Thanks for the reply John. So far I've been running the conjugations at about 2 mg/ml (following what I've seen in some journal articles).  I've also checked out the surface of BSA and there are a bunch of free lysines available and Cys 34 is available.  Occasionally I'll see some very faint bands of higher MW in my gels, but published work seems to imply the conjugation should work well enough to see a significant band. I might try overloading the gel to see if I can get the higher bands darker.

I will definitely check out that book, thanks!

Generally you need large excess of polymer to conjugate to proteins using these chemistries. Generally 15-50 excess of NHS group per amino groups. This is because NHS group hydrolyse quickly in water at neutral or basic pH. I generally use 15 molar excess of NHS groups per amino group and the reaction works well. The reaction was performed for overnight at room temperature in phosphate buffer.   

Something you may want to consider is how your triblock behaves in solution. Are all the blocks equally hydrophilic? Is there any chance the NHS end-group is buried in a unimolecular particle?

You might also consider trying to conduct the reaction in a "dry state". The Maillard reaction - conjugation of reducing sugars to proteins via reaction with lysine, works quite effectively at intermediate temperatures and moderate humidities. Typically when I did these reactions I freeze dried the protein and the polymer together from solution, then placed the powder (in a petri dish) into a desicator at 60oC and 76% relative humidity (saturated KBr) and reacted them for between 3 days and 2 weeks depending on the Mw of the polymer you want to conjugate.  Ref for BSA Maillard: Jung et al. Biosci. Biotechnol. Biochem. 70(9) 2064-2070, 2006.

Elizabeth, since you are talking about a triblock copolymers, I assume this polymer may be amphiphilic and form not only unimolecular particles as John suggests, but actually micelles at such high concentration. This may complicate things, depending on the actual structure of your polymer. Otherwise, I agree with the other comments.

Both Maleimide and NHS groups are hydrophobic, so depending on what the rest of the polymer looks like, as suggested above, they may well be hidden in the core of a polyer micelle. In this case, apart from upping the polymer concentration, you could try adding something to make the solution a bit more hydrophobic. DMSO, ethanol etc. to reduce micellisation.

Is the polymer charged? At pH7 BSA has a net negative charge, so if the polymer is strongly negatively charged that may also not help. In this case incresing the salt concentration might work, though it would not help with the former problem.

These are great suggestions. Thanks!  I hadn't considered the micellization aspect. I will try running some with DMSO added tonight.

The general polymer structure is PEO100-PPO50-PEO100 (It's pluronic F127). Neutral at pH 7.  I had seen other papers conjugate this to proteins previously, almost always in aqueous solution, so I didn't think too much about it. Granted, I haven't come across F127-MAL chemistries yet.

Protein conjugation with pluronics has been pioneered by Alexander Kabanov, you may want to look at some of the attached papers.

Elizabeth, how sure are you about your functional end-groups? End-group analysis of polymers of this size is typically not trivial. NHS is highly reactive and may be easily lost if the work-up or storage was not correct.

An important point Robert! I think that is the main reason NHS-activated compounds are usually added in several fold excess, because many or even most hydrolyse rather than couple. Maleimide is much more stable in this respect, however as the protein typically has many fewer thiol groups for coupling the rate may also be reduced.

The functionality of the polymer is so far only based on proton NMR and IR ( which is just suggestive but not really a conclusive method). I've been integrating the peaks from the NHS or MAL group to the CH3 from the PPO segment. I realize this is not exact, but it at least gives me a ballpark idea of how much functionalization might have happened. I'm looking into secondary means of confirming and quantifying the amount of functionality,  either direct detection of an NHS/MAL group, or possibly indirect methods by titrating to see how much free COOH/NH2 groups are left. 

The attempted conjugations were run quickly after the polymers were functionalized and looked at by NMR so I would expect not much to have changed. That was a few weeks ago though, and I have not re-taken the F127-NHS NMR yet.

Hi Elizabeth, did you confirm whether you have COOH group attached to the polymer before you modify that to the NHS or maleimide groups. It depends on how did you introduce it to the polymer. If that reaction was not efficient you will have issues.

A number of very good answers here.  in our experience protein polymer conjugates do not work well on PAGE gels.  Further we have found that only a small number of the available sites on a protein are conjugates by grafting to techniques and that substantial amounts of unreacted  protein remains.  We have also encountered some solubilization problems following freeze drying.  You did not say how you separate the unreacted polymer from the protein If you are dialyzing you may be losing conjugate at that step as well.  

We have been developing methods for grafting from that results in a more consistent conjugation product.  We have made diblock conjugates with this method.  Feel free to contact me off line to discuss this methodology.

Hi Richard,

I inherited this project from a previous post-doc, and the idea was to utilize commercially available polymers. I agree grafting from would be better, but I'm stuck with this approach based on a proposal that was written : ) 

I've been running the gels on the crude reaction mixtures. So I stop the reaction, freeze-dry them, and then resuspend a portion for the gel. I think part of the problem might be I'm underloading the gel. I had been using 1 mg/ml stock concentrations (loading what I thought to be about 20 ng of protein), but was getting very faint bands so I upped the concentrations quite a bit. I am now seeing 2 very faint higher MW bands ( between the BSA and higher MW gunk), but they're just not as dark as I would expect based on other papers I've seen showing the reaction mixtures. I've attached links with images of the latest gels I've run. I also took MALDI of the freeze-dried mixtures, and I am seeing the weight for the singly conjugated species, but again it's a weak signal compared to all the free BSA that is left.

My plan was that if I could get the cysteine chemistry to work decently, I would run size exclusion chromatography to semi-purify the conjugates. The conjugates should elute in the flow through and the free protein and polymer should remain on the resin that I have.

https://lh3.googleusercontent.com/k0xhWwAULlutz9JVVOOpmHtxahc62-uWuEMqpCwP-4lN-Ffv4ehSqQz7HvEv3w1vDlpZFZFh8hM=w1348-h632

https://lh5.googleusercontent.com/og3YbfN7lbn8EfQaCwUNwfqo2VFWZ2JX_QsxjBpA9D8ftSwNHBhqc5AVOBOhBap1RwLbcUM8JpU=w1348-h632