Sudhir, The manuals always use some standard protein which expresses well. So, before even you worry about the volume of culture, you need to make sure the protein is expressing enough to see by coomassie staining.

I think you might be using ITC or SPR or protein-protein interaction, if so you need good amount of protein. Again, you are working on very big protein. So, I would think you would benefited if you workout the expression of the protein. Daniel has made really good points. for example " A good rule of thumb is that if you can't see the band appear by Coomassie, then you will not have enough material to warrant purification."

1. work on the expression- you can lower the temperature, change the expression strains, changing tag (his to GST or MBP), changing vectors, several other factors could be varied to get this done. If you are getting protein which could be detected only by western blot or silver stain then you could do only pull-downs are the best options.

2. You could even work with domains instead of 238 kDa protein. I dont know if its necessary to use full length protein for interaction studies. Small domains expresses well.

3. once you are sure that you could see the protein induction, then check the solubility by taking small volume and binding with beads. Chose denaturing protocol only if its not soluble.

4. If its soluble, then any manual can guide you through the rest.

Have your tried running coomassie on the flow through? Can you try a western blot for your specific protein? Finally, it may be worth checking into a silver stained gel for low concentration samples. If you run these techniques on eluted and flow through, you should be able to locate where your protein is. Good luck.

1. one point is that what was the concentration of imidazol in eluation buffer? is it high enough? you can try by different concentration of imidazol (from 20 to 500).

2. maybe you did not expressed your protein at all. you can check it before purification. because the concentration is low.

3. you can run a SDS-PAGE gel for pellet, FTh, washed protein and eluated one. then if you can see the process of purification.

it will be better if you check the pallet by SDS-PAGE gel to make sure that you have the desired protein.

Re-fold the protein before passing through the His6 column.

you can optimize the elution buffer composition. Sometimes lowering the imidazole concentration helps.

Your yield is low in elution for this method.

1. check the beads. In denatured purification, some times its hard to elute unless you use low pH as Elena Shersher suggested.

2. I assume that since you had good expression but little soluble protein you chose denature method otherwise you need to work on expression

3.This manual is very nice and might help you. http://kirschner.med.harvard.edu/files/protocols/QIAGEN_QIAexpressionist_EN.pdf

Good luck

Are you expressing your protein in E. coli or another system? How do you know that your protein is induced/expressed? These protein concentrations you report are about 100-500x lower than what I would expect for a well-expressed recombinant protein. If you have any Coomassie-stained gels of your different fractions (non-induced, induced, solubilized fraction, column flow-through, etc), can you please post it? This would help in getting the best possible troubleshooting advice.

I'd suggest to go through step by step procedure and get a sample at each stage (bacterial cells pellet, unpelleted lysate, pellet, supernatant, flowthrough, wash, elutes ) and run on SDS PAGE. This will help you in establishing the protein loss if any. As mentioned by people above, do play around with the wash and elution buffer concentration as well as bead volume.

Can you please mention the buffer concentrations, and bead volume??

I have found the wash beffer concentration to be anywhere between 10mM to 30mM imidazole (you need to standardize). Elution concentration that i use is between 250mM to 300mM.

You may also play around with the buffer.

EDIT:

The method of binding to bead can also be altered. For example, if you are doing the binding by column method, then try batch method, it is generally more efficient in binding of proteins.

run a his-vision in gel

I wud suggest

1) Check whether ur protien is expressing or not? either by western using anti-HIs or SDS-PAGE comparing it with uninduced.

2) If expressing, check whether its binding to beads or not, which u can do by loading a small volume of beads on SDS-PAGE and do a western or compare with flow through and Clear Lysate looking for the band of interest.

Hope this helps...

the concentration you mentioned is beyond the sensitivity of coomassie stain. first check uninduced and induced sample on SDSPAGE before taking your protein to column..

Thank you all for the response. As many of you have suggested I have bought the anti-His antibody so i will come to whether it is expressing at all or not. but this will take sometime. I am adding some more details here;

1. I have run the lysate, wash protein as well as elute on SDS PAGE. I see band in all lanes except the eluted one. since protein conc is very less that might be the reason why i am not able to see band in CB staining so next time I would try silver-staning.

2. I am extracting the protein in denatured state so there is less possibility of hindered His-Tag.

3. I am not adding imidazole in binding or wash buffer not even in elution buffer. Buffer conditions are attached. I used buffer B, C and E.

4. It is an S. aureus protein (~238kD) that I am trying to express in E. coli Bl21 pLysS, cloned in pET21b(+) vector. I am using 1 mM IPTG for induction.

One more thing I should have told that gene contains several repetitive sequences therefore I am inducing at 30oC instead of 37 to avoid gene-instability. I have tried 4 hr as well as overnight induction. But I got the same protein concentration in elute.

Until you can see a clear difference in the Coomassie blue staining for induced vs uninduced samples for a band at the appropriate molecular weight, it is impossible to tell what the problem is, or even whether it is expressed to begin with.

Make sure you are using freshly transformed bacteria for your experiments, and please indicate the culture volume you are using for your purification experiments, since this has some bearing on the expected yields.

Sudhir- how much protein do you need for your experiments? If a few micrograms suffice, you might have better luck with an in vitro transcription/translation system. If you need hundred of micrograms to milligram quantities, then there is little to be gained from more sensitive assays (silver stain or Western blot) since you still have to address why your overall protein levels are so low at all points in your experiment.

@DANIEL Sir, I am using the 30ml culture volume.

Do you mean without column purification i should run the induced and uniduced cell lysate, right? i will try this followed by silver-staining as yield is too low. I have stored the columns 4oC. I will also try with modified elution buffer by adding 300mM imidazole.

Sudhir- typically one would try a few liters of culture to get adequate yields of a recombinant protein on a column purification. 30mL is simply too little starting material. If you run 1-3 OD's worth of induced vs. uninduced bacterial (whole cell lysate) on a polyacrylamide gel (7.5%, since you have such a high MW protein), you should be able to see a band by Coomassie staining. A good rule of thumb is that if you can't see the band appear by Coomassie, then you will not have enough material to warrant purification.

Silver staining and Western blot are time consuming and are not always trivial to interpret, so I wouldn't recommend spending a lot of time on them. The exception would be if very low yields of protein are acceptable for your downstream applications.

my ultimate goal is to study protein-protein interactions. I really dont have idea and havent given any thought how much protein will be needed for this pupose. first of all i need to express in good amount.

litres? Actually I just followed the protocol given in Qiaexpress. where they recommend 30 ml- 50 ml culture to start with. I hope increasing the amount of culture would help.

Most protein-protein interactions occur with micromolar affinity. So to measure binding, you'll need mg/mL concentrations, unless you have an extremely sensitive label such as radioactivity which will allow quantitative measurements even at low levels of binding in dilute samples. Increasing the starting volume of culture will help improve yields, but you likely face challenging conditions with such a high MW protein (yields are typically inversely proportional to protein size).

Sudhir, The manuals always use some standard protein which expresses well. So, before even you worry about the volume of culture, you need to make sure the protein is expressing enough to see by coomassie staining.

I think you might be using ITC or SPR or protein-protein interaction, if so you need good amount of protein. Again, you are working on very big protein. So, I would think you would benefited if you workout the expression of the protein. Daniel has made really good points. for example " A good rule of thumb is that if you can't see the band appear by Coomassie, then you will not have enough material to warrant purification."

1. work on the expression- you can lower the temperature, change the expression strains, changing tag (his to GST or MBP), changing vectors, several other factors could be varied to get this done. If you are getting protein which could be detected only by western blot or silver stain then you could do only pull-downs are the best options.

2. You could even work with domains instead of 238 kDa protein. I dont know if its necessary to use full length protein for interaction studies. Small domains expresses well.

3. once you are sure that you could see the protein induction, then check the solubility by taking small volume and binding with beads. Chose denaturing protocol only if its not soluble.

4. If its soluble, then any manual can guide you through the rest.

Try expression at 18 C overnight with 0.5 mM IPTG induction. If you see protein expression in induced cells as compared to un induced cells, it's a start. If there are several repeats in the sequence try codon optimising.

hi Rucha. yes some people have suggested me to use lower temperature for induction to increase the solubility of protein. I would give a try.

Sudhir, Optimizing expression takes a while but do it systematically to rule out each parameter. Don't do a random testing and follow shortcuts because if it works it will be fine otherwise you will be back in square 1. If you don't have any one in the lab or department to guide you through this tricky situation it might be little frustrating but if you read everyone suggestion you can come up with a systematic way of doing this you can tackle this. This is one time thing and it will become easy for you to work on this even for structural studies. I forgot to remind you to see if there has been some work done on the this protein? Feel free to seek more help as everyone is willing to help. Keep us posted with what worked for you. Good luck

When optimizing your expression conditions be sure to analyze pellets and supernatants (especially after cell lysis) to make sure your protein isn't being produced in inclusion bodies. Changing the temperature for expression is generally a good place to start for optimization, but you may also want to consider the concentration of IPTG used for induction (I've used as little as 10 uM), expression medium (LB vs 2XYT vs Terrific Broth, etc) and OD at which you induce protein expression.

Sudhir,

- you said that you have only about 5 ug/ml of protein in your eluate which is not very much indeed. I do hope that you concentrated your eluted sample before gel loading, unless you will barely see much with coomasie staining (as sugested do a WB with anti-His antibodies).

- As Daniel said, it worth to first check induced/uninduced on whole cells and then of soluble/insoluble extracts.

- Using denaturating conditions usually means that used the protein are expressed in inclusion bodies (unfolded state) and generaly in large amounts. As suggested GST of MBP tags can help to solubilize your protein and avoid refolding step which can be very painfull without enzymatic activity to validate the refolded protein.

- Finally, if you have an elution issue, you can use 0.5M EDTA.

Sudhir- one other factor to consider is rapid proteolysis of your protein. Try measuring induction after IPTG addition at a few different time point (0.5, 1, 2, 4 hrs). Also make sure your lysis buffers are supplemented with protease inhibitors.

Just save an aliquot of all your steps and then do a Western Blot. This will help you find out if you have protein in the end, and if not, it shows you where you lost it or if it even was there in the beginning....

10 micrograms per mL is very low. If you loaded 10-15 microliters of this concentration you might see it on Coomassie. Use an anti-HIS antibody and determine if your protein is in your elusions by western analysis. If it is not detectable, then take a step backwards and perform a western with anti-HIS to determine if your protein was even expressed.

Sudhir, let me add my voice to that which Daniel wrote above, you must always have protease inhibitors present or you wide up with fragments not protein. Micheal is right about Westerns but only after you get protein bands to know it is your protein that is expressing.

Hi Xavier. using EDTA would really help? Somewhere i read that EDTA may chelate the Ni from the column and would end up in less binding affinity of column.

@Danial. yes I did use a cocktail of protease inhibitors.

Almost all advises given here may work, but if not, there is a trick that may. You can add to your culture 30 or 40 minutes after induction one antibiotic, rifampicine to a final concentration of 150 ug/ml. This will block any transcription except for your protein that is been transcribed by T7 RNA polymerase and its not susceptible to this antibiotics as the natural RNA polymerases o E. coli. And you can incubate your culture up to 6 hours after that. This sometimes diminish degradation and solve other problems associated with the expressing system. Have you sequenced your expression clone? Be sure that you have a proper his-tag, another way may be an antibody anti-his-tag. I also use to assay everything (expression, solubility and time after induction, IPTG concentration etc) by essaying in small scale, that's mean 1 ml of culture, lysis with lysozyme and bath chromatography with 50 microliters of Ni-NTA-agarose in eppendorf tubes and by centrifuging and resuspending gently the matrix with a pipette. You can equilibrate, bind your protein, wash it and elute it, just by centrifuge and decant. Centrifugation steps at 5000 g, 2 minutes. Binding steps 15 of shaking in a rotation step. You can prepare a matrix with the conditions that you want to test and run SDS-PAGE to see all and scale up the best condition. Less than two days of work. It is my special shutgun strategy that almost never fail. Good luck!!

Hi Sudhir,

First to answer your question about the flow-through protein concentration, the flow-through protein concentration should be higher than the eluted sample concentration, especially if your protein is not expressed at high levels. But those protein concentrations tell me that you are loading a very dilute sample onto the column. How are you lysing the cells? The cells should turn viscous when they are lysed due to the release of the chromosomal DNA. And then that DNA has to be broken up before the sample can be added to the column, how are you doing that?

You said that you see the protein band in all lanes except for the eluted sample, so that tells me that you are getting decent expression and the protein is not being degraded. So you passed the crude lysate over the column, did you then see the protein band in the column pass fraction? If the protein is not in the pass fraction then it bound to the column. Being such a large protein it may bind to the column using several different mechanisms and low pH elution may not work well. If you still have the column, you could open it up and take a sample of the resin (25 uL) and heat it in SDS/PAGE buffer (85oC) and then run that on the gel to see if the protein is still bound to the resin. If the protein is in the pass fraction, then it did not bind, and that could be due to the lysate/protein not having enough time in contact with the resin. You could try doing the purification in batch mode to give the His-tagged protein more time to bind to the resin (see below).

I have done purifications in batch mode when scouting new conditions. Take 1-2 mL of crude lysate and add 25-50 uL of resin (equilibrated in binding buffer) then incubate at 5oC for 30 min. to 1 hr with rotation to keep the resin suspended. Then you wash the resin and take a sample of the resin and run on SDS/PAGE. You can also try different conditions to elute the protein off of the resin in batch mode and then check by SDS/PAGE to see if the protein eluted, just use small batch elution volumes (100 uL, and repeat batch elution 2 times) that way the protein concentration will not be too low to see on SDS/PAGE.

I want to design his-tagged primer. My purpose is I want to use his-tagged primer to amplify my dna sample which already recombined with plasmid,namely pklac2. Then, I transform into e. coli competent cells and put on LB plate without X-gal/IPTG. After I got colonies, I screen a few colonies and need to confirm which one has positive colony via confirmatory colony pcr. Then, I inoculate the positive colony to culture in lb broth. Next, I purify the culture using purification kit.The purified his-tag dna plasmid will be used for yeast transformation for expressing his-tagged protein. Are all consecutive procedures in a good order?I just want to know whether my work is ok or not.

@Sudhir. About your EDTA question, you have to use it in replacement of high concentration of imidazole for the elution and NOT during the binding. As you said, il will strip the Ni from the resin.

Hi Budak,

I would post your question as a new and separate question/post.

You may need to run western blot to detect if your protein still has his tag attached. Sometimes, the his tag is cleaved from the expressed protein, We also saw buried his tag from crystal structures.

You can do a western blot on a native gel to find whether HIS tag remains exposed and intact. Also, as a checking step before every cloning, Model your protein using online servers and find out whether the region you have added HIS tag to, will be exposed or folded inside. Sometimes, It may be folded in a manner that its not exposed and hence not binding to the Ni-NTA column but still showing a positive result with SDS + Western blot.

Hi everyone,

Last few months I have tried many things as per your suggestions. It took a little time to optimize the protocols.

Now I am getting expression of my protein, which i have confirmed by both SDS-PAGE as well as western blot using Anti-His antibody. I am facing problem when I try to purify the protein using Ni-NTA column. Actually it remains insoluble and going into pellet. For SDS PAGE, I can solubilize it by increasing the SDS concentration but when I will have to do interaction studies i will have to purify it in native condition. How should i go about it?

Please suggest how can I improve its solubility ?

Thanks in advance.

Sudhir

You can try adding different buffers with varying amounts of reducing agents like TCEP or BME. And you can add arginine or detergent to improve solubility. I would recommend getting a refolding buffer kit, and diluting your protein into each buffer and then see what concentration of soluble protein you can have by running SDS PAGE. So is any of your protein soluble, at first, and then it crashes out, or does it go right to inclusion bodies? If it goes right to inclusion bodies, ie a white precipitate after lysing the cells, then you will likely need to solubilize in things like urea or guanidinium HCl, purify, and then refold. If there is some soluble material after lysis, then you can play with the buffers to optimize the solubility, ie, pH, detergent possibly, glycerol, reducing agents, arginine.

If these things don't work, then you need a new construct

Good to hear you got the protein expressed. I would try to solubilize the protein in 6M GuHCl, if that works then load it onto the column and see if it binds, if it binds then wash the column with your standard wash buffer plus 6M GuHCl until you reach baseline. Then do a "gradient" buffer exchange, while the protein is bound on the column, to get rid of the GuHCl. If things go well, your protein will refold on the column and then you can elute it using a standard elution buffer. I have done this procedure and it worked fine.

Refolding on the column works well as Gary recommended. However, if the protein tends to precipitate to begin with, you will likely need to know more about what buffer to refold in, before you can do the buffer gradient on the column. And when you add the guanidinium, it is important to also add a reluctant like 5 mM TCEP or DTT.