionic strength as well. Thus, two buffers at (nominally) pH 7.5 could have VERY different ionic strengths.
And, here I can help you out. Many years ago I wrote a calculator for construction of thermodynamically corrected buffer recipes. It has now been used to prepare three quarters of a million litres of buffers. I am aiming for a Megalitre before I retire!
Link:
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I am having trouble in the last step of membrane preparation of COS cells.The final membrane pellet does not seem to dissolve in the buffer.What should I do?
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Which protocol do you use?
I once prepped membranes from COS7 and other cells using this protocol:
Buffer A
250 mM sucrose
1 mM EGTA
10 mM HEPES-KOH (0.11915)
Protease inhibitors
pH 7.5 using KOH
Buffer B
1M Na2CO3
Buffer C
250 mM sucrose
1 mM MgCl2 (0.010165)
10 mM HEPES-KOH
-wash cells, scrape, resuspend in buf. A (approx 0.5 mL per 6w plate well)
- homogenize (Dounce-homog. 25x or -20°)
- centrif. 2x 500 x G 15 min 4°C (keep supernatant)
- supernat. + 100 µL buf. B per mL buf. A
- shake 45 min 4°C
- 100.000 x G 15 min 4°C
- resuspend pellet in buf C
never had a problem with this protocol...
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I have been purifying a protein (enzyme). In gel filtration I see two peaks all the time, one of the peak falls in the end of the void volume. Protein pI:6.1; buffer used Tris HCl pH 7.5.
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Hi Ambika. If you have a good assay for your enzyme then you really don't need to see a sharp peak to know where your enzyme comes out.
Is the high MW peak active? If so, you can rerun that fraction and see the lower MW activity appear, if the HMW peak is oligomers. If that peak is inactive, then it is more likely aggregates that will not yield an active enzyme.
Protein aggregation does not have to be concentration dependant, although it is more observable in concentrated samples. In general, protein aggregation occurs when the native protein structure is less energetically favorable than the unfolded protein structure. The reason for an energetically unfavorable protein structure is mostly dependent on its interaction with its solvent.
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Finding it difficult to get an active protein purified from inclusion bodies. So is there any protocol by which i can get the protein in soluble form.
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You are asking a million dollar question, there is no answer other than trying different conditions,host,etc..
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I have been trying to purify some proteins to use in a biochemical assay, I need at least 5mg total yield in ~2ug/ul concentration. They all have STREPII tag, I'm using 1ml Strep tactin column in an FPLC system. Concentration is very low and not very pure either, I need over 90% purity.
Does anyone have any advice? Is there any commercial source where I can send plasmids or transfected leaf tissues and they will purify the proteins for us?
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GE site says your column has a dynamic binding capacity of Approx. 6 mg Strep II-tagged protein/ml medium when they tested with GAPDH-Strep(II), Mr 37 400. Even if you get 50% of that you still could good amount needed for your studies in 2-3 purification.
But, if your yield is too low then you should optimize the expression. You could look up some excellent suggestions made in a parallel question @ http://shootingcupoche.com/post/How_do_I_know_my_His-tagged_protein_is_binding_to_the_Ni-NTA_column_or_not
You could additional ion exchange and gel-filtration step to remove contamination.
I dont have first had experience with Strep tag, but some times adding second tag makes life easier. Eg; Adding a his tag at c-terminal to MBP-tagged protein made purification lot easier for me.
I have suggested these thinking you are expressing protein E coli.
Good luck
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Whenever i m performing enzyme assay test on nitrate reductase it usually gave very less activity but i got god results in crude extract, so i m curious that after ion exchange chromatography how to protect my enzyme from degrading?
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Which type of nitrate reductase are you working with? Soluble or membrane associated? Are you assaying using methyl viologen?
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It is a cationic peptide of 40 aa. I used solid phase extraction as a concentration method, after a cation exchange that yields more than 97% purity, determined by RP-HPLC.
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hi, it depends on the aggregation maybe? is the peptide very hydrophobic? have you tried formic acid, ethanol, DMSO etc..?
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We make many gels and it is time consuming to mix acrylamide, pH buffer, SDS, and water together each time we make a few gels. Is it possible to mix these ingredients (No APS or TEMED) beforehand and store them for a few weeks at 4 C?
Are TEMED and APS stable when mixed together and frozen at -20?
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not recommended. TEMED induces the radicalization of APS to crosslink Bisacrylamide and acrylamide. Therefore, you want this reaction to take place as quickly and efficiently as possible. It only takes about 3-4 minutes to mix all the reagents together and 45-60 minutes for polymerization. If you want to expedite things and know you'll be needing a few gels in the upcoming days, simply pour a few plates the day or two before, let them polymerize and wrap the glass plates (layered in wet paper towels) in saran wrap and store them at 4C with the combs in-place! This will prevent the gels from shrinking and can be stored (with success) up to 4 days prior to use.
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Hi Joe,
In principle, the plasmid itself hasn´t so much effect on the expression level - however, sometimes the copy-number of the plasmid affects expression - so you might want to test both - low and high copy-ones. For expression of membrane proteins it is much more important to find optimal expression conditions (medium, temperature, induction time and concentration) and of course to find the optimal expression host: A lot of membrane proteins require certain lipids or need to be posttranslational modified (glycosylation,...) thus, it might be necessary to test expression in different hosts. For bacterial proteins often E.coli works but for mammalian proteins you might also want to test e.g. insect cells, yeast (also special yeast type like pichia pastoris) or human cell culture. Unfortunately, membrane proteins are not as easy to handle as the nice soluble proteins and it takes some effort to obtain reasonable expression. It might also help to check some literature like e.g. "Production of Membrane Proteins: Strategies for Expression and Isolation" (a wiley book). Hope this is not too depressing...
BR Christian
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I am doing protein purification, I need to ask about iptg induction using E coli BL 21 strain. Is it efficient or BL21(DE3), is there any difference between using BL21 and BL21 (DE3) for protein expression?
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E. coli BL21 (DE3) has the following genotype; F–ompT hsdS(rB– mB–) gal dcm λ(DE3). Here λ(DE3) means these E. coli cells have prophage (Engineered phage) that has T7 RNA polymerase controlled by Lac regulatory construct. So it will be active only if the media contains IPTG. But, one should remember that Lac repressor mechanism is not a tightly regulated system (Trp regulatory systems are tightly regulated). Hence, one can expect a basal level expression of T7RNAP. Plasmids like pET vectors are transformed into these cells for protein expression. pET system will have GOI controlled by T7 promoter. So, theoretically, the target protein will be expressed by these cells only if they are transformed and the media contains IPTG to induce the expression of T7RNAP, which in turn transcribes GOI in the plasmid. Almost all kinds of proteins can be expressed by this system except those proteins that are toxic to these cells (e.g. CRP). Because, you don't want these cells to express the toxic proteins all the time which will kill them and otherwise, you will end up getting low or no yield. As mentioned above, this system is not tightly regulated. Therefore, the basal expression of T7RNAP will transcribe your target protein and accumulation of these toxic protein will kill the cells. So, to express these kinds of toxic proteins, one should co-transform E. coli cells with pLys S or R, which is a small plasmid that encodes for T7 Lysozyme. T7 lysozyme forms a complex with T7RNAP by binding to the C terminal domain of the T7RNAP and makes it fall off the T7 promoter by creating conformational change. But the T7 lysozyme can only control the basal level expression of T7RNAP. When IPTG is added to the media, there will be a sudden rise in the level of T7 RNA pol which will not be controlled by the T7 lysozyme. Therefore, the toxic proteins can be expressed whenever you want simply by adding IPTG.
As mentioned above by Mr. Twachtmann, you can use both E. coli strains for protein expression if they have been cloned with Lac regulatory construct. But, if you are using pET vectors, then you have to use DE3 strains only. Also for toxic protein expressions, you can use DE3 strains.
All the best for your experiments..
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I am planning to isolate total proteins of gram positive bacteria. At final stage, I will resuspend the pellet in protein sample buffer that includes 40 mM Tris/HCl (pH 8) , 4 mM EDTA, 8% SDS, 40% Glycerol and sonicate it. Is it convenient for total protein isolation or should I change the buffer for better efficiency?
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I would include NaCl (150mM or more) to maintain the ionic strength. and glycerol seems a bit higher to me, normally we use 10% at max. In addition you can consider adding PMSF/protease inhibitor cocktail to kill the proteases. edta and sds are kept to be stringent since it may disturb the protein you are interested in!
Protocol for expressing soluble T4 lysozyme?
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Basically, I did try expressing T4 lysozyme (with a few amino acid deletion in the C-terminal end) in BL21 DE3 cells. The protein has an N-terminal HIS tag and I did HIS purification using nickel column, but never really get any protein from the lysate. Is T4 lysozyme generally insoluble and hard to produce or are there certain protocols (i.e. induction temperature, IPTG concentration) that I have to strictly abide to?
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Gene Volume 38, Issues 1–3, 1985, Pages 259–264 Non-toxic expression in Escherichia coli of a plasmid-encoded gene for phage T4 lysozyme L.Jeanne Perrya, Herbert L. Heynekerb, ∗, Ronald Wetzel, a a Departments of Biocatalysis and Genentech, Inc., 460 Point San Bruno Boulevard, South San Francisco, CA 94080 U.S.A. Tel. (415)952-1000. Ext. 6268 b Departments of Molecular Biology, Genentech, Inc., 460 Point San Bruno Boulevard, South San Francisco, CA 94080 U.S.A. Tel. (415)952-1000. Ext. 6268
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When we want to remove the N-terminal affinity tag from the purified protein, we often use the Tev protease, Thrombin protease, ULP1 and so on. After cleavage, one or two amimo acid will be left in the protein sequence. On crystal growing, two amimo acids may not have significant impact. But if the tag is linked at the C-termini, the protease above seems not so suitable, which will leave at least 4 amino acids in the protein sequence after cleavage. Could you give me some suggestion to solve the problem? (By the way, I can't use the N-terminal tags to purify my protein for some reasons.)
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HI Tengfei,
In our lab we use C-terminal fusions to an intein tag. This is a protein that self-cleaves in the presence of reducing agent. The benefit of this tag is that it doesn't leave any residues behind! You could also try to obtain a SUMO-his6 vector. SUMO protease recognizes the tertiary structure of the tag, and does not leave any residues behind!
Best of luck!
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18 answers
I want to degas a small volume (about 400 µl) of my precious protein sample. I thought about speedvac at RT, ultrasonic bath and bubbling helium through the sample. Does anybody know a good way to do this without harming or spilling the sample?
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Place the protein sample in a small eppendorf vial and place this in a septum vial. Close it with a rubber lid. Put a vacuum on the sample, which should be placed on ice, for 5-10 sec followed by argon. Repeat this a few times (5-10 times) and you will have an anaerobic protein solution. The ice will prevent bubbling of the sample.
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I try to find sufficient parameters such as Hz of sonicator, sonication time, cycles and breaks...etc
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I agree with Tobias and David. Try and error would help you to find the best protocol. Good lucks!
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I purified my protein using Phenyl Sepharose Fast Flow as a second resin. I have a problem with high binding of the protein to the resin. I have used dd H20 to elute the protein. Binding buffer is 40 mM Tris pH 8,0 100mM NaCl, but I can't change this buffer because it is my refolding buffer. My problem is I have a really poor yield, approximately 1% yield. Almost everything binds to the resin (app. 90%), but I can't elute it. I tried using low sub instead of high sub resin, but then my protein doesn't bind to the resin at all. I tried to use isopropanol to elute protein (10, 20 and 30%) but it doesn't work as well. What should I try?
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Did you try binding to the low-sub resin with ammonium sulfate (e.g. start with 1 M ammonium sulfate added to the sample and to the equilibration buffer)? As you said, it's not binding to this resin without ammonium sulfate; so, a decreasing ammonium sulfate gradient may work fine in the classical way ;-)
Is it possible for gene cloning of a toxic protein to be impaired by leaky T7 expression on DH5-alpha E. coli?
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I have been trying to clone a family of bacterial proteases into an expression vector under T7 promoter. After various protocols, the effective ligation/transformation is still to be achieved. Maybe T7 promoter may lead to a weak expression in DH5-alpha without induction and impair the growth of positively transformed bacteria? Or lead to the loss of the plasmid? What can I do to overcome this? Change the vector? Change the E. coli strain for transformation after ligation? It is worth saying that after ligation, the control (digested plasmid ligation) does not result in any colonies on the plate. The ligation with the inserts result in a few colonies, all negative. Sometimes strange things happen - when I digest the apparently positive clones, two bands appear, but both have lower lenghts than I would expect for vector and plasmid. Also, the higher the insert ratio I add to the ligation reaction (1:3 or 1:5), the more negative colonies I have!
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Just because DH5a lack the T7 polymerase doesn't mean they don't transcribe the locus. Other sequences in the plasmid can serve as promoters; indeed other bacterial promoters are present to drive antibiotic resistance markers. If a gene is highly toxic, even this low level of transcription (T7-independent) can be problematic. Though it would be convenient if we could exert total control of gene expression merely by using a strain-specific promoter, the observed biology is never that clear cut.
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When I purified my protein with Bradford's assay, it comes to 0.22 mg/ml with standard curve of BSA (macro assay). When I counter-checked with known concentration of lysozyme (i.e 1mg/ml) it tells 0.87 mg/ml. Should I move forward with this value of amount of my protein?
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Have you tried precipitating your purified protein out of detergents before running the assay?
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I wanted to perform IMAC with Cu-imidodiacetic acid using 50 mM Tris, 0.15M NaCl and 15 mM bME, but the color vanished. I'm not sure what color Cu(I) is (now I found that CuCl is white, so it could be only reduced), but I found that IDA can be striped under reducing conditions. Now I tried 5 mM bME, but it's just the same obviously.
I found a chapter from Methods in Enzymology (see the link), where they write that IDA is actually quite useless for this. Is there any way to do IMAC under reducing conditions without buying NTA?
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Tomas, I have never been able to get IDA Cu-imidodiacetic acid or Fe-imidodiacetic acid to work under reducing conditions. I have also not heard of anyone else using it successfully under those conditions. For this reason, even though IDA is cheap and plentiful, the more expensive NTA is probably the best way to go.
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I performed purification on FPLC hydroxyapatite column (CHT5-I from Bio-Rad) last week. I attached the chromatogram. Running conditions were as follows:
buffer A: 5 mM KH2PO4; pH 8.0; 0.15 M NaCl; 5 mM mercaptoethanol; 0.5 mM CaCl2
buffer B: 0.75 M KH2PO4; pH 8.0; 5 mM mercaptoethanol
the sample was in 50 mM Tris/HCl; pH 8.0; 0.15 M NaCl; 5 mM mercaptoethanol.
I managed to collect separately the very first peak, than the three smaller peaks together and the main peak at the beginning of the gradient. The activity was both in the three peaks and in the peak at the start of gradient. I'm looking for any suggestion, how to improve the binding, because it obviously eluted everything very early.
My ideas:
1) Use lower pH of the buffers, but I'm a little scared of that since my protein likes much more higher pH (9-10). And I'm not sure, if it would really help, but it seems that everybody is using pH between 6.5-7.5
2) exchange the buffer of the sample. However, I tried to keep the Tris buffer with other enzyme once and there was no big difference.
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Proteins have multiple different interactions with hydroxyapatite including cation exchange, metal chelation, anion exchange and hydrogen bonding. This was systematically examined with a library of proteins. The results suggested synergistic binding with both metal chelation and cation exchange interactions play a major role for protein binding. So I would take a look at Anal Chem. 2011 May 15;83(10):3709-16. doi: 10.1021/ac103336h http://pubs.acs.org/doi/pdf/10.1021/ac103336h. For phosphate buffer titration - because phosphoric acid has multiple dissociations you might try a phosphate Buffer Calculation (Javascript) - Fuse Home Pages
home.fuse.net/clymer/buffers/phos2.html‎ - A Javascript that calculates the amount of monosodium phosphate and disodium phosphate necessary to achieve a buffer at a given pH and buffer strength.
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I am trying to run my samples on a 4-12% Bis Tris plus (BOLT) gel from invitrogen, and am unable to get rid of some smeared pattern that I find at the lower half of the gel. I have tried dialyzing and filtering the sample through a 0.22micron filter. Also tried cleaning up the samples using a 2D clean-up kit from GE. Still the smear is appearing. Though it has faded now, it still appears and masks my protein bands. Any inputs on this?
Also, When am trying to clean-up 100ug (for example) of my sample, what is the quantity of protein I recover from the pellet? Should I account for any loss of protein?
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Problem: Band smearing
Problem:The voltage used is too high
Solution: Decrease the voltage by 25-50%.
Problem: The concentration of the
protein is too high
Solution: Reduce the amount of protein loaded on the gel.
Problem: The salt concentration is too
high
Solution: Dialyze the sample, precipitate the protein with trichloroacetic acid
(TCA) or use a desalting column.
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I'm trying to express and purify a recombinant chemokine in Drosophila S2 cells. I've already confirmed expression by ELISA. The protein is secreted and should be directly purified from the supernatant. If necessary, concentration of proteins could be adjusted prior to purification. I'm now looking for a cheap and efficient method for its purification. So does anyone have experience with custom made columns filled with anti-flag beads? And does anyone have a good protocol for the reconstitution of the beads, so that I could use them several times? Any help is appreciated!
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Due to the cost of the M2-agarose anti-FLAG resin, I wouldn't describe any purification scheme using the FLAG tag as necessarily cheap.
That said, you can certainly pack bulk resin into a reusable column. We typically bind in batch format, for e.g. 60 min, in a tube or bottle and then pour the resin into a column, generally an Econo-Pac Column or Glass Econo-Column from Bio-rad - although any will do. Elution is done under gentle conditions with 100 µg/ml 3x FLAG peptide. We normally do 3-4 elutions of 1 column volume each, incubating each elution with the resin for 30 min prior to collection - this ensures efficient release of the protein from the resin.
Regeneration can be done by applying 3 x 1CV 100 mM Glycine pH 3.5 and then re-equilibrating in TBS. Sigma recommend storage in TBS-buffered glycerol containing 0.02% sodium azide and storing at 4ºC or -20ºC -see the attached manual. You can certainly re-use several times without significant loss of efficiency.
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I wanted to do negative staining of polyacrylamide gel, using zinc sulphate or zinc acetate salts. The procedure I followed was 5 minutes incubation with 1% sodium carbonate, followed by a long incubation of 30 min in 200 mM imidazole made in 0.1% SDS after which I incubated the gels in 0.1 M zinc sulphate and also tried with zinc acetate. no staining could be observed even in 5 minutes or 10 minutes whereas the protocol says it should have been stained in few seconds. what could be the possible reasons? Can anyone please give me a tried and tested protocol and also explain me the reasons behind using them? Zinc acetate made a very opaque solution, should it be dissolved in ethanol?
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Others have sent similar, but there is a protocol in Current Protocols in Protein Science (2007) "page" 10.6.3 2007. It uses Zinc sulphate & imidazole to stain. I have used this and can see the negative bands where a lot of protein is loaded. It is considered reversible because the white precipitate can be removed using 0.3% citric acid. However, I have not tried that part.
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I have a problem regarding the purification of his-tagged protein. My protein is expressed in E.coli. My protein is about 50kDa and I have successfully purified out the protein using HisTrap FF column from GE healthcare. And the band is thick. The problem is, sometime (about a week) I have seen some white precipitates formed in my eluted protein. I ran SDS-PAGE again to check and found a lot of other contaminant bands in my purified fraction.
The condition for my binding buffer is: 50mM NaH2PO4, 500mM NaCl and 20mM imidazole pH7.4
For the elution buffer: 50mM NaH2PO4, 500mM NaCl and 500mM imidazole pH7.4
Is my protein degraded? Should I add protease inhibitor? Or there is other reason?
Any suggestions please?
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Do you have EDTA in your protein storage buffer? Nickel ions can leach off your column and promote aggregation of his-tagged proteins after purification. Maintaining your protein in an EDTA containing buffer can avert this problem.
In addition, protease contamination can never be completely avoided. In my experience it is best to always include protease inhibitors even after purification, unless you have a downstream application that precludes their use.
From what you've said, it is not clear that the precipitation problem and the potential degradation of your protein sample are related problems. You may need to troubleshoot them separately.
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I'm going to express a protein in the membrane surface of a gram-negative bacteria and I wonder which should be the best protocol to obtain the membrane protein fraction for western blotting purposes. I understand there are in house protocols based in ultracentrifugation separation and also a few commercial kits available. Any recommendations?
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This method that I am going to describe is tried and tested and works really well in my lab.
1. Once you have harvested your bacterial cells (over-expressing your membrane protein), wash the cells PBS to remove residual media. Re-suspend yor bacterial pellet (approx 4-5g wet cell weight) into ~ 30 mL lysis buffer*.
2. Lyse your cell suspension either via probe sonication (30 sec ON, 30 sec OFF, 12 cycles, keeping the suspension cool, i.e on ice) or French Press (3 passages).
3. Add appropriate levels of DNAse, RNAse, and protease inhibitors (I use the Roche inhibitor cocktail tablet). Centrifuge the cell slurry @ 27,000 xg at 4 deg for 45 mins.
4. Remove your supernatant and centrifuge at 120,000 xg at 4 deg for 90 mins (obviously using an ultracentrifugation (UC)). Remember to keep your samples cool (i.e. on ice at all times).
5. After UC, you should see a "dark brown/orange" residue at the bottom of your tube, this is your membrane fraction. Carefully remove the supernatant (cytoplasmic fraction) from the tube.
6. you now want to re-suspend your membrane fraction, this is the tricky part. If you mix this fraction up and down it will bubble due to the detergent effect of the lipids, this is not such as great idea, your activity will be affected. I find the best way is to completely remove all the supernatant using a P200 tip. Then add back to your membranes fresh buffer (what ever that might be, i.e most appropriate for your membrane(s) protein(s)). I usually add ~300 uL to the tube. Cover the top of the tube with some parafilm and leave on ice or at 4 deg overnight. The membrane residue will naturally "lift" into the solution and you will find it easier to re-suspend into solution via very gentle/mild pipetting action (remember not to introduce bubbles).
* remember, selecting the correct lysis buffer will dramatically affect the outcome of your experiment. This is something only you can answer.
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I am looking for small peptides or proteins soluble in water. I need them to study the influence of some co-solutes on protein stability in water solutions. I cannot produce them in my own lab, so they have to be commercially available. Does anyone have recommendations? I have already found beta-thymosin.
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Thermolysin from Bacillus thermoproteolyticus should be easy to find and cheap, it is a zinc proteinase and was used often for structural analysis
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I would like to concentrate and purify FMD virus preparation (to get rid of non-structural proteins of less than 100 KDa) . My idea is to initially concentrate the virus by PEG precipitation, then use this preparation for further purification and then concentration by adapting a methodology, which members of RG suggested.
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The manufacturers of amicon spin filters state that for a protein to pass through the filter it has to be 1/5 of the MW cut off. So you would need a 500k NMWCO filter to efficiently filter out 100K proteins. However, my experience is that this is optimistic at best. I have attempted to filter histone H4 14kDa through a 100K NMWCO filter and almost no histone H4 filters through. Using SEC as suggested by Joachim is the best choice. Try to find a resin that has a cut off such that 100K or more comes out in the void volume. Collect your protein in the initial fractions and dump everything else then concentrate. Good Luck
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I have eluted my protein with 200mM imidazole and got some non specific bands in the elution but concentration is good. Elution with 300mM imidazole leads to the reduction of non-specific protein elution and decrease in the concentration of protein of interest. I have not used any imidazole in the wash buffer, so how much concentration of imidazole do i have to use to reduce the non-specific protein elution? No protein elution with (20mM,50mM imidazole).
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There are some steps that you might take to overcome this problem:
1) Add up to 20 mM of Imidazole in your sample and wash buffer. It can avoid weak interactions of His from other proteins
2) Make a linear gradient of imidazol in FPLC equipment. Try a "slow" (0,5 mL/min) 0-500 mM of Imidazole gradient
3) Adding 500 mM (or even 1M) of NaCl can avoid weak electrostatic interactions of other proteins with your protein or the column.
And keep in mind that:
1) The more the purity degree, the less yield you may achieve
2) This affinity purification is usually a first-step purification process. You can get higher purity using ion/size exclusion chromatography later (don't forget to dialyse your sample to remove Imidazole!).
Hope it helps! =)
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I have some confusion about the amount of sample normally required to load on gel during Western blot.
I will be loading 50ug of sample in total volume of 20ul (including loading buffer) . Therfore my question, should it be 50ug/20ul or 50ug/ul?
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I understand it is not your question but I have another question for you. What type of gel are you using. The gels I use have a loading capacity of 20 ug. These are Nupage Bis tris precast gels. So I'm really surprised you are loading 50ug. The amount sounds too much for any get! Just a suggestion to look into it! Also our wells also only need 20 uL because like Kenneth pointed out you don't want overflow!!
BTW Kenneth is correct it is 50ug/20uL!!! GOOD LUCK!
I'm a western queen considering i'm in a proteomics lab and its all I do. :P
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Does anyone know a good starting point/generic Triton X100 (or any detergent) protocol?
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Check this
Biochimica et Biophysica Acta 1798 (2010) 1926–1933
it may help you..
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I'm going to work on polyphenol oxidase (PPO) extracted from a fruit peel, and from what I read recently I came out with a series of purification steps involving ammonium sulphate fractionation, dialysis and gel filtration chromatography (Sephadex G-200), done accordingly after crude enzyme extraction (using sodium phosphate buffer).
I'm not sure of which substrates to use for PPO extracted from this fruit peel yet, so I might choose 2 of the best substrates with lowest Km values out of 4 substrates that are going to be tested on this enzyme.
So my main question is: Is my current protocol enough to obtain a partially purified PPO? At least 20 times pjavascript:void(0);urified?
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Hi Allan,
In addition to former answers, all of them helpful, you have a lot of literature describing classical PPO purification of a number of fruits. See for instance:
Isolation, purification and physicochemical characterization of polyphenoloxidases (PPO) from a dwarf variety of banana (Musa cavendishii, L)MAM GALEAZZI, VC SGARBIERI… - Journal of Food …, 1981 - Wiley Online Library
Purification of polyphenoloxidase from coffee fruits P de Fátima Pereira Goulart, J Donizeti Alves… - Food chemistry, 2003 - Elsevier
Polyphenoloxidase from apple, partial purification and some properties
A Janovitz-Klapp, F Richard, J Nicolas - Phytochemistry, 1989 - Elsevier
Strawberry polyphenoloxidase: purification and characterization
P WESCHE‐EBELING… - Journal of Food …, 1990 - Wiley Online Library
And many others. A diferent point is the appropriate substrate to follow the activity. Sometimes, the affinity is not the more determinant parameter. A high Vm and a simple colorimetric methods coulb be great although the Km is not very low.
If you need more details. just say. Cheers.
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I am purifying a protein, initially I used to add only serine protease inhibitor (PMSF), now I want to use a protease inhibitor cocktail (from Calbiochem- 539132), where amount to be added is not given - how do I use it in an effective proportion? Does anyone have any suggestions?
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Prem, Since I work on a tight budget, I tend to make my own protease inhibitor cocktails. The tablets that are available from BD Scientific, Roche, or Sigma work terrifically well, but they are pricey. Therefore, I buy my own protease inhibitors and then make frozen aliquots of a common stock solution. You need to use a mixture of protease inhibitors, since no one protease inhibitor will inhibit all available proteases. I make a stock solution of 100 mM PMSF that I usually dilute 1/50-1/100. I also use a stock solution of 100 mM benzamidine HCL that I dilute the same. My stock solution of Pepstatin is at 100 µg/ml that I dilute 1/20 and 100 µg/ml stock solution of Leupeptin that I dilute 1/10. With this combination of protease inhibitors and by keeping your extracts ICE COLD, you will usually be able to inhibit any problematic proteases. Oh - EDTA (pH 8) at a final concentration of 1mM is also important for metalloproteases.
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I am trying to purify E.coli protein tagged biotin on its N-terminal, using FPLC system. Very few amounts of my protein was eluted from the column and a large quantity was released after washing the column by the regeneration buffer, Acetic acid 10%. Why I couldn't elute the protein using the elution buffer and it is still stuck in the column?
The same protein has its C-terminal strep tag, I got the same problem when I purified it using strep-column. The protein was not eluted using the elution buffer and was released from the column by NaOH 0.1 M, the column regeneration buffer. NOTE : the protein is DNA-interactant.
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What is the real question?
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I have taken my sonicated sample and kept it for centrifugation (10000rpm for 15min) to separate the pellet from supernatant. After that, I centrifuged my supernatant @15000rpm for 30 min and separated it from the pellet. Now I am running pellet1, pellet2 and supernatant on the gel. However, after staining on the gel, there is no protein at the corresponding molecular weight in the case of the supernatant, but there is a good amount of protein in pellet1 and pellet2.
Can anybody explain why this happened or whether my protein is insoluble?
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hydrophobic interactions are the most prominent factor in folding of proteins and help them attain their energy minima during folding. however, whenevr a "violent" method like sonication is used with cells or cell extracts, the energy minimum of a protein is overcome and it unfolds, sometimes exposing a significat proportion of its hydrophobic regions, and leading to aggregation. treatments like ammonium sulfate bring about the same unfolding but prevent aggregation due to their kosmotropic nature. Inclusion of a small amount of a kosmotropic salt during release of proteins from membrane compartments definitely helps. if you dont want salt in your extract you can try something like glycine. inclusion bodies as suggested by others here are definitely a problem, but i have consistently found that if you can control aggregation during homogenization, most of your worries are done away with. you can alternatively use a bead based homogenizer instead of a sonicator. as for the first thing you asked, yes, if proteins are aggregating, they would definitely be found in the pellet instead of the supernatant. i would like refer you to a research paper that deals with these issues quite extensively:
Bondos and Blicknell (2003): detection and prevention of protein aggregation before, during and after protein purification. Analytical Biochemistry, 316: 223-231.
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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?
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Just a thought if you're interested in trying something different: in my own work, I prefer repeated freezing/thawing in lysis buffer over sonication for the very reason you mentioned, as well as that sonication produces transient pockets of very high heat, which could denature your prep to some extent. After 1-2 freeze/thaw cycles, my crude pellets are a single color (no light brown opaque bottom layer = unlysed bacteria) and my protein is intact and soluble. Your result may not be the same, but you may find better lysis without risking protein denaturation.
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Is there any special centrifuge filter or another instrument for the purpose of tiny volumes?
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Easy: There are a lot of dialysis filter devices that can be spun in centrifuges. Amicon/Millipore is a good provider of nicely working centrifugal filters. They are, however, nicely expensive, too.
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Yesterday, I salted out my proteins (basically crude extract from maize) and dissolved in 1/10th volume buffer. I wanted to desalt it on HiTrap column so I filtered it through 0.22 um filter. When I measured activity, it was there after the dissolution, but was gone after desalting. I doubt the desalting would be responsible (although the ABS280 behaved weirdly, see the attached picture), plus I saw that almost everything stayed on the filter. Is it possible that 0.45 um filter would not cause such lose of activity? I don't think so. What would you recommend to concentrate and desalt proteins quickly?
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Acetone precipitation is very likely to cause loss of all activity. There are three simple options for desalting your protein after re-suspension of the ammonium sulfate pellet: (1) gel exclusion chromatography, which can be done with a hand-operated desalting cartridge, (2) repeated diliution with buffer and concentration in a centrifugal ultrafilter, or dialysis against several changes of salt-free buffer. For any method, you should clarify the re-suspended pellet by centrifugation before proceeding. Ultrafilters should be PES, not cellulose acetate or nylon, as these may strongly bind certain proteins, and may be the cause of the loss of activity. We typically do #1 for our protein preps where salt precipitation is a purification step.
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I am conducting 2DE of hippocampal proteins. The protocol is as follows:
1. Sample preparation: hippocampus were solubilized in the lysis buffer (7M Urea, 2M Thiourea, 4% Chaps, 30mM Tris-cl, pH 8.5) and sonicated for 10s. Then the homogenates were centrifuged at 1,4000 rpm for 15min. The supernatants were added with ice-cold 100% TCA to a final concentration of 15% and incubated on ice overnight. Then the solution were centrifuged at 1,4000 rpm for 15min at degC to remove the supernatants. The precipitation were solubilized with three volume of ice-cold acetone. (Since the precipitants could not resolve in acetone, I have to sonicate them until the precipitants solubilize in acetone) Then the solution were centrifuged at 1,4000rpm for 15min at 4 degC. Repeat for three times. Then the precipitants were left on ice for 30min for the evaporation of acetone. Then the precipitant which is removed of acetone were solubilized in lysis buffer(7M Urea, 2M Thiourea, 4% Chaps, 30mM Tris-cl, pH 8.5) again.
2. IEF: The purified protein solution were added with rehydration buffer (8 M urea, 2 % CHAPS, 0.2% DTT, 2% (v/v) IPG buffer, pH 3–11 NL, 0.002% bromophenol) to the volume of 450μL. The IEF conditions: Strips were actively rehydrated at 20 degC for 18 h at 50 V, focused at a constant temperature of 20 degC beginning at step 300 V for 2 h, step 500 V for 2 h, step 1000 V for 2 h, gradiently 8000 V for 8 h, and finishing at 8000 V for 10 h step.
3. SDS-PAGE: After the IEF, the strips were equilibrated in 2% SDS, 30%Glycerol, 6M urea, 75mM Tris-cl 6.8, 0.002%Bromophenol blue with 1% DTT and subsequently 2.5 % IAA for 15 minutes, respectively. Then they were transferred to the 12.5% SDS-PAGE. The cathode and anode buffers were both the Tris-glycine-SDS (5X: 125mM Tris, 960mM Glycine,0.5% SDS, pH 8.3, what I used was 1X)buffer. 1W/Gel for 50min and 11W/ Gel for 6 hours.
The results showed heavy Horizontal streaking in the high molecule weight part and heavy vertical streaking in the basic part. I think it is due to the wrong sample preparation. I wonder if someone can suggest me the possible solution. Thank you for your help.
Attached is the new result of coomassie blue result.
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Hi Xiaojing Sui,
I have performed many 2Ds with protein extracts from rat hippocamus, but never have to use precipitation methods or any Clean up kit. Please take a look at my publications for the recipes i ve used and the amount of proteinloads per IPG/Strip. I would really omit clean up methods since they will add recovery issues to your 2D methodology.
good luck,
Murat
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I'm trying to express an antibody fragment that is covalently held together by cysteine residues that I have added to the sequence. After running the western (non-reduced) to assess expression, I obtain two bands. One band represents half of the fragment, the other is the covalent version at double the size (cys-fragment). I'd like to increase the amount of covalently bound fragment.
Does anyone have experience with the addition of copper sulfate to the media to promote thiol bonding? I've found some literature from Chaderjian et al, which was helpful, but I'm also curious if the copper will inhibit purification via 6-his tag?
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@Matthew - I gave your suggestions some thought, and what about adding copper to the media after harvesting it from production? Since in my case the protein is secreted into the supernatant, the cells have long been filtered away prior to purifying on the Ni-NTA column. At this point I don't need to worry about cell viability because I'm only concerned with the supernatant. Considering I am am using serum-free media during production, there might be a benefit to adding copper to induce disulfide formation once the supernatant is harvested. The copper could then be removed by dialysis prior to purifying the supernatant on an NI-NTA column. I could be way out in left field though, as I have no experience with this.
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Protein should be purified from HCP and other impurities with a vol productivity of 2gm/ml
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With a high isolelectric point like this you can run your protein through a strong anion exchange directly on to a strong cation exchange. At neutral pH your protein is unlikely to bind to anionic column and most likely will bind to cation column. You will remove a lot of proteins this way.
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I am trying to purify my protein (protein implicated in the DNA recombination process) which was overexpressed in benthamiana plants using the FPLC system, so, I grind my infiltrated leaves and extract the total proteins using the buffer: Tris 100mM Tris ph 8.0, 5mM EGTA, 5mM EDTA, 150mM NaCl, 10mM DTT.
After centrifugation at 20,000g 2 times (to clarify the supernatant) I notice that my precipitate is gelatin like (could be DNA). A western-blot test of the precipitate showed my protein is in the majority of the precipitate.
- Could anyone suggest a method to decrease the viscosity of my protein extract, keeping the complex protein-DNA intact?
- How can I show, by a quick experiment, the interaction of my protein with DNA (protein is tagged) and have an idea about the nature of DNA interacting with my protein?
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To reduce DNA associated viscosity, you can use sonication to fragment DNA, or DNAse (MNase for example) treatment (your protein will still be associated to DNA as there will be a "footprinting", protection of the DNA associated to your protein against the DNAse activity). You can look at the first steps in protocols for ChromatinIP that should give you some informations about such treatments.
If your protein is ossociated to DNA, during/after purification you can monitor the absorbance at 280nm(protein) vs 260nm (DNA). A strong 260nm absorbance will assess the presence of DNA together with your protein.
You could also imagine to perform some gel shift assays between your protein alone (after a high salt wash for example) and your protein associated with DNA
To determine the nature of DNA associated to your protein is more complicated, the only way I see to have a real identification is a CHIPseq like method.
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The protein is prone to aggregation
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Dear Laura,
Ok, so at least you have soluble protein - it is not in inclusion bodies, and you can purify it, to some degree. It seems to me you have a couple of problems - low yield/weak binding to the Ni column, and aggregation following elution.
Regarding binding to the his column:
I would suggest increasing the ionic strength of your binding and washing buffers - 0.5-1M NaCl can help reduce non-specific contaminants.
Secondly, you should include a small amount (5-10mM) imidazole in your binding buffer, and if binding is very weak bind either in batch, to resin, or by passing your lysate over the column multiple times.
I would also suggest, if you have not done so, to run some beads on a gel after elution to confirm that your protein is not remaining bound to the column (if it is aggregating, this is not uncommon).
Lastly, another way to get rid of non-specific contaminants is to remove your his tag, dialyse into a non-imidazole buffer, and then pass several times over a his column - your free his tag should bind to the column, as should the contaminants, but your protein should remain unbound.
Regarding aggregation:
There are a number of additives that can help to suppress aggregation of your protein. Probably the first one I would try, as it has worked very well for we in the past, is a sub-CMC concentration of detergent - maybe 10-15mM beta-octyl glucoside. I would particularly suggest trying this in your case, as it sounds like the aggregation you observe may be due to exposed hydrophobic patches on the surface of your protein (which the octyl glucoside will shield).
Different additives can be screened by including them in either/both of the IMAC elution buffer and the gel filtration buffer. 5% glycerol can by very effective, as can an equimolar mixture of arginine and glutamate (20-50mM of each).
Also screening different pH buffers for the gel filtration is worthwhile, as often your protein will be happy at one pH, but very prone to aggregation at another.
Lastly, it may be that the aggregation is due to improper folding of your protein, in which case it may be worthwhile modifying your expression conditions - reducing the temperature of expression to 16-20 degrees celcius and expressing overnight would be one of the first things I would try.
Coexpressing chaperones may also help (there is a set of plasmids for this purpose available from Takara).
A couple more questions:
When you say only 10% of the loaded protein binds to the His column, I assume that is determined by western of the clarified lysate?
When you say the protein elutes in fraction corresponding to a MW of 270kDa, is that fraction in the void volume of the column? or in a peak after the void volume? Because if it is the latter, you may be dealing with a specific oligomer of your protein, rather than a non-specific aggregate.
Good luck,
Oliver.
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I am purifying a protein from a cell supernatant with a His-trap column from GE. I get quite low yield and it seems as if the protein doesn't bind very well since I can obtain protein when applying the flowthrough on the column again. I have seen different advices regarding buffer optimization. Some say that one should increase the imidxol concentration to approximately 50 mM to reduce unspecific binding (which may be a problem when applying a cell culture supernatant) and others say that the imidazol concentration should be low (5-10 mM). For me it appears as i the protein binds better at lower imidazol concentration. Does anyone have an idea of why? I have a buffer with 0.02M phosphate and 0.1M NaCl (besides imidazol) pH 7.4. Should I increase the salt concentration?
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there is no "magic" imidazole concentration - depending on the oligomeric state of your protein and the accessibility of the His-tag optimal binding, wash and elution concentration vary. My advice:
- use as little Ni-resin as possible to avoid impurities binding in the first place. It is better to lose some protein that can't bind than get less pure protein in the end. We usually use 0.5 ml resin suspension per litre of bacterial culture.
- do the binding step in batch, not in a column
- then pour in a column and elute with a step gradient, for example: 10, 20, 50, 100, 200, 500, 1000 mM imidazole. Run eluates on a gel and decide which fractions to pool.
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I've asked a question about this topic last week and got very useful responses. Thank you all for that. So would running a native gel give me any information about this behaviour of the protein. If yes, at what step in purification should I do that and how?
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Native gels with membrane proteins can be tricky. Have you run the SDS-PAGE and Western first to see if the peaks are really your protein of interest? If you have enough protein in your peaks, I would run them on a size exclusion column to check for aggregation/oligomerization. Of course, a functional assay is vital if you have one developed.
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If we have two test tubes containing protein sample, is there any way to know weather the two proteins are same or different (unknown proteins) without using any instrument?
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First, hydrolyze the peptide bonds. Then conduct 2D Thin Layer Chromatography and lastly use the AA's pattern and the relative spot intensities to compare samples.
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I have separated a crude mixture of protein by ammonium sulphate precipitation method (70%). I now want to separate a specific protein of about 66 kDa in size. I am a bit confused about the use of a dialysis membrane. Can someone suggest which type of dialysis membrane I should use to separate the protein(66 kDa)?
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Assuming you want to keep your protein native I would use gel filtration and ion exchange to further purify your protein. Success of these downstream purification procedures will depend on how complex your 70% ammonium sulfate precipitation mixture is. If your 66kD protein is a prominent band by SDS-PAGE, dialysis using a 30K MWCO membrane into a buffer such as PBS should allow you to do gel filtration using a column similar to a Superdex 200 column. You will then need to transition fractions eluted containing your protein of interest into a low salt buffer and further purify using cation or anion exchange depending on the isoelectric point of your protein. This approach has the potential to yield protein that is quite pure.
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Is there any protocol or kit to do this?
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Hi, I've extracted protein from native PAGE simply by crushing the gel through syringe few times and then overnight incubation in buffer. I think that should work also for SDS-PAGE.
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I want to analyse the expression patterns of my protein of interest in various tissues such as brain, liver, intestine, kidney, spleen etc. Does anybody know a good protocol, which is suitable to get comparable total protein extracts from all kinds of tissues?
I have been looking for protocols, but all I found was limited for certain tissues or cell types. Because I want to have a first overview, I would like to include as many different tissues as possible.
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The protocol which I have used for different tissues - heart, liver and kidney -
Homogenized (~ 200 mg) in protein extraction buffer (Tris-HCl, NP-40, NaCl, EDTA, NaN3, and PMSF at pH 7.5) with freshly added protease inhibitors (DTT, leupeptin and aprotinin) using mortar- pestle or using sonication method and centrifuge lysate at 27,000g for 20 min. Collect the supernatent and determine protein concentration to
using protein assay e.g. Pierce Protein Reagent Assay BCA Kit.
We have found this method consistent and seems to work for majority of tissues.
How to purify chromatin binding protein complex that easily degrades in 100mM, 150, or 250mM NaCl?
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I am using agarose beads and Ab against FLAG-tagged protein during O/N incubation of cell extract in buffer with Roche protein inhibitor tablets and PMSF. At higher salt concentrations I can recover protein itself and see some degradation but no binding partners and at lower salt concentrations protein degrades. There are alternative approaches but I want to try working out this approach first. Any suggestions? I thought of decreasing O/N incubation to 3-4 hrs and adding more protease inhibitors.
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Thank you Binal!
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I have a protein solution in PBS with 50mM DTT and I would like to get rid of DTT in order to have my proteins in PBS. Any suggestions?
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The likely easiest ways are either dialysis, or alternatively spin desalting columns (e.g. http://www.piercenet.com/objects/view.cfm?type=productfamily&id=c6b5ebdf-7e66-4279-bb9e-2140ec349b3e).
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I'm trying to isolate proteins to perform an IEF of Leishmanial proteins, so I have to eliminate all the DNA present in the lysate. The problem is that 10 min after the incubation with DNAse (20 micrograms/ml DNAse + 5 micrograms/ml RNAse) at 37 ºC all the proteins contained in my lysate get precipitated. I would greatly appreciate a solution to this problem. Thanks in advance
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Do you experience protein precipitation without DNAase after 10 min incubation at 37 C as well? It might be, that the proteins precipitate at 37 C in aither case. Try to keep your sample at low temperature and as I mentioned use Benzonase.
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Normally I see that if you use 100ul slurry volume beads, then use 100ul SDS loading buffer at the final step to boil the proteins off the beads. I wonder if I can use less volume (say 30ul) to boil the sample so I can load everything in a single well of SDS-PAGE gel?
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100 ul is quite a lot, If you are experienced gel runner even 30 ul is quite a lot, generally to get sharp crisp bands the lower the loading volume the better, you may make an argument about the stacker, still lower volumes are good. after boiling spin the tubes, so all settle well down at the bottom. Again if experienced go for 20 ul, if not 30 is good. after boiling you would lose some in evaporation anywhere from 5-10 ul. Good Luck with results.
Mary
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I have a PVDF membrane with the proteins blotted onto it. I placed the membrane in blocking solution (2.5% non fat milk) overnight but I don't have the right antibody to probe it with. Can I just store the membrane as such (i.e. dry it at this point and store at -20oC) until I get the right antibody to probe it with or should I strip it of the blocking solution before storage?
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agree with elizabeth
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I am looking for a precipitation protocol for protein samples before doing the iso-electric focusing (IEF). I have tried a TCA/Acetone protocol but the protein pellets I get at the end are very difficult to dissolve even in rehydration buffer for proteomics.
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When doing TCA precipitation, re-solubilization of the resulting pellet is often easier when adding 10% glycerol in sample prior to precipitation. If you have enough sample to play with, maybe this is something to test out...
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I'm trying to stabilize my protein by use of different buffers. It obviously loves alkaline pH, because below pH 6 the activity was gone in few days even when stored at -80°C. However, even at pH 8 and stored at -80°C the activity decreases (it's little weird, because until day 3 it stays at about 100% and on day 5 it drops down). However, I wanted to try other buffers, eventually with higher pH. For pH 9 I want to use Tris with either HCl or citric acid. I was thinking also about pH 10 with borate buffer, but I found out, that borate can be used only up to pH 9.2 (see the link attached). What I found was either bicarbonate-tetraborate or glycine-NaOH. Which one do you think would be better?
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Hey,
Are you sure that your protein behaves well with the said pH? I would suggest you to use ProtParam tool from ExPasy (online) and find out about your protein parameters using the sequence data. You can find out what is the theoretical pI (iso-electric point) for your protein and then store them in buffers with pH farther than your pI. Your protein will be least soluble if the pH of your buffer is close to pI value.
I believe this factor is important.
Best Regards,
Sagar Sridhara
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His tagged protein eluting at 2 or more seperate peaks on AKTA
What could be the reason(s) for having a His-tagged TM protein (Cx50) to elute in multiple peaks at different imidizole concentrations?
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This could reflect different oligomerization states of the protein or that a portion of the protein is not properly folded.
I would suggest to test these different peaks on a gel filtration column
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I am trying to purify my protein using Nickel column after expressing it using a Baculovirus expression system in Insect cells. I use detergent to lyse the cells. Spin it down to remove DNA and other cell debris and then apply the sample to a Nickel column. The problem is, the sample runs through the column very slowly and the whole purification process (gravity flow) takes me several hours. I use 5mM and 20mM of Imidazole to wash and elute using 500mM Imidazole. The flowthrough and wash steps take several hours. Since I am not seeing my protein on the SDS gel, I believe it might be getting denatured due to the length of the procedure. What am I doing wrong?
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One thing you may think about trying in order to minimize the time of each step, is to do your purification in a "batch-wise" fashion rather than on the column. I do this for my protein which I have to keep in detergent as well. Basically, after rocking the clarified lysate with the resin for an hour in the cold room, I spin down the resin and carefully pipette off the "flowthrough" i.e. unbound protein. Then, I add the appropriate wash buffer, let that incubate with rocking for 10-15 minutes and then collect the resin by centrifugation again, pipetting off the solution above i.e. the "wash". I do this for each step. I do this more because of limited time (kids need to be picked up etc) and one has to be very careful to ensure all the buffer is removed before adding the buffer for the next step. You may want to increase the number of washes if you are not confident in your ability to remove all the unbound portion. For 5 mL of resin, I generally use 25 mL buffer for each step so the "trace stuff' really is trace. Also, when you are done, be sure to clean your resin of detergent.
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The proteins will be extracted from tissue samples obtained from human brains embalmed with formaldehyde/glutaralldehyde mixture. I found a couple of published protocols that have been used to reverse formaldehyde crosslinking by heating tissue samples at 70oC for 30-45 min prior to protein extraction, but I did not find any protocols to reverse protein crosslinking with glutaraldehyde.
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I do not recomend that, due formaldehyde fix the tissue but at the same time degradate some proteins
Membrane Fragments Purification
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I need to get purified membrane fragments (e.g. liposomes) from cell culture cells with my desired membrane protein. Does anybody have a protocol?
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Ok...I'll get you a protocol that will essentially involve differential centrifugation and then membrane isolation and washing. You'll need to decided how you lyse your cells. And in general washed membranes can be mixed with glycerol and frozen. Check your inbox. And if whole cell ELISA works, perhaps you're releasing a cross reactive protein or something that is interfering with the ELISA. Out of curiosity, do you have a way to track your POI? At each of these steps it's ideal to take an aliquot and assess enrichment by at least performing a western (if there is say no activity to track).
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I am working on purifying membrane protein expressed in native organism and the purification process requires total anaerobic condition. I use french press to break cells.
The problem I am having is my whole cells show good activity but once I break the cells the activity in cell free fraction is lost. The lysis buffer I use is 30mM MOPS, pH7.3 and I tried protease inhibitor Benzamidine which did not help. Should I add salt to the lysis buffer?
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I haven't done this under anaerobic conditions, but I would explore low temperature, protease inhibitor cocktails and consider gentler membrane lysis such as nonionic detergents or digitonin....
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I have purified one membrane protein for protein crystallization. I am refining the conditions for crystal. What else can I do with this purified protein? I am searching for some new approaches or ideas.
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Wow! A very open ended question, the answer to which depends massively on what you want to know about your protein, and what your protein actually is!
With a TM protein, one might not be able to simply throw it at cells and expect it to integrate with the membrane properly, so sadly this option is perhaps less useful.
However - In vitro functional tests, such as ligand binding assays perhaps?
Biophysical characterisation maybe? SEC-MALLS, AUC, CD spectroscopy, may help you better understand the protein.
You may get a more focussed answer if you provide a little more information about your TM protein - it is a receptor? A transporter?
Does anyone have experience with Millipore's Direct Detect system?
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this could potentially make my work more enjoyable, since I'm doing a lot of Bradfords, but I'm not sure if it's worth the money.
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Hi David, no, unfortunately I haven't bought it yet, since there's noone I know using it and thats a bit odd. It just comes out as an overpriced machine without real contribution...
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I have a cell-free extract in K phosphate buffer pH 7.0 and I want to run Ni-NTA column to purify my protein. The protocol says to use buffers of pH 8.0.
Is it necessary? Can we try a different pH?
What if my protein is not stable at pH above 7.5?
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I would recommend that you use a higher pH than 7: pH 8 is normal for running nickel affinity columns.
The reason for this is that the nickel affinity column relies on the binding of neutrally charged histidine to the nickel ion (via a lone pair on one of the nitrogens). The pKa of histidine (i.e. the point at which 50% of the histidine is protonated on both nitrogens) is 6.0. Therefore, if you run at pH 7, between 5-10% of histidines will be protonated, and will repel from the positively charged nickel. Result: more of your protein will flow through. At higher pH, this proportion will be reduced, and you get far more specific binding.
It is also worth considering the pH of your buffer at the temperature that you will run the experiment at. Most of us make buffers at room temperature, and purify protein at 4 C. Almost all buffers change pH with temperature, and usually pH increases as the temperature drops (this is especially the case with Tris, a common choice for IMAC buffers). You may well find that, at the temperature that you are running the experiment, your buffer is at a higher pH than you think. Many people who use Tris pH 8.0 (RT), like I do, are really purifying at pH 8.6!
As a result, it might not be so surprising that your protein is pH sensitive if you are working at low temperatures.
Hope this helps....
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Does anyone has a good advice how to enrich the lysine acetylated proteins prior to the mass spectrometry analysis? I will be using primary T-cells thus good enrichment step will be essential.
Anti-lysine antibody enrichment is one of the possibilities, but I was wondering whether there is more efficient way to do it? Additionally, which antibody manufacturer would you recommend?
On the other hand, would enrichment using bromodomain linked beads be an option as more efficient way to pull down the acetylated proteins/peptides? I have heard it might be possible, but did not manage to find any paper using this technique...
Any advice or comment is welcome. Thanks!
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I think Immunoprecipitation (IP) with Anti-acetyl lysine antibody (Cell Signaling) may solve your problem.
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Protein purification
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The best would be to make recombinant proteins with a linker for biotinylation. This linker would contain a specific cleavage site for a protease.
After your purification on the beads, "cut" the protein to relase it and to have it ready for concentration, analyses and other controls processes.
Sorry, I know that it's not an easy solution. I hope that you will have a simpler and cheaper way...
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I am looking for a good protocol to purify my proteins extracted from muscles before 2Dgel.
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1:9 sample to -20C acetone. Mix very well and keep for 1 hour to over night at -20C. Spin in the appropriate tube at ~17000xg. Easy if your sample is no more than 150ul so that you can use a 2ml microfuge tube. IF you have a large sample then pick the tubes that you will spin in and do the precipitation in them. I found that TCA wasnt as complete as acetone and damaged glycosylation (if that's important to you). I would wash the pellet 2 times with -20C Acetone to remove contaminating lipids and detergents. I had good luck solubilizing my pellets in DeStreak Rehydration solution from Amersham. This publication describes some of the process "Global analysis of community-associated methicillin-resistant Staphylococcus aureus exoproteins reveals molecules produced in vitro and
during infection" Best of luck.
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I am extracting Lentinan from Shiitake and the method says that I need to remove protein usin the Sevag method. I have searched and the only thing that I have found is the usage of chloroform and octanol, but I haven´t found the exact method
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The exact method is simple. Take your crude lentinan extract and shake hard with 1 volume of butanol/chloroform (1/5) mixture in a 50 ml bluecap tube. Centrifuge the emulsion at 2500 rpm 10 min and take the upper layer. Interphase contains denatured protein. Precipitate the lentinan by adding 2 vol. ethanol and keep at -20 for 1 hr. Centrifugation will give a pellet of partially purified polysaccharide. Pure lentinan is white.
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Our lab purified some soluble protein and kept them at -80, but the concentration of the protein varies when we evaluate them again. Some of them increased while some of them decreased. I wonder if the changes in concentrations resulted from protein solubility? And what factors could potentially affect it?
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Sometimes a freeze/thaw cycle will cause proteins to crash out of solution or form aggregates- both can affect concentration measurements. Adding a cryo-protectant such as glycerol, sucrose, or DMSO can help. You can do a test, freezing small aliquots of your protein in a range of glycerol, sucrose or DMSO concentrations (say 0-20%), thawing the next day and measuring the concentration. This will show you what condition works best for your protein. Also, I always did a high-speed spin after thawing to remove any aggregates that may have formed before I used the protein. Another thought - do you flash freeze in liquid N2 or allow them to freeze slowly? For fragile proteins, flash-freezing may work better than simply placing them in the freezer. Having said all this, I had a protein that definitely behaved better when stored at 4degC. Sometimes you just have to try different conditions until you figure out what makes your protein happiest. Good luck!
Concentration of secreted bacterial proteins using DOC-TCA precipitation?
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I'm trying to analyse the secreted proteins from a culture of bacteria. At the moment I'm trying to precipitate protein from a 50ml bacteria culture supernatant, using a DOC-TCA precipitation. In short, I incubate with DOC (0.2mg/ml) for 30 mins on ice, then precipitate the solution with 10% TCA overnight. Spin down the entire 50ml solution, and then wash with 2ml acetone. I then transfer the acetone wash into a micro-centrifuge tube. Spin down again, and then re-suspend in 50-150uL of 0.1M sodium phosphate buffer + 1% SDS. I run the samples on a 12% SDS-PAGE gel, and the results are often mixed, sometimes the protein bands are pretty good, other times its awful (just 2-3 bands, compared to multiple bands). Here's where I hit a problem: 1. Sometimes after the first centrifugation I find a white precipitate collected at the bottom after centrifugation, but this wont pellet properly. I've tried centrifuging at 20,000 rpm, but it still happens. 2. Another problem I have is that it seems I'm losing a lot of protein when I transfer the 2mL acetone into a micro-centrifuge tube for the 2nd centrifuge step. 3. Also sometimes the pellet wont dissolve very well. Can anyone help with these problems? Or suggest alterations to my above protocol? I've thought about doing the acetone precipitation and chloroform/methanol precipitation methods, but these require larger volumes of liquids. As I have a large starting volume of culture supernatant, it gets awkward when centrifuging.
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If still do you have any doubt, do not hesitate to contact me. Best of luck
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Our company needs to decide which protein purification equipment we will buy that will allow us to develop methods and upscale them for the production of pharmaceutical grade proteins for use in pre-clinical and clinical trials phase 1. We would ideally only buy one machine without the need to change to something else at a later stage.
It is important that this machine and software must comply with the FDA requirements and should be able to produce enough protein for the trials. We are currently looking at the GE healthcare AKTA systems.
Is there someone here with experience purifying proteins for clinical trials? What are you using and what would you recommend, what is important to look at when we buy this?
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HPLC is almost always injurious to proteins, pressures too high; GPC (gel permeation chromatography) is lower pressure but loses the fab advantages of regular HPLC (speed, tight peaks).
You’ll have to follow the US Federal regulations (CFR) for manufacturing a pharmaceutical. In brief, anything that ever touches the compound or is used to formulate it must be assessed, assayed and kept on record. Even for preclinical data you need to validate the equipment before any process. cGMP (current good manufacturing practice) procedures are detailed in CFR Title 21 Part 110 (http://www.accessdata.fda.gov/scripts/cdrh/cfdocs/cfcfr/cfrsearch.cfm?cfrpart=110), and Part 211 (http://www.accessdata.fda.gov/scripts/cdrh/cfdocs/cfcfr/cfrsearch.cfm?cfrpart=211)details production and manufacturing controls; knowing the rules thoroughly before making any purchases is wise. If the system you buy can’t be validated, it won’t be acceptable. For instance, ultrafiltration and column chromatography have their own special considerations; and computerized systems are another hurdle (21 CFR Part 11).
The point is, you can’t just clean up a product and sterilize it at the end: the entire process needs to be GMP. When you’ve narrowed your preference of a good method of purification, ask the equipment vendor if any pharmaceutical companies use that piece of equipment for manufacturing a like product. Do what those who have already succeeded before the FDA do-- your approval speeds through if there is a predicate compound already approved that was made the same way.
The ÄKTAready manual states it fills several regulatory standards including some FDA, and “the ÄKTAready system is biocompatible and hygienic, and meets all GLP and cGMP demands for Phase I-III in drug development and final-scale production.” Coming out of the people who gave us Pharmacia of course promotes confidence. Buying any machine designed with the CFR in mind should mean fewer potential pitfalls for you, so you can develop a shortlist between vendor assurances and the equipment used in other successful approvals. Your regulatory manager or consultant is a good partner in making this choice.
You’re right about the scale-up issue. Using a large-volume protein derived from nature, we used a separation process and proprietary industrial GPC we were able to sterilize and validate every step of the way, never had to switch. But if you don’t need large amounts for trials, the smaller scale may be more economical in the long run—the full-bore system is such a headache and time-intensive to validate all the time for small amounts. And if your compound fails trials…?
Any tannin considerations? A trial period for the system before purchase might be astute.
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I am trying to express a plant membrane protein using pGEX vector in BL21 strain. I could express the protein but all went into the pellet fraction. Should I purify the protein from the pellet fraction (inclusion bodies) or express the protein in another strain like C41(DE3) orC43(DE3)? I need the protein for protein interaction studies.
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Hi Navjyoti,
You can get the recombinant protein to the soluble phase by lowering the induction temperature and induce the protein with less amount of inducer. Lowering the temperature and the inducer concentration can allow the bacteria to grow slowly and can give time for your protein get folded properly and come to the soluble phase. Changing strains can also help you to get your protein to soluble fraction.
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I will be using RIPA lysis buffer for other cells as mouse brain tissue has a lot of fats so my worry is will RIPA buffer work best on brain tissue? Another thing is if someone has any idea which option will be best if I use sonicator or dounce grinder for homogenization of brain tissue lysate?
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I often extract embryonic brain lysate using
50 mM Tris-Cl pH 7.4 , 150 mM NaCl, 1% Triton Buffer + protease and phosphatase inhibitors. I then pipette up and down and spin in a tabletop centrifuge cooled to 4 degrees C at full speed for 10 minutes. This lyses both the cytoplasmic fraction and nuclear fraction.
For adult brain tissue, I first flash freeze the tissue and then use the same process as above.
I have also used RIPA buffer in the past, and this works well. Alternatively, you can use the Urea lysis buffer recipe attached below. This works well for phospho-proteins and other large sized proteins.
Good luck!
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I am purifying a GST-tagged protein, and cleaving the tag with TEV protease. My buffer contains 20 mM HEPES, pH 7.5, 300 mM NaCl, 5 mM DTT. There is 75 mM reduced glutathione around from elution, and I add 0.5 mM EDTA for digestion. I incubated overnight (ratio 1 mg enzyme to 10 mg protein) at 4 degrees.
In the morning, the solution was cloudy. So I spun out the precipitate, and took samples to analyze by SDS-PAGE. Some of my protein precipitated, but most (90%) stayed in solution. The 27 kDa TEV protease precipitated, and this was evident because most of my fusion protein was not cleaved. Another lab member used the same prep of protease (different protein), and had no problems.
Anyone have any ideas as to what is going on here? Has anyone ever heard of TEV protease precipitating?
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As Ladislav said, TEV has been bioengineered to increase solubility/stability. If you know your sequence you can check for the 3 most commonly used mutants. S219V eliminates autolysis. L56V and S135G increase the solubility greatly (up to 40mg/mL as a double mutant). However, without these mutations it will begin aggregating at 1mg/mL. Also, you have 300mM NaCl in your original conditions which will make it precipitate out even faster. So check your sequence and remove the salt. If you only have the S219V mutation, you should keep your TEV stocks at 1mg/mL or less with ~50% glycerol. 
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I've purified a protein through FPLC, but the concentration is lower than what I need to set up a reaction. Any advice about a good kit to concentrate my protein would be of great help. The protein is 53KDa.
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Depending on your sample volume, centrifugal filter concentrators from Millipore, Corning, Pierce etc are a good choice. For larger volumes I prefer to dialyse my sample using Pierce's SnakeSkin tubing followed by lyophilisation (freeze-drying). That way you can resuspend your protein powder in the desired buffer and buffer volume. Moreover, I often observe an additional purification effect through freeze-drying as many contaminating proteins won't resuspend again.
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I am looking for methods to delipidate protein solutions, or protein & cell-suspensions after cell-disruption. These methods should be scalable for use in a large-scale protein purification process. Is there someone who can help me?
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You can try TX-114 phase separation.
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I tried Abcam, but they do not have that. Any suggestions?
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Thanks a lot Dr Stefansson. I was able to get what i was looking for from the link you have provided. Thanks again.
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I am looking at biotinylating a protein in vivo in insect cells where the insect cell media will be supplemented with 4 uM biotin. The protein accumulates in the cytoplasm. The question is: has anyone purified a biotin tagged protein from insect cell lysates using Pierce Monomeric Avidin Ultralink resin (or similar resin)? And did you have to dialyze the cytoplasmic lysate to get rid of the free biotin? I wash the insect cells in PBS to get rid of media associated biotin before lysing them and would prefer not to dialyze the cytoplasmic lysate.
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We purify biotin labeled proteins several ways: dialysis against avidin-containing buffer, ultrafiltration, or column chromatography using affinity labeled support. All depends.
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The antibody (and exact protocol) that I'm using worked in the recent past, but now I get very high background, along with a "ladder" of negative ghost bands against the dark background. My protein of interest still faintly shows up as a positive (dark) band, despite all the non-specific ghost bands. But from my reading, ghost bands supposedly indicate the binding of too much secondary antibody. Does this imply that my primary antibody is somehow preventing binding of the secondary? By the way, the secondary is working fine with other primaries.
I've attached an image of the scanned film: lane 1=protein positive control, lane 2=knockout tissue, lane 3=wildtype tissue, lanes 3,4=experimentals.
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I recently had a similar problem. As is standard with western blotting, there are a few potential causes for the non-specific bands. Unfortunately there is no "quick fix" and it really is all down to trial and error to see what will work best for you and the antibody in question.
1. The membranes aren't sufficiently "blocked" prior to incubation. Do you include any type of blocking agent in with your antibodies? I usually include 2.5-5% non-fat milk or BSA in both my primary and secondary antibody.
2. The concentration of one or both of your antibodies is too high. If I ever suspect this, I first try diluting the secondary antibody. It might also be worth doing some dot blots to find out exactly what the optimal concentrations are for your antibody combination. You never know, the manufacturers may have changed something, or if you are using a different brand of secondary antibody that might also affect the binding properties.
3. You are not washing the membranes for long enough in between incubations and/or before exposing the blots. After my secondary incubation I usually allow the membranes to wash for a couple of hours in TBST at room temp. (changing the TBST every 10-20 min). If I still find that I have lots of non-specific bands, I will leave the membranes overnight to wash at 4 deg C. That usually cleans the blot up enough to get a clean image. If the antibody is particularly problematic, I'll also wash for an hour in between primary and secondary incubations.
Personally, I'd begin by giving the membranes a good wash and if this isn't successful, start again and this time make sure everything is blocked and washed sufficiently. If you are still having problems after that, try reducing the concentration of your secondary antibody.
Lastly, even if the antibody has worked a hundred times before AND with the exact protocol you are using, the western blot god is a cruel mistress. Maybe you're not wearing the right socks on the days it hasn't worked? ;) Good luck!
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My protein is not precipitating but can it still aggregate. Aggregation is not observed when I do size exclusion chromatography (SEC), that is I see a single monomeric peak for my protein. Can aggregation somehow escape SEC? My protein is around 23 kDa.
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1. Protein-protein complexes can dissociate in the column and will elute as a single peak if the exchange is rapid. This usually manifests as a single peak with a trailing edge that shifts position with concentration if the binding is weak. An example can be seen here from Protein Sci. 2002 April; 11(4): 875–882. Amyloid beta is another well known example of this phenomenom Biochemistry, 2012, 51 (17), pp 3694–3703.
2. If your aggregates are very large they will not enter the column at all and the monomer will appear as single peak.
3. Do you have some reason to expect you have an aggregate?
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I have tried to purify membrane protein in HEPES buffer but still there are some other buffers there. Which buffer and other biochemical parameters should be followed for better protein induction and purification. For membrane protein purification, which buffer is the better choice so that I can maintain my membrane protein folding and membrane hydrophobic part?
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Tris or phosphate buffers are useful at physiological pH, as is HEPES. HEPES is a useful alternative to tris and phosphate buffers if you need to add divalent cations such as calcium, or perform labeling chemistry that involves primary amines. However from your question I wonder if you should focus less on buffers and more on the detergents you might try to stabilize your protein. Detergent choice can be quite complicated and a lot will depend on your goals downstream. For instance, digitonin is an excellent detergent for maintaining membrane proteins in a folded state for functional experiments, but terrible for NMR as it is purified as a natural product. It is also pretty expensive. CHAPS and NP-40 (aka Triton-X-100) work for some proteins and not others. Ionic detergents tend to be more denaturing. There are dozens of detergents to chose from and in the end the choice will likely come down to empirical observations on how your protein behaves in each, rather than an ability to predict what will work and what won't.
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I purified my recombinant protein His tag with His-Nickel gel and eluted them with 0,5M imidazole. I confirm with SDS and Western my protein is there and clean. However, whenever I try to do buffer exchange with ultra filtration filter and dialysis I always lose my protein, either by sticking to the filter column or forming aggregates and also sticking to the dialysis membrane. I check that my protein were contaminated with Nucleic acid. Does DNA/RNA contamination in sample protein interfere with protein purification?
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To get rid of DNA/RNA if you got a His-Tag :
- Bind your protein with 20 mM Imidazole (to avoid unspecific binding of proteins).
- Wash with a buffer without imidazole at low salt concentration, about 150mM NaCl (strengthens the interaction with nickel, to avoid protein elution in the next step)
- Wash with high salt buffer without imidazole (usually up to 1 M NaCl) : DNA/RNA are eluted
- Wash again with buffer devoid of imidazole and with 150 mM NaCl
- Make the elution of your protein
This often works very well, but your protein loss problem can come from elsewhere...
Is your protein having a high pI ? Because these proteins are often sticky on most of the surfaces...
Did you try to change your buffer with a SEC-like column (Desalting, PD10...), because this is a more rapid way of buffer exchange, and therefore could reduce the loss...
Hope this will help
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I would like to purifiy my crude protien and protein concentration too low (25ug/ml). I dont know where to start my work.
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Proteins are unique and beautiful--like snowflakes, but probably with a longer half-life. Whenever we purify a protein, we use its intrinsic properties to help us in its purification--sometimes this is easy (I work with folate enzymes, and we can use a folate-based column matrix for our purification) or it can be very difficult (where we then make the protein recombinantly and engineer a certain TAG to the protein). If you let us know the precise protein (or type of protein), we can probably point you in the best direction to go.
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I need to check transcriptional activation of NFkB in rat brain.
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You can try a differential isolation by centrifugation. It´s less expensive a may be adecuate for your purpose.
Homogenate,
Centrifugation 500 x g 2 min, 4ºC. Take the supernatant
Centrifugation 500 x g, 5 min 4ºC. Save the the pellet (nuclear fraction). and take the supernatant
Centrifugation of supernatant 25.000-30.000 x g 45-60 min. Save the supernatant (cytosolic fraction).
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I want to separate rabies virus glycoprotein. Can anybody suggest purification method and protocol?
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Separate rabies virus glycoprotein from what? Do you mean " how to purify rabies virus glycoprotein?"
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We have a recombinant protein (plant virus) that binds RNA. We would like to crystallize this protein, thus we would like it free of nuclei acids. Absorption spectra indicates a significant amount of nucleic acids bound. How can we remove this- we need a protocol that is cheap as the volumes are large?
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Hi Arni,
What is the pI of the protein? I could remove nucleic acids from my samples by Ion Exchange (normally Anion Exchange using HiTrap Q column)! Then the nucleic acids, which are strongly negatively charged, would bind tightly to the column and it would normally elute only in high concentration of salt (say more than 1M salt) whereas the proteins would elute earlier.
It worked very well for me!
Cheers,
Izabella
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Any electrolyte has it's own ionicity - either it's cationic or anionic, also sometimes non-ionic. So how one can find out its ionic charge?
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I don't really understand this question. In solution, both cations and anions from any electrolyte are present. Whether the electrolyte is a strong or weak electrolyte in solution will depend upon its dissolution constant, among other factors.
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Purification of cellulase from a source
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There are many c-sources like u can use banana peels, banana leaf, sugarcane bagasse, ground nut shell, wood chips, corn cob, wheat straw, coconut coir etc. I am working on all the above substrates and the best one is banana peels for bacteria and sugarcane bagasse for fungi. Hope u will be benefited from this.
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For the first time, I am facing a strange problem with a purified PCR product. It is around 1.3 kb and I am extracting it from gel. After extraction, when I reanalyze the purified PCR product, it shows two bands - one at its normal position and the other almost at the dimer position. Can anybody tell me how to prevent this unwanted band? Or can I proceed with this mix population?
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I also suggest you to incubate the sample to 70-90 degrees before applying to the gel. Your amplicon has a lot of GC %?
Can anyone suggest ways to optimize my recombinant protein purification?
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I have been working on protein purification, my protein is induced by IPTG in E. coli but the problem is that it is in inclusion bodies. I have used lysozyme and sarkosyl and Triton X100, I have solubilized the proteins but during its purification, the affinity to Ni-Agarose is affected, so my efficiency is poor. I would really appreciate any suggestions.
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I agree with previous answers - often the solubility of overexpressed protein is the matter of fine tuning of growth and expression conditions. I would too suggest you to try expression at lower temperatures (16 C over night often gives good results). Also, sometimes it's better to use lower IPTG concentration for induction, so overexpression is slower and therefore protein folding better. You can separate your bacterial culture before induction into several aliquots and vary the IPTG concentration (for example 0.1-1 mmol dm-3). 0.2 mmol dm-3 of IPTG usually gives good results. Another thing that could be very helpful is to add a small amount of NP-40 in your lysis buffer - 0.2 % (v/v). Once, this made a miracle for my protein, when even the low temperature and lower IPTG concentration weren't helping.
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Methods in Enzymology
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I agree with Mr. Malih... The best way will be to do a SDS-PAGE (Tris-Glycine or Tricine, depending on your resolution requirement) and stain it with Silver staining method. Since silver staining is sensitive than the coomassie staining, you will get to know even the minor contaminants. I hope this will help you.
How to use alkaline phosphatase to remove the P groups in the protein samples run by Western Blot analysis?
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Western blot analysis
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I have found that Lambda phosphatase works much better than alk phos when dephosphorylating proteins (http://www.neb.com/nebecomm/products/productp0753.asp). It can be used in lysis buffer prior to the addition of the SDS/LDS loading buffer.
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I try to purify a his-tagged protein which is either secreted and glycosylated or intracellularly expressed and not glycosylated (using Pichia pasotris). I am using the Ni-NTA resin from M&N. The unglycosylated protein binds perfectly under denaturing conditions (8M Urea) to the resin (no signal in the flow through fraction), but the glycosylated protein does not (roughly 90% in the flow through signal). I already decreased the imidazole in the binding and washing buffer - no improvement. It seems that the decrease in binding is due to the glycosylation. Any suggestions, how the binding could be inhibited by the glycosylation and what to do to circumvent that? The glycosylated protein has an n-terminal signal sequence and the his tag is fused to the c-terminus. I thought of some kind of stercial hindrance and therefore also used a derivative, that has a FLAG tag as some kind of "spacer" between the protein and the c-terminal His-Tag. That seemed to improve the binding a bit, but still much of the protein is in the flow through fraction. Does anyone have suggestions on how to quickly optimize the binding on a physiochemical basis without any further cloning?
Wow, thanks for all your input so far!
Some additional information:
the glycosylated protein and the unglycosylated protein are exactly the same except of the a-MF signal sequence on the n-terminus for the glycosylated protein. Both proteins have the his-tag at their c-terminus and could be detected with a his-antibody by WB... though i normally monitor the purification with a FLAG-AB. The FLAG tag is at the N-terminus (after the signal siquence for the glycosyalted protein). As mentioned obove i also tried it with the FLAG tag bewteen his-tag and the protein.
I tried binding with 10 mM imdiazole first and after i saw that strong signal in the flow through i decreased it to 5 and finally 0 mM. The wash buffer was 10 mM and 20 mM (there is no signal in the wash freactions). Elution was performed with 250 mM imidazole.
the basis for the buffers tested were all possible combinations of either 50 mM Tris or Phosphate buffer, 0.3 M NaCl, with or without 10 mM 2-mercaptoethanol, with or without 8M Urea (and with the beforementioned imidazole concentrations). pH was 8 and of course i checked lysate and buffers pH values right before their application.
All in all I don't think that it is the imidazole competing for the binding. I was rather wondering if the glycosylated proteins might somhow stick together and prevent themselves from binding to the column.
Deglycosylation is no option, i need it glycosylated as well as unglycosylated to compare both ;)
concerning interfering substances: my protein is to large to pass the yeast cell wall and therefore i am not working with culture medium but cell pellets like i do in the case of the intracellular protein. I allready performed an ammoniumsulfate cut to get a purer starting material and than changed the buffer by extensive dialysis. That didn't help.
I tried batch purification from 1h to o/n, without any success... I am using a 6x his tag
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As I understand you need to purify a glycoprotein. So, change the paradigm of purification: instead of Ni-NTA resin use lectins concanavilin A (ConA) and wheat germ agglutinin (WGA) resins, respectively, as ligands to bind specific classes of glycoproteins. ConA lectin recognizes alpha-linked mannose and terminal glucose residues, while WGA lectin selectively binds to N-Acetyl glucosamine (GlcNAc) groups and to sialic acid.