Protein Purification - Science method

The purified protein is exhibiting a trimeric state based on my NATIVE PAGE gel and SDS PAGE shows monomeric configuration. Can you recommend detergents to destabilize the H-bonds mediated by sulfate ions (based on literature)? I have already tried BME and DTT and they don't work.

Hello,

We are trying to purify proteins using a secretion system but do not have TFF cartridges for our device. Does anyone know where we can purchase these? Or is there a better replacement? We want to downscale the volume from 3 L to 100-200 mL.

I've attached a picture for reference.

Thanks,

Thomas Newton

I use the refolding buffer with the following specifications for the refolding of brolucizumab protein.

l-Argenine, Sorbitol, EDTA, Tris, and fresh Cystein and Cistine.

I have already used these compounds to refold this protein and I was getting a proper protein refold. Currently, although I use the same compounds and with the same concentrations, the answer I get at the end is not the same. In fact, the protein disappears after being solubilization and added to the refolding buffer.

The issue really confuses me and I don't have an answer for it. Can anyone have an answer for me?

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.

Dear All,

I am trying to purify a protein secreted by HEK293 cells. My target protein fails to specifically bind on a nickel/cobalt column probably because of the presence of BSA coming from the serum. The protein is eluted at 50mM imidazole along with a huge amount of BSA.

Moreover, my target protein has very similar isoelectric point and MW than BSA so it is getting very difficult to separate them using classical methods.

Any advice ? Is there a specific resin that is binding to BSA ? Or something that can be added to reduce the non-speficic binding of BSA ?

With many thanks,

Gianluca.

I do an 8L induction at 18C for 16-20 hours in TB media. I routinely harvest ~50-80g of bacteria. I freeze my pellets at -20 for > 3 hours after harvesting, and have been using a 1kW blender to resuspend and homogenize them in my lysis buffer. However, I have been noticing that the blender causes a lot of air to be entrained in my lysate, which seems to be affecting my protein yield. Is there a better method to resuspend and homogenize my quantity of bacteria?

I can provide more information upon request.

Ammonium sulfate (40%) was used to precipitate proteins from calf thymus extraction. After centrifugation, the pellet was dissolved in PBS. The pellet can’t be dissolved completely (still had lots of pellet after centrifuging at 12000g for 10 min).The supernatant was cloudy and couldn’t be filtered through 0.22 um NC membrane. We further centrifuged the supernatant at 12000g for 1 h, but the supernatant was still cloudy and could not get through 0.22 um membrane. To get clear sample through 0.22 um NC membrane is the first step to further purify the proteins. Now it’s totally stucked here. Is there any recommendations? THX.

What are the steps in the regeneration of 'fresh' DEAE Sephacel, which comes in the swollen form, in 20% ethanol. And what is the significants of each chemicals in these steps, can anybody explain?

I have a trouble with my dialysis bag

Once protein has been extracted and quantified, and before protein digestion, I even concentrations to 1 mg/ml using U/T buffer (7M urea, 2m thiourea, 30mMTris). Then I re-quantifiy protein to make sure that protein concentration is 1mg/ml. For some reason that I do not know, buffer and protein do not mix homogeneously; the more concentrated the original aliquot is and the more buffer is added,  the less concentrated the final aliquot is. I did repeat the protocol 3 times with fresh buffer and I always get the same. However, I did not have this problem when sample was diluted in water. Any idea or suggestion? Thanks!

I'd like to perform a SEC step prior to on-column refolding of a protein I am expressing/purifying but am worried about the extended time the protein will be in the presence of urea (and subsequent protein carbamylation) in the current refolding workflow I have.

Is there any concern of protein modification with a 4-6 hour denatured protein purification workflow using 6M GndHCl?

Hello.

I am working on heterologoulsy expressing and purifying a protein using the Bac-to-Bac method. The protein expresses as verified by western blotting. I typically do a 4.2 L expression using P2 baculovirus. I lyse the cells via sonication in the following buffer (5 mL per gram of wet cell mass): 20 mM Tris pH=8, 500 mM NaCl, 10 mM imidazole, 5% glycerol, DNAseI, and protease/phosphatase inhibitor tablets. After sonication, I centrifuge the lysed cells for 1 hour at ~54,000 x g. After centrifugation the supernatant is not viscous and has a clear yellow tint. This clarified supernatant will clog the IMAC column (I have tried both HisTrap and self poured columns). Only when I filter all of the clarified supernatant using a 0.22 um syringe filter can I load without clogging. Does anyone know a better way to solve this issue of column clogging?

Hi everyone

I have a problem with Histag protein purification. I have this construct with his tag on N-terminal. I went for small-scale expression and I had some good bands. However, on a large scale, the protein didn't bind with the HiTrap TALON cobalt. My protein contains an Mg2+ ion coordinated by the side chains of Asp, His. How to overcome this metal ion issue, or Should I change the column to a Ni-Trap?

I appreciate your help

I am purifying a protein by denaturation method(8M Urea). After solubilizing and running a His TRAP with 1M NaCl Wash, and elution. I refold by dialyzing to 2M urea and then desalting the column to remove Urea and refold. At lower concentration till 1 - 1.5 mg/ml, I have a normal 260/280 of around 0.63 but when I concentrate further the protein starts showing weirdly higher 260/280 (like <1 ). I see no precipitation also( 350 nm seems fine and visually also protein is not cloudy). I am quite confused with this trend. Does anyone have any similar experience?

How can I overcome this problem?

When measuring protein concentration with a spectrophotometer, is the use of Coomassie Brilliant Blue necessary? Protein (BSA) has a peak in UV, so can I prepare different concentrations of it and determine the unknown concentration?

Currently I am doing protein purification from my transformed E. coli by using Histrap FF column. The protocol says, to prepare the sample I can dilute it with binding buffer but not using strong bases/acids due to precipitations risk. In my lab we only use NaOH and HCl for pH adjustment. What weak bases/acids usually used for pH adjustment?

I am trying to express the soluble fraction Membrane Protein SARS Cov2 in E. Coli. I am getting only monomers while it's the dimer form that is active. Literature that I have come across have used yeast and mammalian expression systems only. I want to know if it can be expressed in E. Coli. If not, why?

Why does the initial protein elution (elution No 2)(Each elution volume 500microL) in maltose binding protein purification exhibit high concentrations of the target protein on SDS-PAGE, but fail to demonstrate high activity compared to the lateral elution fractions with lower target protein concentrations? Can the activity of the target protein be disrupted by the expression of other proteins related to the vector (PMALC2X)?

I am currently trying to express Turritopsis dhornii TET enzyme with a 6x HIS tag in a BL21 E.coli host. Every time I purify my protein it shows double bands on SDS-PAGE, when testing its activity on ELISA there seems to be no activity. My lysis buffer contains 50mM HEPES ph 7.5, 30 mM imidazole, 500mM NaCl, and 1mM DTT. I purify with 800uL of Nickel Sepharose beads. Are there any adjustments I could make to stop this double band from showing?

By keeping rest of the protocol same does centrifugation speed will have any effect on protein extraction from XL1-blue colonies

What is the Maldi-tof technique?

Is it necessary to perform Maldi-tof technique after protein purification?

My N -terminal GST tagged protein fail to purify using this condition: 30ml sample (3mg of crude protein) injected at flowrate (0.5ml/min), 40ml wash (PBS pH 7.3) and 5ml elution (50mM Tris HCl + 10mM reduced glutathione, pH 8). For washing and elution, the flowrate was 1ml/min. Buffers were prepared all according to the manual. For your information, based on SDS -PAGE analysis, the crude has decent amount of GST tagged protein. Therefore, I suspected that the GST tag was hidden in the conformation. The protein structure was not discovered yet so I can only hypothesized. Based on previous structural analysis, the N -terminal was predicted to be at the cytoplasmic region. Does this means, whatever tag that I put in the N -terminal will be forever hidden in the structure? Should I cleave the tag and purify the protein using other means (IEX/ SEC/ HIC)?

suppose: we have a purified protein with us; after this step what are the methods and procedures that can we use to see the protein is intact- KINDLY HELP

The purpose is cell based functional assay for the recombinant protein not protein purification.

I have laryngeal cancer samples from which I am to extract DNA/RNA and Protein. However, the kit I have only extracts DNA and RNA (Qiagen AllPrep DNA/RNA Kit), and does not extract protein. How do I go about protein purification without having to order another kit, keeping in mind that the samples are quite limited in size. Thanks.

Theoretically, if the value of A260/A280 goes high it shows DNA contamination in protein sample, and if goes down (0.6-0.8) it shows good purity of protein. In this way, a value less than 0.6 should show a more pure protein. but lower value of A260/A280 of protein <0.5 is considered not good for protein purity. Why

I am expressing a human RNA binding protein in E. coli and purifying it using ammonium sulfate precipitation followed by heparin, butyl, and size exclusion chromatography. While the first batch of protein purification did not show any RNA contamination, subsequent batches consistently exhibited RNA contamination. I have tried to maintain the same purification protocol but cannot seem to eliminate RNA contamination.

Have other researchers experienced this issue while purifying RNA-binding proteins, and if so, do you have any suggestions for troubleshooting or improving protein purity? I would appreciate any insights or recommendations to help me achieve a pure sample of my protein of interest.

Dear researchers, I am now trying to express several proteins from bacteria（Agrobacterium Tumefaceins) in one of its biosynthetic gene clusters. However, some of the proteins can be easily expressed and purified but many of them can not even be expressed. I also checked the pellet by SDS-PAGE and found that there were no over-expression bands in the pellet.

I want to ask how can I possibly get these proteins? Here are my protein expression conditions:

vector： pET28a

host：E.coli BL21

Tag：6xHistag

Culture until OD value reaches 0.6 and 0.5 mM IPTG was added(working concentration) to induce over expression.

Some of the proteins in the cluster can be expressed very well using this common protocol, yet others remain even unexpressed. I do not know what had happened, since there were no over expression bands in the pellet, suggesting over expression did not even take place.

I tried other recombinant tags like SUMO, MBP and GST, but none of them helped.

I am really stuck in this situation now. :(

Hello

I'm sure this question has been asked a lot but the protein I am purifying is not as clean as I would like and all the potential solutions I have read about have not worked for me.

I am purifying a protein that forms inclusion bodies. The construct is 14x His tag - Tev cleavage - protein of interest. I use BL21 Star (DE3) cells and TB media for growing the bacteria and induce at OD 0.6 for 4 hours at 30 degrees.The purification protocol is based on a previously published paper. Following sonication, lysis buffer and washing the pellet the protein is extracted from the inclusion bodies using 6M Guanidine Hydrochloride, applied o/n at room temperature to a Ni NTA agarose column. It is then eluted in 4M Guanidine. I then pass it through a RP-HPLC. As you can see by the attached image although I am getting a high amount of protein it has a large smear which I am not sure if it is contaminants or degredation. I overloading with sample when trying to judge purity but I think it's better to get an accurate representation of what’s going on rather than kid myself I am working with a pure sample.

The HPLC makes no difference so I think optimisation on the Ni-NTA purification is needed but nothing thus far has worked, I have tested different induction temperatures, leaving it on the Ni column at both 4 degrees and for a few hours (rather than O/N at RT like the protocol says), protease inhibitors and washes/step-wise elutions with different concentrations of imidazole (as in attached file) - yet none of this has worked. I've even thought about making the His tag smaller (14x seems quite large, but I cant see how this would impact purity other than needing more imidazole for elution).

If anyone has comments on the purity and has any tips that I have not tried it would be greatly appreciated!

Thanks!

Hey, I need help! I'm trying to purify two proteins (26 kDa and 96 kDa) using a Ni- NTA resin. Both enzymes have a 6- histidine tag (bioinformatic models shows that the tags are correctly exposed), and the expression occurs in E. coli (strain BL21). The problem I'm having is non- specific binding, the elution come out just as dirty as the original sample or the flow through.

I have tried using two different brands of resin (Quiagen and Cytiva), currently using the last one. I already tried using different buffers (20 mM Tris-HCl, 300 mM NaCl + 300 mM imidazole for elution, pH 8.4 or 20 mM sodium phosphate, 500 mM NaCl + 500 mM imidazole for elution, pH 7.4), just as indicated in the manuals.

I tried two methods of cell disruption (lysozyme + freeze and thawing cycles or sonication), thinking that maybe the lysozyme was interfering with the resin's maximum capacity. I've tried doing both -batch and column purification-, and got the same result in both cases. It puzzles me, because my mentor purifies many proteins from E. coli lysates using the same resin and conditions, and she gets good results. We don't know what's failing, I'm going crazy! Will it help adding a nonionic detergent to the elution buffer (e.g., 0.2% Tween- 20)?

These are today's results. I resuspended the bacterial pellet in 50 ml of binding buffer and sonicated it. I centrifuged and passed it through a 5 ml column, with a flow rate of 0.5 ml/min and usind an imidazole- gradient elution. (I already tried increasing the flow rate). I got a good peak elution, but when I load the collected fractions on a SDS- Page.. There are a lot of proteins, and they all elute together in one unique peak! (Sorry for the SDS- Page pic, I took it with my phone. Colected fractions are the last five ones).

Thank you in advance, Julieta.

I use 6x-his tag purification to purify my protein.After binding to the Ni-NTA beads, I use 6M urea washing buffer which contains 60mM imidazole to compete with the non-specific protein. However, my protein was washed out when I was washing the beads. Because of that, the elution quality of my protein are very poor. Does someone know the reason why?

I was looking for any recommendation for protein purification system for my research. We dont have much budget for akta and are looking for affordable alternative. We do have HPLC in the lab, can we attach column to the machine and purify our protein successfully?

Hi all,

I am a beginner in cryoEM field.

I know sometimes we must remove his, flag, strep, MBP, eGFP in X-ray crystallography study.

However, when I read papers, I found that these tags were not necessary to be removed in cryo-EM study. So I am wondering if we can keep all the tags for cryoEM study, especially the big tags such as MBP and eGFP tag, but will they affect the protein real conformation I mean in vivo conformation? I didn't remember who told me that MBP connects to proteins through a flexible linker, so I am not sure if we can see the big tag density in this case.

On the other hand, these big tags can help to increase the protein particle size, and may also help to improve the orientation preference in cryoEM study.

Are there any cases or publications that solved cryoEM structure containing big tags e.g. eGFP or MBP tag?

Any comments or suggestions are welcome, thanks!

I purify my recombinant his-tagged protein (40 kDa) using Akta Pure 25.

However, I have been through a couple problems.

I use Lb medium, but when expressing using TB medium (only 250mL pellet resuspended in 20mL lysis buffer), my 1mL HP column (Ni+) seems to get clogged and the back pressure increases by the end of sample application, specially after only a couple uses.

I am aware of the need to use DNAse as it helps with viscosity, but I have never used it to my cultures. I usually prefer LB, but I am trying out TB medium for higher cell concentration.

Questions are:

1. Bypassing flow restrictor helps decreasing the pressure, but I have bypassed it once and the column seemed to crash even under pressure limit. The flow restrictor module seems to be increasing a lot of the pressure, cleaning it up would be enough?

2. Has anyone experienced any limit for protein concentration of the lysate (not his tagged protein) or maximum volume of culture pelleted for 1mL HP columns?

3. Besides filtering lysate and buffers, any recommendations to avoid high pressure? Such as avoid using TB medium, high volume cultures…

4. I’ve been able to reuse 1mL HP columns for 5x using LB medium before it crashes (the beads seem to collapse and a “void” is visible in the column). Any recommendations to increase column lifetime?

I have solubilized my protein with 0.3% sarcosine and purified by using Ni-NTA,during purification most protein is going into flow through.

I have diluted my sonicated sample to 0.1% sarcosine but still I am unable to get binding of protein.

I saw this product https://www.thomassci.com/scientific-supplies/Tube--o--dialyzer, do you have any products like this that you are more comfortable with? Will it compromise the quality of the purification?

Hi everyone,

I have been working on purifing Tn5 transposase for quite a long time. Basically following the protocol from http://genome.cshlp.org/content/24/12/2033.full. Occasionally I got one batch of transposase having activity, but after around 3 months, activity was not detectable any longer. Other than that batch, I got no more active protein. According to protein gel, the protein size is correct, roughly 53.3 kDa. Is there any tricky step that I missed? Any suggestion or help would be appreciated!

Best!

Hi,

I've been routinely using Dialysis for removing salts after we do protein purification using Ni-NTA or any other chromatography. But I'm curious to know how economically and operationally feasible this process is when we move to Industrial set ups?

Since we have lower volumes (15-20mL) of protein in R and D lab it's easier to perform in beakers, how would it affect the process when we move to 15-20L of the same protein?

I'll be happy to read if anyone has any literature in this area.

Thanks

Hello,

I expressed a protein with a his tag (65 kDa) and purified it by Ni-NTA affinity chromatography. The amount of protein that I got was huge, so I didn't wonder about the aggregation in the fraction tubes overnight. The next day I spun them down and made an SDS PAGE. I repeated the SDS PAGE several times with different amounts of protein to make sure I was doing it the correct way. Everytime I get an additional band (and smear) right above my protein (the thickest band is my protein). I then diluted my protein 1:10 and purified it again by Ni-NTA affinity, but I got exactly the same result.

This puzzles me a lot, because my colleague expressed this protein many times and purified it the same way, and he never got this additional band. So now I am wondering, what could be the reason for the sudden appearance of the band?

Could it be somehow that the amount of protein is too much for the column? Was the expression too long (around 16h at 16°C)? I am not sure how to proceed now. Repeat the expression (but what should I change) or rather purify it further, e.g. with IEX?

Hello, In order to speed up our protein production process, we would like to use an affinity purification TAG. Some well-known TAGs work very well (strepTag) but identifying a cGMP compliant column is a challenge, others like His TAG have cGMP compliant columns but are difficult to use due to the risk associated with nickel and imidazole. We have identified C-Tag. Do any of you have recommendations or other suggestions for us? Thank you

Triton X 100 is used to increase the solubility of the protein when used in the lysis buffer for bacterial lysis. But I am not sure how much percentage can be used so that it increases the solubility of my his-tagged protein and doesn't effect the binding of the protein into the Ni-NTA beads. If, someone is using Triton x-100 in their lysis buffer, I would like to know how much final percentage is safe for Ni-NTA protein purification.

Hi all,

Recently I am working on protein purification.

1. affinity column(GST)

2. Add PSP enzyme to remove GST-tag(O/N)

3. GST-tag off

4. gel filtration

However, the protein tends to form large amount of soluble aggregate shown as the gel filtration figure (first peak). I need the native proteins for futher assay so how can I avoid such a problem?

I am attempting to do Triton X-114 temperature induced phase separations as an endotoxin removal step on my proteins. I have no issues reaching the cloud point and performing phase separations when my protein is in PBS, pH7.4 (or similar buffer). However, when I attempt to do the separation on a protein denatured in 8M Urea, 50mM Hepes, pH8.0 buffer I cannot reach the cloud point and Triton X-114 does not separate into a distinct layer after heating/centrifugation.

I assume that a strong denaturing agent is affecting the phase behavior of the Triton, however, Triton X-114 separation protocols that I have found seem to have little issue using 8M Urea or 6M GdHCL (or at least do not mention any relevant problems). 

Does anyone have experience that could help shed light on my situation? Thank you.

am trying to purify a human DNA binding protein (normally localized in nucleus and mitochondria). I am currently optimizing the conditions I tried 2 different approaches:

The expression was performed in BL21 (DE3) E.coli, 2YT media at OD600 = 0.8 induction was performed with either 0.5 or 0.1 mM IPTG at 16oC for 12 Hrs.

In all the conditions my protein wasn't pure, after doing mass spectroscopy I figured out that the additional bands are from E.coli (the host).

Can you please suggest me how to get rid of the contaminants?

I have attached the gel for more information, the protein of interest is in red rectangle

I'm using Ni-Nta beads to purify the His-tag recombinant protein. After sonication, I haven't found any dimers upon SDS-PAGE. But after that load and the next purified eluted protein formed dimers. I used 3 mM beta-mercaptoethanol in all lysis and wash, elution buffers. my actual protein is 36 KDa but I'm obtaining a band at 72 KDa. My protein has a total of 11 sulfur molecules in protein atomic composition. How to reduce this dimer formation.

Hello all,

I am facing an annoying problem wherein I am attempting to remove my GST tag from my (glutathione eluted) chimeric protein using a Thrombin Cleavage kit (Biotinylated Thrombin), but however after 10 hours of cleavage reaction at 20 degrees I am not being able to get the protein in the supernatant , but however it is remaining bound to the glutathinone sepharose beads.

I need it in the supernatant for use. The protein w/o GST tag is of ~17kda.

NB: The vector I am using is pET42 which has a thrombin cut site upsite of my protein.

I have attached an image of sds where there is LtoR : marker , supernatant post incubation of eluted protein with thrombin , and glutathinone sepharose beads incubated with thrombin treated chimeric protein. sorry for bad quality image

I conducted gravitational His tag protein purification manually and observed that the A280 of my crude is similar to the eluent post sample application. How is this possible? Does the His protein eluted out along with the unwanted ones? For your information, the purification graph shows no significant peak post His protein elution buffer application

I have a couple of proteins (95-130 KDa) to make them clean. What are the best microcentrifuge tube based column filtrations you guys have tried and worked best. Any suggestion with direct product link would be great help. SO my retention range is (80-130KDa) and everything else below 95KDa (+/- 5) I want to get rid off.

I have purified a membrane protein via affinity purification using detergent OG. I was trying to dialyze the protein with ST (NaCl-150 mM and Tris-20 mM) buffer(pH 8). Unfortunately, during dialysis, the protein gets precipitated inside the dialysis bag.

The predicted pI of the protein is five, so I also tried the pH of the buffer adjusted below and above the PI. But no luck; it is still being precipitated. Any suggestions and answers will be helpful

Hi there.

My protein was found to have a high nucleic acid peak when it was further purified by exclusion chromatography. How can I remove the nucleic acids when purifying my protein?

I'm struggling to purify a full-length His-tagged recombinant protein with GFP domain and S-tag.

I produced the protein in E.coli batch culture and purified it using Ni-NTA affinity agarose beads from QIAGEN where I incubated the beads with the protein overnight at 4° then used gradient imidazole to elute the protein. I performed a WB using Abs against the His-tag, GFP, and S-tag where I could see only fractions of the protein and never the full length!!

Can anyone provide insight? would love to know your suggestions.

I am trying to express and purify nanobodies from a nanobody library in E. coli. I am using a pBAD vector to which I added the pelB sequence for periplasmic localization. The nanobody is fused to YFP with a His tag.

I induce the expression with arabinose and I can see on an SDS-PAGE gel that the nanobody is being expressed (although the expression seems to be lower than for some of the other proteins that I am expressing using the same vector), but I am having trouble with the extraction and purification steps.

I have tried to extract the nanobodies using lysozyme with PMSF following a protocol that usually works for me and I have also tried the osmotic shock protocol ( ) but the nanobodies seem to be stuck in the cell pellet even after lysis.

I do not have any experience with nanobodies so maybe there is an important step in the protocol that I am missing or not doing properly. I would appreciate tips or good protocols for expressing and extracting nanobodies.

I just need keywords to search for.

Thanks for time.

Does anyone have a good way to quantitatively measure the amount of nuclease in a purified protein?

Hello,

I am trying to express and purify recombinant nuclear pore proteins in E. coli cells. All the published protocols I have found use 8M urea in the lysis buffer and subsequently during purification using Ni-NTA column. Is there any other role of urea in the buffer, besides the solubilization of nuclear pore proteins?

Thank you.

I want to purify Peptide-Fc Conjugate through Affinity purification, So I tried using Mab Select Sure which is a Protein A column used for purification of mAb's, But during elution there was no peak observed, So I would like to know which column should be preferred for purification of Peptide-Fc Conjugate.

Also I'd like to know what are the possible reasons for un binding of peptide-Fc conjugate to the column ?

I performed protein purification through his-tag and Ni-Agarose. The result of this process is quite good but the problem is that I need to use this protein for the test with the substrate. Too high a salt concentration (Imidazole) will affect the activity of the protein. I did a semi-permeable membrane dialysis and was hoping the salt levels would come down.

The problem is is there any method that will help me to determine the concentration of this salt in my protein solution?

I have a protein labeled with a fluorescent dye (fluorescein) and I am hoping to get a quick measure of purity using TLC after dialysis. TLC is quick and easy so I am hoping to start there.

What types of plates and solvents should I purchase? Normal phase or reverse phase? I know I can use ninhydrin to develop the proteins but that's about it. I have had trouble finding information elsewhere...

I used the fluorescently labeled oligonucleotides (RNA:DNA hybrids) as substrates. After incubating helicase with RNA:DNA hybrids, there are decreased RNA:DNA substrates but I can not detect the increased displaced products (single-stranded RNA or DNA). I can not understand these phenomenon.

The typic helicase activity is in an ATP-dependent manner. I am confused that the RNA:DNA substrates seem to be removed more quickly in the absence of ATP from my results.

Here is the procedures and buffers in helicase assay I performed, can anyone give some suggestions to solve these problems above ?

Reaction buffer: 20 mM HEPES pH 7.0, 50 mM KCl, 2 mM MgCl₂, 1mM DTT, 5% glycerol, 0.1% Triton X-100, RNase inhibitor, 10 nM RNA:DNA hybrids and 50 nM cold RNA or DNA oligomer. Total 10μl in system. 37 ℃, 60min.

Stop buffer: 20 mM HEPES pH 7.4, 300 mM KCl, 10 mM EDTA, and 200 ng/μl proteinase K. Add 5μl to each reaction. 37 ℃, 60min.

Hi All:

I am trying to purify an endogenous protein. But the protein is known to have many other interacting proteins in the cell system I am working on. Is there any way I can decrease protein-protein interactions during IP, remove its interacting proteins, and purify my protein of interest alone? Thanks a lot for your help!

So I would like to isolate RNA and Protein from frozen mouse brain tissue, and I was curious to know with which available kits you have had the best experience? Thank you in advance.

while i am doing dialysis for protein purification the dialysis membrane breaks and the protein sample mixed with the buffer(sometimes it was happening not every time). Kindly share your suggestions

I have expressed human granzyme A with His tag on N’ in Pichia pastoris system, the protein was exported to the medium, which was collected and stored in -20°C. The presence of the protein was validated by Western Blot. It seems like after >1 week protein disappears from the medium. Similar case is observed when protein is purificated by Ni-NTA resin and stored in 50mM Tris, 100mM NaCl, 500-550mM imidazole, pH 8 (imidazole concentration gradient was used to elute protein from resin and in this concentration sample contained granzyme A). Can you suggest how to treat samples to not loose protein? What I’m missing? I need to work under conditions that retain granzyme A in native form, as next steps are removing His tag to obtain active enzyme for enzyme kinetics assays. Also the trouble is with concentration because after expression the protein is only detected by Western Blot, not by Coommasie staining. Can I concentrate protein before purification? I’m wondering if medium composition may impact centrifugal concentrators membrane.

I’ve doubt that after the final step of CIP Protocol (washing with Binding Buffer) should I directly pass 20% Ethanol or was washing with water fallowed by passing 20% Ethanol ?

The steps I’ve fallowed are :

1) Equilibration buffer

2) 0.1 to 0.5M NaOH

3)Equilibration buffer

4)Water

5) 20% Ethanol.

Is this process is correct ?

Every member of the lab I am currently part of uses different centrifugation speeds (5000xg to 8000xg) and times (10mins to 30mins). They all seem to work for downstream protein purification from these cultures. As a precaution I tend to use the highest centrifugal force (8000xg) and for the longest reccomended period (30mins) to try and maximise the size of my cell pellet. Is it possible to over centrifuge the cells, which may cause problems I am unaware of?

Hi,

I need a suggestion regarding simultaneous extraction of RNA/Protein extraction kit

We want to use one of these kits to obtain both RNA and protein in one process.

Are they all equally good quality and easy to use? Does anyone have any concerns about any of them?

Thank you

Hi all,

it's my first question here on ResearchGate. My question is: what are your yields(i.e. __ mg from __ volume of E.Coli cell extracts) from His-tagged recombinant protein purificaction using Ni-resi? And what's the cell rupture method you have been used (i.e. sonication, Bugbuster, or homemade buffers)?

A little bit about why this question matter to me. I am a Ph.D. student in biology and currently working on an expereriment that need a lot of purified His-proteinX. The cloning was succesful and I am able to obtain ~100 microgram(ug) of "purified" His-proteinX from cell pellet collected from 100 mL E.Coli broth by His•Bond Resin (Cat. No. 69670, Novagen). I was following the small batch method on the manufacutuer's manual.

However, while the current yield is acceptable for my need, I am wondering whether the yield I have falls into a reasonable range and is there anything I can do to optimize the workflow and increase the yield of purified His-proteinX? That will be sweet if you have any related experience to kindly share.

Thank you!

Key words: Ni-NTA/Resin/protein purification/His-tag/biochemistry

Pls help me to choose an appropriate entry level protein purification system? Infront of me two machines are there 1. Akta start and2. BiologicLP. When I compared 1 has pressure upto 5 MPa and flow rate 0.5-5 ml/min. At the same time 2 have 0.05-40ml flow rate2 MPa pressure. What are the othee important things to compare? Which one would you suggest me ?

I need a suggestion regarding simultaneous extraction of RNA/Protein extraction kit

We want to use one of these kits to obtain both RNA and protein in one process.

Are they all equally good quality and easy to use? Does anyone have any concerns about any of them?

I am working on TAT-tagged protein transfection in mammalian cells. Cells are transfected with TAT tagged akt1 protein and akt1 phosphoprotein. Akt1 is phosphorylated at threonine and serine positions. When I transfect TAT-akt1 protein, cells are growing normal but when I add TAT-akt1 phosphoprotein cells become stop growing and dormant. but phosphorylation of akt1 increases cell growth and proliferation. During protein purification, I added phosphatase inhibitor in the buffer which I did not add for TAT-akt1 purification. I am adding a very small amount of protein 0.5 or 1uM , it should not be toxic to cells as TAT-akt1 transfected cells grow well.

1. Does phosphatase inhibitor-containing TAT-akt1 phosphoprotein not good for cells?

I'm trying to express and purify a 40 kDa his-tagged protein from E. coli BL21. Because of prior difficulties obtaining a sufficient amount of soluble protein, I have begun purification attempts from inclusion bodies. This entails washing the insoluble lysis pellet with a buffer containing Triton-X100, then solubilization with 8 M urea. I then try to refold and purify the protein using IMAC (4 mL of Ni Sepharose 6 Fast Flow resin in a gravity flow column) by linearly reducing the urea concentration to 0 over 15-20 bed volumes, then elute the protein by increasing the concentration of imidazole. However, attempts have resulted in the vast majority of my recombinant protein coming out in the flow through, implying poor binding.

Before trying to purify the protein from inclusion bodies, I was purifying a small but detectable amount of protein from the cleared lysate. In those cases, virtually all of the protein bound to the column. I only started having issues with binding after adding urea to the buffer.

Some technical details about the chemical environment:

Binding buffer: 50 mM sodium phosphate, 500 mM NaCl, 30 mM imidazole, 8 M urea (pH 8.0)

Example total protein in sample: 500 ug

I have tried reducing the urea concentration (I could get it down to 6.8 M urea to get most of the protein to dissolve), increasing the sample volume and reapplying the sample to increase the amount of time the sample contacts the resin, and using a freshly regenerated column. After talking with tech support, they suggested I reduce the imidazole concentration to 5 mM in the binding buffer to see if that helps, but barring that next step, does anyone have any other ideas? Your help will be greatly appreciated.