Hi! Usually I work with larger targets, however, the principle should work as well with your protein / peptide: I am employing spin filters with 10 kDa to remove dye (approx. 1kDa) from an approx 8 MDa complex. You add eg 400 uL of your buffer to the filter, weigh in 100 uL of your labelling solution, spin to remove approx. 450 uL of buffer (+ dye, you should still see a coloured region on the filter corresponing to your labelled target tough), replenish the buffer, repeat spinning, etc. In my case 2 - 3 spinning steps are enough to remove the dye. However, you can also do more. You just have to be careful that your protein doesn't stick to the membrane. In your case 3 kDa filters might work. The problem is that the pore size for 3 kDa filters results in long spin times and high rcf values... (For an example you can have a look at my publications concerning gelatin nanoparticles, ammodytoxin or rhinovirus.) Alternatively, you can opt for dialysis, which will however result in sample dilution.

I have had success separating unincorporated dye from peptide using chromatography on Sephadex LH-20 in distilled water.

Hi Tesmine, it all depends on the properties of your protein but I would suggest RP-HPLC (even FPLC might work) as this is usually done in our laboratory when we purify peptides. I would be a bit afraid of using spin-column method suggested by Victor, as you might experience significant loss of your peptide without removing enough contaminants [e.g. 3kDa column has 42% retention of vitamin B12 (1250 Da) after 30 minutes spinning].

Thank you all for your suggestions. 

Hi Tesmine.

The method of choice I will usually follow is using a C-18 rpHPLC column and perform a separation in 0.1% TFA (in aqueous buffer) against Acetonitrile (HPLC grade). Separation based on hydrophobicity is much better in this case than is a separation based on molecular weight. First perform an analytical run for the dye only and identify its elution peak. Then, in the labeling mixture, you will hopefully have two different elution peaks. If the free dye and the labeled peptide have the same elution time, try ion exchange HPLC. Make sure the peptide can handle low pH that exists in such separation. Also make sure Alexa 647 doesn't change its absorption spectrum in low pH (I don't remember if it does).

I would be cautious in using fel filtration (such as Sephadex) columns as the molecular size vs. elution times are not linear and an elution peak for a molecular weight of ~3 kDa might have some overlap with ~0.5 kDa molecules (which is the molecular weight scale for these organic dyes).

I would not try separation using any membranous methodology (filters, dialysis bags,etc) - I have some experience with that using proteins labeled with Alexa 647 and the dye sticks a lot to membranes and you lose the majority of your product.

Separation on Sephadex LH-20 is based on hydrophobicity rather than size. The name is misleading. LH-20 is modified with hydroxypropyl groups.

Hi, as we want to avoid any loss of our expensive protein, have low amount so our choice instead of column is dialysis cassette and it works fine and easy to use for anybody :)

Thank you all for your suggestions. For HPLC, what kind of column would be better for the separation of the dye from labelled peptide - C4, C8 or C18? 

The best choice of reverse phase column depends on the particular substances you are trying to separate. Consult the manufacturers for advice.

You might consider the following:

According to my information, Alexa Fluor 647 is highly sulfonated and hence very hydrophilic. This might make an RP separation difficult.

Depending on your application, you might avoid a separation altogether. In some protocols, only a 1.1x excess is used. If your application involves some washing steps, the excess of dye will be washed away. You only have to make sure that your conjugation did not fail completely.

Please find a supplement of a publication in the link below.

http://www.pnas.org/content/suppl/2011/05/23/1101929108.DCSupplemental/sapp.pdf

http://www.atdbio.com/content/34/Alexa-dyes

Adam B Shapiro Hello Sir!

Can You please send the links of few papers in which they used Sephadex LH-20 for separation of unlabeled dye with Peptides.

Thanks in Advance!

I describe the method in this paper:

Article A High-Throughput, Homogeneous, Fluorescence Resonance Energ...

It was just an isocratic gravity elution in water with a tall, thin column.

Hello!

Can someone guide me please.... I am labelling my peptide with MW 768.46 with FITC in DMF/TFA/H2O solvent mixture. After 12 hours stirring i get my product. It is a liquid mixture, i want to get solid product. Which method is suitable to get solid product? Or can i add some solvent which can produce precipitates of my product and I get solid product by paper filtration. Please reply me ASAP.

Thanks in Advance!

Dear Faheem Askari,

I fear that your labelling failed. According to my knowledge, FITC needs quite basic conditions for conjugation. Trifluoroacetic acid (TFA), however, is highly acidic.

To get your peptide in a solid form is highly dependent on the structure of it. But in any case, you should remove (evaporate under vacuum) DMF and TFA.

You may find some useful hints here:

https://www.researchgate.net/post/The_best_protocol_for_FITC_labeling_of_proteins

They suggest a carbonate buffer of pH 10!

Michael G. Weller Thank You Sir for your kind response. Sorry, by mistake I wrote TFA, it's TEA. I am using 10uL TEA in 2mL DMF solution and starting pH is 11 pH. At this pH, I allowed to stirrer the mixture overnight (12h) under N2 air. After completion of reaction I adjust the pH (3-4) using neat TFA.

When I evaporate the DMF/TFA using rotary evaporator, it damage my peptide and I get two peaks in HPLC (C-18 column; ACN:water) while without evaporation I get only one peak in HPLC (same protocol).

Is it possible that I will add some suitable solvent in the mixture which produces precipitates of FITC-peptide, so that I may get solid product by simple filtration??

You should not use a rotary evaporator, but a gentle nitrogen stream or freeze drying to get rid of your solvents. I am not aware of any solvent, which might be suitable for this purpose. In some cases, diethylether was used for precipitation.

Michael G. Weller Thank you Sir.