- Marcia Moss added an answer:Can I do a pull down of my protein from cell line overexpressing my protein and check whether it interacts with cholesterol?
My cell line is HEK-293
I would use any commercially available cholesterol detection kit after the pull down. Abcam sells one. They use cholesterol oxidase to detect the cholesterol. Ab65359.
Others sell ELISAs. For ease, I would use the a ABCAM kit.Following
- Vipul Kumar added an answer:What is the optimum amount of protein that should be loaded on SDS PAGE to exactly quantify it?
If I load high amount of protein on SDS PAGE and do densitometric analysis (I am using BIO RAD's Image Lab tool) the relative intensity/amount of other bands increases to higher extent than the desired protein. However if I load a low amount, other bands almost disappear giving me higher percentage of my protein. So what is the standard amount that is loaded to do a relative quantification.
Thank you all for your useful comments. I will follow them and get back if I still get some confusion.Following
- Sophie Briggs added an answer:How can I correct unequal loading in Western Blots?
I am currently doing western blots with BTIC cells (Brain tumour initiating cells) which grow as neurospheres. I have done western blots before (with monolayer BTIC's) and had no issues whatsoever. However, I cannot seem to get equal loading on any westerns at the moment. I have changed pretty much everything about my technique bit by bit and still no luck. I was just wondering if anyone has any suggestions as I am at a compete end now and not sure what to try.
So you have a rough idea of my technique, I wash my pellets in PBS, lyse and sonicate, spin down and measure protein using bradford assay (biorad). Then add 40ug protein with ddH20 and 4xSDS loading dye (40uL total), spin breifly, boil for 5 mins at 95 degrees, spin breifly again and load onto my gel.
Any help would be greatly appreciated. Thanks.
Variations are quite substantial, often more than 50% difference.Following
- Arthur Wing Hang Li added an answer:Any advice with my His-tag protein purification with talon affinity column giving lower yield at larger sample volume?
We have encountered something strange. Our group is purifying an N-tagged 6xhis protein with Talon 5ml prepackaged column. When we first tried with 2ml sample volume, everything worked great. Then we tried 8ml sample volume (4x), and we expected a 4 fold increase in protein yield. Indeed the result was as what we thought.
However, when we apply 15ml sample volume, the protein yield is about the same as what we get with 8ml sample volume... It was about 1.5mg of protein, Theoretically, Talon column is able to bind 20mg protein per ml resin, so there is no way we were reaching that limit.
Our binding buffer: 50mM PBS, 300mM NaCl, which was used as cell lysis buffer with additional PMSF.
Wash buffer: 50mM PBS, 300mM NaCl, 5mM Imidazole
Elution buffer: 50mM PBS, 300mM NaCl, 150mM Imidazole
No EDTA was involved at all. And we don't think it's the buffer issue otherwise it would not work with 2ml and 8ml sample volume, right?
Please help! Thank you so much!
PS we really don't want to purify under denatured condition, so let's just skip the detergent part! :)
Like what the others said, protein yield varies from prep to prep. For one of my proteins, I went from 200 ml to 2 l, and still only got very little increase in the yield. What are the volumes of your culture? As you go up in volume in a fixed container, the surface area to volume ratio decreases, and it affects the aeration. Grow your cultures in a number of small flasks instead of one big one would help.
- Olivier Danot added an answer:Can I use TCA precipitation to concentrate proteins for SDS PAGE ?
I want to concentrate my proteins using TCA, and do 1-D SDS PAGE. I have a few concenrns.
i) Will the TCA remove salts or concentrate them as well
ii) For resoulubilizing precipitated proteins, is the SDS loading buffer the only thing i could use. And if i use it then what should be the volume. Is there any other SDS containing buffer to resolubilize proteins
iii) how will i quantify protein concentration to be bale to load equal amounts in each well
iv) Would any other precipitation method , (e.g Acetone) be a better option
Kindly let me know. And if possible provide references for proper protocol
Thank you so much :)
Yes you can use 16% TCA to preciptate your proteins (15 min at full speed in an eppendorf centrifuge at 4°C is generally enough). It won't precipitate the salts (it can be used to get rid of salt that would perturb the migration).
In some instances (when you precipitate too much protein at a time) you can have trouble to get rid of the TCA that is caught within the pellet, so that when you redissolve your protein, your sample buffer will turn yellow because of the acidity of TCA (if it contains bromophenol blue) because of the acidity of TCA (even after the acetone wash). Then you should add 10 mM NaOH µl by µl to neutralize the acid until the samples turns blue. Also drive your sample buffer all over the pellet several times to redissolve it.
Also, I have observed that once precipitated by TCA, proteins have a tendency to degrade, so load your gel on the day you do the precipitation, and do not reuse samples once they are precipitated (aliquot before precipitation if you think you will need to load the same sample several times)
I am not sure I understand your question about protein concentration. Either you can normalize your samples from the beginning of the experiment (i. e. load quantities of extracts corresponding to the same amount of cells) or you measure the protein content by Bradford assay, the method of Lowry, or similar methods. If your concern is that the efficiency of the precipitation might be irreproducible, you can just add the same amount of a purified protein like BSA in all your precipitation. Anyway, the TCA pellet sticks quite well to the tube, even if it is not visible, so if you work carefully the precipitation should be reproducible.Following
- JG Tapia added an answer:Will cross-flow filtration be also suitable directly after crude extract for protein purification and concentration ?
I have come across TFF systems and MWCO cassettes being used to purify and concentrate proteins. Is this an accepted method of protein purification and concentration of proteins directly from crude extract supernatants ?
Use a depth filter is a better option to cell culture clarification and impurity reduction, but chromatography is a right method for the purification.Following
- Hana Ensair added an answer:How you would express and purify a recombinant His-tagged protein from E.coli that has been cloned into a lac-based expression plasmid?
Any suggested readings about this particular topic or your expeience
Thanks a lot Gaurav Chhetri. I followed your steps and I had a god result. I have another question for you. Do you know about quantitative sialidase assay to test enzyme activity ?Following
- Lily Oguh added an answer:Has anyone observed large sediments after centrifugation cell lysates extracted in Laemmli buffer?
I treated some cancer cells with inhibitors, lysed them with laemmli buffer (containing protease and phosphatase inhibitors) by scraping on ice. They were left on the rotor for 16hrs in a cold room to allow proper lysis. After centrifuging at 16000xg I notice that there are large sediments at the bottom of the tubes including the control (no treatment). I am worried because when I carried this same protocol previously I did not notice any thick (mucoid) white sediment has anyone else notice this?
Thanks to you all. There was definitely less protein to soak up SDS.Following
- David Schwarz added an answer:Elution of FLAG-tagged protein with Enterokinase?
The protein has an N-terminal tag (single copy) and is being purified from insect cells (Sf9). We have been unable to elute it from FLAG affinity beads using either 1X FLAG peptide or pH3 glycine (estimate 0.5 mg bound to 1 mL bed volume). The protein is destined for structural studies so a mild elution is necessary (and retention in pH3 glycine is a concern - leading me to question potential efficacy of 3X FLAG peptide). Has anyone ever tried to elute captured protein via enterokinase treatment? I suspect steric occlusion will limit success - but happy to hear from anyone that has tried.
Thank you for the thoughtful responses. There is no crystal structure available for our protein, but we can make some reasonable assumptions as it belongs to the kinase superfamily. To clarify, there is a FLAG tag (DYKDDDDK) at the amino terminus, so the enterokinase cleavage site (DDDDK) is built into the tag and should liberate an untagged protein following treatment. Becasue we have shown that all of our protein is now "stuck" to the anti-FLAG antibody/resin, enterokinase would need to essentially "compete" with the antibody in order to recognize and cleave the protein. In afterthought, this is very unlikely and would (likely) minimally require us to engineer a spacer into the recombinant protein (as suggested above). I prefer not to take this route so we'll explore some other options, including alternative affinity resin provider(s).Following
- Neelakant Varma added an answer:Can anyone help me, why my SDS-PAGE gel smears like that?
It especially concerns 9th well. Will diluting my SB sample with water help?
A likely explanation is that each time there was a delay between loading the samples and actually running the gel might cause this problem. please go through the link below for troubleshootingFollowing
- Phan Y. X added an answer:How can I increase the binding affinity of GST-tagged protein to GST sepharose column?
I am purifying a protein having GST tag. I am using 20mM tris pH 7.5, 200mM NaCl, 5mMDTT and 5% glycerol as the equlibration buffer. Column is of 2ml bed volume (I have hardly 3mg of my protein). Protein is not completely binding to the column and nearly 30% I can find in the flow through. Please suggest me how to increase the binding capacity of the protein.
Thanks in advance..Following
- Gary Laco added an answer:How can I confirm streptavidin - peptide binding and concentration?
I have been looking at formation of a peptide multimer bound to a streptavidin scaffold. The peptide is biotinylated, so should bind inherently with the streptavidin backbone.
Currently, I have been quantifying concentrations using absorbance values from the unreacted substrates followed by the reacted product after passing through a zeba column to remove small particles, whcih should include any unbound peptide.
The concentrations look OK, and product concentrations are similar to what I would expect, knowing the stock concentration of streptavidin and absorbance produced.
However, when I am applying the construct in downstream cell treatments, the results are quite variable between batches. I wondered if anyone knows of some QA type tests I could run to confirm the multimer is forming correctly?
Thanks in advance for any help :)
How are you determining the concentration of your biotinylated peptide? I am assuming it was chemically synthesized with the biotin and then HPLC purified, but the purity would depend on what was specified (higher purity means higher cost and lower yield). The weight of a dried peptide cannot be used to determine concentration since the dry weight includes salts that can be up to 40% of the dry weight, and then if the peptide is only 80% pure your net biotinylated peptide could be less than 40% on a per weight basis.Following
- Matthias Körling added an answer:How can I separate peptides from small molecules?
Our lab is interested in the effects we have seen across different cell based and bacterial assays using supernatants of clinically relevant bacteria. We have mapped the activity to a fraction less than 3kda in size by ultrafiltration and testing, but we do not know now if it is a small molecule/metabolite or a peptide. What would be the best next technique to use to elucidate what is causing the observed activity? I was thinking of performing some kind of organic extraction and testing the aqueous components but I do not know how to go about this.
I would also recommend HPLC/MS. Maybe dialysis may help to separate small molecules?Following
- Faizan Ahmad added an answer:How do I interpret the results for CD Spectroscopy for my protein?
I have got CD results for my protein. However I do not know how to interpret the data for CD. I have uploaded my results in word format. Kindly download to take a look at the results.
I understand that each of the three structure of a protein gives a characteristic CD Spectrum. Based on my CD spectrum results (in Figure 1), what I can interpret is that my protein is predominantly in the form of alpha-helices (Correct me if I am wrong). However, what confuses me here is that the table (Table 1 in the word file ) that shows my protein consists of almost ~47% of Random coil?
Does the presence of high random coil indicate that that my protein is unfolded?
Awaiting for your help.
Almost all have been said that you must know about the analysis of CD spectrum of a protein. I hope, by now, you would have been able to know the contents of secondary structure of your protein. (I hope you must have gone through the literature suggested to you.) I would like to add the following that one must consider, 1, Does your protein contain any prosthetic group that contributes to the observed CD of the protein? 2. Have you subtracted contribution of the blank solution from the observed CD of the protein? Is the signal to noise ratio at each wavelength in the recommended range?
In the end I must say that the far-UV CD is an excellent technique to monitor change in the overall peptide backbone orientation. However, it MAY NOT be that good in the qualitative analysis of secondary structure of protein due to obvious reasons.
- Esen Dogan added an answer:Can I use the PBS buffer or TEA buffer as a solvent to concentrate my protein of interest (31.4 KDa) using a vivaspine 20 ?
The best wash buffer concentration I was optimized to purify my protein of interest was : Tris-base (Ph 8.0) 50 mM, NaCl 650 mM and Imidazole 100mM; while, the best elution buffer vs Tris-base (Ph 8.0) 50 mM, NaCl 275 mM and Imidazole 400mM
Thank you very much in advance.
If you are going to aliquot your protein, flash-freeze and store then I do not recommend having imidazole in your buffer while concentrating your sample since it causes precipitation once you thaw your aliquot. However if you are concentrating your protein sample in order to apply it to an AEKTA column for further purification or buffer exchange then the buffer you are concentrating your sample with does not matter.
By the way, imidazole concentration in your wash buffer sounds a little bit too high but then again it is optimized for your protein yet don't be surprised if your protein precipitates next time you thaw your aliquot if you freeze it in that buffer..
- Daniel Lee Adams added an answer:Where did my high molecular weight proteins go during transfer?
I used a small pre-cast gradient (4-15%) gel from Biorad and transferred it in a med. size box @90V 1.5hrs in a cold room with an ice pack. The high mol weight markers were gone (not in gel, membrane, or filter paper) at the end.
I thought there was something wrong with my buffer so I made new buffer and tried again at 75V and had a slight improvement, but still lost my big proteins.
I didn't think a large protein could pass through the membrane and they are not in the gel where did they go?
I'm glad you fixed your problem. To simply answer your question, its a combination of size and charge in relation to you transfer conditions. Semi dry is a different transfer medium, so different results. Keep in mind small proteins move faster out of the gel, but large proteins move faster because of charge. All of the parameters above have different effects on different proteins based on different transfer conditions.Following
- Frederick Crane added an answer:Does anybody know plasma membrane isolation protocols?
Does anybody know plasma membrane isolation protocols?
I need to isolate plasma membrane, to purify all proteins present, I have seen some commercial Kits, but I would like try first some non-commercial methods. And I would like to try different protocols, as it is known that not all proteins are extracted from membrane using only one protocol.
I need it for 2D gels and mass-spectophotometry among other characterising methods.
I will appreciate very much if you can help me with different protocols to extract the whole plasma membrane organelle and proteins.
Have a nice day. Thank you.
try D.J.MORRE at PURDUEFollowing
- Ziguo Zhang added an answer:What special attention should I pay for purification of recombinant type II restriction EcoRV?
Hi experts, I need a lot EcoRV restrict enzyme for routine usage. I prepare to make by my own. As restricts are not normal protein, do I need special tricks to deal with the over-expressed enzyme for its activity toward coli genomic DNA and the recombinant plasmid? would the leakage of the T7lac promoter matter? Any established expression and purification protocol for type II restricts will be helpful. Thank you.
Thank you, Roger. You are very helpful. ZiguoFollowing
- Pal Gahlot asked a question:If an enzyme is supplied in ammonium sulphate solution, then is it necessary to dialyse?
which buffer? this enzyme is to be used for a coupling assay?Following
- Robert Lindner added an answer:Any advice on immuno precipitation from crude/unclarified lysate?
I am stuck in this situation at the moment, trying to immunoprecipitate (IP) and Co-immunoprecipitate (CoIP) partners of an ion channel that have totally different solubility requirements.
N.B. - I have functional evidence (patch clamp recordings) that this channel is present fully assembled in my cells. Also, the two partners of this channel are part of other channel complexes (<10% form the channel I am interested in).
Protein 1: Super hydrophobic (17 transmembrane segments), I can detect it by western blot when solubilizing the PELLET in LDS sample buffer after lysate clarification. I have tried Triton-X100, CHAPS, DDM, Pierce IP lysis buffer and FivePhoton Transmembrane protein lysis buffer - none of these effectively solubilize this protein (even with additives - spermidine, glycerol). Sonicating the crude lysate helps somewhat, but this appears to disrupt the delicate interaction between protein 1 and protein 2 - i.e., I cannot see a CoIP band, but I do see a faint IP band. I can see a darker IP band when resolubilizing the pellet in lysis buffer + sonication to break up the pellet. But still no CoIP band (pellet will not re-dissolve without sonication).
Protein 2: A little more co-operative. I can detect this by western blot and IP from CLARIFIED LYSATE. I have not tried IP from the pellet for this protein. I do not see a CoIP band from clarified lysate.
My question is this: Is it necessary to clarify the lysate? I've been trying to come up with the "perfect lysis buffer" to solubilize everything, but not disrupt the interaction - but maybe I'm wasting my time? Could I just try my IP/CoIP from crude lysate? Has anyone ever tried this?
Happy to give more info on this. Thanks in advance for the help!
I also would suggest to try crosslinking on cells. Perhaps a disulfide-cleavable, water-soluble x-linker might be optimal for this purpose. This would allow you to use really dissociative lysis conditions afterwards, as long as you avoid cleaving the linker by reducing agent (include iodoacetamide or NEM in the lysis buffer). After IP, you can cleave by reduction and should be able to detect your two bands of interest. Good luck!
- Vishal Srivastava added an answer:How can I stop the protein coming out in flow through while doing purification?
I am trying to purify protein having molecular weight ~50Kda under denaturing condition through Ni-NTA chromatography and got following profile. Any suggestion to stop protein coming in flow through and not binding with column.
protein dissolved in buffer-A (tris 50mM Urea 8M, Nacl-300mM)- for loading the sample on column.
column washed with same buffer supplemented with 60mM Imidazole.
and then eluted using imidazole linear gradient from 60-800mM Imidazole in Buffer-A
Lane 1- marker
Lane 2-10 is elution during gradient
Lane 1- marker
Lane 2-8 is elution during gradient
Lane-9 flow through collected at the time of sample loading on column
Lane-10 -washing flow through of column by buffer A supplemented with 60mM Imidazole.
Image -3 FPLC profile
Thanks to all suggestions. I would try to follow the suggestions and report the results back.Following
- Patrizia Di Crescenzio added an answer:How can we increase the binding capacity of our protein while using Ni-NTA column (The protein of my interest contains 13 Cysteine residues) ?
While doing protein purification a lot of my protein is getting lost in the crude extract which clearly indicates that it is not able to bind properly to the Ni-NTA resin. Even after much standardization, the loss of protein does not seem to decrease at all. Also, significant contamination of protein bands is observed along with required protein even after using suitable reducing agent. How can these problems be overcome considering the above mentioned factors?
Hi Rashmi! I agree with previous suggestions, if His tag is somehow "hidden", you may try with a longer His tag, or directly use a different tag. What's the pH of your binding buffer ? His needs pH higher than 7.5 to be charged, otherwise it doesn't interact with Ni. You may also try to replace Ni with cobalt or copper, His has a higher affinity for both of them. Good luck for your experiments!Following
- Revathy Krishanmurthi added an answer:Any suggestions on protein aggregates during buffer exchange in size exclusion?
Hi fellow researchers, am an newbie in protein purification and hence this question. I am working on a novel protein whose size is 17kda.it does not have cysteine. It's PI is around 7.83. I did a his tag to purify it. I did able to purify.. But when I want to remove imidazole it aggregates I guess. Last time I could able to recover 2mg of my protein for a litre of culture. This time nothing I got.. When I diid sec using s200 column, everything gets eluted close to void volume (2ml away from void). But its single mono dispersed peak. I use ph 8.5 tris buffer . could anyone tell me what could be the possible problem?
Thanks for your input. I have increased sodium chloride concentration in my buffer and it solved my problem.
- Krzysztof Bielikowicz added an answer:How can I optimize my pepsin cleavage for a monoclonal antibody?
I am currently working with a monoclonal mouse antibody (IgG1). I have been using PNGase F to prime the antibody for pepsin cleavage. However, my antibody is relatively dilute (0.2 mg/mL). I can only use about 200 uL to optimize my cleavage due to the monetary aspect involving antibodies. I tried optimizing the pepsin cleavage twice - the first time I did not see any bands on the SDS-PAGE (used 20 uL aliquots from the 200 uL). The second time I tried visualizing using a size exclusion column and a UV detector, however, the peaks were extremely small and not very definitive.
Is there a way I can concentrate my antibodies prior to optimization? Is there a way to optimize the pepsin cleavage so that I can see what is happening better (whether it be on the SDS-PAGE or the size exclusion column doesn't matter).
- Muralidhar Reddivari added an answer:Lysis of SF9 cells infected with baculovirus to extract target protein for further purification?Recently I started to work with insect cells to produce protein samples for structural determination. I have experience of protein purification from E.Coli but I have a few doubts about the extraction of the target protein from insect cells. In a typical purification, starting from a pellet of insect cells obtained from 2.5 L of culture, I suspend the cells in about 70 mL of lysis buffer (50 mM TRIS pH7.7, 300 mM NaCl, 5% glycerol, 0.2 mM PMSF, protease inhibitor tablets) the lysis is performed adding Triton-X100 to a final concentration of 0.1% and incubating for 15 minutes on ice, mixing gently every 3 minutes. After that a mild sonication is performed (2 seconds “on” 10% amplitude/60 seconds “off” repeated 15 times). The mix is then spun down at 20000 rpm for 30 minutes at 4C to remove insoluble particles. At this stage my lysate looks very cloudy and it is difficult to filter it with 0.45 um syringe filters. Moreover, a layer of white “stuff” is also visible on the surface of the lysate. Eventually, after changing lots of filters, I managed to filter the supernatant and the protein is successfully purified. I believe that I lose about 30% of the total sample in the filtering process of the clarified lysate. Am I doing anything wrong? Could anyone suggest how to fix this problem?
2 cycles in a microfluidizer should be sufficient. please don't add detergents unless you need them as they are impossible to remove especially for structural workFollowing
- Joe C Polacco added an answer:What is the best protocol to extract 7s, 11s and 2s proteins?
Thank you for your answers.
It's been a long time since I played with seed proteins, but they are separated, usually, by their solubilities. The 7S and 11S of soybean are globulins, and are usually soluble in saline, while albumins, probably your 2S, are water soluble. Prolamins tend to require base, or perhaps organic solvents, but I don't think you are going after those.
See the following:
Peter R. Shewry,' Johnathan A. Napier, and Arthur S. Tatham (1995) The Plant Cell, Vol. 7, 945-956, Seed Storage Proteins: Structures 'and BiosynthesisFollowing
- Vivek Hamse added an answer:How can I purify the protein fraction from a plant sample containing mostly polysaccharides?
Trying to do some proteomics from a very viscous plant sample containing mostly a mixture of high molecular weight polysaccharides. Traditional methods to precipitate the proteins like TCA/acetone or salting out result in co-precipitation of some of the polysaccharides too. I thought on SEC but it is tedious... maybe some kind of filtration around 300 Kda? Any other alternative?
Hi, If you boil the sample, protein may get denature. The best method what we practice is that. The filtrated sample (May be in water or buffer) can be passed through molecular filters like Centricon after ammonium sulfate precipitation, however this would depend on your knowing the sizes of molecules in your sample in order to choose filters.
Eduardo's method worked quite well
0.1% cetavlon can ppt most of polysaccarides
You can use Ethanol to ppt Polysaccarides, But make sure that you dont loss activity of the your protein of interest.
- Elizabeth Baldwin added an answer:How can I reduce splaying of an SDS-PAGE gel?
I have consistently found my gel splays outwards at one end of the gel (light end). I am using a pre-cast gel, and have been running this according to instructions at 160V. I placed the gel in the fridge to attempt to reduce the splaying. This did work (although still some splaying seen) but the bands appeared to diffuse slightly. Any advice of what to try would be great.
Thank you all for your responses. I will give these ago and see if it improves.Following