- Rimma Bachmann added an answer:Please suggest me why I am getting very less amount of my protein of interest compare to tagged GST protein after cleavage with Prescission Protease?
I am purifying a 8 KDa protein tagged with GST and cloned in pGEX 6P 2 vector. When I cleaved 8 KDa protein from GST by prescission protease I got very less amount of 8 KDa Protein compare to tagged GST protein. Cleavage reaction conditions are as follows- cleavage Buffer (50 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1 mM DTT, pH 7.0), Incubation time 4 Hours.
I am also attached gel photo for the same.
Please tell me why it happens?
How I will get more amount of 8 KDa protein?
It would be interesting to know how the western blot of your protein lysates before purification look like. If your Protein restriction would be inefficient, you would expect a third band for your fusion protein. Which you don’t have!
Of course it might be, that the affinity for your fusion protein is lower, as GST alone. If you have chosen higher expression temperature, it might happen that your protein gets degraded. Make an expression test by different temperatures to make sure the conditions you have chosen are the best for your protein.
Another problem of which I might think is, it happens rarely but it happens that you have picked a mix clone, a clone with a plasmid for only GST and one for your fusion protein. Usually the Expression for the GST alone is much higher. This would explain your result, either. If you don`t have any antibody, check your Plasmid DNA by restriction.
Best of luck!Following
- Anthony Harrington added an answer:Can anyone help with purifying crude protein extracts?
Hi, I have a question about crude protein lysates (extracts). A fellow lab member in my lab is trying to isolate a 27KDa protein from here crude protein lysate. She mentioned that this is the smallest protein in her lysate (confirmed with SDS-PAGE). Right now, she is planning on running gel-filtration to isolate it but has yet to start. She did ask other labs at the school that have HPLC/UPLC about running her samples but it seems that something always comes up right before its time to run the sample. I mention that she could use a membrane 30KDa MWCO filter along with a vacuum filtration set-up to isolate her protein. I was curious if this is possible? She is ignoring my suggestions and I just want to know if its because my suggestion makes no sense. Any advice will greatly appreciated and will be passed on to my lab mate.
Thanks for all the replies it really is helping her. A little about her situation, she has cloned a holin gene specific towards a certain bacterium (cannot reveal) into a plasmid that has an arabinose inducer (araC). Yes, I agree, she should have thought it out more and included an affinity tag which she is now considering. Right now, she is using what she has because she will be attending the National ASM meeting and does not have time to start from scratch (It would not take her more than month in my opinion). I guess the gel-filtration method is better and now I know why she has ignored my suggestion about the MWCO filter. Does anybody have any suggestions regarding what type of sephadex she should use (order), medium or fine? I will forward your info to her and maybe she will take the advice and tag her protein in the future. Thanks again, especially to Dr. Ariza and Dr. Halavaty.Following
- Santosh C M Kumar added an answer:How can I get rid of the chaperon?
After Nickel column and mono Q column, there is still chaperon binded to my protein. My protein pI is 5.5, people say Heparin can get rid of the chaperon. But I don't known in a buffer (pH ~6.5), whether my protein can still bind the heparin. My protein is a RNA helicase, I would like to do some ATPase assay, that's that's why I would like to get rid of the chaparon. Thank you.
As a guy working on chaperones, I suggest to use very low salt in your lysis (or loading) buffer and wash buffer. I would not keep any salt. Just use, 50 mM Tris (pH: 8.0 + appropriate imidazole concentration). Salt induces hydrophobicity and thereby chaperone binding.
- Dawid Deneka added an answer:What is a “gentle” method for antibody-antigen release (ELISA)?
Please could anybody recommend a method how to release a protein, which is bound to an ELISA capture antibody? The bound protein is no enzyme (so activity can be neglected), but its structure and posttranslational modifications shouldn`t be changed by the elution procedure. Do you think that change of pH or the addition of detergents, etc would also release the capture antibody from the bottom of the ELISA well? Thanks a lot in anticipation!
If your protein doesn't have any disulphide bridges you can use reducing agent. Antibodies' structure will be altered and your protein will be released.Following
- Dawid Deneka added an answer:Why nikel ion does not bind to Ni-NTA column after APS treatment?
I treated Ni-NTA column with 10% APS because I purified a protein using a buffer containing DTT and it tourned brown. After APS treatment I wash the column with water and then I stripped it with EDTA, wash it again and finally used NiCl2 to recharge column, but when I washed again with water all the nickel was eluted. Is the column damaged?
Adam and Guarav: APS(ammonium persulphate) is a way to remove reduced nickel. When it's brown and reduced it is not possible to elute it simply with EDTA.
It is quite possible that you damaged the resin. I am not sure but I've heard about using 1 or 0.1% APS to clean it. Make sure that you washed away all the EDTA before loading a column with Ni2+. If it didn't work the resin may be simply damaged.Following
- Arshad Jawed added an answer:What is the Extinction coefficient of GST?
I am purifying 8 Kda Protein tagged with GST. My 8 Kda protein don't have any tryptophane. So for calculation the concentration of my fusion protein I need extinction coefficient of GST. Please tell me the extinction coefficient of GST in 50 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1 mM DTT, pH 7.0 Buffer.
Simply use a spectrophotomter with your purified protein.
Tyrosine and tryptophan have Abs maxima ~278 to 280 nm
If you dont have these and still have phenyl alanine try 265 nm
If you even dont have Phenyl alanine read the absorbance at 215 - 220 nm as its Abs Maxima for peptide bond.
Since you already have GST Tag, use purified fusion protein, GST alone and check / screen the difference at various concentrations and buffer strengths.
Do let me know if you have problems.
All the best
- Antonio Ariza added an answer:Concentrating protein using Vivaspin Concentrator takes longer than usual. Why?
I have carried out a His-tag assisted affinity purification of a protein (43kDa) and now I am trying to buffer exchange and concentrate down my protein sample using a Vivaspin 20ml concentrator (30K MWCO) to concentrate down my protein. However it takes too long for the buffer to pass through the filter (1h-1h30min). Does anyone know whether this signifies that something is wrong with the protein sample?
Thanks in advance for your time!
P.S. I know I should have used a concentrator with lower MWCO but I wanted to get rid off a 27kDa protein that co-purifies with my protein.
We produce large amounts of protein for structural studies and they usually need to be concentrated to anywhere between 4 to 30 mg/ml. Concentration runs of one to two days are not uncommon for us, so I wouldn't worry about spending two hours concentrating your protein. We set our centrifuges to 4oC and spin them at 4000 rpm.
As has already been mentioned, resuspending the protein every 15-30 mins is helpful as this stops it from forming a gradient. The protein at the bottom of the concentrator will eventually become much more concentrated than the portion at the top and it can easily crash out of solution.
Also, to reduce the probability of your protein sticking to the concentrator membrane, you should choose concentrators with nylon membranes or at least PES (polyethersulfone) membranes, but stay away from any membrane that has regenerated cellulose in it.Following
- Shufen Cao added an answer:Can anyone help with protein purification?I have purified a protein of 21 kDa by Ni-NTA. The purified protein shows a single band on SDS-PAGE but three bands on Native-PAGE. The protein is a monomer, as confirmed by FPLC. The protein also seems to be aggregating as it does not even cross the half-way mark after the dye has run out on native gel. I need the protein for NMR and immunological in vivo studies and am worried of the two extra bands in further experiments. My protein is not degraded by trypsin hence cannot do MALDI. Please advise.
According to my personal experience, sometime the His tag also has an impact. The truncation of His tag sometime can help avoid oligomerization.
Also the buffer matters, like the PH value, salt concentration and reducing agents lick DTT and TCEP will definitely affect protein state and stability.
- Antonio Rosato added an answer:How can I extract protein from an Agarose gel?
Hi to all,
Is there any method mass-spec compatible to extract proteins from an agarose gel and purify them so that I can identify them by a direct injection in a ESI-MS?
For example if I dissolved Agarose by heating and I performed a zip-tip purification on the dissolved selected band, Would I achieve my goal?
thanks very muchFollowing
- Priyankar Sen added an answer:Can anyone help with an unexpected protein loss during the final steps of purification?
I am attempting to purify a large, His-taged, protein (~300 kDa) that is being expressed in SF9 cells and am experiencing problems in the latter part of the purification procedure with unexpected protein loss (the entire procedure is given for reference).
The SF9 cells are crushed and the supernatant is run through a Ni-NTA column (in 10 mM Imidazole). The supernatant is incubated on a rotary shelf for 45 minutes at room temperature (ideal binding conditions for this particular resin and protein). The column is then washed 3 times with 20mM Imidazole to elute contaminates. The target protein is then eluted using 100mM Imidazole.
At this point, a coomassie blue gel indicates that we successfully rid the sample of most contaminates and have minimal target protein loss. After this confirmation, the protein sample is then incubated with TEV protease or 48 hours at 4ºC to remove the His-tag.
The previous part works well, herein is where the issues begin:
First, the buffer is exchanged to get rid of the TEV protease using 10K Amicon Ultra filters (the new buffer is 5mM Imidazole Native Binding Buffer). The sample is centrifuged down and then diluted two times for this. The now ~30 μL sample is diluted to 5 mL and 0.2 % Triton is added to prevent the target protein from non-specifically binding with the resin and then runs through a reverse Ni-column 3 times.
The collected flow through shows on coomassie blue gel that the protein loss is still minimal, and the purity is ~95-97 % (with a maximum concentration of ~7 mg/mL). However, as crystallisation is the ultimate goal, we need to remove the Triton.
To do this we add 0.03 g of SM2 beads per 500 μL of sample (as experimentally determined) to remove as much triton as possible with as little protein loss as possible. This mix is incubated for 2 hours at 4ºC on a rotary shelf.
Afterwards, the protein sample undergoes a buffer exchange (from 5mM Native Binding Buffer to TBS) is then concentrated by centrifugation with 10K Amicon Ultra filters to a final volume of ~30 μL (this process usually takes about two-20 minute sessions).
Yet, even with a 7 mg/mL stock, the maximum concentration we have been able to reach is ~1 mg/mL. So far, we have eliminated the triton-absorbing beads from being responsible for this unexpected protein loss. We also detect minimal loss of the target protein in the flow-through of the 10K filters. At the moment, we suspect that the target protein is interacting with the filter’s membrane and becoming stuck there.
Any suggestions of how to circumvent, or fix this problem would be greatly appreciated.
have you checked the presence of protein after each step by SDS-PAGE? sometimes BCA or spectroscopic analysis may give wrong result. do you know the pI of the protein?Following
- Ruben Salcedo-Hernández added an answer:How can I purify protein from lysate containing 1M arginine-hydrogen chloride?
My protein is almost insoluble without high concentration of arginine-hydrogen chloride adding to the cell lysate in the expression test. It cannot bind to the Nickel affinity column with high concentration of arginine-hydrogen chloride. Are there any other protein purification methods which can be used in this case?
If your protein is insoluble without arginine, it is possible to do a lysate dialysis using a buffer arginine free, with this condition your protein is going to precipitate (the rest of “normal” proteins will be soluble), after recovering the precipitate you can try to recuperate the protein solubility in your buffer with arginine high concentration. If some material remains insoluble just remove it by centrifugation. Other possibility is the use of a detergent (v.g. Triton X-100), ), in this condition your protein could be soluble without arginine, the niquel column works fine in the presence of neutral surfactants.Following
- Magdalena Schacherl added an answer:Why is my MBP-fusion protein forming soluble aggregates?
I am working with a protein which becomes soluble thanks to MBP. I do not see much expression with GST and also see the contamination of GroEL when I overexpress with GST. However the protein solubilizes with MBP tag but very less fusion is obtained from BL21. In shuffle cells, I see a lot of protein come out in the superntant but they elute in the void volume in size exclusion. When I attempt to cleave the tag, the protein precipitates. I have tried different glycerol and salt concentrations but it does not seem to help. One of the suggestions I got was the use of a eukaryotic expression system. But since it would be convenient with bacteria I thought I should post this question.
The fusion does not bind the affinity column efficiently. There are 13 cysteines in the construct.
I apologize, I did not mean to talk MBP down.
In our hands and in the hands of some colleagues it just did not work.
- Dina Morshedi added an answer:Does anyone purify recombinant alpha synuclein?
When I look at the SDS-page pattern of alpha synuclein in some studies its band appears higher (around 17 KD) than the real place of its molecular weight (14.4 KD). However, our purified protein appears at around 15 KD. The sequence is OK and N-terminal also was analyzed and it is OK too. Dr.Sidhu in "Abnormal migration of human wild-type -synuclein upon gel electrophoresis" showed that ASN migrated in SDS-PAGE unusually, so why would our protein migrate and appear in its real position?
Thank you for answering the question, but my problem is that our protein (purified alpha synuclein) migratesnormally but in other studies it migrates less and localize in upper position( near 17 KD)Following
- Anna Tan-Wilson added an answer:Why does my protein concentration decrease after dialysis?I am purifying a 35 kDa protein, its temperamental and oddly, it crashes in low imidazole rather than in a higher concentration of imidazole. Anyways, I wanted to dialyse it for SAXS analysis at an initial concentration of 5mg/ml. However, after dialysis, when I took it to SAXS, and measured the concentration, it was 0.5 mg/ml. I can't really compute as in to why this happened. Anyone with ideas? Is this normal?
As an alternative to the good suggestions already given, I find the Slide-a-Lyzer cassettes sold by Pierce, now Thermo Scientific, quite useful since it does not require tying dialysis tubing. They come in different mol wt cutoffs and different sizes. http://www.piercenet.com/cat/slide-a-lyzer-dialysis-cassettesFollowing
- Pawan Singh added an answer:Why am I getting two light bands on SDS page along with my protein of interest?I am trying to purify mammalian HSP90 from E.coli Bl21DE3 cells. They are histagged. I use Ni-NTA, followed by Anion Exchange chromatography and then GFC in Sephadex 10/300. My protein has a mol wt of 85 Kda. I am always getting two light bands which are very close to each other and are pretty close to Hsp90 in the gel. I have used Sod. phosphate 50mM and 0.2M nacl pH-7.8 in all the three steps. In the Ni-NTA step, I have done step gradient elution in manual column for 50,100.150, 200mM Imidazole. I get it around 100mM most of it. I pool them and ran AEC using again a step gradient from 0.1-1M Nacl in gaps of 0.1M(0.1,0.2,0.3...) on a manual column. The i concnentrated the samples,ran a gfc but still the two bands are there the very same position. Is there some problem in SDS PAGE running technique or the purification itself? Lysis and Equilibriation buffer contain 20mM imidazole. Washing bufffer also contains 20mM imidazole! Please suggest something. Thanks again!
Hi, I have faced the same contamination problem as some E.coli protein co-purify with the your tagged His-tagged protein. Try Co2+ bead (from pierce) it is more specific than Ni2+ bead. I have tried it personally it works much better than Ni2+. One thing I would like to highlight that yield may get compromise as compare to Ni2+ but that you can manage by purifying with more induced culture.
Hope it work for you.Following
- John M Flanagan added an answer:Can anyone help me figure out why I am losing protein during ion exchange chromatography?I am purifying a protein from a bacterial system and I was able to successfully purify the protein through an affinity column. My protein has a pI of 5.9 and it is stable at higher pH. After affinity I started with IEC using HiTrap Q FF column from GE and my protein is in a buffer of pH 7.5 (50 mM tris, 200mM NaCl, 5% glycerol and 10mM beta ME). However I could not recover my protein after loading the sample in the column. I followed gradient elution from 20-100% of Buffer B (2M NaCl) unfortunately, I could not see any peaks after flow through. I repeated my experiment twice but found no changes in my results (I checked all the samples on an SDS-PAGE). Please let me know if I am going wrong anywhere in the protocol.How well expressed is the protein? If it is at low level and you are using too larger a column you may not see as it will be diluted significantly. You may want to try alternative resins. For example, the EM science tentacle type medias have a very high capacity and allow you to use smaller columns to capture the same amount of protein. Is this a published or a well worn protocol that you are repeating, or are you working this up fresh. Did you try taking running your protein, after the affinity column on a SEC column? It would give you some ideas on whether it was well behaved or not. If it elutes as a sharp peak from SEC (and not in the void volume) it is likely to be monodisperse and you may want to test your IEC column with a known protein since they do not generally disappear if they are in sufficient quantity and well be haved. They may not stick, for example the RSV CA protein, which is highly acidic does not stick to any anion exchange column unless it is unfolded (i.e. pI is not the only determinant in binding) although it elutes as a single symmetric peak from an S75 column. Further, I have several proteins with a low pI but they bind better to cation exchangers then anion, presumably they have a highly organized positively charged patch that interact well with the column.Following
- Hong Yu added an answer:Which buffer should I use for solubilization of hydrophobic proteins?I used a phosphate buffer but protein not dissolved. I attach 1D GEL scan. Bands are not resolved. There is lots of smearing.
The glycerol can be added to sample buffer and gel buffer in 10%, use protease inhibitor in sample buffer .Following
- Romain Kapel added an answer:How to prevent protein aggregation during purification?I am purifying a protein having molecular weight around 65KDa. I want to use it at a concentration of 5mg/ml but at this concentration it gets aggregated. What should I do to prevent its aggregation?
@Bhupesh: thank you for commenting back !
1- Yes, you're right it's a risk...But as long as you don't know the ammonium sulfate concentration required for protein desorption you can't tell. Some proteins are adsorbed at 5-6 M ammonium sulfate and desorbed at 0.5-1 M...It is just hypothetical, it will depend on protein hydrophobicity, stationary phase polarity,...
2- Ultrafiltration through microporous membranes. With an appropriate membrane cut-off, proteins can be concentrated (without loss if they are totally retained by the membrane) without changing buffer composition (if microsolutes composing it aren't retained).Following
- Nipuna Parahitiyawa added an answer:How to determine nitrate reductase activity in bacterial supernatant?How do I perform an enzymatic assay?
Redox sensor Green is a reliable marker of bacterial reductase activityFollowing
- Nattha Ingavat added an answer:Could imidazole make protein crash out?I purified my protein using Ni-NTA as a first step. I eluted my protein with 25 mM bis-tris buffer, 50 mM NaCl, 300 mM imidazole, 0.5 mM TCEP and 10% glycerol. My protein was soluble in that final solution. However, it crashed out after I concentrated it down using a concentrator (centrifugal unit). I wonder if the imidazole could cause protein precipitation when the protein gets more concentrated or when the concentrating process is harsh? Before this, I tried dialyzed and concentrated this protein using the same buffer mentioned above, except no NaCl and no imidazole, and no precipitation was observed.
Thank you so much for all comments..Following
- Valeriya R Samygina added an answer:Can anyone help with buffer exchange issues?
My target protein is extracted in PBS, is eluted from column in DAP and neutralised with Tris-HCl, then dialysed against PBS resulting in 4mg/ml. However as I want to do an initial crystal screen I used a pre washed centricon to exchange buffer to Tris and subsequently have a much reduced concentration (1.4 mg/ml) and have lost activity. Any suggestions please? pI is 4.5, Tris & PBS are @ 7.2 pH, could it be to do with the molarity of the tris (10 mM)?
additional remark: you can not exlude protein loss by unspecific binding of centricon filter membrane. Certainly, buffer is very important as well as salt concentration, but may be you can try to use dialysis on the last step, just to be sure...As far as I understand you used centricon only at the last step?Following
- Ananda Ayyappan Jaguva Vasudevan added an answer:Does using Urea in the purification of a Strep tagged protein from an insoluble fraction influence the binding to the resin?
I would like to purify a Strep tagged protein from the insoluble fraction and I would like to know whether using urea could influence the binding to the resin.
HIS tag might be the better option...?Following
- Keneth Iceland Kasozi added an answer:Protocol for total protein extraction from various mouse or rat tissuesI 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.
I am impressed by the feeds because i have to extract RNA from the myocardium for my research. Greetly appreciate all the above feedz...Following
- Mohammad Asif Shah added an answer:Can anyone help me with low protein recovery in gel permeation chromatography?I have been trying to purify a protein (an enzyme - Mw: 59K Da) by gel permeation on FPLC. This protein was pre-purified by affinity chromatography, then injected into FPLC at the conc. of 1 mg/ml (injection volume = 1 ml). Then, I collected the eluted fractions which contain the interested protein, next, I re-concentrated these factions back to initial 1 ml, then reinjected into FPLC. This time, the UV detector (at 280 nm) showed sharply reduction in detector response (150-200 times) comparing the first injection. Enzyme activity measurement by spectrometer of re-concentrated protein also showed massive reduction comparing with the sample before inject. Can anyone give me tips or explanation to solve this problem. Thank you very much in advance
Loss in protein recovery in gel permeation chromaatography is not unusual. We have loss of around 40-50% in a typical gelfitration experiment owing to diluation during the run. The only remedy is to do more runs/load more concentration of the protein and get enough of protein for the desired study. Also calculate the specific activity of two runs and compare the fold purification.
Best of Luck..Following
- Deepti Agrawal added an answer:I am unable to decipher the gel side of the IPG strips purchased from Bio Rad. Can Anybody help in this regard?Earlier I had used Serva IPG Strips in which it was easy to decipher the gel side of the strips but recently we shifted to Bio Rad and here we are encountering this problem. Your valuable suggestions are required.Thanks Dinesh .....Following
- Prasanta K Ray added an answer:Can anyone please help me in interpretation of an ion exchange chromatogram?During my experiment, I remove a 7Kda protein from affinity column after its cleavage using prescission protease in a buffer containing NaCl ,Tris and DTT. After running the removed protein on SDS-PAGE gel, two protein bands are detectable, one narrow band belong to GST and the other sharp band belongs to 7Kda protein. But after injection to cation exchange chromatography (elution buffer NaCl 1M, Tris 50mM), two peaks are observable (attached file), one shows my protein but what about the first one? Can anyone please tell me what it belongs to?Please elute the first peak material and test for GST separately and individually and see what happens.Following
- Aparna Mitra added an answer:Too many bands in western blot for Egr1 protein, what should I do?I am trying to identify Egr-1 protein in H9C2 cells following hypoxia and reoxygenation. The band of interest should appear at 82KDa. But every time I run the cell lysate in western blot, I get too many bands for Egr-1 protein. However, the other proteins on the same membrane are fine and gives single band ( I cut the membrane). I lyse cells in RIPA buffer with protease and phosphatase inhibitors. I lyse on ice for 30 mins, vortexing in every 10 minutes and store the lysate in -20C. Prior gel run, I add laemmli buffer and BME, mix by vortexing and then boil the samples for 10minutes at 90C. Then I vortex again for 2 seconds. Then I rapidly cool the samples on ice and then spin them. I tried 5 minutes of boiling at 95C but results didn't change. My primary antibody is rabbit polyclonal IgG from Santa Cruz and I tried dilutions from 1:200, 1:500, 1: 1000, but in all occasions it gave too many bands. My secondary is goat anti rabbit from Biorad and I use 1:5000 dilution. Both primary and secondary antibody is diluted in 5% skim milk TBST solution. I block for 30mins with 5% skim milk TBST solution at room temperature. I checked non specific secondary binding by using a secondary antibody control (without primary antibody incubation) but there is no non-specific binding from secondary antibody. Please suggest a solution. Also, I loaded 25ug of protein in the gel.I suggest you block for 30 mins after incubating with your primary antibody and then wash and add secondary antibody.Following