- Vincent P.K. Titanji added an answer:Why does the specific activity of my enzyme decrease at each step of purification?
Can anyone provide possible reasons why the specific activity of my cellulase (from bacteria) keeps reducing after each step of purification? So far, the crude enzyme has the highest activity after ammonium sulphate precipitation and dialysis.
- The most common reason for loss of activity is enzyme instability.you should therefore spend some time testing conditions to render the enzyme stable during purification.First of all eliminate the obviouS factors like proteolytic activity that could destroy the enzyme as
- Selva Chemist (Bio) added an answer:Do you have any suggestions to overcome this membrane protein purification (expression) problem?
We are trying to purify a membrane protein expressed in BL21. We tried both N and C terminal his tag, but the protein doesn't bind well to the Ni column. Along with the protein of interest, many bands are appearing in the elution. Strangely, the protein doesn't have (or very poor) UV absorbance. Also, I don't see any color change when I add SDS sample buffer to this protein, but still see a band in SDS-PAGE. I tried Co instead of Ni, and got a band smaller than expected, but it didn't show activity. I think something is happening during the purification because I see enough protein in the membrane suspension. I use DDM to solubilize it from membrane because we want to crystallize the protein. Proteins pI is 9. 2, buffer pH is 8 in all cases (from cell lysis to purification). I am new in membrane protein field. Any advice would be appreciated. Thank you.
Dear Puunaa Dashnyam,
Te main problem because of it's -N-terminal blocked during your process, means the terminal reactivity - the active functional group reactivity is neutralized by aggregation. so that it have very less binding. and also UV absorption not efficient.
Your protein is in membrane are you sure about this, you just try to purify it by using urea and then reload it. and make sure about it solvation before starts. if solvation analysis give good result then you do. it posses strong aggregation and functional group terminal reactivity blocked so that you have receive such a problem concentrate in this.,
Hope You Got.,
Good Luck & Regards
- Chanakya Nugoor added an answer:How do I get good yield of eukaryotically expressed proteins?
i wish to express a protein eukaryotically in good quantity?
which system i can go for? adeno viral vectors are good choice whether these vector have fusion tags to purify?
(except yeast expression systems)
there are plenty of systems to express. the question is which eukaryotic system even other wise than Yeast, how much quantity you need and what is the application . depending on that the question can vary. I assume you want a mammalian protein and express in mammalian system. then Life tech has those CHO vectors which can you give you good expression up mg quantities. if you need industrial scale, there are ways to improve it especially with MAR elements in the vector which can give you gram quantities. TO come to the purification, there are several purification but most popular is His tag owing to its ease in purification. there is no need to have a vector these days to go for tag incorporated vector. one can simply synthesize it these days especially Gen script or Gene art does it .Following
- Adam B Shapiro added an answer:Hi, can anyone suggest how to stabilize a plant protein in purification steps?
I am trying to purify a plant dehydrogenase, at 65% ammonium sulphate saturation I am retaining all the activity in pellet. But I am loosing all enzyme activity after dialysis. and I am not able to see any bands in SDS-PAGE also. Can any one suggest the solutions for the above problems? As it was unstable at 40c I have 100mM KCl+ 30%glycerol+ 1mM DTT+1mM EDTA+1mM PMSF, pH 9 in my lysis buffer. I have tried 7.5 % to 15% SDS gels. I am getting bands with positive controls . I don't know why no bands are in my sample.
To skip the dialysis by using gel filtration to remove the salt, keep the sample volume as small as possible. Since you don't know the mass of the protein, you may want to try gel filtration media for low and high ranges of molecular weights. Do everything at 4oC and keep fractions on ice.Following
- Mohd ZIAUDDIN Ansari added an answer:Can anyone help me with the problem in Protein expression and lysis buffer?
I am expressing 6xHis tagged 8.5KDa human Protein (pI~4.19) which cloned into pET21b expression vector and express in BL21 DE3 Codon plus strain. I see expression when I mix bacterial pellet with SDS Gel loading dye(Gel loading buffer contains 0.05% bromophenol blue , 40% sucrose, 0.1M EDTA (pH 8.0) and 0.5% SDS) and heat at 95°C for 5 miutes and then load on 15%SDS PAGE. But when I lysed it with lysis buffer (500mM NaCl, 20mM Tris pH 8.0, 2mM Benzamidine and 10 µg/ml Soyabean Trypsin inhibitor) and then sonicate it with pulse 2 Second ON and 12 second OFF For 6 Minutes with 32% amplitude, I am not able to see expression in supernatant as well as in pellet.
I also purified it with Ni-NTA beads and elute with 150, 250, 350 and 450 mM Imidazole but nothing was detectable on gel.
I also monitored OD of Induced and Uninduced culture for see whether my protein is toxic to cell or not and Uninduced culture grow at double rate compare to Induced culture.
Please suggest me-
1. Is my protein toxic to bacterial cell?
2. Is there any relation with pI of protein and pH of lysis buffer? As my protein pI is 4.19 and I am using lysis buffer with pH 8. If yes then what will be suitable pH for lysis buffer and its composition?
3. Whether my protein is express or not?
All gel picture and OD data is attached with word file.
@Arbind Kumar Sir I had already checked induction with 1, .5 and .2 mM IPTG conc. and OD at 25 degree and 16 degree. Gel image also attached with this for the same. please have a look on that and then please give me suggestion. I also tried purification of same protein with GST tagged but after tag removal I don't got my protein thats why i cloned it in pET21b VectorFollowing
- Jonathan Elegheert added an answer:What is the best tag to purify membrane protein?
I'm trying to purify membrane protein from HEK293 cells for x-ray structural determination.
I added the his-tag at the c-terminus of the protein, but the eluted protein from the resin contains huge amount of non specific proteins.
I confirmed that the protein is active.
I asked about his-tag purification on this website before and understood that his-tag is not suitable for purification from human cells.
Now I'm wondering what is the next tag I have to try.
The tag must be small peptide because I don't want to digest the large tag with protease.
Please give me suggestions.
there are many tags that are suitable for membrane protein purification. We have had good results in our lab with the "1D4" tag (C-terminal TETSQVAPA nonapeptide) and anti-1D4 Ab purification. See e.g.;
I like to use the N-terminal Twin-strep tag with Strep-tactin resin as well, however this tag would have to be cleaved off;
The His-tag is not unsuitable, it is my workhorse tag for purifying soluble proteins from eukaryotic DMEM medium. Stripping of the matrix was always a problem though. But GE now markets a Nickel Sepharose "Excel", that is totally resistant to stripping by small molecule chelators in the medium.
You could also use e.g. a deca-His (His10) tag instead of the normal hexa-His (His6) tag. This would allow you to do more stringent washing of the beads before elution. Also, Cobalt resin (e.g. Talon) retains much less contaminants than Nickel resin.
Hope this helps,
- Mona Al-mugotir added an answer:What is the best way to measure the concentration of a 111 kDa protein-25mer DNA complex mixed at 1:1 ratio?
The protein is the heterotrimer replication protein A (RPA)
- Adam Zlotnick added an answer:How can I remove bound DNA from purified protein?
We have purified a human protein by expressing it in E. coli. We recorded spectra and it looks like it has DNA contamination. How can we run the purified protein on agarose gel so we see some bands? When we digest with DNaseI, its not getting digested but when we digest first with proteinase K followed by DNAseI, its getting digested. So how can I get rid of the bound DNA during protein purification itself? Can I conclude the interaction is non-specific?
High NaCl will weaken electrostatic protein-nulcleic acid interactions. However, if you don't separate the two components, they will re-associate. Thus binding the complex to an affinity column and washing with high salt can work. Another strategy is to displace the DNA/RNA from the protein by binding it to something else. High MgCl2 will bind and precipitate DNA and RNA so that you are left with nucleic acid depleted protein left in the supernatant; for precipitation you want long nucleic acids, do not nuclease treat before purification.Following
- Adam B Shapiro added an answer:Is there any method of protein extraction wich does not require dialysis?
I have worked with salt precipitation but in that we need to go for dialysis. I want to ask if any other method is available that would help me skip the dialysis and still get lower contamination in terms of salts. can TCA or Cold Acetone Precipitation Work?
The method you choose depends on what you want to be able to do with the protein after extraction. TCA and acetone precipitation will denature the protein. You could use it for SDS-PAGE, Western blots, or as an antigen, for example, but not to measure enzyme activity.
If you want to purify the protein in an active form, starting with an extract precipitated with salt (are you talking about ammonium sulfate or a chaotrope like NaBr?) but without using dialysis, start by centrifuging the precipitate, resuspend the pellet in a buffer, and perform gel filtration chromatography, using the same buffer to elute the column. This will separate the protein from the residual salt and also fractionate the proteins in the extract by size.
If you are not interested in purification but just want to get rid of the salt, you can use gel filtration spin columns (also called desalting columns) instead. Another way to do it is by ultrafiltration, where you use a membrane to concentrate the protein. In this case, you concentrate the dissolved protein, then re-dilute it with a buffer. You can repeat the process to further dilute the residual salt.Following
- Shridhar Chougule added an answer:How can I express recombinant protein with molecular weight lower than 10 KDa in E.coli?
I have been trying to express a 8 KDa recombinant protein in BL21, But so far has not been successful. If possible. Please guide me.
if your gene contains some rare codons (for which t-RNA is not present in your host), then try expressing your protein in another host which facilitates translation of that codonFollowing
- Yakup KOLCUOĞLU added an answer:What is different between N or C terminal Histidine tag for protein purification with Ni column? which one is good for protein purification?
N terminal or C terminal?
İf you have whole sequence your protein and can be modeling, prefer the terminus that outer part of the protein or close. Otherwise you should be tested.Following
- Mansoor Azeem Siddiqui added an answer:Can someone help me with Protein Purification using Ni-NTA?
I am working on a protein, that I tried to express in BL-21 cells. This protein is an integral membrane protein, so I tried harsh buffer conditions to take out this protein in soluble form. The gene responsible for this protein has been cloned in pET28 vector. I am using Ni-NTA to purify this protein, but I am not able to purify it. After purification, I am getting three more proteins of 45, 47 and 47 kDa, with my protein, that is 41kDa. I have used 500uM imidazole for elution, but I didn't get any protein, so I tried 250mM imidazole for elution. Please suggest me, what else I can do to purify it. I can not go for western blot, as we lost this protein primary antibody.
First you must try to optimise conditions of Culture growth and induction etc.Try to induce at lower OD values (.5), Use .1 mM IPTG final conc. and low temperature post induction 18-20 and pots-induction period should also be shortened.
Check if your protein predominantly goes in soluble fraction or in IB .Also ensure proper sonication.
Second, use high concentration of NaCl ( you can use anything between 350mM to 500mM in binding buffer to minimise protein protein interactions .Do equilibrate Ni-NTA resin with 5-6 CV of binding buffer before loading protein.
Do through washing with Binding buffer beforeFollowing
- Shaban Ahmed Ali Abdel-Raheem added an answer:Can tyrosine intrinsic fluorescence be detected at an emission wavelength either than at 307nm?
I'm doing fluorescence spectroscopy on my protein expecting to see the intrinsic fluorescence of tyrosine and also its effect with addition of cu(i). The excitation wavelength was 280 nm while the emission wavelength were from 300-500nm. From my reading, it supposed that tyrosine will have an emission peak at 307nm. However, in my case, an emission peak appeared at 380nm. None at 307nm. What can that be? Is it still tyrosine or something else? My protein is in MOPS buffer pH 7.
- Steingrimur Stefansson added an answer:Does anyone have a suggestion to remove LPS from recombinant proteins, If the recombinant proteins bind to LPS?
All proteins are not in the inclusion bodies (all of them in the soluble fractions). Proteins were previously purified by His-tagged cobalt resins. We have used the Pierce High Capacity Endotoxin Removal Resin to remove LPS from three recombinant proteins, but the recovery of the proteins appear to be proportionally dependent to LPS concentration in the final sample (LPS measured by LAL Chromogenic Endotoxin Quantitation Kit) - much low recovery of recombinant protein (Quantification of protein by BCA protein assay).
Hi Fernando, the Pierce High Capacity Endotoxin Removal Resin is immobilized poly-lysine. It essentially is an anion exchange column. LPS is negatively charged and binds to this resin.
If your protein binds lipids, and if the only lipids in your prep is LPS, then you will lose a lot of your protein.
Possible solutions: if your protein binds lipids, add octyl glucoside to it before putting it through the LPS removal column.
If your protein is negatively charged and does not bind LPS, you can try TX-114 phase separation.Following
- Muralidhar Reddivari added an answer:What are some strategies to stabilize purified proteins?
I am planning to purify resuscitation-promoting factors produced using an expression plasmid in an E. coli expression strain. The proteins inactivate shortly after purification.
I have explored techniques such as non-aqueous co-solvents and glycerol additives but was wondering if there is an optimal strategy to stabilise the protein for long term storage whilst retaining enzymatic activity.
try avoiding gel filtration step till you actually are going to use the protein. in our experience gel filtration results in loss of co-factors which stabilize protein or activity. just do affinity [tagged protein] followed by rapid ion exchange. also set up a 96 well plate with different buffers and check protein for activity under different buffers.
you are sure to find the right bufferFollowing
- Tami Coursey added an answer:How can I elute my protein from a native antibody without inhibiting protein activity?
I have an antibody against my protein of interest that I would like to use to help with purification. Unfortunately I haven't been able to find a lot of information about how to separate the protein from an antibody without damaging the protein itself. I need to purify this protein for further activity analysis. I've tried tagging it and following it up with elution from an HA column, but I haven't been able to successfully elute it from the HA column without using harsh conditions. This is an alternative I'm looking into. Any suggestions?
Thank you all for your suggestions! I think I'll try my hand at a few of these and leave an update on the outcome. Thanks again!Following
- Borbala Gesser added an answer:Can you recommend useful protocols for removing bound DNA that co-purifies with recombinant proteins?DNA enzymes and binding proteins expressed in E. coli often have bound DNA or RNA that can be troublesome to remove completely for biochemical experiments and crystallizations.
You can also from the beginning dissolve the nuclear fraction in Tris buffer with 7 M Urea. You can centrifuge down DNA (14.000g) and thereafter dialyze the supernatant for pure water. After 5-6 changes of water, you can freeze dry the sample and redisolve the proteins in a new buffer for gel electrophoresis. You can also try to separate the sample in the Tris buffer with 7 M Urea, on a gel filtration column from the DNA.Following
- Peter D G Dean added an answer:How can I detect or remove Nickel Hydroxide from my protein sample?
I cleave my protein from its vector using Ni sulfate under high temperature and PH, after lyophilizing. My protein came out yellowish, and it should be white, so I assumed that there are some traces of Ni hydroxyde that might have been made under the harsh basic cleaving conditions. Now, I want to get rid of it, or even prove my theory. So I'm looking for a way to detect Ni hydroxyde, or to remove it from the sample.
dialysis after adding EDTAFollowing
- Savita Shah added an answer:How can I fix unknown lines in SDS-PAGE electrophoresis?
I take a SDS-PAGE electrophoresis gel. It appears some lines which I don't know what they are. The line connect all wells of gel and they are more than two.
What's it and How to fix my trouble?
Please help me.
Did you wash your cells/plate (if the cells are attached) before you lysed the cells to isolate your protein. Since we use pretty high conc. of FBS (10%- which is really BSA) in the media, if you do not wash your plate with PBS, you will always carry over this protein in your cell extract..When you repeat this expt. , wash you cells/plate throughly with PBS and then proceed for protein extraction..You should see this band (almost)goneFollowing
- Calvin S Leung added an answer:For Co-IP experiments, is it more ideal to have more bait or prey protein?
I have been trying to do a Co-IP between Protein X and Protein Y. I tried pulling down for X, which is highly expressed in the cell, and probing for Y, which is not expressed as much, and I am getting a very weak signal in my western blot. Would it make a difference if I did the reverse IP?
Thank you for all the helpful replies! I am doing the reverse IP and will know the results soon.Following
- Florencio E. Podestá added an answer:Can anyone suggest how to remove imidazole from the purified protein samples?
There are two protein which i am trying to purify for my in-vitro assays.
1. A his tagged protein which i have eluted in 50 mM NaH2PO4, 50 mM NaCl and 250 mM imidazole. Now what buffer combinations should i use to dialyse these protein which i plan to use in subsequent assays.
2. A S- protein tagged which was eluted in 3M MgCl2, what buffer combination should i be using to dialyse the protein for downstream applicability.
If anyone has a better protocol can you kindly share it with me. Thanking you for the help.
that depends on your sample size.
We have been using as a routine the Penefsky method for desalting small volume samples. You can desalt 500 microliters per columnin a matter of minutes.(Penefsky, H.S., 1977. Reversible binding of Pi by beef heart mitochondrial adenosine triphosphatase. J Biol Chem 252, 2891-2899.) If this is your case and you have any doubts regarding the method you can contact me whenever you want.
Higher volumes will require the columns mentioned above or one you can prepare yourself with Sephadex G50 or G25, depending on the size of your protein. The higher the G number the faster you´ll go. A pressurized membrane concentartor will do the job as well. I prefer any of these methods to dyalisis.
- Adam Wei Jian Soh added an answer:How are conditions for bacterial cell sonication determined?
Hi guys, I am currently optimising the expression of a protein that is found predominantly in the pellet. However, I am unsure how the sonication condition can be adjusted such that I have a larger amount of that protein in the supernatant, which contains about 10% of my protein of interest. May I know if the dry weight matters? By the way, I am using a sonicator that relies on a certain percentage of sonication output. The condition I have been using is 70% output for 10 sec pulse and 10 sec rest. (4times). Not sure if this helps but thanks in advance for any suggestions.
Hi Shridhar, I do see dark brown speckles when I pelleted those sonicated cells. Will play around with the above mentioned suggestions. Thanks!Following
- Yasser Gaber added an answer:Which laboratory offers protein purification service and sequence identification for a fee?
What are trustable commercial laboratories doing protein identification and purification.
thanks J. J. Fung , interesting!Following
- Sandip Rath added an answer:Can we use protease inhibitor instead of PMSF for protein extraction from FFPE tissue samples ?
Can we use a cocktail of protease inhibitor instead of PMSF for protein extraction from FFPE tissue samples ?
Thank you all for the answers.Following
- Victoria Shingler added an answer:Why do I get a smear when using a TNT cell free expression system to express my plasmid?
I am trying to express my protein of interest in a TNT cell free expression system (Promega, L1170). My protein is tagged With V5 and His tags. The problem is when I run the samples in a western blot, though I can see a band of the proper molecular weight (about 37 kDa), I see a smear under it. I attach a picture so you can see what I mean. In the lines 2 and 3 I am using the anti HA ab, and in 4 and 5 the anti His. (The first one is a marker and in the last two I am using an antibody against the protein that does not seem to recognize my tagged protein very well).
I followed the instructions of the manufacturer. I also tried adding protease inhibitors, but I get the same results.
Anybody has any ideas?
The double band may also indicate that you have an internal methionine that serve as a start point for translation.
This could only work as an explanation if the both your antibodies give double bands ad your tags are C-terminal fusions.Following
- Tengfei Liu asked a question:How can I elute the chitin bind protein from chitin beads?
Thanks in advance.Following
- Nausheen Jamal added an answer:Protein folding: How can I refold proteins from inclusion bodies in a single conformation?
I want to purify a recombinant protein from inclusion bodies. After the re-folding and dialysis step, the protein elutes in two distinct peaks on ion exchange chromatography. First peak has poor yield but good purity (@ low salt concentration) and the second one has good yield, but the fractions are contaminated because this elution is at high salt concentration.
Can anyone suggest how to resolve the issue?
Thank you everyone for your careful responses. Sorry I did not get back to you with my observations earlier.
Trying long incubation hours after the refolding step worked well. Much protein was collected at low salt concentration, and it is good in quality as well, ie; i have better purity, activity and yield. Perhaps sufficient incubation time after the refolding step can make a difference.Following
- Nazlı Ezgi Özkan added an answer:Has anyone encountered different results when extracting a membrane protein using a hypotonic buffer and NP-40 or a needle-based lysis?
We recently encountered a weird difference between two extraction methods that should yield the same results, and we could not find an explanation:
1. Cell lysis was performed using a hypotonic buffer with 0.5% NP-40 followed by centrifugation at 16,000g for 15min (in a cooled centrifuge).
2. Cell lysis was performed using a needle (25G) followed by centrifugation at 16,000g for 15min (cell lysis was verified).
In both methods we get no detection of our protein in the supernatant (cytosolic fraction). However, when we dissolve the pellet in triton-based buffer our protein is extracted only in method #1. We know it is in the pellet in method #2 since it can be extracted using SDS.
We could not explain why triton only extracts our protein in method #1, so maybe someone here would have some idea.
Thank you in advance.
As I understood, you don’t have any detergent in method #2. In my opinion, you might be lysing the cells but, without detergent, membranes might stay intact and form micelle-like structures, in which you hydrophobic protein is protected from triton.Following
- Fortunate Mufunda added an answer:Which way can the presence of a di-sulfide bond in a protein be detected?
I want to isolate the protein and look for disulfide bond in that protein?Is it possible to do it on SDS-PAGE without b-ME? Or are there any assay for it?
Mass spec, NMR and IR spectra are the easiest and look for the relevant peaks against standardsFollowing