I have some queries regarding preparation of cyanine dye encapsulation with DOPC lipid, Anybody can help me to clarify these.
I would like to follow this procedure for preparation of dye encapsulated liposomes preparation.
1) 6 mg of DOPC (100 %) lipid was diluted in 1 ml of chloroform solution (8 mmol stock).
(is this concentration is fine for dye encapsulation ??)
2) The solvent was evaporated to obtain a thin lipid film. Residual solvent was removed under vacuum overnight.
3) The lipid films were solubilized in HEPES (1 ml) with cyanine dye (25 uM) (30 mM pH 7.6).
In this step I have some queries
1) Actually my dye is weekly soluble in buffer solution it would be require small amount of methanol so can I use small amount of methanol to dissolve the dye in buffer.
2) How much dye would be normally requires for encapsulation. Is 25 uM is sufficient enough.
3) In re hydration step How much time I have to incubate the cyanine dye with DOPC lipid and also what temperature and time would be ideal for this incubation.
Regards
Prathap
Hi Prathap,
I guess you can try adding the required amount of buffer dissolved in organic solvent (chloroform, methanol or a mixture of chloroform and methanol) to the lipid solution, and then removing the solvent to make a thin film.
Regarding the incubation temperature, it should be ideally above the gel-liquid phase transition temperature of that lipid.
Simply add the cyanine dye in chloroform to the lipid mixture and rotavap to encapsulate the dye directly into the membrane without the need to transfer it from aqueous phase to the lipids.Check out one of my publications for a step by step procedure.
Depending on the dye, in my experience you can go up to several mol% with respect to the lipids, but you have to check it out. 25 µM should work out. If it's too much, it will either crash out in solution (hazy solution) or you will find the dye on your extrusion filter, in case you use one.
If your liposomes are not PEGylated, 8 mM is quite high and I suspect that they will aggregate quickly over time. 1 mM concentration is a little bit more safe.
Hi Prathap,
1) The ideal concentrations and preparation protocols depend strongly on what is the purpose of your liposomes - what will you use them for, by what techniques you will study them
2) I am a bit confused by the term "encapsulated" dye. Under that term I would imagine that you want the dye solution to fill the interior of the liposomes such as is used in liposome leakage assays, e.g.: https://www.researchgate.net/publication/262148761_Peripheral_and_Integral_Membrane_Binding_of_Peptides_Characterized_by_Time-Dependent_Fluorescence_Shifts_Focus_on_Antimicrobial_Peptide_LAH4?ev=prf_pub
You are not going to get this with a dye that is poorly soluble in buffer. You are most likely to get liposomes containing the dye in their membranes. In that case the easiest way how to incorporate the dye is to mix it with the lipid solution in chloroform as recommended by Robin.
3) If you only want to do fluorescence imaging of your liposomes, you can use pretty high dye/lipid ratios, as Sven said. If you want to use any single-molecule techniques such as FCS or particle tracking you'll need to reduce the dye concentrations to something like dye/lipid = 1/10^5 or lower (depending on your imaging setup).
Article Peripheral and Integral Membrane Binding of Peptides Charact...
Hello.
1) 6 mg of DOPC (100 %) lipid was diluted in 1 ml of chloroform solution (8 mmol stock).
Dear all,
I am extremely sorry, I could't notice your reply till today. Thanks for your valuable suggestions.
Regards
Prathap
Dear members,
Can you please help me out, how to visualize cyanine dye encapsulated liposomes. As my dye is weekly fluorescent nature in lipid vesicle will it be useful to take fluorescence or con focal microscopy pictures?? Is there any other techniques available to know whether my dye was encapsulated in liposomes or not?? How to confirm it?
I would like to encapsulate my weekly fluorescent dye in interior of liposome and I have to confirm the dye encapsulation with any of the know technique then would like to see leakage experiments. Please suggest how to confirm this type of dye encapsulation by using know techniques. Is there any standard procedures for this?
Regards
Prathap
Dear Prathap,
I can offer some considerations:
1) Having even 20 mM total lipid in LUVs is no problem in my experience if the lipid is most stable in the lamellar phase. I would advise against storing the solution for more than a few days, though.
2) If your dye permeates the membrane very well, you cannot get any kind of encapsulation.
3) If your dye is lipophilic/amphiphilic, it will mostly reside in the membrane. If it nevertheless (for an amphiphilic dye) has charged groups or some other polar group that makes flip-flop from one leaflet to another very, very slow, you can get 50% encapsulation efficiency by adding the dye already in the lipid mixture from which you prepare your liposomes from. This means that the dye will be evenly distributed in both leaflets of the LUV bilayers.
4) If your dye is water soluble, then unless it is charged and permeates the bilayer well or it is an acid or base, so that electrical potential difference or pH difference can be used to enrich the dye in the intraliposomal compartment, then you can typically expect up to 8 % encapsulation efficiencies, since that is the intraliposomal volume for high lipid concentration extrusion. You can then use size exclusion chromatography to separate the dye from the liposomes. Yet, if the dye leaks fast, it won't work.
5) I wonder why you insist on using weakly fluorescent dye (since it must be weakly fluorescent both in water and organic solvents, if it does not fluoresce in liposomal solutions). The only thing I can think of is that it is reactive and turns into more fluorescent due to some chemical reaction. Then you can use that reaction (unless the other reactant permeates bilayer very well) to quantify the amount of dye inside the liposomes, i.e. add the reactant on the outside, then wait until the reaction starts to near equilibrium/steady state, then add Triton X-100 to break the liposomes, and you should see an increased rate again. With NBD dyes it is possible to use dithionite to form a non-fluorescent species.
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