Recombinant Protein Expression and Purification - Science topic

Hi All,

I wanted to purchase Rosetta™ 2(DE3)pLysS Cells (Product No# 71403-M )from Merck but I was told that this product has been discontinued. In the circumstances, could you kindly suggest/recommend me an alternative strain to Rosetta™ 2(DE3)pLysS Cells?

This strain is intended to be expressing the LbuCas13a protein (at 16 degrees Celsius) and the capsid protein of tobacco mosaic virus (TMV).

Thank you for your time and consideration.

I look forward to hearing from you.

Subha

Purification protocol for human PARP1 with good yield.

I'd like to know that what are the different ways to know/identify whether a particular Gene is expressed or not ?

Few points from my side are :

1) identifying it's corresponding m-RNA transcripts level.

2) identifying the protein that was produced by the expression of that particular Gene.

Any other points ?

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.

Suppose you are trying to express a protein in Bacteria but that protein is coming in Inclusion bodies but not secreted out. So is it possible if I express that same protein in mammalian cell with signal peptide to make it secreted out in the supernent so that I can purify it easily ?

I've few queries regarding bacterial and mammalian plasmids for expression of Gene of Interest. What plasmid elements/components that are differ between bacterial and mammalian Plasmids to express a gene of Interest.

According to me :

The elements/components that are common between bacterial and mammalian Plasmids are :

The elements/components that are differ between bacterial and mammalian Plasmids are:

I'd like to know is there any other differences?

Thank You.

I'd like to know the Maximum Yield we can achieve with CHO cell line (Irrespective of CHO-s/ExpiCHO/CHO-K1 e.t.c and also the mode of operation like batch, Fed-batch and Perfusion) ?

Hello! I'd like to ask a question about protein expression. My question is how I can remove N-terminal formil-methionine from E.coli recombinant protein if the next amino acid after methionie is phenylalanine. It is known that methionine aminopeptidases such as MAPs from Pyrococcus furiosus are depent on a penultimate residue of substrate and they do not react with Cys, Asp, Asn, Leu, Ile, Gln, Glu, His, Met, Phe, Lys, Tyr, Trp, Arg amino acids. So, How i can deal with this problem. What type of enzyme will be applicable for M↓F removal ?

I am planning to use the Lambda Red recombineering system to insert an insertion in the following format depicted in the picture to E.Coli genome.

My questions are

1. Is the T7 promoter suitable for this purpose ? If not what promoters are better ?

2. Can I include the lac operator also in the insertion?

I really appreciate any guidance from you.

Dear all,

Attached is the image of expression profile of my protein of interest. Starting from the left after molecular weight marker, are total cell lysate (uninduced), total cell lysate (induced), supernatant (Uninduced) and supernatant (induced). IPTG used for the induction was 1 mM. The expression system used was BL21 DE3. Here, the prominent band is in the right range of expected molecular weight. But I am worried I see almost identical expression in both uninduced and induced. Western blot could answer definitely if it is my protein of interest ( need to be done). But is it possible with normal BL21 DE3 cells ? Please give your insights into it.

Thank you

With kind regards

Prem

HI,

I cloned a bacterial protein (19kDa) in pET15 vector, with 6x his tag. After overnight induction at 30 degree C, I sonicated the cells and purified the protein on fresh NiNTA column.

After immobilizing the protein onto the column for 15-30min at 4 deg C, I wash the column 5 times without imidazole (protein got eluted during wash with 20uM imidazole in pilot expt). Even though my last wash is clean, as soon as I add 100uM imidazole, I see proteins start to come out of the column, everything gets eluted by 300-400uM imidazole. Surprisingly I see more than 10-12 bands in my 100uM imidazole eluate, ~5 bands in 200uM eluate and 2-3 bands in 300uM eluate. The intensity of my protein also decreases with each eluate, hence I cannot use the subsequent eluates for my assay.

Kindly help.

Regards,

Kasturi

I want strong, constitutive expression of my protein in m. smegmatis.

I have a strong, constitutive promoter. However, the RBS is very weak.

I want to exchange the RBS for a new one that is better.

However, I worry that I will "break" the promoter when I try to add the new RBS.

How do I determine the important parts of the promoter, so I can replace the RBS without destroying the promoter?

Hi,

we silanized a glass column with NTA and afterwards tried to couple nickel onto the column to obtain a nickel-NTA-column for 6x-His-Tag purification.

Unfortunately first experiments yielded 0% protein.

I added our coupling protocols to the question.

We already coupled proteins onto the columns with the same protocol (protein A instead of NTA in the last reaction step) and got very good results. Now we wonder, why it's not working with NTA. Or is our coupling protocol of the nickel to the NTA not a proper way?

Anybody have a clue on what point there may be a mistake?

Thanks a lot and best wishes,

Marco

Hello,

I am expressing different proteins in bacteria with c-terminal mCherry-3xflag fusions. There is an unknown product there I can't identify. It appears to have the same flag tag as my constructs, so it must be a truncated/degraded version of my proteins, but why? It's not nonspecific, as my "empty construct" and "mcherry-flag only" constructs showed no bands. I worry about this unknown product as I needed the mcherry to specifically indicate the protein. The experimental details are as below:

I did a western blot of the lysate, staining for flag.

The constructs and their *predicted* band sizes are below:

What I actually saw on anti-flag blot was:

Essentially, all bands are expected, but there is an unknown 45 kDa band.

I don't understand where the 45 kDa band is coming from.

It's exactly the same size and intensity for proteinA, NTD of protein A, proteinB, and proteinC.

But the 4 fusion constructs don't share enough common sequence to generate a 45 kDa product.

Does my mCherry just run "large"? But why us the "mcherry-3xflag" construct only 30 kDa?

Where could this 45 kDa product be coming from?

I am thinking on expressing non-mammal eukaryotic protein in mammalian expression systems but I am not quite sure if it is something doable or not.

Based on this paper

,TEV protease can cleave between the Gln and several amino-acids (besides Gly/Ser) with acceptable efficiency in its recognition site.

Therefore, it's practically possible to purify many proteins (without an extra residue at the N-terminal end), by using affinity chromatography.

I was wondering if anyone could share their experience/knowledge using TEV protease to cleave between Gln and Met?

I have retinal samples from transgenic mice. Beclin1 and P62 were over-expressed compared to WT samples, but LC3b-I and LC3b-II both showed normal intensity. In addition, Lamp1 (lysosomal marker) seemed to be down-regulated in transgenic samples. This is quite puzzling. I cannot rationalize how LC3 levels can be normal as both Beclin and P62 are clearly over-expressed. Does anyone have an idea can this be even possible, and if yes what could be the functional meaning of this phenomenon?

I would like to screen for intracellular expression of recombinant protein production in S. cerevisiae by FACS. I am aware that many yeast researchers screen for intracellular protein expression by Western Blot. However I have a large number of samples to screen and need a more high throughput method. The recombinant proteins have His, Xpress, and V5 tags for detection, and antibodies for these are easily available. However yeast is not easy to permeabilise so I'm wondering whether standard protocols using a fixation buffer with 1-4% paraformaldehyde and a permeabilisation buffer containing detergent will work? 

I am trying to purify a plasma membrane protein from ecoli.

I think the standard procedure for membrane protein extraction is like this:

However, sometimes proteins are in the pellet after the low speed spin (step 2). I think this is referred to as insoluble fraction typically.

For proteins in this pellet, does this guarantee the protein is in inclusion bodies?

Or can properly folded, hydrophobic plasma membrane proteins naturally be here without inclusion bodies?

Hi folks. My protein is about 260 kDa in size and I was wondering if there are concentrators out there with MW cut-off larger than 100 kDa. Of course I could use the 100 kDa ones, but thought it might help polishing the low MW impurities (especially that the protein doesn't come out very pure after gel filtration -S200) if I use one with a larger MW cut-off concentrator. Thank you!

Will be part of curriculum development at community college biotech department. If anyone knows of a reliable source where I could get this I would appreciate it. Thank.

I have a problem with my research about cellulase purification.

1. When I precipitate the cellulase with crystal ammonium sulfate 20% and 40% saturation. I couldn't get the precipitat/pellet. What is the problem?

2. Then, I tried to precipitate the cellulase with crystal ammonium sulfate 60% saturation and I got the precipitat. After that,I centrifuge 4000 rpm 30 mnt and dissolved with buffer phospat ph 9 1:1. But, when I analyzed the cellulase activity with DNS (pH 9,50 degree Celcius), the result was negative. I have tried again and again, but still negative. Although, the activity of crude enzyme positive.

What is the problem of my research ? I have tried to change the dissolved buffer from pH 6.8 to optimum pH (pH 9) and I have incubated 24 hours after add the ammonium sulfate. But, the result still bad. Can you help me,please? Thank you.

I want to incorporate two proteins (6 and 2 TMDs) as a complex into nanodiscs. 

I am wondering if it would be better to express and co-purify them together (either from a single plasmid or with two different ones). I tagged them differently (C-term with His/Strep) to be sure I have both in my discs and tried to express them from pETDUET1 using both MCSs (0.1mM IPTG; diff. temperatures & diff. strains C41/C43/pLysE/Rosetta).

In this case only the His-Tag version in the 2MCS is there but expressing them separately works – more or less – fine, no matter if I use the 1st or 2nd MCS or His/Strep tag. Should I enhance IPTG level or clone both into a single MCS (as a kind of operon or even a fusion protein)?

The alternative would be to purify them in parallel and combine them during the nanodisc step. But in that case, I assume the detergent may inhibit complex formation.

This protein used to express beautifully in large cultures as well as small. It is a his tagged small protein in pET21a. I hadn't needed to make more in a few months and when I tried again it would not express at all. I retransformed, nothing. I sequenced to be sure, still nothing. I tried different strains of BL21, nothing. Then I re-cloned it back into the pET21a, and I managed to get it to express again in a 5ml culture! two different clones worked. Then I scaled up to 500ml, and again nothing! I used an overnight culture of 20ml into 500ml and induced at an OD of 0.5. Any help would be greatly appreciated! I have tried everything! 

It seems that it can do the same purifications as the higher end Akta Purifier/Pure/Avant, it's just that the pumps are peristaltic instead of piston based, but the price is much better!

I am using E.coli BL21 Plyss for a recombinant protein production. The cells debris becomes too sticky after lysis. I spin down the pellet at 13000 g (that is the maximum available here). The problem is I could recover only 45-50% of the supernatant. What can i do to recover most of the supernatant?

Protein was expressed in SHuffle cells in presence of 0.5%glycerol in the LB media(230ml).

pellet was dissolved in 20ml lysis buffer(50mM Tris pH 8.0, 150mM Nacl, 10% glycerol).

i couldnt see any difference to resuspended pellet after applying sonication to the pellet.

(20 mins, 20 % amp, 30s ON, 30s OFF). then i further extended the time to 10 mins i saw little difference in the solution. Then again i extended the time about 10 mins.

In the purification, most of the protein went in pellet. I doubt that cell lysis process is not done effectively or its too much sonication.

-

Before further repeat this experiment, what are the things we can tweak?

Am trying to purify my protein in Tris-Hcl pH 8.0 with DTT. I am keeping the protein for overnight dialysis. I found that, around this pH 8.5 in phosphate buffer stability of DTT is about 1.4 hours only.

I am planning to express a recombinant protein in Ecoli and want it secreted extracellular (into the media). I have been reading that Gaussia's sec signal would work in prokaryotes as well. I wonder if the secretion signal is cleaved or not.

Also, do we know if it follows sec or tat pathway?

Any help would be appreciated.

Thank you

I have been getting problem recently in BL21 DE3 strain for expression of my protein. I co transformed with pLYSs and PGEX-6p2 containing my two insert. Cells were very much competent,as observed after successful co transformation. Do you guys have any explanation on why the BL21DE3 is not expression my proteinat all?

I changed the strain from BL21 to Bl21 codon plus and it started working. Any possible explanation?

I have cloned a gene in pET28a with a protein size of 47KDa, I am obtaining fairly good yield after induction and purification using the Ni-NTA Agarose (Qiagen). I would like to know the easiest and a most rapid method of His Tag removal. I don't think Thrombin can be of help since its slow and cleaves randomly.

Please help.

I have a clone of 9kDa protein in pET28a, at EcoRI and HindIII. Can anyone suggest me, how can I remove the His tag?

Hi All,

I have a very basic question to ask. We generally inoculate a single colony in small culture for overnight growth following transferring them into big conical and wait for the OD 600 to reach 0.6 to 0.8. Now, if you inoculate a single colony early in the morning(lets say 8 am) and by 8 pm to 9 pm you see the OD 600 is already 0.6, can we start our induction? Is it always necessary to do small inoculation first? I know this way is very convenient but I am just asking for the sake of argument. If I am growing a single colony to 0.6, the culture is still fresh right?

The pGEX vector system does not include a termination signal sequence. The termination signal sequence has an important regulatory role in translation.

Does anyone have good experience with the use of pGEX vector?

Thank you for your answers.

My protein of interest was cloudy when it was purified in high concentration. But after overnight dialysis with storage buffer (20mM Tris, 150mM Kcl, 0.1mM DTT, 0.1mM EDTA, 50% glycerol), it becomes transparent with no sign of cloudiness. The protein was centrifuged at max rpm to check if there is any precipitation. There was no precipitation and activity of protein was really good. After a month of storage at -20dC, the cloudiness was seen again. What can be done to avoid such changes in protein.

(NOTE: The PI of protein is same as the pH of elution and storage buffer. Still, many literature including the crystal structure of protein, reported the same buffer conditions).

I have to ship a batch of purified proteins at ambient temperature to a collaborator. The functional state, activity, and folding of the proteins after shipping are not important as they will be used for proteomics analysis. The proteins have been His-tag purified and precipitated with the methanol-chloroform-water method. I am wondering whether it would be better to ship the proteins as precipitates or freeze-dry the precipitate before shipment. Maybe the protein will be less stable as a precipitate compared to as freeze-dried during shipment, thus causing degradation and loss of material. On the other hand, re-solubilisation of freeze-dried proteins could be challenging, which might also lead to loss of material from incomplete re-solubilisation.

I know that each protein could behave differently but I would very much appreciate any comments on whether in general shipment as precipitate or as freeze-dried at ambient temperature could be better, i.e. lower degradation and loss of material?

Thanks a lot in advance!

We want to add a 10xHis tag after the signalling peptide followed by the protein and then design primers for the beginning of protein expression. Can anyone help?

Hello.

I subcloned a gene In pet-28 (his tag in N-terminal) and i have low expression, but the same gene in pet-23 (without his) give high levels. The sequence in both are good and they are in frame...What could be the problem? His tag?

Thank you so much.

I have 9 recombinant GST-tagged proteins. Maximum proteins formed inclusion bodies. I use one kit for solubilization and renaturation. It works well but problem is in elution. Proteins not elute from the Glutathione Sepharose beads. Here I attach a picture of my work. After addition of Glutathione Sepharose beads and O/N incubation, elution procedure is as follows

1. Centrifuge 9000 rpm/500 g for 5 min at RT. Discard supernatant.

2. Wash beads three times (9000 rpm/500 g for 5 min at RT) with 1X PBS containing 1% Triton X-100.

3. Add 10/20 mM reduced glutathione. Mix gently to resuspend the gel. Incubate at room temperature (22-25°C) for 10 minutes to liberate the fusion protein from the gel.

4. Centrifuge at 9000 rpm/500 g for 5 min to sediment the gel, and remove the supernatant.

5. Repeat elution and centrifugation steps twice more. Pool the three eluates.

I did a trial run of a his-purification and found that my protein would be a little dilute for my purposes. Any thoughts on the best way to concentrate a denatured protein?

This is my buffer and protein...

10mM Phosphate pH 7.4, 500mM NaCl, 500mM Imidiazole, 8M Urea

~35kDa

I'm leaning to ultra-filtration since it will likely leave most of my protein in solution. I'd rather not let it fall out if I can help it. I will be going to dialysis next, so maybe I could perform a two in one!

Thoughts?

Dear Everyone,

I’m expressing human recombinant proteins in CHOK1 cells. The expression level is high and the proteins are secreted efficiently to medium. But the molecular weight and N-terminal protein sequencing of one of the proteins suggests that the protein which is present in medium still contains the signal peptide.

Can anyone tell me if there is a method to persuade cells to cleave the signal peptide? I have no idea why they don’t do it naturally. The signal peptide which is used is a native peptide for those proteins.

Thanks in advance,

All the best,

Anna

I'm trying to express a small protein domain that contains two disulfide bonds; I thought I'd try targeting the protein to the periplasm of BL21 E.coli cells to allow these disulfide bonds to form in the oxidising environment. However following cell lysis, when I run a sample of soluble and insoluble fractions on SDS PAGE (and confirmed with Western Blot), my protein is mainly in the insoluble fraction. 

I induce at 16C overnight with 0.1mM IPTG. Of course it's always protein dependant, but I've read that lower temperatures and lower amounts of IPTG are effective, as well as aeration; for example using non-baffled flasks. Does anyone have any further suggestions on what can be going on, and how to improve solubility?

Thanks in advance! 

Thrombin is supposed to cleave the correctly sequenced and in-frame LVPR'GS site in the middle of a ~70kDa extracellular protein known to homotetramerize. The goal is to use the N-terminal MBP for expression and cut it off with thrombin prior to crystallizing the target protein, which is only a 25kDa fragment of a larger construct, but, even at 37ºC, thrombin fails to cleave it (determined by SDS-PAGE). I included my analysis of the possible causes, but I would like for someone who has experienced similar obstacles to tell me how he or she solved them. I've put way too many hours into designing a special vector and optimizing thrombin cleavage conditions to drop this project, so I'm open to any suggestions.

THROMBIN. The thrombin itself should not be an issue since, over a year ago, I successfully used many aliquots of the same batch, which, being frozen in the -80ºC in 2012, should have an almost indefinite shelf life.

HYDROPHOBIC INTERACTIONS. The cleavage site could be buried inside the construct's 3D conformation, rendering it inaccessible to thrombin. I've considered adding a mild detergent, such as a low concentration of SDS, to counteract that, but my PI said no.

REDOX. Since thrombin consists of two subunits bound by a disulfide bridge, it is super sensitive to reducing agents. While I've added no reducing agents to the mixture, the construct itself could have a redox ability. I wouldn't normally consider this due to the infinitesimally small probability; however, I've observed that, when I purify the construct from the lysate using nickel resin, the construct turns the nickel resin and elution product a faint greenish brown color, although, after elution, the resin returns to being good old nickel blue. Generally, color changes + nickel resin = some redox going on.

I am trying to express and purify a human protein in E.coli (His6-SUMO fusion tag). The total protein size is 67kDa with the fusion tag and the protein of interest is 47kDa. The expression is great and about 50% is in the soluble fraction. I usually follow the Ni-NTA purification, size-exclusion, and TEV-protease cleavage. The final buffer is 50mM Tris pH 7.5, 500mM NaCl, 10mM MgCl2, 0.5mM TCEP and 10% glycerol. For some reasons, the final product is not >95% purity. I am seeing clipped products of my protein (~15%) in SDS-PAGE. I use EDTA-free complete protease inhibitors (Roche) during cell lysis and Ni-NTA purification. I have also tried C-term His tag (to check if this is a protein degradation issue) but unfortunately, the expression is very low and I still see the degraded products. Has anyone come across this issue before and what is the best possible way to go about this? Any suggestions would be really helpful.

I isolated my protein complex from mammalian cells. Purify it using affinity purification methods. but after running chromatography the concentration became very low to go further structural study. How can I I increase concentration without affecting the protein complex? 

I need a practical experiences to choose  a strongest promoter that works in cho cell line-DG44.

thanks

Amir

I am expressing ODC protein in E.coli rosetta DE3. After induction with 1mM IPTG, I pellet down 50 od cells. Always I observe that the pellet size of the strain containing empty vector is bigger than the strain with ODC protein. Also, when lysis was carried out @ 4 degrees with 1ml lysis buffer(50mM Tris, 2mg lysozyme, PI ), ODC protein containing strain gets completely lysed while the strain with empty vector doesn't get lysed in 45min. I have tried increasing incubation time to 90min for the mock cells but only 60-70% lysis was observed. I don't understand why there is an abnormality in cell concentration and cell lysis of the mock? 

Hello Everyone,

I am trying to develop a protein expression and purification system. Hence, can anyone tell me about "Intein Tags" or related papers, articles or suggestion? 

Please Help

Thanks

Hey everyone,

I am trying to do protein expression with BL21(DE3) cells and I am growing my cells in TB. I ordered the TB from RPI and added 4mL glycerol before autoclaving for 30 min. After innoculation with starter culture, the cells seem to be doubling fine (~20-25 min doubling) and I induce at OD of 0.6. I then come back to my cells and each time the OD is around 2-2.5 after 3 hours of induction at 1mM IPTG. I did a control where I did not add the IPTG and the cells only made it to OD of 2.8 after 3hrs induction. I am not sure if there is something wrong with my media, but it doesn't feel like a toxicity problem because the cells seem to be dividing fine during the earlier growth phase (leading up to OD 0.6). Do you guys have any idea what could be causing this issue?

I am growing 1L at 37C in a baffled 4L flask with rotation speed of 250 rpm.

Any suggestions are appreciated!

Best,

Joe

Dear members,

I am trying to express and purify a mouse-derived enzyme (rotamase A) having 7 X His-TEV cleavage site.

The protein overexpression at 37C (0.8M IPTG induction for 3 hrs) shows a clear band in SDS-PAGE. However, the purified protein (after Ni-NTA) shows autodegradation resulting two closely placed band in SDS-PAGE with identical intensity. I tried cell lysis with and without protease inhibitor cocktail + PMSF. Unfortunately, there is no difference and two bands are observed. 

It will be a great help if someone could help me in finding the problem of such autodegradation. I look forward to your comments and suggestions.

Thanks and regards,

Bikash

Dear all, 

We would like to check the purity of a GST-tagged recombinant protein by SDS-PAGE. We purified the protein using Baculovirus expression system.

We run the reducing SDS-PAGE (GST was used as a positive control) to check for purity. But unfortunately, we can only see a small band of our target protein (the expected molecular weight: 65 kDa) except for GST and some cleavages. 

I don't know whether it was due to the reducing SDS-PAGE that we used or the protein purification has failed? If it is possible, I hope to receive any of your comments or suggestion. 

Thank you, 

Yours sincerely, 

Tu

Hello, I'm trying to see the overexpresion of a protein about 8KDa in P. aeruginosa or E. coli. The protein is cloned in a pHERD vector and it has a 6His-tag at the C-terminal but I have not succeded in visualising it either in SDS-PAGE tris-tricine (staining with silver, Schägger 2006) or by WB.

I'm starting with a 500mL culture, but it rarely defines bands below 10 kDa (according to the marker).

Does anybody have a better protocol to follow in this case? or any piece of advice?

I am working on recombinant protein (Ig's) expressed in form of inclusion bodies. Upon giving certain treatments there is a up shift in protein band. If my protein is degrading during the process; there must be "downshift" but here the scenario is different. I am unable to find out what may be the possible causes? What changes might occured in protein since protein is in the form of IB's.

Image: All lanes have same samples of same batch with different treatments. Lane 9 is old previous ref sample; where protein should be.

Thank You.

I use the E.coli system to express the protein.It's a heat-stable protein and it's only 9KDa. I use the HiTrap Q HP column to purify it but I find a huge contamination.I guess it is nucleic acid.The UV absorbtion is so high and I can't identify the peak of protein.So, who can tell the what the contamination is and how to get rid of it?

I am trying to purify a recombinant protein for its functional study and I want to do some biochemical assays. Due to hydrophobic nature of the target protein most of its fraction forming inclusion bodies. I have tried various experimental conditions to increase the protein concentration in soluble fraction like induction at different temperatures and IPTG concentrations, tried with different tags ,e.g., His, GST, MBP. With MBP tag the protein concentration in  soluble fraction increased to very little extent. But this increased concentration with MBP tag is not sufficient to subject the protein to second column purification like Gel filteration or ion exchange. Solubilization and refolding of protein from inclusion bodies is a tricky process and the protein may not be functionally active. So I am thinking of in vitro transcription translation system. Is it possible to purify the target protein after its synthesis by in vitro translation system so that it can be used in biochemical assays. Thanks in advance.

Dear colleague

does anyone know how to purify the secreted protein from leishmania into the superntant. I used leishmania as expression system for my protein and it should be in the superntant, I want to determine the protein by WB then purify it

can anyone help me in this issue or if there is a protocol from recent papers that can I fellow ?

I am working with a small protein using instant TB media. I grow bacteria at 18 degree for 36 hours, OD reaches 15-16 at harvest. There is induction, but induction is very low and n-terminal sumo tag has been mostly cleaved. Any suggestion?

I have been trying to express my recombinant protein (15 kD) for some time now without success. I have tried @ 37 C, OD600 0.6 for 5 hours with 1mM IPTG and I have tried 32 C, OD600 0.5 for 4 hours with 0.5 mM IPTG. Does anyone have any suggestions as to the conditions to express such a small protein. 

My protein has HisTag x6.

I'm performing IMAC to purify my his-tagged protein (enzyme) from e. coli lysate. It seems I can get a good amount of my protein of interest but the problem is that I also get alot of coeluates, see attached file.

My protocol (everything at 4C):

Spin down 1L bacterial culture and sonicate the pellet in 50ml binding/wash buffer (20mM Tris, pH8, 20mM Imidazole, 500mM NaCl) --> spin down --> incubate lysate using 2ml nickel chelating resin from G Biosciences (786-281) overnight --> wash 4x 20ml --> elute in 9x 1ml increments in elution buffer (20mM Tris, pH8, 500mM Imidazole, 500mM NaCl)

After this I tried troubleshooting as indicated by G Biosciences: use less amount of resin, elute step-wise by increasing imidazole, reduce lysate-resin incubation time.

So I tried another experiment: 500ml bacterial culture sonicated in 25ml binding/wash buffer --> incubate lysate in 0.25ml resin for 1hr --> wash 4x 20ml --> elute in 7x 0.5ml increments in elution buffer with imidazole ranging from 100mM to 500mM (also incubated the flowthrough O/N with another 0.25ml resin and performed same experiment)

However, I still get background proteins. Any thoughts on this will be greatly appreciated. 

I just want to check if the transcription factor exists or not. I don't have access to antibodies so Western blot can't be used. But I was wondering how can I determine  the presence of just some protein based on SDS page. 

Experiment:

There is a host gene expression, 5 minutes after infection with pathogen. For negative control we have a host cells with no infection at time 0 and for positive control we have host cells that die 15 minutes after infection. 

I am trying to express a functional protein in pet28b in BL21. When I run my SDS-page gel I see my protein in the soluble and insoluble fraction. However, when I perform a Western Blot I only see the protein in the insoluble fraction.

How do I go on about to recover my protein as functional?

When I run an SDS of my Ni-aff purified protein I get 2 bands, one at approximately the correct mass and one 2kDa higher. I confirmed these masses by MS and also ran an assay where I attach a prosthetic group to the protein. This results in both the proteins increasing by the expected mass, suggesting they are both ACPs.

I've tried different strains of E.coli, gel filtration (but the masses are too close), using a cobalt column instead of nickel, running the Ni-aff on an FPLC with a gradient and can't overcome this problem. Another member of my lab has come across the same problem, but hasn't managed to solve it.

Can anyone suggest ways this could be resolved? I've seen suggestions about changing the loading buffer of the SDS-PAGE gel, but this wouldn't get rid of this second band.

Thanks.

I am trying to reproduce an old purification protocol from 2004 that claims the use of a pseudo-affinity chromatography based on Green A Dyematrex gel (Amicon). The product is discontinued from Amicon catalog. Any alternatives around ?. Thanks

Though i followed the expression and purification conditions exactly in the paper, i could not see the expression. What could be the reason? Did you face same problem in reproducing the results of protein purification?My vector is pET28a expressed in BL21(DE) at 0.4mm IPTG at 16C for 20 hours. The sequence is in frame.

Your assistance will be highly appreciated.

Hi!

I have been having trouble in expressing his-tagged streptavidin by following Mark Howarth's protocol,"Imaging proteins in live mammalian cells with biotin ligase and monovalent streptavidin" in Nature Protocol 2008. This paper calls for dissolving inclusion body in 6M GnHCl, then refold in 1x PBS. After that, ammonium sulfate precipitation is carried out and his-tagged streptavidin will be pelleted out by centrifuging at 17700g, 4 degrees, 15 mins. 

I followed the above steps but could not get a pellet after centrifugation. I have then tried to increase the concentration of ammonium sulfate because I thought the ammonium sulfate concentration might be too low. But then, still no pellet.

Is the precipitation step necessary? Could I skip this step and purify the refolded protein in Ni-NTA column?

Here's the link to the nature protocol: http://www.nature.com/nprot/journal/v3/n3/abs/nprot.2008.20.html

Thank you!!

I'm working with E3 ligase human RNF125. I've trouble to get expression using RNF125 antibody. In the market different RNF125 antibody available but I'm confused which one is better to get western blot expression. Is there anybody having experience using RNF125 Antibody and got Western blot expression fine? plz suggest me. Thankx in advance

i have change an amino acid codon asparagine to aspartic acid in my recombinant protein plasmid then transformed in bl21(de3) and used it for expression of my new mutant protein.I expressed it using 0.6 mMIPTG. WHEN  I run in SDS PAGE ,its band was not that clear compared to the wild type recombinant protein. Is it because of the mutation I have done? 

Hi,

I am trying to get plasmid profiles from clinical isolates. I extracted them and visualized them in the gel electrophoresis. I tried to cut them with a 6bp enzyme XbaI to have more ideas on the size of the plasmids (anyway, I chose an enzyme randomly, since I don't know the sequence of the plasmid). I have some difficulties in the interpretation of the data and I attached the file (sorry the image is not great). They are 6 isolates and for each I run the cut and the uncut plasmid. I don't know how to determine the size of the plasmid and how to determine if I have more plasmids in my extract.. For example, as to the first isolate (K1), the cut  produces 2 bands, one of which is larger than the only one band visible in the uncut product.. in your opinion, could the bands be the two pieces after cut of the plasmid? If yes, why are they larger than the single band visible for the uncut plasmid? does it mean that the uncut product is a supercoiled form e migrate faster?...the other example, the cut of the plasmid extract of K5 produces different bands, again,  larger than the K5 uncut. Are they pieces of the same plasmid after cut or different linearized plasmids?Again, the fact that I have bands larger for the cut plasmid DNA can suggest that the uncut plasmid is in the supercoiled form? I used the 1kb ladder

I performed my PCR of genes with sizes between 1000 to 2000pb to volumen of 60uL. Later, I purified with Ethanol-Acetate precipitation (or sometimes with PCR purification Kit), I made ligation with TA cloning kit (Invitrogen) and I transformed in competent cells (made in laboratory). I have verified my PCR and purification products in gels. Besides, when I have transformed I get a good positive control with a high quantity of colonies. But, my samples do not have colonies or have very few colonies. I have had some experiments with good results using this flow, but these do not function now.

Could you send the protocol for detection of recombinant protein on the surface of bacteria by FITC anti His tag antibody. My E.mail: hamedir2010@gmail.com

I have been trying to express a kinase recombinant protein in BL21 DE3 and C43 but couldn't get any expression.

I used 2L LB culture and 0.5mM IPTG induction.

I have even tried with colonies from fresh transformation into fresh competent cells still couldn't observe expression.

I would like to know which steps could have been gone wrong?

I constructed a pET-53 to express yeast gene in E. coli BL21(DE3). The culture condition was culture the BL21(DE3) until OD600 = 0.4 

Hi All,

Is 8M Urea compatible with Q and SP sepharose resins? Please let me know.

Thanking you in advance,

Jyothish 

How should I choose an effective protease inhibitor which can can be used for the purification of a range of proteins to be purified by Q-sepharose or Ni-NTA  or SP-sepharose?

I started to work with different fragments of a 500 kDa human protein in E. coli. We got the X-ray structure of one small domain of 30 kDa (after expression in E.coli).

I am trying now to express different construct around 100 kDa in E.coli but it's not working really well. I got a weak expression visible on Coomassie but when I follow with a gel-filtration, it goes out in the void, so I think I have aggregation... I tried different plasmid (pCold and pGEX), strains (Bl21*, Lemo21, Gami, Rosetta, C41, C43), expression conditions (temperature induction from 37°C to 16°C, time of induction and concentration of IPTG), co-expression with E. coli chaperones (GroES, GroEL, dnaK, dnaJ, grpE and tig) and different buffers during purification (with DTT, betamercaptoethanol, HEPES, Tris, Bicine, MES, different salt concentration, different pH) but nothing seems to improve it.

I also recently try one of the construct in insect cell, but again the construct aggregate.

So I want to continue on these fragments with any ideas, you could have? I also will like to try to express the full-length protein but I don't know what will be a suitable strategy for it?

Many thanks in advance to anyone that can help!

I am trying to express a membrane protein that, in the literature, was fused to a TrpLE tag. I am having trouble finding the actual sequence or a plasmid that encodes the TrpLE leader sequence. I would appreciate any information on how to track down the sequence or a plasmid that encodes the sequence. 

Thanks!

Chad

I am working to isolate and purify a protein that I have inserted into an IPTG inducible plasmid in BL21 cells. Most of my reading has said to induce with 1mM or less IPTG, but thus far I have had low concentrations of isolate. Will it cause any issues to use a higher IPTG concentration for induction and are there any other factors that I should keep in mind when optimizing?

recently i learn some knowledge about co-expression.

the EMBL website said, in the case of co-expression from different vectors, To ensure plasmid stability, the vectors should have:

(1)different selectable markers, usually antibiotic resistence markers.

(2)different origins of replication.

But I check some publications, they did co-expreesion from 2 vectors with different resistance markers but same replication origins. like pET16b-geneA and pET28a-geneB. or  pET22b-geneC and pET28a-geneD. and both were succeed to express two protein from single colony.

so whether it means the ''different origins of replication'' is not necessary?

and in my experiment, protein PCID2 are inclusion body in several different plamid, I try to obtain soluble protein PCID2, and then i did co-expression with its binding protein Y, i co-transformed pET16b-PCID2 and pET28a-Y to E.coli, the replication origin are both pBR322, pick up single colony, Y is soluble, protein PCID2 is still inclusion body. I do not know if my co-expression succeeded or not.

I am working on pBADmyc HisA and pFLAG-CTC vectors for expression of my gene of interest. My construct is like this in pFLAg one ATG of the pFLAG vector after RBS and then there are 5 amino acids and then ATG of my gene of interest). I am wondering to ask that are two ATGs making problem in my expression? as im not getting any signal on western blot?

We are expressing thousands of proteins in 96 well monolayer culture. Does anyone have experience to share about optimal vectors/cells? Serum-free conditions are not needed, but membrane protein yield is important