I want to see the interaction between lipid NPs and proteins, but I am not sure which solvent is the best for both of them.
Shiva Montasseri
You study proteins and lipids in order to later draw conclusions about their connection with the human cell. In a cell, proteins and lipids are connected to each other by hydrophobic interaction with the help of water. Any other solvent will disrupt this interaction. So you need to explore in the water.
For NMR experiments involving proteins, deuterated water (D2O) is a common solvent. It provides a suitable environment for protein solubility and stability.
Chloroform/methanol mixtures are often used for lipid NMR studies, but it's crucial to ensure that the solvent conditions are compatible with the lipid properties and the goals of your experiment.
Please check solubility before doing any experiments so it will save your worthy sample.
I agree. Water is the best to start as this is the most abundant solvent in living organisms. I also add about 5% DMSO to increase the solubility of lipophilic compounds. I have not published anything in NMR, but used it to confirm my other data. You may read about solubility in my paper below. Good luck!
https://pubmed.ncbi.nlm.nih.gov/35406720/
David Salehi Thanks for your answer. 5% DMSO should be a good idea. it can increase the lipid NPs solubility.
Dear Shiva,
the answer depends on what you define as "soluble" for LNPs. In aqueous/organic environment you might be able to obtain a "suspension" with high-MW LNPs essentially invisible in NMR.
In my experience it is extremely crucial to really compare the amount of LNPs "seen" in NMR with what you have put in the tube (in terms of ug's). Otherwise you are "fishing in the dark"...
Alfred
P.S. we very very often have difficulties in studying the interaction of hardly soluble compounds with proteins. The sensitivity of NMR is simply magnitudes of order lower compared to other methods in use to study interactions.
Alfred Ross
Dear Alfred,
Thanks for your reply. I am new to the NMR instrument.
I have 2 LNPs which are made from Monoolein (MW: 356 g/mol) l and Phytantriol (MW: 330). By using MRI, I want to confirm the interaction of them with proteins (mucin). But I am not sure, about the solvent. Can I disperse lipids in D20 water by using a Sonifier? I know that Sonifier can create air into samples.
also, I couldn't find any relevant articles related.
I would appreciate it if you give me a suggestion.
Dear Shiva,
I agree to the answers above.
I have only experience with proteins not with lipids in solution:
Use Water with 5% D2O in order to perform the lock (which uses the deuterium channel). If you use pure D2O, you will lose all water accessible, exchangeable protons. Like the HN and HO at the protein surface. If that it no problem, you can also use pure D2O.
What is also relevant, is the pH of your water. If it is too high, you protein can aggregate. So you should use a puffer with specific pH (usually acidic). This is crutial if the sample should last longer.
If you use any other solvent (or add solvent to the buffer), like was mentioned above, the structure of your protein might be disturbed (or will be disturbed) which influences the binding. Sonication can also influence the protein structure.
I would recommend trying to collaborate with a Bio-NMR (liquid state) specialist. Or at least ask for his advice in detail. Since your measurements are not trivial.
Many Greetings,
Kristof
Our LNP expert suggest to prepare a dispersion of LNPs in aqueous buffer. I expect to have this extremely low concentration. You can then measure the protein to see whether amplitude disappears or diffusion changes in comparison with buffer only ... if you are however only interested to prove Bindung of Protein and LNP there must be way more simple methods (e.g. DLS or Ultracentrifugation) which work at much lower protein and LNP concentration..
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