I've used a number of different methods to try to reproducibly quantify phosphorylation. First off, it's tough. Any of the gel based techniques are tricky - they take a lot of attention to detail, careful execution, experience and a little bit of luck. I've used the Marston phos-tag protocol (Messer, AE, et al. Proteomics Clin App, 2009) and I've been able to get it to work reproducibly. One of the most important factors I've found, is that your acrylamide mix is REALLY fresh. We were using a biorad premade acrylamide but, even though it wasn't expired, we had variable results based on the age of the solution. We now make our own acrylamide (and bis) and use it for no more than one week. The other thing that we've found critical is temperature regulation. Depending on the salt concentration in your sample, there can be local heating of the gel, so it's important to have similar sample compositions in each lane (including unloaded lanes - at the very least, use sample buffer) and to use low enough voltage to keep the gels from heating.

Good luck. This is an important issue that is often overlooked. You might want to look into one-dimensional isoelectric focusing to separate phosphoproteins - but that's a tricky protocol as well.

Hi Daniela!

The Mn -based Phostag is usually not comparable with the Zn- Based Gel. For every "new" protein we test Zn vs. Mn and usually the resulted pattern is highly reproducible, respectively - at least with purified protein. You said, that you treat your protein with chemicals - are there any chelators in your reaction mixture? high concentrations of divalent ions ?

I've used a number of different methods to try to reproducibly quantify phosphorylation. First off, it's tough. Any of the gel based techniques are tricky - they take a lot of attention to detail, careful execution, experience and a little bit of luck. I've used the Marston phos-tag protocol (Messer, AE, et al. Proteomics Clin App, 2009) and I've been able to get it to work reproducibly. One of the most important factors I've found, is that your acrylamide mix is REALLY fresh. We were using a biorad premade acrylamide but, even though it wasn't expired, we had variable results based on the age of the solution. We now make our own acrylamide (and bis) and use it for no more than one week. The other thing that we've found critical is temperature regulation. Depending on the salt concentration in your sample, there can be local heating of the gel, so it's important to have similar sample compositions in each lane (including unloaded lanes - at the very least, use sample buffer) and to use low enough voltage to keep the gels from heating.

Good luck. This is an important issue that is often overlooked. You might want to look into one-dimensional isoelectric focusing to separate phosphoproteins - but that's a tricky protocol as well.

Lori thanks a lot for your suggestions: do you mind to give me your working protocol, please?

I don't know the details of the protocols, but maybe there is a difference in use of acidic conditions. If your protein is phosphorylated e.g. on histidine residues this modification is labile in low pH.

Hi, Daniela. I am Eiji Kinoshita, who is one of developers of Phos-tag SDS-PAGE, at Hiroshima University in Japan. Thank you for your question and using our Phos-tag reagent. And I apologize for your inconvenience in using our Phos-tag Acrylamide. I will answer your question.

Although there are some differences in the electrophoresis migration patterns of certain phosphoproteins between the alkaline Laemmli's buffer system and neutral Bis-Tris buffer system, I think the the profile will be basically identical in case of a protein with only one single phosphorylation site.

If you allow, please describe more details information on your troubles.

I can introduce you our Lab's protocol and recent publication on both approaches.

Please access the following URL site for download!

Lab's protocol:

http://home.hiroshima-u.ac.jp/tkoike/protocols.html

(Click at Phos-tag Acrylamide, this URL is a Japanese site, but you can read the term of Phos-tag Acrylamide.)

Recent publication:

http://www.springer.com/biomed/neuroscience/book/978-1-61779-823-8

(Click at Download Sample pages 1 (pdf, 632 kB), our article is free as a sample.)

Dear Eiji,

I am following already your protocol as reported in the website and the problem is that I get always distortion of bands. So the purpose of using this system is lost.

If you can help me, I will be very glad.

I can tell you the details of my procedure:

I work with 293T cells (treated with 0.5 uM Thapsigargin for 0, 1h and 8h). Then I lyse the cells with RIPA buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1% NP-40, 0.5% Na-Deoxycholate, 1mM EDTA) and I measure the protein concentration and I prepare for each sample a solution of 1.3 mg/mL (diluted in RIPAbuffer and containing sample buffer 3X). Then I boil them 5' and I load 60 ug into a 6% Phos-tag gel (50uM) (MnCl2 recipe) MINIGEL. I run the gel very slowly over night and over day , I wash 5 times with 1mM EDTA and transfer on a nitrocellulose membrane with a tank system over night at 6V and over day 24V (but the transfer is always PARTIAL).

The bands are distorted even if I load the protein standard marker far from the samples.

I tried also the Zn recipe but in my hands the results were even worst.

So I know that EDTA can cause distortion of bands therefore I tried to quench it by adding the same amount of MnCl2 but still BAD!

Have you an idea where I am doing wrong, please???

I really need this system working!

Thank you

Dear Daniela,

Thank for your reply and information on the details of your procedure.

There are some points I can advice. I hope they will help you.

Your procedure is almost OK, but the amounts of the sample you applied and concentration of Phos-tag you used are maybe large for the Phos-tag method.

Because the efficiency of protein transfer from the Phos-tag gel is not so good, I can understand your strategy. However, it is recommended in the case of procedure using the mini-slab gel that tests should be conducted by using smaller amounts of the sample and lower concentrations of Phos-tag for complex cell lysates that contain various phosphorylated and nonphosphorylated proteins, and low-molecular-mass biomaterials. We usually use less than 10 µg of total proteins and 20-25 µM of Phos-tag when analyzed cell lysates. Please try such conditions and determine optima for your target.

In order to increase the electrotransfer efficiency, furthermore, the addition of SDS to the transfer buffer just before loading is recommended. The optimal percentage (w/v) of SDS (e.g., 0.05–0.2%) to achieve adequate electrotransfer should be determined. However, the efficiency is dependent on proteins. So, please notice that some hydrophobic (or small) proteins may pass through the blotting membrane If you use more than ~0.2% of SDS. We usually select 0.1% of SDS in universal experiments.

And also, the use of PVDF membrane is strongly recommended for the Phos-tag method.

Good luck for your further works!

Thank you for your answer, I will test your suggestion.

But actually my problem is not the transfer but the distortion of bands (see the gel attached).

And I am wondering if the problem is due to the composition of lysase buffer.

So can you tell me in details how do you prepare cell lysates for phostag gel?

Thank you

About the gel:

the protein is IRE1 which runs around 100 kDa and the treatment has been made with 0.1 uM of Thapsigargin (the conditions have been tested by another assay and are right), so the protein has one phosphorylation site and after 1 h of treatment we should see a shift.

But unfortunately I have been trying many times phos-tag gel and I could not get any improvement!

Dear Daniela,

I have got little to no experience with phos-tag SDS-PAGE, but your bands look like you have a problem with the so called "smile effect". Most of the time this problem appears when the center of the gel is getting too hot during the run. I read that you run your gels really so the temperature is unlikely your problem. Another reason for smile effect can be a insufficient pH gradient between the stacking and the resolving gel.

I once overcome the smile effect by using 0.5M Tris pH 6.8 for the stacking gel and 1.5M Tris pH 8.8 for the resolving gel, maybe this could (if not already applied) solve your problem with the distortion of bands.

Good luck!

Dear Daniela,

Thank you for your data.

The protein bands are exactly smiling in your Phos-tag gel, but I think it is not so problem. Maybe, this phenomenon will be solved by decreasing amounts of the protein sample or concentrations of Phos-tag. We have the same experience as you do.

The real problem in your result is that there is no observation of the gel-shift band in the drug-stimulated sample. In the case of conditions that the bands are disordered (waving or smiling), mobility shift bands are also disordered as shown in my attachment. In your gel, however, there is no mobility shift band. This means no phosphorylation of the target or no function of Phos-tag in your gel.

This gel is made by using Mn(II)-Phos-tag or Zn(II)-Phos-tag?

I forgot to attach the example data of waving or smiling gel-shift bands of phosphoprotein species.

Dear Daniela,

EDTA! You have 1mM EDTA in your lysis buffer. How high do you estimate the final concentration of EDTA in your SDS-PAGE sample? The phos-tag gel contains only microM order of divalent cation. According to our own experiences, EDTA in the lysis buffer almost completely abolishes the separation of phosphorprotein.

The distortion of the bands seems to occur more frequently than in conventional SDS-PAGE. Phos-tag seems to somewhat affect polymerization. Longer polymerization time or higher APS and TEMED may improve the distortion. We usually let the gel polymerize for 1.5 hr (>1 hr) even though we use 8x8 cm mini-gel (Bio-Rad Mini-PROTEAN). Even so, we sometimes see distortion of bands.

Dear Daniela,

We usually prepare cell lysates by using the following two procedures. In both of them, adherent cells are used.

Preparation by using RIPA buffer:

To terminate certain stimulation, culture medium is removed, and the remaining cells (10^6 cells in a 30-mm plate) are rinsed with a TBS solution (10 mM Tris–HCl (pH 7.5) and 0.10 M NaCl) at room temperature. After the saline was removed, the culture plate is placed on ice. The cells are exposed to 50 µL of a cold RIPA buffer consisting of 50 mM Tris-HCl (pH 7.4), 0.15 M NaCl, 0.25% (w/v) sodium deoxycholate, 1.0% (v/v) NP-40, 1.0 mM EDTA, 1.0 mM PMSF, 1 µg/mL aprotinin, 1 µg/mL leupeptin, 1 µg/mL pepstatin, 1.0 mM Na3VO4, and 1.0 mM NaF. The plate is gently rocked for 15 min on ice, and the adherent cells are then removed from the plate with a cell scraper. The resultant suspension is transferred to a microcentrifuge tube (1.5-mL tube). The plate is washed with 50 µL of the RIPA buffer, and the washing solution is combined with the first suspension in the microcentrifuge tube. The mixed sample is incubated for 60 min on ice and centrifuged at 13,000 × g for 10 min at 4 ˚C. The supernatant fluid is used as the cell lysate. The concentration of the solubilized cellular proteins is usually adjusted to 2.0 mg/mL with an appropriate amount of an RIPA buffer. Each sample is mixed with a half-volume of 3 × SDS-PAGE loading buffer (195 mM Tris-HCl (pH 6.8), 3.0% (w/v) SDS, 15% (v/v) 2-mercaptoethanol, 30% (v/v) glycerol, and 0.10% (w/v) bromophenol blue) and is heated at 95 ˚C for 5 min before SDS-PAGE analysis. As described previously, we usually apply less than 10 µg of total proteins to Phos-tag SDS-PAGE. I think on the basis of our experience that the contents of the sample will not affect the electrophoresis imaging in Phos-tag SDS-PAGE if the sample is applied within the range of this amount (less than 7.7 µL).

Preparation by using 1 × SDS-PAGE loading buffer:

After certain stimulation, each culture (10^7 cells in a 90-mm plate) is washed twice with the TBS solution. The cultures is then lysed in 0.50 ml of a 1 × SDS-PAGE loading buffer, which consisted of 65 mM Tris–HCl (pH 6.8), 1.0% SDS, 5.0% 2-mercaptoethanol, 10% glycerol, and 0.03% BPB. The lysate sample solutions (about 1~2 mg/mL proteins) were sonicated briefly, and boiled for 5 min.

If you have any other questions, please feel free to reply.

We use it an a regular basis, let me know in case you don t manage to have good results.

cheers

a

Concerning the Lori's comment, I also have some experiences on the age of materials.

We checked a variety of premade gels from various suppliers. Regardless of the expiring date, transfer efficiency is dramatically reduced within one or two weeks.

Electrophoresis itself seems okay even when using expired gels. Proteins can be sufficiently resolved and stained with CBB. But only the protein transfer to the membrane is greatly affected. This phenomenon occurs even when you use premade gels that expire 6 months later!

So, you should use acrylamide gels in a week after you prepare or purchase them if you would like to do western blotting, as Lori recommended.

Cheers

I have had a similar problem and the distorted bands turned out to be because of my molecular weight markers. In general I have been told that markers as not needed since the phos tag can massively distort the predicted running speed.

I now just leave the markers off and just stain the membrane with panceau s solution if in need to cut the membrane. This solved the problem for me.

So I prepared this gel according to Chen paper (because they reported phosphorylation of IRE1, the same protein that I am trying to study) and I did not load the marker because I thought that it could distort the bands and this is the result:

I loaded the same series of samples twice.

Daniela, how much reducing agent (DTT or b-ME) is in your sample buffer?

we use 15mL of b-ME for 100 mL of 3X Sample Buffer (so 15% that becomes 5% as final concentration).

That should be enough. You might want to try to include a little b-me in your inner gel buffer (about 600uL / L). That might help your distortion.

I tried again, changing only the sample buffer (which is 5X SDS-saple buffer) and I got a smiling gel!

Oh goodness. That's worse than before. I think that looks like a salt concentration problem - try lowering the salt in your final sample. That should help.

Dear Daniela,

Thank you for your data.

You have continued troubles in your experiment. Please accept my apology for your Phos-tag blue.

As suggested by Lori, the smiling in this case maybe is caused by salts, especially EDTA, which contains in the lysate solution.

I recommend to perform the procedures of 'buffer substitution and desalination'.

In our cases under the similar trouble, we conduct Acetone precipitation of all lysate samples, and then dissolve the precipitated cellular proteins by using 1X Sample Buffer consisting of 65 mM Tris–HCl (pH 6.8), 2% LDS (lithium lauryl sulfate), 100 mM DTT, 10% glycerol, 0.01% BPB, and 6 M urea. Urea should be added just before use.

Because it is generally difficult to dissolve the precipitated proteins, the buffer with urea described above are used instead of Laemmli's Sample Buffer. Please note that the sample boiling should be avoided in the case using the buffer with urea.

I do hope this will help you. Good luck

Eiji

Sorry to intrude in this thread, but given that you all seem to be Phos-Tag experts...

Wouldn't it be easier, having a single target protein, to immunoprecipitate it, then Western blot it and probe with Phos-tag-biotin + streptavidin HRP, rather than a complicated Phos-tag gel? Or am I being naive? Has anybody tried that?

Hi, Olivier. I know some users, who have tried the procedure as you suggested (see PMID: 22001259). By the strategy, however, the ratio of phosphorylated and nonphosphorylated forms (or multiple phosphorylated forms) of a certain protein cannot be measured. For obtaining more detailed information on phosphorylation of a specific protein, Phos-tag SDS-PAGE would be best.

Anyway, it should be necessary for me to develop an improved Phos-tag method to succeed under any sample conditions.

Hi, Quan, please find my answers described below. I hope they will help you.

Many thanks!

1. For your target with molecular masses of more than 60 kDa, please determine an optimal concentration of acrylamide of 6–8% (w/v).

2. You don’t need to stop running. Please determine an optimal running time for your target separation. I think that the condition of the current is OK.

3. Improvement by the procedure of overnight transfer is not so expectable. As described previously in this thread, it is effective to add SDS in the transfer buffer you usually used. Please determine an optimal concentration of SDS of 0.05–0.2% (w/v) for your target. However, the efficiency is dependent on proteins. So, please notice that some hydrophilic (or small) proteins may pass through the blotting membrane We usually use 0.1% of SDS in our experiments.

I am having problems of inconsistency of PhosTag gels.

1. Bromophenol blue front migrated very quickly and was eluted from gel in just 10 min after start of electrophoresis (running conditions were 20 mA constant current per gel, gels had 10% acrylamide/bis and were supplemented with 50 microM of phostag/Mn2+)

2. We also used prestained markers (we know they are not recommended but I never saw an image to understand how they behave). The markers stayed pretty much packed at the top of the gel with little separation. So the problem is that if bromophenol blue migrates too quickly and MW markers too slowly, how are we going to follow the procedure to know when to stop?

3. We find the washes with 1 mM of EDTA insufficient. We even tried 3 washes under such conditions and the transfer was still very poor. However, transfer was dramatically increased with one wash with 10 mM EDTA and inclusion of 1 mM of EDTA in the transfer buffer during the transfer.

4. Most serious problem has to do with batch-to-batch inconsistency. Using 2 different PhosTag batches keeping all the rest the same (i.e., acrylamide sol., APS/TEMED/ buffers, etc) yielded two gels that gave markedly different behaviour. The first gel (with the old batch of PhosTag) did not interfere with protein marker migration, althoug it induced a slight smiling effect. The second one showed the problems described above.

5. Any response will be appreciated

Hi George,

I also have problems with transfering. How much sds contain in your tansfer buffer? I use carbonate buffer containing 0.8% sds. What about yours?

Looking forward to your reply.

could you please let me know where did you buy a phos-tag chemicals?

Thanks

Hi, Thirupugal,

Thank you for your interests in Phos-tag chemicals.

Please check the web site of Wako. I describe below the URL.

http://www.wako-chem.co.jp/english/

Many thanks!

Phos-tag does not always work. No shift CANNOT exclude the possibility of protein phosphorylation. i have tested a couple of proteins that we have commercial phospho-specific antibodies for them. We observed phospho bands using specific antibody but didn't see protein shift in some proteins we are interested in. I thought it might be related to high basal level of phosphorylation but their technical teams in both USA and Japan don't know the reason. When you use this reagent, you need to be very careful to interpret your data. Their tech team is not helpful!

I think the concentration of Phos-tag is too high.

Steve,

      It depends on the size of the protein you're interested in but I found that a 7.5% acrylamide gel with a bis-acrylamide:acrylamide ratio of 37.5:1 gave me great bands and ran in a reasonable amount of time.

       Let me know if that helps. If you look on these forums (or just google accuracy phos-tag), you'll find a great protocol by Peter that really helps.

 There is no problem for your procedure. Good luck!

Hallo Eiji,

for my bachelorthesis I establish a Phost-Tag agarose enrichment for phosphopeptides. I have read your protocol for the solutions. My Question ist about Sol. G. the concentration for the MES doesn´t fit to the mass of MES for 100 mL. I would like to know wicht is right, the mass or the concentration of the MES?

Thank you for your help.

Niklas

Hi Niklas,

I am sorry for the delay of my response.

As you pointed out, I found a type error in our protocol. The correct concentration is 10 mmol/MES. So, this means 0.2 g/100 mL.

More recently, the buffer systems are improved for enrichment of phosphopeptides. Please find the attached protocol.

Actually, we are preparing an article on Phos-tag Tip, a novel tool for phosphoproteome study. In near future, we will introduce it to you.

Many thanks.

Eiji

Hi everyone, it seems that the problem we are experiencing has not happened with anyone here.

When we try to release the Zn-Phos-Tag from the gel casting modules (plastic modules), the gel sticks to the walls of the plastic module, resulting in gel residuals in both walls of the casting module. It is important to note that this happened only when using Zn-Phos-Tag gels, and it did not happened with the Mn-Phos-Tag gels.

Please note that judging the consistency of the left over gel in the tube where it was prepared I would say that the polymerization of the gel turned out to be OK... Also, the gel run seemed to be OK, the gel was not hot. I did notice anything "weird".

Has anyone experienced this problem and/or have any suggestion?

Ps. We are planning to use casting glass modules this coming week.

Thanks a lot!

Can I stain SDS-phos tag gels with CBB staining  

Dear Kishore,

Yes, staining with CBB works well. You can also perform negative stain, silver stain, and fluorescent stains as they are all compatible with the Phos-tag system. If you have any other technical questions please contact Wako Chemicals (USA or GmbH).

Elliott

Dear Elliott Blackwood,

Thanks very much for the quick reply

Hi, Dehong,

Thank you for your question.

Recently, we have published a paper on phosphorylated forms of Asp and His.

Please see the paper (http://www.ncbi.nlm.nih.gov/pubmed/26151934).

Good luck!

Eiji

Hi, Dehong,

I described bellow my suggestions for your questions.

>How did you lyse the bacteria, with the ultrasonic? 

For analysis using Phos-tag SDS-PAGE, a 2 mL aliquot of culture was centrifuged and the pelleted cells were washed with 1 mL Tris-HCl (pH 6.8) and then lysed with 0.20 mL of 1× sample-loading buffer for SDS-PAGE containing 5 units of benzonase. The lysates were immediately analyzed by Phos-tag SDS-PAGE or stored at –20°C; stored lysates were used within two days (see Materials and Methods in the above paper).

>Did you boil the sample before loading?

No, we didn't. As you suggested, boiling destroys the phosphorylation of Asp/His. We demonstrated that in Fig. 8B (100 C, 3 min) of the above paper.  No up-shifted band was observed after boiling .

Good luck!

Eiji

Hello all,

      Thank you for dispensing such great advice. Since this thread has turned into a de facto technical support page, do you mind if I ask another question? 

      We love running Phos-Tag gels and do so routinely using the improved Zinc Phos-Tag gel protocol. Normally, we get great bands and clear separation, but once in a while I notice that a particular protein doesn't run as cleanly. The biggest offender, and the most important to our research, seems to be MYC. I've run at least half a dozen Phos-Tag gels analyzing MYC phosphorylation, and this is the cleanest I've ever gotten. I've tried diluting the salt in the lysis buffer with a 1:1 ratio of Laemmli buffer to MilliQ water, I've tried running more or less protein, adding 1 mM zinc nitrate hexahydrate to the sample, but nothing seems to make a difference. Reducing the Phos-Tag concentration actually did help a fair bit. 

       I was wondering what other optimization steps you may have tried and found to be effective. Thanks for any help you can give and take care!

First of all, thank you so much for your fantastic advices.

I am currently using phos-tag gels with MnCL and I use the old school Tris-Glycin (10% Methanol) transfer buffer for blotting on the nitrocellulose membrane. Fortunately, I had really good results for proteins around 60 KD, but I can not see any transferred proteins on the membrane which stained with CBB R250 under 35 KD (they do not present in the membrane too). I also tried the same transfer buffer including 0.1 % SDS and it only causes that big proteins go through the membrane, but nothing improved on the membrane for small proteins.

Since I am not able to use precast phos-tag gels as Eiji mentioned, do you have any other suggestion to improve this blotting before switching to Tris-tricin buffer?

Thank you

Hi Siavash,

recently colleagues published a very clever combination of standard gels with some phostag poured in, allowing the gel shift.Then the separating gel proceeds "normally" We tried it in our lab - works nicely. No blotting problems anymore.

Check this out: http://www.plantphysiol.org/content/169/4/2874.short

Cheers Lars

http://www.plantphysiol.org/content/169/4/2874.short

thank you Lars,

I am gonna try it very soon. 

Hi lars,

can you please post here the full protocol you use the recipe of the gels, and so on..they are not using a classical laemli gels at first sight.

thanks

Just going to hop in here and see if anyone has an answer to this question. In the Phos-Tag acrylamide handbook recipe list for the Mn system the gels are poured with SDS. However, in the recipe for the Zn system, there is no SDS mentioned in the gels. SDS is only added to the running buffer.

Is this correct? What is the reason for excluding the SDS from the pH 6.8 Zn system but not the pH 8.8 Mn system?

Thank you for your message.

When we developed the method of the Zn system, there was almost no differences in the migration images of Phos-tag gels with or without SDS.

This is the reason, just simple.

Important thing for Phos-tag SDS-PAGE is that you should use the running buffer including SDS.

As for the Mn system, the procedure were developed according to Laemmli's method, which has been widely accepted. In his method, the gel including SDS has been conventionally used. We only followed the procedure, but we also note that commercial precast gels for SDS-PAGE do not have SDS generally.

Good luck!

Eiji

Eiji,

Thank you so much for your prompt answer. I ran two 6% gels that I cast with the Zn system with 50 uM and 75 uM Phos-Tag reagent for 4 hours at 120 V. My proteins of interest are all ~130 kD so I've had to run the system for much longer than recommended to even try to get separation.

However, both of my gels turned out as shown in the attached image, with all the bands forming a smile. Are they pulling on each other to create the distortion? How many lanes separation with 1X loading dye would I need to prevent this distortion and get interpretable results? Do I need to completely eliminate all possible EDTA from the system to get things to run normally?

Thanks!

Dear Elizabeth,

Thank you for your return message.

I describe below my possible suggestions.

1) Strong signals are visualized in the two middle lanes. The distortion may be caused by overloading of samples in the lanes.

2) Your samples are complex biological samples like cell or tissue lysates? If so, the concentration of Phos-tag you used may be too high. Please optimize the condition with lower concentrations of 5 to 25 uM.

In your data, no upshifted bands were detected. I wonder if your targeted protein is phosphorylated under the conditions. The phosphorylation has been confirmed by other experiments? I recommend using some positive controls, which are upshifted in your Phos-tag experiments.

Good luck

Eiji

Eiji,

Thank you for your additional answer. I really appreciate the feedback about the system.

1) The middle wells are from a sample with a higher expression of the protein of interest. The same amount of overall protein was loaded into all 6 wells.

2) Yes. My samples are full cell protein lysates. I was not aware that a lower concentration of Phos-Tag might be better for that system, but I will definitely try lower concentrations during optimization.

We have been trying positive controls that we know are phosphorylated under our condition (as confirmed with phospho specific antibody). However, we have not been able to see a shift even in our positive control blots. I will try and change the Phos-Tag reagent concentration and see if that helps with our positive control protein.

Dear all and dear Eiji,

I would be very curious to know whether any of you got like me a phosphorylated form of its protein of interest migrating lower than the non phosphorylated one.

You'll find a photo of my WB using Mn2+ phostag (75 uM) . The protein is ROCK 160 kDa on a 8% gel.

+CIP sample: HUVEC were lyzed in Hepes 50 mM pH 7.5 NaCl 120 mM 1% triton+ Protease inhibitors , sonicated +1 ul CIP. Incubated on ice 30 min then 30 min RT.

-CIP sample: lyzed directly in 2X laemli buffer supplemented with Phosphatases inhibitors.

I must say that phosphorylations open up the protein so big changes in protein conformations are expected.

Are the lower bands the phosphorylated forms of ROCK ?

Thanks for your help,

Eva

Dear Eva,

Thank you for your message.

Before consideration of the migration image, I am worried about your experimental conditions. I describe my suggestions.

1) Please use the same buffer composition between the two samples in Phos-tag SDS-PAGE, like below.

+CIP sample: HUVEC were lyzed in Hepes 50 mM pH 7.5 NaCl 120 mM 1% triton+ Protease inhibitors, sonicated +1 ul CIP. Incubated on ice 30 min then 30 min RT.

-CIP sample: HUVEC were lyzed in Hepes 50 mM pH 7.5 NaCl 120 mM 1% triton+ Protease inhibitors, sonicated +1 ul water. Incubated on ice 30 min then 30 min RT.

Please prepare them at almost the same time. After sonication (before treatment with CIP), the samples should be centrifuged to remove insoluble fractions.

2) Your target is relatively large (160 kDa). So, please optimize the conditions as follows:

The concentrations of the gel: 5–6% w/v (T:C= 79:1)

The concentrations of the Phos-tag: 5–25 uM

Many thanks!

Eiji

Dear Eiji,

thank you very much for your very precious suggestions. I am on the way to test them. I'll let you know,

Best,

Eva

Hi all,

I hope someone can help troubleshoot my PhosTag problems, thanks in advance for any assistance. I am trying to identify phosphorylation of the C5a receptor on primary human neutrophils. The cells have been lysed with 0.5 % SDS, 0.1% TEAB + HALT protease inhibitor (contains EDTA) and sonicated. I denature the protein with BME/Laemmli's, boil at 95C for 5 minutes, load 25 ug protein and run in a 12 % gel with 100 uM Phos-Tag and 100 uM Mn2+.

Please see below my most recent blot, which shows a band corresponding to the C5aR at the correct molecular weight (approx 46 kDa) and a smear of what I think is degraded protein below it. Some degradation is expected in human neutrophils unless you have massive concentrations of protease inhibitors/SDS present. The PhosTag gel has a similar appearance (beware of the sample order not being the same!) but everything is shifted upwards. I do not think the 2 bands represent separated phosphorylated vs non-phosphorylated proteins as the bottom band looks like degraded protein and is also present on the normal Western.

Reading this thread I think I might change a couple of things:

1. Reduce PhosTag concentration to 25 - 50 uM

2. Remove EDTA from the lysis buffer

3. Potentially load less protein (maybe 10-15 ug)

My questions are:

1. Should I reduce the protein with BME/DTT prior to loading, or will this interfere with the action of PhosTag? If we don't reduce the proteins won't this affect their migration through the gel?

2. Do I need to reduce the concentration of Mn2+ in the PhosTag gel if I reduce the concentration of PhosTag reagent, if so by how much?

3. If anyone has other tips on using PhosTag or Westerns in general I would love to hear them!

Thanks for assistance, Eiji Kinoshita Lori Walker

Eva Faurobert yes , some phosphorylated protein run faster than the non phosphorylatd..the typical cases are S160 phospho CDKs (fesquet et al, 1993) more recently FBW7 bonne andrea (PLoS One. 2017; 12(8): e0183500...the reason is unclear ..conformationall changes?

Hi Alex,

We have spent a good deal of time optimizing PhosTag gels. Based on your experiment, I would recommend the following changes:

1. Remove all EDTA (Including sample buffer). It will cause strange looking bands (they typically look U-shaped to me) and may interfere with your gel running.

2. I would try the Zn2+ protocol. Note you will need to make up an entirely different set of buffers for this, but we've noticed that sometimes the Zn2+ PhosTag system will work when the Mn2+ does not.

3. Re: titration of PhosTag - In my experience, lower amounts of PhosTag (e.g. 20 uM) give better separation of some targets, especially out of a complex lysate. I almost always run 20 uM PhosTag gels, but I do optimize 20, 50, and 100 uM gels for each target before settling on a condition. To answer your question about the divalent cation: it should always be 2x of your PhosTag gel (so, for 20 uM PhosTag add in 40 uM Mn2+ or Zn2+). I also typically run ~20 ug of lysate, so I think you're probably in the correct range here.

4. I would run a lower percentage gel. The protocol states that your protein of interest should run to about 1/3 of the way from the bottom of the gel, so I think 12% is too high. I have been working with proteins in the ~40 kDa range and have found good success with a 10% gel. For a 55 kDa protein, I need to go down to a 7-8% gel for good resolution. (Note: this will be different for all targets, but in my experience this is a good starting point).

I hope that gives you some ideas as to how to proceed. If you have further questions or would like to chat, feel free to message me or send me an email ([email protected]). Good luck!

Best,

Natalie

Fantastic thanks so much for your help Natalie Niemi, I'll give it a go.

Thanks everyone for all the information. It is really helpful. I am wondering what is the best way to lyse the cells. Which buffer is the best? should I use proteases and phosphateses inhibitors in the lysate? Is a TCA precipitation recommended before running the sample in the gel?

Thanks for your time!

Best,

Carmen

Hi Carmen,

Thank foy your message. That depends on proteins you target. I think the Laemmli's sample loading buffer for SDS-PAGE is the most simple. We often use it. Regarding the strategy of TCA precipitation, I can intoroduce a paper described by Dr. Takeya, who is one of the users of Phos-tag for a long time. I attached it. Good luck!

Eiji

I am sorry. I attached the paper mentioned above.

Hi Eiji,

Thank you very much for your response and the paper. Now I have more questions :). Do you usually use proteases (or/and phosphatases) inhibitors in the lysis buffer (laemmli's buffer)? Do you sonicate? How do you lyse the cells (incubation, centrifugation)? I know it depends on the target protein, but I would be great to have a protocol for the lysis to start with. Thanks a lot for your time.

Best,

Carmen

Hi Carmen,

Thanks for your questions. I described below my suggestions.

1) Do you usually use proteases (or/and phosphatases) inhibitors in the lysis buffer (laemmli's buffer)?

No. In case of using the laemmli's buffer, we do not use these inhibitors.

2) Do you sonicate?

Yes, or we often treat with nuclease such as Benzonase.

3) How do you lyse the cells (incubation, centrifugation)?

Incubation → sonication (treatment with Benzonase) → centrifugation

I think the above procedures are general sample preparations for SDS-PAGE without Phos-tag.

Good luck!

Eiji

Hi Eiji,

Thanks a lot for your suggestions. I will follow them.

Best,

Carmen

Hi Eiji,

I'm try to use the Phos-tag system for detect the phosphorylated form of my protein. My problem its that I work with plants, with Arabidopsis system plant. I try to detect the phosphorylated protein in vivo, using transient expression in Nicotiana benthamiana. When I run the gel, my protein its from 45 KDa, I use 8% and 50 uM Phos-tag, during 2h at 35 mA, in these conditions practically don't see nothing of difference. I don't know if any people have experience with phosphorylated proteins in plants that use the Phos-tag system and could help me. Thanks.

César.

Hi César,

Thank you for your message.

I am sorry, but I have never experinenced using plant's lysate samples.

Some goups have reported the mobilty shifts of phosphoproteins in the same plant samples as yours. I descrbed below.

I recommend using lower concentrations of Phos-tag (

yeah