J.W.S. is an Investigator of the Howard Hughes Medical Institute. This work was supported in part by a grant (290363) from the Simons Foundation to J.W.S. Both A.E.E. and K.P.A. acknowledge support from University of Minnesota startup funds.

1. Vesicle preparation
<ol>
	<li>Preparation of thin films
	<ol>
		<li>Use gas tight syringes to transfer certain amount of lipids as described in Table 1.1 to chloroform in a glass vial.</li>
		<li>Evaporate the resulting solution under a stream of nitrogen or argon in the fume hood.</li>
		<li>Subject the resulting thin film to house vacuum for at least 2 h to remove any chloroform residue. The lipid can be left under vacuum overnight at this point.</li>
	</ol>
	</li>
	<li>Rehydration of vesicles
	<ol>
		<li>Prepare 10 mL of 5 mM calcein 250 mM tris-HCl pH 8.0 hydration buffer by dissolving 31 mg of calcein powder in 2.5 mL of 1 M tris-HCl pH 8.0 buffer and adding another 7.5 mL of deionized (DI) water.</li>
		<li>Pipette 250 &#181;L of above hydration buffer into an empty tube and add 1.875 &#181;L of 10 M NaOH for a final 75 mM base (&#189; equivalent of total fatty acids). Transfer the solution to the thin film of dry lipid to form vesicles with a total lipid concentration of 0.15 M .</li>
		<li>Use high-speed (&#62;3000 rpm) vortexer to vortex the resulting mixture for 4-5 s.</li>
		<li>Leave the lipid-buffer mixture on a low speed (ca. 30 rpm) rotating shaker to rehydrate, at least overnight.</li>
	</ol>
	</li>
	<li>Sizing of vesicles (optional)
	<ol>
		<li>Using tweezers, apply one filter support to each inner surface of the syringe ports of the extruder.</li>
		<li>Wet each filter support with 250 mM tris-HCl, pH 8.</li>
		<li>Using tweezers, apply one track-etched 100 nm polycarbonate membrane to one of the extruder O-rings and filter supports. Taking care to not tear or puncture the membrane, gently push the membrane into the O-ring so as to make good contact between the two surfaces.</li>
		<li>Assemble the extruder, taking care not to displace the membrane and filter supports.</li>
		<li>Fill an extruder syringe with ca. 0.5 mL of 250 mM tris-HCl, pH 8. Insert this syringe into one side of the extruder.</li>
		<li>Insert an empty syringe into the other side of the extruder.</li>
		<li>By hand, push the plunger of the syringe containing buffer slowly (&#8804; 50 &#181;L/s), and verify that resistance is felt, indicating the track-etched membrane is in place and intact. It is helpful to practice this step without a membrane in place in order to gauge the expected level of resistance.</li>
		<li>Remove and empty the two syringes. It is not necessary to clean the two syringes at this stage since they contain buffer of identical composition to the liposome preparation.</li>
		<li>Load the liposome preparation into one of the two syringes.</li>
		<li>Reassemble the extruder with syringe containing vesicle sample on the left side and empty syringe on the right side.</li>
		<li>By hand, push the plunger of the syringe containing vesicle sample very slowly (10-25 &#181;L/s).</li>
		<li>Carefully observe the syringe on the right side of the extruder; clear buffer will initially enter the right side of the extruder (ca. 50 &#181;L, this is due to the inner dead volume of the extruder), followed by a small plume of extruded liposomes. Immediately stop pushing at this point, remove the syringe on the right side of the extruder, and discard this solution.</li>
		<li>Replace the syringe on the right side of the extruder and extrude until the left syringe is empty.</li>
		<li>Reverse the orientation of the extruder and repeat step 13. Continue for the desired number of cycles (typically 7, 9, or 11); an odd number is always used to ensure un-extruded liposomes are not collected from the syringe initially containing the pre-downsizing liposome preparation.</li>
		<li>Gently dispense the contents of the right syringe into an Eppendorf tube and place the tube on a low speed (ca. 30 rpm) rotating shaker for around 0.5 h.</li>
		<li>Proceed with vesicle purification, as described in Section 2. Extruded vesicles should be used within 24 h or re-extruded before next time use.</li>
	</ol>
	</li>
</ol>
2. Vesicle purification
<ol>
	<li>Preparation of vesicle purification mobile phase
	<ol>
		<li>Prepare 5 mL of 250 mM tris-HCl pH 8 hydration buffer by adding 1.25 mL of 1 M tris-HCl pH 8 buffer, 3.75 mL of DI water to a 15 mL Falcon tube. Then add 37.5 &#181;L of 10 M NaOH (&#189; equivalent of base relative to unesterified fatty acid) to the hydration buffer. Pipette 235 &#181;L of pure oleic acid directly into the Falcon tube resulting a vesicle solution with 0.15 M total lipids.</li>
		<li>Use high-speed (&#62;3000 rpm) vortexer to vortex the mixture for 4-5 s, then tumble on a low speed rotating shaker for at least 2 h. The lipid preparation can be left overnight on the rotating shaker at this point. Filter the mobile phase through 0.22 &#956;m syringe filter unit before use to remove any potential aggregates.</li>
	</ol>
	</li>
	<li>Purification of vesicles on Sepharose
	<ol>
		<li>Remove ca. 5 mL of ethanolic Sepharose 4B slurry from the bottle using a pipette. Apply this to a disposable 10 mL chromatography column.</li>
		<li>Allow the slurry to settle and ethanol flow-through until ethanol approaches top of the resin bed.</li>
		<li>Apply 5 mL of deionized water to the top of the resin and let flow-through to wash away ethanol residue.</li>
		<li>Apply 250 mM tris-HCl, pH 8 in 1-2 mL portions to the top of the resin, applying a new portion each time the liquid level approaches the top of the resin bed. Repeat for 3-5 times.</li>
		<li>Clamp the column on the retort stand, then connect the tip of column with stopcock connector and tubing to the fraction collector. Add another portion of buffer to flush the tubing, close the stopcock when the liquid level in column approaches the top of resin.</li>
		<li>Apply the extruded vesicles from Section 1 to the top of the resin using a 200 &#181;L pipettor, taking care to apply the vesicle preparation as evenly as possible to the resin without touching the resin bed or column wall.</li>
		<li>Open the stopcock to begin the flow and start collecting fractions into a 96-well plate. Apply vesicle purification mobile phase to the top of the resin bed in 0.5-1 mL portions as the buffer depletes, taking care not to allow the resin bed to dry out. Collect in five-drop fractions, collecting at least 36 wells (three rows of a 96-well plate).</li>
	</ol>
	</li>
	<li>Fluorescence characterization of purification fractions
	<ol>
		<li>Take the 96-well plate from the previous section and read it on a plate reader (&#955;ex=485 nm, &#955;em=515 nm).</li>
		<li>Plot the resulting fluorescence data as fluorescence vs. fraction number. Vesicles elute first, followed by the unencapsulated fraction (Figure 1).</li>
	</ol>
	</li>
	<li>Repurification of vesicles to monitor vesicle leakage
	<ol>
		<li>Repeat the purification and characterization steps in Sections 2.2 and 2.3. Vesicles that have retained their contents will exhibit no peak of unencapsulated solute (or a very small one of ca. 5-10% of the intensity of the vesicle peak).</li>
	</ol>
	</li>
</ol>
3. Use of Vesicles in the Presence of Magnesium
<ol>
	<li>Use of unliganded magnesium
	<ol>
		<li>Prepare and purify vesicles as described in sections 1, 2.1 and 2.2.</li>
		<li>Prepare 1 mL of 50 mM MgCl2 solution in 250 mM tris-HCl pH 8.0 buffer by adding 250 &#181;L of 1 M tris-HCl pH 8.0 buffer and 50 &#181;L of 1 M MgCl2 to 700 &#181;L of DI water. Vortex to mix well.</li>
		<li>To give the desired magnesium concentration of 5 mM, mix 0.9 mL of purified vesicles and 100 &#181;L of magnesium solution and stir rapidly to minimize vesicle disruption by transient exposure of vesicles to high local concentration of magnesium. 
		NOTE: Rapid mixing of magnesium solution is critical to vesicle stability. If magnesium and vesicles are not quickly mixed by immediate vortexing, some vesicles will be exposed to higher concentrations of magnesium, resulting in inconsistent results from sample to sample.</li>
		<li>Leave the vesicle sample on a tumbler for at least 30 min before repurification as described in 2.4 to check for contents leakage. Add the same concentration of magnesium as in vesicle sample to the repurification mobile phase.</li>
	</ol>
	</li>
	<li>Use of liganded magnesium
	<ol>
		<li>Prepare and purify vesicles as described in sections 1, 2.1 and 2.2. 
		NOTE: Use &#189; equivalent KOH instead of NaOH to deprotonate fatty acids; it has been found that this produces more stable liposomes in chelated MgCl2 systems.</li>
		<li>Prepare 2M potassium citrate solution and adjust pH to 8.0 with KOH.</li>
		<li>Premix MgCl2 and potassium citrate at the specified ratio (ca. 1:4 for stable oleic acid vesicles) in 250 mM tris-HCl pH 8.0 buffer.</li>
		<li>Add premixed magnesium citrate solution to the vesicle sample, briefly vortex. 
		NOTE: Always premix magnesium and ligand solution. Never expose vesicles to unchelated magnesium solution alone.</li>
		<li>Leave the vesicle sample on a tumbler for at least 30 min before repurification as described in 2.4. Add the same concentration of magnesium and citrate as in the vesicle sample to the repurification mobile phase.</li>
	</ol>
	</li>
</ol>
4. Non-enzymatic RNA Copying in Vesicles
<ol>
	<li>Preparation of monodisperse RNA encapsulated vesicles
	<ol>
		<li>Prepare a dry oleic acid film as described in section 1.1.</li>
		<li>Prepare vesicle rehydration buffer with 50 &#181;M fluorescent labeled RNA primer, 150 &#181;M RNA template and 250 mM tris-HCl pH 8.0 containing &#189; equivalent KOH relative to oleic acid.</li>
		<li>Add 250 &#181;L of rehydration buffer to the lipid film and follow step 1.2.2 and 1.2.3 to make vesicles with total 0.15 M lipid concentration.</li>
		<li>In order to make small unilamellar monodisperse vesicles, follow section 1.3 for vesicle extrusion.</li>
	</ol>
	</li>
	<li>Magnesium-citrate addition and vesicle purification
	<ol>
		<li>Premix magnesium and citrate stocks, and then mix this with the vesicle sample to a final lipid concentration of 0.1 M.</li>
		<li>Purify vesicles on Sepharose 4B size exclusion column, with 250 mM tris-HCl pH 8.0, 0.1 M lipids and given magnesium and citrate as mobile phase.</li>
		<li>Collect vesicle fractions by following section 2.3.</li>
	</ol>
	</li>
	<li>Primer extension reaction
	<ol>
		<li>Prepare 200 mM 2-MeImpG (guanosine 5&#39;-monophosphate 2-methylimidazolide) stock solution with 250 mM tris-HCl pH 8.0. (NOTE: Follow previous published protocol10 for 2-MeImpG synthesis.)</li>
		<li>To initiate primer extension reaction, transfer 150 &#181;L of 2-MeImpG stock solution to 450 &#181;L vesicle sample to reach a final concentration of 75 mM lipid and 50 mM activated monomer. Start the timer and keep the reaction sample tumbling all the time.</li>
		<li>(Optional) For continuous fresh monomer feeding, transfer 300 &#181;L of reaction solution to one chamber of a lab constructed small-volume liposome dialyzer11 and put 300 &#181;L feeding solution in the other chamber. The feeding solution should contain all the ingredients at the same concentration as in the reaction solution, except for replacing RNA containing vesicles with empty vesicles. Each round of dialysis takes 1-24 h, depending on the analyte and membrane used.</li>
		<li>For kinetic studies, take a 100 &#181;L aliquot at each time point, and repurify the aliquot through a Sepharose 4B size exclusion column with 250 mM tris-HCl pH 8.0 as the mobile phase.</li>
		<li>Collect the vesicle fractions in a 1.5 mL eppendorf tube.</li>
		<li>Pipette Triton to the vesicle fractions to a final around 0.1% v/v.</li>
		<li>Add 0.5 mL of cold ethanol to the tube and incubate at -20 &#176;C for at least 2 h.</li>
		<li>Centrifuge all the samples at 16.1&#215;1000 rcf (16.1 &#215; g) for 10 min and gently pipette out the liquids. Wash the RNA pellets with 70% cold ethanol and centrifuge again for 5 min. Discard the liquid and put the tube with RNA pellets on an 80 &#176;C heat block for 2 min to evaporate residual ethanol. Dissolve the pellets in 50 &#181;L of 8 M Urea with 1xTBE loading buffer.</li>
	</ol>
	</li>
	<li>PAGE analysis
	<ol>
		<li>Prepare 20% PAGE gel by mixing 200 mL of UreaGel system concentrate, 25 mL of UreaGel system diluent, 25 mL of 10xTBE buffer, 80 &#181;L of TEMED and 250 mg of ammonium persulfate. Quickly pour the gel mixture between clamped gel plates (35 cm &#215; 45 cm) and insert the comb. Wait for at least 30 min till complete polymerization.</li>
		<li>Assemble the gel plate onto the gel stand and fill the gel boxes with 1xTBE gel running buffer. Take off the comb and flush the wells with syringe. Pre-run the gel with 60 W for 30 min.</li>
		<li>Heat the samples on 80 &#176;C heat block for 2 min and load 5 &#181;L of each sample per well.</li>
		<li>Turn on gel power box and set gel running with constant watts at 60 W for 2.5 h.</li>
		<li>Disassemble the gel plate from the gel stand, wipe clean the glass and put the whole plate in gel scanner to start the gel scanning.</li>
		<li>Quantify the gel with ImageQuant TL 1D analysis. Plot the natural logarithm of the ratio of the amount of primer remaining at a given time point to the initial amount of primer vs. time, fit to a line, and calculate the pseudo-first order reaction rate (Figure 2).</li>
	</ol>
	</li>
</ol>
5. Hammerhead RNA Self Cleavage in Vesicles
<ol>
	<li>Preparation and purification of ribozyme encapsulated in vesicles
	<ol>
		<li>Prepare a lipid thin film (OA: GMO=9:1) as in section 1.1 with components as in Table 1.2. NOTE: warm GMO to at least 60 &#176;C for complete melting before using.</li>
		<li>Prepare vesicle rehydration buffer with 5 &#181;M of each hammerhead ribozyme strand and 250 mM tris-HCl pH 8.0.</li>
		<li>Add 250 &#181;L of rehydration buffer to the lipid film and follow steps 1.2.2 and 1.2.3 to make vesicles with 0.15 M total lipid concentration.</li>
		<li>In order to make small unilamellar monodisperse vesicles, follow section 1.3 for vesicle extrusion.</li>
		<li>Purify vesicles on a Sepharose 4B size exclusion column, with 250 mM tris-HCl pH 8.0, 0.15 M mixed lipids as mobile phase.</li>
	</ol>
	</li>
	<li>Hammerhead ribozyme self-cleavage reaction
	<ol>
		<li>Prepare magnesium solution and mix with purified vesicles as described in section 3.1 to initiate the self-cleavage reaction.</li>
		<li>For kinetic studies, take 100 &#181;L of the mixture at each time point and directly repurify this aliquot through a Sepharose 4B size exclusion column with 250 mM tris-HCl pH 8.0 as mobile phase. Collect the vesicle fraction.</li>
	</ol>
	</li>
	<li>PAGE Analysis
	<ol>
		<li>Follow steps 4.3.6 to 4.3.8 to prepare RNA loading sample.</li>
		<li>Place commercially casted 15% TBE-Urea gel into gel box and fill the gel box with 1xTBE gel running buffer.</li>
		<li>Heat the samples on 80 &#176;C heat block for 1 min and load 5 &#181;L of each sample per well.</li>
		<li>Run the gel with constant 200 V for around 1 h.</li>
		<li>Scan the gel.</li>
		<li>Quantify the gel with an analysis software, like ImageQuant TL 1D, to measure the extent of cleavage by comparing the intensity of remaining substrate RNA band and cleaved RNA fragment band (Figure 3).</li>
	</ol>
	</li>
</ol>
6. Giant Fatty Acid Vesicles for Microscopy
<ol>
	<li>Preparation of giant fatty acid vesicles
	<ol>
		<li>Follow section 1.1 and Table 1.3 to prepare an oleic acid thin film with 0.2 mol% of fluorescently labeled lipid for better membrane observation.</li>
		<li>Rehydrate lipid film with 500 &#181;L 250 mM tris-HCl pH 8.0 to make the vesicle sample 10 mM lipids in total. The buffer may contain fluorescent dyes or RNA molecules if desired. Leave vesicle sample tumbling overnight.</li>
		<li>Prepare 150 mL of pure fatty acid dialysis buffer with 10 mM total lipids by mixing 470 &#181;L of oleic acid and 150 mL of 250 mM tris-HCl pH 8.0 containing &#189; equivalent NaOH.</li>
		<li>Extrude the vesicle sample through a 10 &#181;m polycarbonate membrane to remove large aggregates.</li>
		<li>Transfer the vesicle sample into 1 &#181;m large-pore dialysis cassette, as previously described.12</li>
		<li>Place the dialysis cassette into a 100-mL beaker containing 30 mL dialysis buffer. Agitate the beaker gently on a table-top shaker at 80-100 rpm.</li>
		<li>Change the dialysis buffer every 30 min for 5 times to remove free dyes or RNA and small vesicles.</li>
		<li>Carefully remove the vesicle sample from the dialysis cassette with a syringe and transfer to an Eppendorf tube.</li>
	</ol>
	</li>
	<li>Microscopy Observation
	<ol>
		<li>Pipette 10 &#181;L of vesicles onto a clean standard microscope slide and place a glass cover slip over the sample. Seal cover glass with transparent nail polish.</li>
		<li>Observe the vesicles with a 60X oil-dispersion or similar objective by confocal microscopy using the appropriate laser source for the fluorescently-labeled membrane, RNA or encapsulated dye (Figure 4).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Hanczyc, M. M., Fujikawa, S. M., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+Models+of+Primitive+Cellular+Compartments:+Encapsulation,+Growth,+and+Division.">Experimental Models of Primitive Cellular Compartments: Encapsulation, Growth, and Division.</a> Science. 302, 618-622 (2003).</li><li>Mansy, S. S., Schrum, J. P., Krishnamurthy, M., Tob&#233;, S., Da Treco, ., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Template-directed+synthesis+of+a+genetic+polymer+in+a+model+protocell.">Template-directed synthesis of a genetic polymer in a model protocell.</a> Nature. 454 (7200), 122-125 (2008).</li><li>Apel, C. L., Deamer, D. W., Mautner, M. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Self-assembled+vesicles+of+monocarboxylic+acids+and+alcohols:+conditions+for+stability+and+for+the+encapsulation+of+biopolymers.">Self-assembled vesicles of monocarboxylic acids and alcohols: conditions for stability and for the encapsulation of biopolymers.</a> Biochim. Biophys. Acta. 1559 (1), 1-9 (2002).</li><li>Walde, P., Wick, R., Fresta, M., Mangone, A., Luisi, P. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autopoietic+Self-Reproduction+of+Fatty+Acid+Vesicles.">Autopoietic Self-Reproduction of Fatty Acid Vesicles.</a> J. Am. Chem. Soc. 116 (26), 11649-11654 (1994).</li><li>Sulston, J., Lohrmann, R., Orgel, L. E., Todd Miles, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonenzymatic+Synthesis+of+Oligoadenylates+on+a+Polyuridylic+Acid+Template.">Nonenzymatic Synthesis of Oligoadenylates on a Polyuridylic Acid Template.</a> Proc. Natl. Acad. Sci. U. S. A. 59 (3), 726-733 (1967).</li><li>Adamala, K., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonenzymatic+template-directed+RNA+synthesis+inside+model+protocells.">Nonenzymatic template-directed RNA synthesis inside model protocells.</a> Science. 342 (6162), 1098-1100 (2013).</li><li>Uhlenbeck, O. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+small+catalytic+oligoribonucleotide.">A small catalytic oligoribonucleotide.</a> Nature. 328 (6131), 596-600 (1987).</li><li>Chen, I. A., Salehi-Ashtiani, K., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+catalysis+in+model+protocell+vesicles.">RNA catalysis in model protocell vesicles.</a> J. Am. Chem. Soc. 127 (38), 13213-13219 (2005).</li><li>Adamala, K. P., Engelhart, A. E., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collaboration+between+primitive+cell+membranes+and+soluble+catalysts.">Collaboration between primitive cell membranes and soluble catalysts.</a> Nat. Commun. 7, 1-7 (2016).</li><li>Joyce, G. F., Inoue, T., Orgel, L. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+Template-directed+Synthesis+on+Random+Copolymers.">RNA Template-directed Synthesis on Random Copolymers.</a> J. Mol. Biol. 176, 279-306 (1984).</li><li>Adamala, K., Engelhart, A. E., Kamat, N. P., Jin, L., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Construction+of+a+liposome+dialyzer+for+the+preparation+of+high-value,+small-volume+liposome+formulations.">Construction of a liposome dialyzer for the preparation of high-value, small-volume liposome formulations.</a> Nat. Protoc. 10 (6), 927-938 (2015).</li><li>Zhu, T. F., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+large+monodisperse+vesicles.">Preparation of large monodisperse vesicles.</a> PloS one. 4 (4), e5009 (2009).</li><li>Zhu, T. F., Budin, I., Szostak, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vesicle+extrusion+through+polycarbonate+track-etched+membranes+using+a+hand-held+mini-extruder.">Vesicle extrusion through polycarbonate track-etched membranes using a hand-held mini-extruder.</a> Methods Enzym. 533, (2013).</li></ol>

The authors declare that they have no competing financial interests.

Liposomes formed from fatty acids have been suggested by many as potential models for primitive cells due to their high permeability and dynamic properties. The carboxylic head group of single chain fatty acids only allows self-assembly into membranes in a restricted pH range, and the resulting membranes are quite sensitive to the presence of salts. As a result, fatty acid vesicles require different preparation and handling methods compared with phospholipid vesicles.
In this protocol, although we use oleic acid as an example for liposome formation, other long chain unsaturated fatty acids (C14) and their derivatives (ca. myristoleic acid, palmitoleic acid, and the corresponding alcohols and glycerol esters) also form vesicles following the thin film rehydration method as long as the total lipid concentration is above the cmc and the hydration buffer pH is close to the pKa of the fatty acid within the membrane. Other than tris-HCl buffer used in this protocol, other buffer systems (ca. bicine, phosphate, borate) were reported to support fatty acid vesicle formation, though vesicles formed in phosphate or borate buffer are usually quite leaky13. The resulting fatty acid vesicles after rehydration are polydisperse and multilamellar, but are easily converted into small monodisperse unilamellar vesicles by extrusion as described. Compared with sonication as an alternative method for generating small vesicles, extrusion provides more options for the control of vesicle size by applying different pore size membranes. Vesicles after extrusion are usually slightly bigger than the membrane pore size, but by increasing the number of extrusion cycles, vesicles with a narrower size distribution and an average size close to the membrane pore size can be obtained.
In order to synthesize functional protocells, fatty acid vesicles need to host specific biochemical reactions resulting from the encapsulation of RNA or other building blocks. The thin film rehydration method provides an easy way to form vesicles containing desired encapsulated materials. However, the encapsulation efficiency is relatively low and a large fraction of precious materials such as RNA are typically lost during the purification process. In some cases the encapsulation efficiency can be modestly improved by repeated freeze-thaw cycles before extrusion. Microfluidic methods for the high yield preparation of phospholipid liposomes allow for almost 100% encapsulation efficiency, however analogous methods have not yet been developed for fatty acid vesicles.
When handling protocells with either chelated or free Mg2+, purification after magnesium solution addition and repurification before each time point ensures the removal of leaked encapsulated material that might affect the accuracy of reaction rate measurements inside vesicles. Since each purification takes at least 10 min to achieve good separation and to collect vesicle fractions, the analysis of fast reactions is difficult, and the reaction must be stopped prior to column repurification.
The protocol we present here is well suited for the construction of fatty acid liposomes that host reactions mimicking those that might occur in primitive cells. Our protocols also enable potential applications in the development of biomedical delivery systems and bioreactors for other biochemical reactions.

Liposomes, or vesicles &#8211; compartments bounded by bilayer membranes comprised of amphiphilic lipids - have found use in numerous biomedical applications as delivery vehicles for pharmaceuticals, models of cell membranes, and for the development of synthetic cells. We and others have also employed liposomes as models of primitive cell membranes in early life.1,2,3,4 Typically, in such systems, we employ single-chain amphiphiles containing only one lipid hydrocarbon tail (e.g., oleic acid), as these molecules are simpler to synthesize without the benefit of the coded protein enzymes modern cells employ.
Liposomes comprised of single-chain lipids are similar to those formed from diacylphospholipids (e.g., 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, or POPC) in that the boundary is composed of bilayer membranes. Liposomes formed from either class of lipids can retain a dissolved payload, and can be downsized and purified by different techniques. Several important differences result from the unique chemical characteristics of single-chain lipids. Vesicles formed by phospholipids are stable over a broad pH range, while fatty acid vesicles are only stable at neutral to mildly basic pH (ca. 7 - 9), which requires certain pH buffer for vesicle preparation. Most of the time, this buffer may also contain specific soluble molecules for vesicle encapsulation, which can be either functional materials (e.g., RNA) for compartmentalized biochemical reactions or simple fluorescent dyes (e.g., calcein) for vesicle characterization.
The presence of only a single hydrocarbon chain produces a membrane that is both more permeable to solutes, as well as more dynamic. Additionally, the carboxylic acid head group present in fatty acids results in vesicles that are more sensitive to the presence of salt and divalent cations (e.g., Mg2+). Magnesium is one of the most important divalent cations for catalyzing biochemical reactions in protocells for origin-of-life studies. In early life, prior to the evolution of sophisticated protein enzymes, RNA may have been the dominant polymer, due to its dual capability to self-replicate and perform catalysis. One representative example of a magnesium-requiring RNA related reaction is non-enzymatic RNA copying, first demonstrated in 1960s.5 When chemically activated RNA nucleotides (i.e. 2-methylimidazolide nucleotides) bind to a preexisting primer-template complex, the 3&#39;-hydroxyl group of the primer attacks the 5&#39;-phosphate of an adjacent activated monomer to displace the leaving group (i.e. 2-methylimidazole), and forms a new phosphodiester bond. This RNA copying chemistry requires a high concentration of Mg2+, which needs to be chelated in order to be compatible with fatty acid protocells.6 Another Mg2+-dependent reaction is that catalyzed by the hammerhead ribozyme, which is perhaps the best-characterized catalytic RNA. This ribozyme, which can be reconstituted from two short oligonucleotides, performs a self-cleavage reaction that is convenient to monitor by a gel shift. As such, it is frequently employed as a model ribozyme in origin-of-life studies.7 Due to a requirement by this ribozyme for unliganded magnesium, liposomes are typically constructed by a mixture of fatty acid and fatty acid glycerol esters, which are more stable to magnesium.8,9 In this protocol, we present techniques we have developed for the preparation, manipulation, characterization of these vesicles and demonstrate application of these vesicles as protocells to host non-enzymatic RNA copying and hammerhead ribozyme catalysis.

<table><tbody><tr><td>sephorose 4B</td><td>SIGMA-ALDRICH INC</td><td>4B200</td><td></td></tr><tr><td>calcein</td><td>SIGMA-ALDRICH INC</td><td>C0875-10G</td><td></td></tr><tr><td>tris-HCl pH8.0 1M</td><td>LIFE TECHNOLOGIES CORP</td><td>AM9851</td><td></td></tr><tr><td>citric acid</td><td>SIGMA-ALDRICH INC</td><td>251275-500G</td><td></td></tr><tr><td>sodium hydroxide</td><td>SIGMA-ALDRICH INC</td><td>71690-250G</td><td></td></tr><tr><td>potassium hydroxide</td><td>Sigma</td><td>30614</td><td></td></tr><tr><td>oleic acid</td><td>Nu-Chek</td><td>U-46-A</td><td></td></tr><tr><td>glycerol monooleate</td><td>Nu-Chek</td><td>M-239</td><td></td></tr><tr><td>Liss Rhodamine-PE</td><td>LIFE TECHNOLOGIES CORP</td><td>L1392</td><td></td></tr><tr><td>magnesium chloride</td><td>Fisher/Thermo Fisher Scientific</td><td>AM9530G</td><td></td></tr><tr><td>sequagel concentrate</td><td>National Diagnostics</td><td>EC-830</td><td></td></tr><tr><td>sequagel DILUENT</td><td>National Diagnostics</td><td>EC-840</td><td></td></tr><tr><td>15% TBE-UREA GEL</td><td>Thermo Fisher Scientific</td><td>EC68852BOX</td><td></td></tr><tr><td>urea</td><td>Sigma Aldrich</td><td>U6504-500G</td><td></td></tr><tr><td>titon-100x</td><td>SIGMA-ALDRICH INC</td><td>T9284-100ML</td><td></td></tr><tr><td>RNA primer</td><td>IDT</td><td>5&#039;Cy3-GCG UAG ACU GAC UGG</td><td></td></tr><tr><td>RNA template</td><td>IDT</td><td>5&#039;-AAC CCC CCA GUC AGU CUA CGC</td><td></td></tr><tr><td>hammerhead substrate strand</td><td>IDT</td><td>5&#039;Cy3-GCG CCG AAA CAC CGU GUC UCG AGC</td><td></td></tr><tr><td>hammerhead ribozyme strand</td><td>IDT</td><td>5&#039;GGC UCG ACU GAU GAG GCG CG</td><td></td></tr><tr><td>vesicle extruder set</td><td>AVANTI POLAR LIPIDS</td><td>610000</td><td></td></tr><tr><td>fraction collector</td><td>Gilson, Inc.</td><td>171041</td><td></td></tr><tr><td>96-well plates</td><td>Fisher</td><td>NC9995941/675</td><td></td></tr><tr><td>plate reader</td><td>Molecular Devices</td><td>SpectraMax i3</td><td></td></tr><tr><td>confocal microscope</td><td>Nikon</td><td>Nikon A1R MP Confocal</td><td></td></tr><tr><td>gel scanner</td><td>GE Healthcare Life Sciences</td><td>Typhoon 9410 scanner</td><td></td></tr></tbody></table>

preparation, purification, and use of fatty acid-containing liposomes

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of these amphiphiles. In particular, fatty acid liposomes enhance dynamic character, due to the relatively high solubility of single-chain amphiphiles. Similarly, liposomes containing free fatty acids are more sensitive to salt and divalent cations, due to the strong interactions between the carboxylic acid head groups and metal ions. Here we illustrate techniques for preparation, purification, and use of liposomes comprised in part or whole of single chain amphiphiles (e.g., oleic acids).

We typically perform liposome purifications on size-exclusion columns. Typical liposome preparations contain a fluorophore of some kind. When liposomes are generated and extruded, the species to be encapsulated are present both inside and outside of the liposomes. By purifying liposomes on a size-exclusion resin (Sepharose 4B), unencapsulated solutes are retained within the pores of the resin, while the larger liposomes are not and elute first (Figure 1A). Collecting fractions and plotting fluorescence vs. fraction number (Figure 1B) typically yields a two-peak trace, with the early-eluting fractions corresponding to the liposomes, which are then collected and used in subsequent applications.
We frequently examine nonenzymatic primer extension reactions, which were a likely means of RNA replication prior to the emergence of ribozyme and protein-based RNA polymerases. These reactions typically employ a fluorescently labeled primer (Figure 2A), which is extended by activated monomers. These reactions can be monitored by gel electrophoresis (Figure 2B) and the resulting electropherograms integrated to obtain rate constants for a given reaction condition (Figure 2C).
To demonstrate that RNA could function inside protocells, we employ hammerhead ribozyme self-cleavage (Figure 3A) as a model RNA catalytic reaction. This reaction requires free Mg2+ to facilitate catalysis, and therefore we used OA/GMO vesicles since they are stable in the presence of 5 mM Mg2+. Similar to primer extension reactions, the hammerhead ribozyme self-cleavage reaction can also be monitored by gel electrophoresis (Figure 3B) and later analyzed to acquire the rate constant under specific conditions (Figure 3C).
We image liposomes employing both fluorescence and transmitted light. Liposomes can be labeled using fluorescent lipids, which give a membrane label (Figure 4A), or using a fluorescent solute within their lumen (Figure 4B). Transmitted light can also be used to observe vesicles (also shown in Figure 4B).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Table 1.1 pure oleic acid in chloroform</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">Component</td>
			<td>Stock</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td colspan="2">Oleic acid</td>
			<td>&#62;99%</td>
			<td>11.7 &#181;L</td>
		</tr>
		<tr>
			<td colspan="2">Chloroform</td>
			<td></td>
			<td>1 mL</td>
		</tr>
		<tr>
			<td colspan="2">Table 1.2 oleic acid and glycerol monooleate (9:1) in chloroform</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">Component</td>
			<td>Stock</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td colspan="2">Oleic acid</td>
			<td>&#62;99%</td>
			<td>10.5 &#181;L</td>
		</tr>
		<tr>
			<td colspan="2">Glycerol monooleate</td>
			<td>&#62;99%</td>
			<td>1.4 &#181;L</td>
		</tr>
		<tr>
			<td colspan="2">Chloroform</td>
			<td></td>
			<td>1 mL</td>
		</tr>
		<tr>
			<td colspan="2">Table 1.3 oleic acid with 0.2mol% Rhodamine-PE in chloroform</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">Component</td>
			<td>Stock</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td colspan="2">Oleic acid</td>
			<td>&#62;99%</td>
			<td>1.6 &#181;L</td>
		</tr>
		<tr>
			<td colspan="2">Rhodamine-PE in chloroform</td>
			<td>10 mM</td>
			<td>20 &#181;L</td>
		</tr>
		<tr>
			<td colspan="2">Chloroform</td>
			<td></td>
			<td>1 mL</td>
		</tr>
	</tbody>
</table>
Table 1. Fatty acid chloroform solutions.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57324/57324fig1.jpg" /> 
Figure 1. Vesicle purification and fluorescence characterization of purification fraction. A. Separation of vesicles containing calcein from free calcein on a Sepharose 4B column. B. Vesicle and free calcein peak detection by plotting the fluorescence in each well vs. well number after fraction collection. <a href="https://cloudflare.jove.com/files/ftp_upload/57324/57324fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57324/57324fig2.jpg" /> 
Figure 2. Non-enzymatic RNA replication inside OA vesicles. A. Scheme of non-enzymatic RNA primer extension. B. PAGE image of a primer extension reaction inside pure oleic acid vesicles, with conditions as in section 4. C. Linear fit of the natural logarithm of ratio of amount of primer remaining at given time point to the initial amount of primer vs. time over 30 h. Reaction rate, calculated from the slope of ln(P/P0) vs time, is 0.058 h-1. <a href="https://cloudflare.jove.com/files/ftp_upload/57324/57324fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57324/57324fig3.jpg" /> 
Figure 3. Hammerhead ribozyme cleavage in OA/GMO vesicles. A. Scheme of hammerhead ribozyme cleavage of fluorescently labeled substrate strand (top). B. PAGE image of hammerhead ribozyme cleavage inside OA/GMO vesicles with 5 mM Mg2+. C. Ribozyme activity inside vesicles. Linear fit of natural logarithm of ratio of amount of substrate remaining at given time point to the initial amount of substrate vs. time in first 4 h. Reaction rate, calculated from the slope of ln(S/S0) vs time, is 0.36 h-1. <a href="https://cloudflare.jove.com/files/ftp_upload/57324/57324fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57324/57324fig4.jpg" /> 
Figure 4. Giant Fatty Acid Vesicles for Microscopy. A. Confocal microscopy image of a Rhodamine PE labeled oleic acid vesicle, scale bar 10 &#956;m. B. Confocal microscopy image of oleic acid vesicle containing Alexa488 labeled RNA with membrane shown in transmitted detector (TD) channel, scale bar 5 &#956;m.

Watch this Scientific Journal Video about Preparation, Purification, and Use of Fatty Acid-containing Liposomes at JoVE.com

Preparation, Purification, and Use of Fatty Acid-containing Liposomes

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of single chain amphiphiles. Here we describe techniques for the preparation, purification, and use of liposomes comprised in part or whole of these amphiphiles.

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due ...

preparation-purification-and-use-of-fatty-acid-containing-liposomes

Microsoft Bing

Research

JoVE Journal

Bioengineering

41.0K Views.  Massachusetts General Hospital. Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of single chain amphiphiles. Here we describe techniques for the preparation, purification, and use of liposomes comprised in part or whole of these amphiphiles.

Video: Preparation, Purification, and Use of Fatty Acid-containing Liposomes

Constant Pressure-controlled Extrusion Method for the Preparation of Nano-sized Lipid Vesicles

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform to study protein-protein and protein-lipid interactions3, monitor drug delivery4,5, and encapsulation4. Phospholipids naturally create curved lipid bilayers, distinguishing itself from a micelle.6 Liposomes are traditionally classified by size and number of bilayers, i.e. large unilamellar vesicles (LUVs), small unilamellar vesicles (SUVs) and multilamellar vesicles (MLVs)7. In particular, the preparation of homogeneous liposomes of various sizes is important for studying membrane curvature that plays a vital role in cell signaling, endo- and exocytosis, membrane fusion, and protein trafficking8. Several groups analyze how proteins are used to modulate processes that involve membrane curvature and thus prepare liposomes of diameters &lt;100 - 400 nm to study their behavior on cell functions3. Others focus on liposome-drug encapsulation, studying liposomes as vehicles to carry and deliver a drug of interest9. Drug encapsulation can be achieved as reported during liposome formation9. Our extrusion step should not affect the encapsulated drug for two reasons, i.e. (1) drug encapsulation should be achieved prior to this step and (2) liposomes should retain their natural biophysical stability, securely carrying the drug in the aqueous core. These research goals further suggest the need for an optimized method to design stable sub-micron lipid vesicles.
Nonetheless, the current liposome preparation technologies (sonication10, freeze-and-thaw10, sedimentation) do not allow preparation of liposomes with highly curved surface (i.e. diameter &lt;100 nm) with high consistency and efficiency10,5, which limits the biophysical studies of an emerging field of membrane curvature sensing. Herein, we present a robust preparation method for a variety of biologically relevant liposomes.
Manual extrusion using gas-tight syringes and polycarbonate membranes10,5 is a common practice but heterogeneity is often observed when using pore sizes &lt;100 nm due to due to variability of manual pressure applied. We employed a constant pressure-controlled extrusion apparatus to prepare synthetic liposomes whose diameters range between 30 and 400 nm. Dynamic light scattering (DLS)10, electron microscopy11 and nanoparticle tracking analysis (NTA)12 were used to quantify the liposome sizes as described in our protocol, with commercial polystyrene (PS) beads used as a calibration standard. A near linear correlation was observed between the employed pore sizes and the experimentally determined liposomes, indicating high fidelity of our pressure-controlled liposome preparation method. Further, we have shown that this lipid vesicle preparation method is generally applicable, independent of various liposome sizes. Lastly, we have also demonstrated in a time course study that these prepared liposomes were stable for up to 16 hours. A representative nano-sized liposome preparation protocol is demonstrated below.

This protocol describes an extrusion method for preparing lipid vesicles of sub-micron sizes with a high degree of homogeneity. This method uses a pressure-controlled system with controlled nitrogen flow rates for liposome preparation. The lipid preparation1,2, liposome extrusion, and size characterization will be presented herein.

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform ...

Detergent-free Ultrafast Reconstitution of Membrane Proteins into Lipid Bilayers Using Fusogenic Complementary-charged Proteoliposomes.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers are less compatible with detergents.
Here we describe a strategy for bypassing this fundamental limitation using fusogenic oppositely charged liposomes bearing a membrane protein of interest. Fusion between such vesicles occurs within 5 min in a low ionic strength buffer. Positively charged fusogenic liposomes can be used as simple shuttle vectors for detergent-free delivery of membrane proteins into biomimetic target lipid bilayers, which are negatively charged. We also show how to reconstitute membrane proteins into fusogenic proteoliposomes with a fast 30-min protocol.
Combining these two approaches, we demonstrate a fast assembly of an electron transport chain consisting of two membrane proteins from E. coli, a primary proton pump bo3-oxidase and F1Fo ATP synthase, in membranes of vesicles of various sizes, ranging from 0.1 to &#62;10 microns, as well as ATP production by this chain.

Here we present two ultrafast protocols for reconstitution of membrane proteins into fusogenic proteoliposomes, and fusion of such proteoliposomes with target lipid bilayers for detergent-free delivery of these membrane proteins into the postfusion bilayer. The combination of these approaches enables fast and easily controlled assembly of complex multi-component membrane systems.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers ...

Biochemistry

Expression, Purification, and Liposome Binding of Budding Yeast SNX-BAR Heterodimers

SNX-BAR proteins are an evolutionarily conserved class of membrane remodeling proteins that play key roles in sorting and trafficking of protein and lipids during endocytosis, sorting within the endosomal system, and autophagy. Central to SNX-BAR protein function is the ability to form homodimers or heterodimers that bind membranes using highly conserved phox-homology (PX) and BAR (Bin/Amphiphysin/Rvs) domains. In addition, oligomerization of SNX-BAR dimers on membranes can elicit the formation of membrane tubules and vesicles and this activity is thought to reflect their functions as coat proteins for endosome-derived transport carriers. Researchers have long utilized in vitro binding studies using recombinant SNX-BAR proteins on synthetic liposomes or giant unilamellar vesicles (GUVs) to reveal the precise makeup of lipids needed to drive membrane remodeling, thus revealing their mechanism of action. However, due to technical challenges with dual expression systems, toxicity of SNX-BAR protein expression in bacteria, and poor solubility of individual SNX-BAR proteins, most studies to date have examined SNX-BAR homodimers, including non-physiological dimers that form during expression in bacteria. Recently, we have optimized a protocol to overcome the major shortcomings of a typical bacterial expression system. Using this workflow, we demonstrate how to successfully express and purify large amounts of SNX-BAR heterodimers and how to reconstitute them on synthetic liposomes for binding and tubulation assays.

Here, we present a workflow for the expression, purification and liposome binding of SNX-BAR heterodimers in yeast.

SNX-BAR proteins are an evolutionarily conserved class of membrane remodeling proteins that play key roles in sorting and trafficking of protein and ...

Genetics

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Neuroscience

Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense lipidic nanotubes that are, in terms of morphology and dynamics, reminiscent of ER. The networks are derived from phospholipid double bilayer membrane patches adhering to a transparent Al2O3 substrate. The adhesion is mediated by Ca2+ in the ambient buffer. Subsequent depletion of Ca2+ by means of BAPTA/EDTA causes retraction of the membrane, resulting in spontaneous lipid nanotube network formation. The method only comprises phospholipids and microfabricated surfaces for simple formation of an ER model and does not require the addition of proteins or chemical energy (e.g., GTP or ATP). In contrast to the 3D morphology of the cellular endoplasmic reticulum, the model is two-dimensional (albeit the nanotube dimensions, geometry, structure, and dynamics are maintained). This unique in vitro ER model consists of only a few components, is easy to construct, and can be observed under a light microscope. The resulting structure can be further decorated for additional functionality, such as the addition of ER-associated proteins or particles to study transport phenomena among the tubes. The artificial networks described here are suitable structural models for the cellular ER, whose unique characteristic morphology has been shown to be related to its biological function, whereas details regarding formation of the tubular domain and rearrangements within are still not completely understood. We note that this method uses Al2O3 thin-film-coated microscopy coverslips, which are commercially available but require special orders. Therefore, it is advisable to have access to a microfabrication facility for preparation.

Solid-supported, protein-free, double phospholipid bilayer membranes (DLBM) can be transformed into complex and dynamic lipid nanotube networks and can serve as 2D bottom-up models of the endoplasmic reticulum.

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense ...

Preparation and Characterization of Nanoliposomes for the Entrapment of Bioactive Hydrophilic Globular Proteins

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their biocompatibility, biodegradability, non-immunogenicity, non-toxicity, and ability to entrap both hydrophilic and hydrophobic compounds. The classical hydration of thin lipid films in an organic solvent is applied herein as a technique to encapsulate tarin, a plant lectin, in nanoliposomes. Nanoliposome size, stability, entrapment efficiency, and morphological characterization are described in detail. The nanoliposomes are prepared using 1,2-dioleoyl-sn-glycerol-3-phosphoethanolamine (DOPE), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)-2000] (ammonium salt; DSPE-MPEG 2000), and cholesterylhemisuccinate (CHEMS) as the main constituents. Lipids are first dissolved in chloroform to obtain a thin lipid film that is subsequently rehydrated in ammonium sulfate solution containing the protein to be entrapped and incubated overnight. Then, sonication and extrusion techniques are applied to generate nanosized unilamellar vesicles. The size and polydispersity index of the nanovesicles are determined by dynamic light scattering, while nanovesicle morphology is assessed by scanning electron microscopy. Entrapment efficiency is determined by the ratio of the amount of unencapsulated protein to original amount of initially loaded protein. Homogeneous liposomes are obtained with an average size of 155 nm and polydispersity index value of 0.168. A high entrapment efficiency of 83% is achieved.

This study describes classical hydration using the thin lipid film method for nanoliposome preparation followed by nanoparticle characterization. A 47 kDa-hydrophilic and globular protein, tarin, is successfully encapsulated as a strategy to improve stability, avoid fast clearance, and promote controlled release. The method can be adapted to hydrophobic molecules encapsulation.

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their ...

Microfluidic Production of Lysolipid-Containing Temperature-Sensitive Liposomes

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading chemotherapeutic drugs, such as doxorubicin (DOX). To achieve this, an ethanolic lipid mixture and ammonium sulfate solution are injected into a staggered herringbone micromixer (SHM) microfluidic device. The solutions are rapidly mixed by the SHM, providing a homogeneous solvent environment for liposomes self-assembly. Collected liposomes are first annealed, then dialyzed to remove residual ethanol. An ammonium sulfate pH-gradient is established through buffer exchange of the external solution by using size exclusion chromatography. DOX is then remotely loaded into the liposomes with high encapsulation efficiency (&#62; 80%). The liposomes obtained are homogenous in size with Z-average diameter of 100 nm. They are capable of temperature-triggered burst release of encapsulated DOX in the presence of mild hyperthermia (42 &#176;C). Indocyanine green (ICG) can also be co-loaded into the liposomes for near-infrared laser-triggered DOX release. The microfluidic approach ensures high-throughput, reproducible and scalable preparation of LTSLs.

The protocol presents the optimized parameters for preparing thermosensitive liposomes using the staggered herringbone micromixer microfluidics device. This also allows co-encapsulation of doxorubicin and indocyanine green into the liposomes and the photothermal-triggered release of doxorubicin for controlled/triggered drug release.

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading ...

Preparation, Administration, and Assessment of In Vivo Tissue-Specific Cellular Uptake of Fluorescent Dye-Labeled Liposomes

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome formulation, liposomes may be preferentially taken up by different cell types in the body. This may influence the efficacy of the therapeutic particle as progression of different diseases is tissue- and cell-type-specific. In this protocol, we present one method for synthesizing and fluorescently labeling liposomes using DSPC, cholesterol, and PEG-2000 DSPE and the lipid dye DiD as a fluorescent label. This protocol also presents an approach for delivering liposomes in vivo and assessing cell-specific uptake of liposomes using flow cytometry. This approach can be used to determine the types of cells that take up liposomes and quantify the distribution and proportion of liposome-uptake across cell types and tissues. While not mentioned in this protocol, additional assays such as immunofluorescence and single-cell fluorescence imaging on a cytometer will strengthen any findings or conclusions made as they permit assessment of intracellular staining. Protocols may also need to be adapted depending on the tissue(s) of interest.

The goal of this protocol is to synthesize fluorescently-labeled liposomes and use flow cytometry to identify in vivo localization of liposomes at a cellular level.

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome ...

Assembly of Cell Mimicking Supported and Suspended Lipid Bilayer Models for the Study of Molecular Interactions

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial protective barrier for all cell types, separating the internal cellular components from the extracellular environment. These membranes are composed largely of a lipid bilayer, which contains outer hydrophilic head groups and inner hydrophobic tail groups, along with various proteins and cholesterol. The composition and structure of the lipids themselves play a crucial role in regulating biological function, including interactions between cells and the cellular microenvironment, which may contain pharmaceuticals, biological toxins, and environmental toxicants. In this study, methods to formulate uni-lipid and multi-lipid supported and suspended cell mimicking lipid bilayers are described. Previously, uni-lipid phosphatidylcholine (PC) lipid bilayers as well as multi-lipid placental trophoblast-inspired lipid bilayers were developed for use in understanding molecular interactions. Here, methods for achieving both types of bilayer models will be presented. For cell mimicking multi-lipid bilayers, the desired lipid composition is first determined via lipid extraction from primary cells or cell lines followed by liquid chromatography-mass spectrometry (LC-MS). Using this composition, lipid vesicles are fabricated using a thin-film hydration and extrusion method and their hydrodynamic diameter and zeta potential are characterized. Supported and suspended lipid bilayers can then be formed using quartz crystal microbalance with dissipation monitoring (QCM-D) and on a porous membrane for use in a parallel artificial membrane permeability assay (PAMPA), respectively. The representative results highlight the reproducibility and versatility of in vitro cell membrane lipid bilayer models. The methods presented can aid in rapid, facile assessment of the interaction mechanisms, such as permeation, adsorption, and embedment, of various molecules and macromolecules with a cell membrane, helping in the screening of drug candidates and prediction of potential cellular toxicity.

This protocol describes the formation of cell mimicking uni-lipid and multi-lipid vesicles, supported lipid bilayers, and suspended lipid bilayers. These in vitro models can be adapted to incorporate a variety of lipid types and can be used to investigate various molecule and macromolecule interactions.

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial ...

Examining BCL-2 Family Function with Large Unilamellar Vesicles

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate apoptosis1. Following cellular stress (e.g., DNA damage), the pro-apoptotic BCL-2 family effectors BAK (BCL-2 antagonistic killer 1) and/or BAX (BCL-2 associated X protein) become activated and compromise the integrity of the outer mitochondrial membrane (OMM), though the process referred to as mitochondrial outer membrane permeabilization (MOMP)1. After MOMP occurs, pro-apoptotic proteins (e.g., cytochrome c) gain access to the cytoplasm, promote caspase activation, and apoptosis rapidly ensues2.
In order for BAK/BAX to induce MOMP, they require transient interactions with members of another pro-apoptotic subset of the BCL-2 family, the BCL-2 homology domain 3 (BH3)-only proteins, such as BID (BH3-interacting domain agonist)3-6. Anti-apoptotic BCL-2 family proteins (e.g., BCL-2 related gene, long isoform, BCL-xL; myeloid cell leukemia 1, MCL-1) regulate cellular survival by tightly controlling the interactions between BAK/BAX and the BH3-only proteins capable of directly inducing BAK/BAX activation7,8. In addition, anti-apoptotic BCL-2 protein availability is also dictated by sensitizer/de-repressor BH3-only proteins, such as BAD (BCL-2 antagonist of cell death) or PUMA (p53 upregulated modulator of apoptosis), which bind and inhibit anti-apoptotic members7,9. As most of the anti-apoptotic BCL-2 repertoire is localized to the OMM, the cellular decision to maintain survival or induce MOMP is dictated by multiple BCL-2 family interactions at this membrane.
 Large unilamellar vesicles (LUVs) are a biochemical model to explore relationships between BCL-2 family interactions and membrane permeabilization10. LUVs are comprised of defined lipids that are assembled in ratios identified in lipid composition studies from solvent extracted Xenopus mitochondria (46.5% phosphatidylcholine, 28.5% phosphatidylethanoloamine, 9% phosphatidylinositol, 9% phosphatidylserine, and 7% cardiolipin)10. This is a convenient model system to directly explore BCL-2 family function because the protein and lipid components are completely defined and tractable, which is not always the case with primary mitochondria. While cardiolipin is not usually this high throughout the OMM, this model does faithfully mimic the OMM to promote BCL-2 family function. Furthermore, a more recent modification of the above protocol allows for kinetic analyses of protein interactions and real-time measurements of membrane permeabilization, which is based on LUVs containing a polyanionic dye (ANTS: 8-aminonaphthalene-1,3,6-trisulfonic acid) and cationic quencher (DPX: p-xylene-bis-pyridinium bromide)11. As the LUVs permeabilize, ANTS and DPX diffuse apart, and a gain in fluorescence is detected. Here, commonly used recombinant BCL-2 family protein combinations and controls using the LUVs containing ANTS/DPX are described.

Biochemically-defined large unilamellar vesicles (LUVs) are a convenient model system to analyze BCL-2 family interactions with immediate implications in better understanding the mitochondrial pathway of apoptosis. A method to produce LUVs, along with standard BCL-2 family protein combinations and controls to examine LUV permeabilization, are presented.