The authors thank the BMBF for financial support and the Center for Biological Systems Analysis (ZBSA) for providing the research facility. We are grateful to P. G. Schultz, TSRI, La Jolla, California, USA for providing the plasmid pEVOL-pAzF. We thank the staff of the Life Imaging Center (LIC) in the Center for Biological Systems Analysis (ZBSA) of the Albert-Ludwigs-University Freiburg for help with their confocal microscopy resources, and the excellent support in image recording.

1. Design and cloning of amphiphilic elastin-like proteins (ELPs)
<ol>
	<li>Clone and design the constructs as described elsewhere8,20. Plasmids are available upon request.</li>
</ol>
2. Protein expression, purification and preparation
<ol>
	<li>Expression of F20E20-mEGFP and F20E20-mCherry
	<ol>
		<li>Inoculate main expression culture from overnight pre-culture to an OD600 of 0.3. Incubate at 37 &#176;C, 200 rpm in sterile 400 mL LB medium supplemented with appropriated antibiotics in a 2 L flask.</li>
		<li>Prepare IPTG stock solution (1 M) for induction of the expression culture in ultrapure water.</li>
		<li>When OD600 0.5&#8211;0.8 is reached, add IPTG to expression culture to a final concentration of 1 mM and reduce incubation temperature to 20 &#176;C. Allow expression at 20 &#176;C for approximately 20 h at 200 rpm.</li>
	</ol>
	</li>
	<li>Expression of amphiphilic ELP containing UAA pAzF
	<ol>
		<li>Inoculate main expression culture from overnight E. coli pre-culture containing the two plasmids pEVOL pAzF and e.g. pET28-NMBL-(TAG)R40F20-his&#160;to an OD600 of 0.3 (see supplementary information for amino acid sequences). Incubate at 37 &#176;C, 200 rpm in sterile 400 mL LB medium supplemented with kanamycin and chloramphenicol in a 2 L flask.</li>
		<li>Prepare 100 mM pAzF stock solution in ultrapure water. For 10 mL of pAzF stock solution, weigh 206.2 mg pAzF and resuspend it in 8 mL of ultrapure water. To dissolve the pAzF raise the pH of the solution with 3 M NaOH and mix vigorously. When pAzF is dissolved, carefully lower the pH to 10.5 and add ultrapure water to a final volume of 10 mL. Use a sterile filter (0.22 &#181;m) and aliquot the solution in 2 mL reaction tubes.</li>
		<li>Prepare 1 M IPTG stock solution in ultrapure water and 20% arabinose stock solution in ultrapure water.</li>
		<li>When OD600 0.5&#8211;0.8 is reached, add pAzF to the expression culture to a final concentration of 2 mM. Incubate culture for 10 min, 37 &#176;C, 200 rpm to allow for pAzF uptake.</li>
		<li>Induce expression of target protein and expression of the necessary tRNA/t-RNA synthetase via simultaneous addition of IPTG (1 mM) and arabinose (2%) and reduce incubation temperature to 20 &#176;C.</li>
		<li>Allow expression at 20 &#176;C for approximately 20 h at 200 rpm. Harvest expression culture by centrifugation at 4 &#176;C, 4000 x g, 40 min.</li>
	</ol>
	</li>
	<li>Cell lysis and protein purification
	<ol>
		<li>Resuspend the E. coli pellet in lysis buffer (10 mL per liter of culture; 50&#160;mM Tris-HCl pH&#160;8, 500&#160;mM NaCl, 4&#160;M urea, 0.25% Triton X-100) containing lysozyme (0.1 mg/mL) and PMSF (0.1 mM). Incubate for 30 min on ice and freeze and thaw twice afterwards by submerging the sample in liquid nitrogen.</li>
		<li>Sonicate the suspension (30%, 15&#160;times, 30&#160;s:&#160;10 s break) and clear the lysate via centrifugation (4 &#176;C, 10,000&#160;x g for 40&#160;min).</li>
		<li>Purify protein using affinity chromatography (e.g. on a 1&#160;mL nickel column using a FPLC system connected to a fraction collector; see Table of Materials). Elute the protein with elution buffer (50 mM Tris-HCl, 500 mM NaCl, 4 M urea, 250&#8211;500 mM imidazole) and store at 4 &#176;C until further processing.</li>
		<li>Analyze the purification efficiency via SDS-PAGE.</li>
	</ol>
	</li>
</ol>
3. Dye-modification of ELPs via SPAAC
<ol>
	<li>Roughly estimate the concentration of the ELP solution. &#160;A280 absorption for concentration evaluation is not valuable since pAzF-R40F20 sequence is&#160;lacking amino acids absorbing in the UV range. Therefore, a previously lyophilized and weighted ELP amphiphile can be used as a reference for SDS PAGE band comparison. Through comparison of the summed gray value intesity of SDS PAGE bands from ELP solutions with known concentrations and your sample the rough concentration of your sample can be estimated.</li>
	<li>Add 1 &#181;L of fluorescent dye BDP-FL-PEG4-DBCO (10 mM stock solution; 20&#160;&#181;M final concentration) to 500 &#181;L of ELP solution (~20 &#181;M). Incubate the reaction for about 10 h at 15 &#176;C, while shaking and protected from light.</li>
	<li>For further use, dialyze the reaction to remove excessive BDP.
	<ol>
		<li>Equilibrate a dialysis membrane (e.g. 12 kDa cutoff) in ultrapure water for 10 min. Cut the dialysis membrane into the correct size to be placed on top of the opening of an reaction tube containing the clicked ELP solution. To fix the dialysis membrane in the opening, place a reaction tube lid with punched out core on the opening, thus closing the tube.</li>
		<li>Place the reaction tube upside down in the chosen buffer. Exchange the buffer (2&#8211;5 L) twice after dialysis for at least 3 h every time. Remove any air bubbles trapped between the dialysis membrane and the buffer to ensure successful dialysis.</li>
	</ol>
	</li>
</ol>
4. THF swelling protocol
<ol>
	<li>Dialyze homogenous ELP solution against phosphate or tris buffer (10 mM) with stable pH 7.5 to remove salts and remaining compounds from his-tag purification.</li>
	<li>Prepare the lyophilizer and cool down to starting temperature for freeze-drying.</li>
	<li>Aliquot the dialyzed protein solution in 1.5 mL reaction tubes (50&#8211;500 &#181;L per tube) and shock freeze in liquid nitrogen. To avoid unwanted mixing of different protein solutions during freeze-drying, caps with a small hole can be put on top of the reaction tube to seal it partially.</li>
	<li>Take the frozen protein samples out of the liquid nitrogen and immediately place them in the lyophilizer to start freeze-drying. Freeze-drying is finished when the sample is completely dry (approximately 24&#8211;48 h). Subsequently, ventilate lyophilized amphiphilic ELPs with dry N2, then immediately close the reaction tube lids to avoid contact with air moisture.</li>
	<li>Add pure THF to the lyophilized samples (ELP, 5&#8211;10 &#181;M) and place the solution in a water bath sonicator containing ice water for 15 min to allow for swelling of the ELP in THF.</li>
	<li>Preheat a thermocycler to 30&#8211;60 &#176;C for vesicle formation or up to 90 &#176;C for fiber formation and prepare new reaction tubes containing either ultrapure water or buffer (50&#160;mM NaH2PO4/Na2HPO4, 50 mM NaCl, pH 5&#8211;13). Spherical coacervates assemble predominantly at 20 &#176;C within pH 9&#8211;13. Vesicle formation is favored at 50&#8211;60 &#176;C between pH 7 and 9. Fiber formation is predominently induced above 60 &#176;C between pH 5 and 12.</li>
	<li>After the sonication step, place the ELP/THF solution as well as the prepared ultrapure water or buffer solution in the thermocycler and heat up to the desired temperature for 5 min. When temperature is reached the preheated ELP/THF solution should be carefully stratified on top of the preheated ultrapure water or buffer solution. A clear separation of the two phases with a distinct interface should be visible.</li>
	<li>Place the mixture in the thermocycler again and incubate for 20 min to allow for vesicle or fiber formation at the interface. Afterwards, let the samples cool down to room temperature for 10 min before analysis via fluorescence microscopy or dialysis.</li>
	<li>Dialyze solution containing the supramolecular structures against ultrapure water or buffer (50 mM NaH2PO4/Na2HPO4, 50 mM NaCl, pH 7&#8211;10).</li>
</ol>
5. BuOH extrusion protocol
<ol>
	<li>Prepare a 1&#8211;50 &#181;M ELP solution in ultrapure water or buffer (50 mM PB pH 7.5, 100 mM NaCl, may contain up to 4 M urea). The concentration of the amphiphilic ELP F20R20-mEGFP and F20R20-mCherry solution can be determined using the molar extinction coefficients (F20R20-mEGFP A280&#160;= 22015 M-1 cm-1 and F20R20-mCherry A280 = 34380 M-1 cm-1) (see supplementary information for aa sequences).</li>
	<li>Add 10%&#8211;20% (v/v) 1-butanol and immediately mix the solution by pipetting up and down or drawing it up through a syringe multiple times. A common 100 &#181;L pipette or Hamilton syringe equipped with a 0.25 x 25 mm needle can be applied. The turbidity of the solution during mixing should increase, indicating vesicle formation. 1-octanol 5%&#8211;15% (v/v) can also be used for vesicle extrusion instead of 1-butanol.</li>
	<li>In order to achieve a narrow size distribution, extrude vesicles using a mini extruder through a membrane with a pore size of 1 &#181;m at room temperature. The membrane size used for extrusion determines the upper size cutoff of the vesicles.</li>
	<li>Dialyze the vesicles as described above (step 3.3) to remove residual 1-butanol.</li>
</ol>
6. Dye encapsulation with the BuOH extrusion protocol
<ol>
	<li>Mix approximately 40 &#181;L ELP solution in 10 mM Tris-HCl, pH 8 with 1 &#181;L Dextran Texas Red (0.0025 mg/mL final concentration).</li>
	<li>Add 10 &#181;L of BuOH to the solution and extrude 5&#8211;10 times through a syringe equipped with a 0.25 x 25 mm needle.</li>
</ol>
7. Analysis of supramolecular structures using fluorescence microscopy
<ol>
	<li>Place a reinforcement ring on a glass slide and firmly press the adhesive side to the glass.</li>
	<li>Add 5 &#181;L of the sample to the inside of the reinforcement ring and place a cover slip on top.</li>
	<li>Seal the sample with nail polish at the edges of the cover slip to avoid evaporation of the sample during analysis.</li>
	<li>Carry out fluorescence microscopy as previously described8.</li>
</ol>

<ol>
	<li>Elzoghby, A. O., Samy, W. M., Elgindy, N. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein-based+nanocarriers+as+promising+drug+and+gene+delivery+systems.">Protein-based nanocarriers as promising drug and gene delivery systems.</a> Journal of Controlled Release. 161 (1), 38-49 (2012).</li><li>Jang, Y., Champion, J. A. <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+Materials+Made+from+Functional+Recombinant+Proteins.">Self-Assembled Materials Made from Functional Recombinant Proteins.</a> Accounts of Chemical Research. 49 (10), 2188-2198 (2016).</li><li>Timmermans, S. B. P. E., van Hest, J. C. M. <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+nanoreactors+based+on+peptides+and+proteins.">Self-assembled nanoreactors based on peptides and proteins.</a> Current Opinion in Colloid &amp; Interface Science. 35, 26-35 (2018).</li><li>Dreher, M. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temperature+Triggered+Self-Assembly+of+Polypeptides+into+Multivalent+Spherical+Micelles.">Temperature Triggered Self-Assembly of Polypeptides into Multivalent Spherical Micelles.</a> Journal of the American Chemical Society. 130 (2), 687-694 (2008).</li><li>Huber, M. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Designer+amphiphilic+proteins+as+building+blocks+for+the+intracellular+formation+of+organelle-like+compartments.">Designer amphiphilic proteins as building blocks for the intracellular formation of organelle-like compartments.</a> Nature Materials. 14 (1), 125-132 (2014).</li><li>Matsuurua, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rational+design+of+self-assembled+proteins+and+peptides+for+nano-+and+micro-sized+architectures.">Rational design of self-assembled proteins and peptides for nano- and micro-sized architectures.</a> RSC Advances. 4 (6), 2942-2953 (2013).</li><li>Rocklin, G. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+analysis+of+protein+folding+using+massively+parallel+design,+synthesis,+and+testing.">Global analysis of protein folding using massively parallel design, synthesis, and testing.</a> Science. 357 (6347), 168-175 (2017).</li><li>Schreiber, A., St&#252;hn, L. G., Huber, M. C., Geissinger, S. E., Rao, A., Schiller, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Self-Assembly+Toolbox+of+Tailored+Supramolecular+Architectures+Based+on+an+Amphiphilic+Protein+Library.">Self-Assembly Toolbox of Tailored Supramolecular Architectures Based on an Amphiphilic Protein Library.</a> Small. 15 (30), 1900163 (2019).</li><li>Jang, Y., Hsieh, M. -. C., Dautel, D., Guo, S., Grover, M. A., Champion, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Understanding+the+Coacervate-to-Vesicle+Transition+of+Globular+Fusion+Proteins+to+Engineer+Protein+Vesicle+Size+and+Membrane+Heterogeneity.">Understanding the Coacervate-to-Vesicle Transition of Globular Fusion Proteins to Engineer Protein Vesicle Size and Membrane Heterogeneity.</a> Biomacromolecules. 20 (9), 3494-3503 (2019).</li><li>Vargo, K. B., Sood, N., Moeller, T. D., Heiney, P. A., Hammer, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spherical+micelles+assembled+from+variants+of+recombinant+oleosin.">Spherical micelles assembled from variants of recombinant oleosin.</a> Langmuir: the ACS journal of surfaces and colloids. 30 (38), 11292-11300 (2014).</li><li>Bellomo, E. G., Wyrsta, M. D., Pakstis, L., Pochan, D. J., Deming, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stimuli-responsive+polypeptide+vesicles+by+conformation-specific+assembly.">Stimuli-responsive polypeptide vesicles by conformation-specific assembly.</a> Nature Materials. 3 (4), 244-248 (2004).</li><li>Mart&#237;n, L., Castro, E., Ribeiro, A., Alonso, M., Rodr&#237;guez-Cabello, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temperature-Triggered+Self-Assembly+of+Elastin-Like+Block+Co-Recombinamers:The+Controlled+Formation+of+Micelles+and+Vesicles+in+an+Aqueous+Medium.">Temperature-Triggered Self-Assembly of Elastin-Like Block Co-Recombinamers:The Controlled Formation of Micelles and Vesicles in an Aqueous Medium.</a> Biomacromolecules. 13 (2), 293-298 (2012).</li><li>Li, Y., Rodriguez-Cabello, J. C., Aparicio, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intrafibrillar+Mineralization+of+Self-Assembled+Elastin-Like+Recombinamer+Fibrils.">Intrafibrillar Mineralization of Self-Assembled Elastin-Like Recombinamer Fibrils.</a> ACS Applied Materials &amp; Interfaces. , (2017).</li><li>Vargo, K. B., Parthasarathy, R., Hammer, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Self-assembly+of+tunable+protein+suprastructures+from+recombinant+oleosin.">Self-assembly of tunable protein suprastructures from recombinant oleosin.</a> Proceedings of the National Academy of Sciences of the United States of America. 109 (29), 11657-11662 (2012).</li><li>Park, W. M., Champion, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thermally+Triggered+Self-Assembly+of+Folded+Proteins+into+Vesicles.">Thermally Triggered Self-Assembly of Folded Proteins into Vesicles.</a> Journal of the American Chemical Society. 136 (52), 17906-17909 (2014).</li><li>Vogele, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Towards+synthetic+cells+using+peptide-based+reaction+compartments.">Towards synthetic cells using peptide-based reaction compartments.</a> Nature Communications. 9 (1), 3862 (2018).</li><li>Vogele, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+Vesiculo+Synthesis+of+Peptide+Membrane+Precursors+for+Autonomous+Vesicle+Growth.">In Vesiculo Synthesis of Peptide Membrane Precursors for Autonomous Vesicle Growth.</a> Journal of Visualized Experiments. (148), e59831 (2019).</li><li>Huber, M. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Designer+amphiphilic+proteins+as+building+blocks+for+the+intracellular+formation+of+organelle-like+compartments.">Designer amphiphilic proteins as building blocks for the intracellular formation of organelle-like compartments.</a> Nature Materials. 14 (1), 125-132 (2015).</li><li>Urry, D. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elastin:+a+representative+ideal+protein+elastomer.">Elastin: a representative ideal protein elastomer.</a> Philosophical Transactions of the Royal Society B: Biological Sciences. 357 (1418), 169-184 (2002).</li><li>Huber, M. C., Schreiber, A., Wild, W., Benz, K., Schiller, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Introducing+a+combinatorial+DNA-toolbox+platform+constituting+defined+protein-based+biohybrid-materials.">Introducing a combinatorial DNA-toolbox platform constituting defined protein-based biohybrid-materials.</a> Biomaterials. 35 (31), 8767-8779 (2014).</li><li>Schreiber, A., Huber, M. C., Schiller, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prebiotic+Protocell+Model+Based+on+Dynamic+Protein+Membranes+Accommodating+Anabolic+Reactions.">Prebiotic Protocell Model Based on Dynamic Protein Membranes Accommodating Anabolic Reactions.</a> Langmuir. 35 (29), 9593-9610 (2019).</li><li>Chin, J. W., Santoro, S. W., Martin, A. B., King, D. S., Wang, L., Schultz, P. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Addition+of+p-Azido-l-phenylalanine+to+the+Genetic+Code+of+Escherichia+coli.">Addition of p-Azido-l-phenylalanine to the Genetic Code of Escherichia coli.</a> Journal of the American Chemical Society. 124 (31), 9026-9027 (2002).</li><li>Sonnino, S., Prinetti, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+domains+and+the+"lipid+raft"+concept.">Membrane domains and the "lipid raft" concept.</a> Current Medicinal Chemistry. 20 (1), 4-21 (2013).</li><li>Br&#228;se, S., Gil, C., Knepper, K., Zimmermann, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organische+Azide+-+explodierende+Vielfalt+bei+einer+einzigartigen+Substanzklasse.">Organische Azide - explodierende Vielfalt bei einer einzigartigen Substanzklasse.</a> Angewandte Chemie. 117 (33), 5320-5374 (2005).</li><li>Li, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large-Scale+Structures+in+Tetrahydrofuran&#8211;Water+Mixture+with+a+Trace+Amount+of+Antioxidant+Butylhydroxytoluene+(BHT).">Large-Scale Structures in Tetrahydrofuran&#8211;Water Mixture with a Trace Amount of Antioxidant Butylhydroxytoluene (BHT).</a> The Journal of Physical Chemistry B. 115 (24), 7887-7895 (2011).</li><li>Huber, M. C., Schreiber, A., Schiller, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Minimalist+Protocell+Design:+A+Molecular+System+Based+Solely+on+Proteins+that+Form+Dynamic+Vesicular+Membranes+Embedding+Enzymatic+Functions.">Minimalist Protocell Design: A Molecular System Based Solely on Proteins that Form Dynamic Vesicular Membranes Embedding Enzymatic Functions.</a> ChemBioChem. 20 (20), 2618-2632 (2019).</li><li>Raghunathan, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparative+study+on+the+stability+and+structure+of+two+different+green+fluorescent+proteins+in+organic+co-solvent+systems.">A comparative study on the stability and structure of two different green fluorescent proteins in organic co-solvent systems.</a> Biotechnology and Bioprocess Engineering. 18 (2), 342-349 (2013).</li><li>Sallach, R. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+biostability+of+self-assembling+protein+polymers+in+the+absence+of+covalent+crosslinking.">Long-term biostability of self-assembling protein polymers in the absence of covalent crosslinking.</a> Biomaterials. 31 (4), 779-791 (2010).</li></ol>

The authors declare no competing financial interests.

A fault while following the described protocols for&#160;the assembly of defined supramolecular structures mainly leads either to the formation of unspecific aggregates (Figure 2, IV) or to homogeneously distributed ELP-amphiphiles. Critical steps of the protocol are discussed below:
For high expression yield of the amphiphilic ELP, a relatively low temperature of 20&#176;C is optimal. For successful affinity based purification of the amphiphilic ELP an urea concentration of 4 M in the lysis buffer was proven to best solubilize the amphiphilic ELP and increase the protein yield in the soluble elution fraction. If lower urea concentrations in the lysis buffer are desired, affinity purification must be tested for the individual constructs. 2 M urea worked as well for some constructs, especially for those where the His-tag was fused to the hydrophilic domain and therefore still able to bind the resin. &#160;An additional purification step after His-tag purification via size exclusion chromatography can increase the vesicle yield as well.
In case of applying the THF-swelling protocol, the amphiphilic ELP needs to be labeled with a fluorescent organic dye for visualization. Importantly for the BDP labeling of the amphiphilic ELP (see supplementary information for amino acid sequences containing UAA pAzF) via SPAAC is the absence of any reductant such as TCEP, DTT nor &#946;-mercaptoethanol in all purification buffers. This is necessary to avoid the well reported azide to amine reduction of pAzF prior to the SPAAC reaction24.
The exact reaction stoichiometry of dye to amphiphilic ELP (e.g. pAzF-R40F20) is not crucial since it is not necessary to label every single ELP molecule for simple vesicle visualization via epifluorescence microscopy. Therefore, the correlation of a reference SDS gel band and the corresponding weighted lyophilized sample is only necessary once for each protein construct. However, if close to 100% labeling yield is desired a ratio&#160;of 1:1 equivalents dye to ELP molecules is sufficient. Very similar amphiphilic ELPs were analyzed in our lab to be fully labeled at an equimolar addition of BDP (data not yet published).
For vesicle preparation using the THF swelling method, the most critical steps are the swelling of lyophilized amphiphilic ELP and subsequent stratification of this solution on top of the aqueous buffer phase. Therefore, the freshly lyophilized amphiphilic ELP should be as anhydrous as possible, which can be achieved by ventilation of the lyophilizer with dry nitrogen gas&#160;and immediate closure of the reaction tube lids. If available, septum sealed dry THF should be used to increase the vesicle yield, but THF p.a. (&#62;99.5%) without septum works as well. The stratification step upon swelling the amphiphilic ELP in dry THF should be executed very carefully. Successful stratification of the two temperature-controlled solutions leads to a clearly visible phase boundary between organic and aqueous phase. The initial stratification step should be conducted slowly even though elevated temperatures lead to thermal induced mixing of these phases. Emergent turbidity of the solution is due to light scattering of formed vesicles, fibers or coacervates. In control samples lacking the protein, no turbidity appears though small sized structures (up to 200&#160;nm) are reported for the THF water-interface25. The THF stratification step is the most critical and failure prone step of the swelling protocol. After the incubation step the supramolecular structures can be dialyzed against buffer or ultrapure water. Preferentially the same aqueous solution should be used which was applied&#160;for initial assembly in order to maintain the osmolarity and prevent swelling or shrinking of the assembled vesicles. After dialysis, the vesicles, fibers and coacervates are usually stable for at least one week. Depending on the environmental parameters during assembly often a small proportion of other supramolecular structures besides the main structure are present if the THF swelling method is applied8. The described THF method increases the vesicle assembly yield by one order of magnitude while the BuOH extrusion improves the yield by three orders of magnitude compared to our previously published in vitro method5.
The BuOH extrusion method is applied to obtain exclusively stable vesicular structures with high reproducibility, circumventing fibers and spherical coacervates. This method is less error prone and compatible with fluorescent proteins. Therefore F20R20-mEGFP or F20R20-mCherry can be applied as well as BDP-R40F20 or BDP-E20F20. The only critical step is the rapid mixing of the aqueous protein solution after addition of 10%&#8211;20% v/v BuOH. The F20R20-mEGFP or F20R20-mCherry concentration should be around 1&#8211;15&#160;&#181;M. By applying BuOH extrusion method vesicles can be assembled in ultrapure water or buffer containing up to 5 M NaCl or 4 M urea and pH ranging from 5 to 8. Extruded PMBCs in 20% v/v BuOH can be stored for at least 6&#160;months at 4&#176;C while preserving their vesicular structure. To narrow the vesicle size distribution, they can be extruded using a mini extruder through a membrane of 0.2-1 &#181;m pore size. This pore extrusion can be done directly after BuOH addition to the amphiphilic ELP or after vesicle assembly. If PMBCs are too concentrated for imaging, assembled vesicles in BuOH can be diluted through rapid mixing using aqueous buffer containing 10%&#8211;20% v/v BuOH.
The major limitation of the BuOH extrusion method is that PMBC dialysis against aqueous buffers often results in poor vesicle yield. Further, the presence of residual BuOH within the membrane space cannot be excluded since simple fatty acids were able to incorporate into the PMBC membrane21. Therefore, PMBC membranes might be to some extent be composed of protein and alkanol moieties.
Encapsulation of chemically diverse cargo molecules works best using the BuOH extrusion method. Further, DMSO as solvent for the stock solution of the dye to be captured increases the dye encapsulation efficiency. For delicate cargo to be encapsulated, 5%&#8211;10% v/v 1-octanol can be used for PMBC assembly and has been proven to be better compatible, when compared to BuOH, with functional encapsulated enzymes such as DNA-ligase or TEV protease21,26. However, due to the shorter chain length of n-butanol it can be dialyzed against aqueous buffer in contrast to 1-octanol, which is not able to permeate the applied dialysis-membrane. Another method limitation is that the applied temperatures and pH values needed to control the desired suprastructure formation can affect enzyme activity. In future work, affinity purification or size exclusion purification should be established to separate non-encapsulated versus encapsulated molecules without deteriorating vesicle membrane integrity.
In contrast to film rehydration methods16,17 the herein described protocols enable the assembly of vesicles sizes greater than 600 nm. This allows monitoring of real time fusion events through simple epifluorescence microscopy and the observation of membrane phase separation8. Compared to temperature triggered vesicular assembly of amphiphilic ELP9 the protocols described here yield PMBC with a long time stability of up to 6 month. However, the main disadvantage is the need of organic solvent for structure formation. Even though BuOH fully preserves the integrity and function of fluorescent proteins27 (data not shown), the activity of encapsulated enzymes might be restricted by residual organic solvent and must be tested individually. However, catalytic reactions involving DNA- ligase, TEV-protease and lipase have been successfully conducted within the luminal space of the vesicles, assembled by 1-octanol or BuOH extrusion26,21. Additionally, even though THF dialysis after assembly is very unproblematic and vesicle integrity is preserved, the BuOH removal frequently results in loss of vesicle integrity due to unknown reasons.
The described protocols enable researchers to assemble micrometer and sub micrometer sized supramolecular structures with distinct physicochemical properties, good encapsulation properties, and long time stability. These supramolecular structures can be applied for the design of minimal cells26 or artificial cell research21, enzyme encapsulation, or drug formulation. The presented functional PMBCs are further promising candidates for drug delivery, since their building blocks are not immunogenic28, exhibit dynamic fusion behavior, and allow for diverse cargo encapsulation.

The assembly of supramolecular structures for biotechnological applications is becoming increasingly important1,2,3,4,5. For the assembly of functional architectures such as coacervates, vesicles, and fibers with desired physicochemical properties it is important to understand and control the physicochemical and conformational properties of the components. Due to the molecular precision of molecules found in nature, building blocks for supramolecular structures are increasingly based on lipids, nucleic acids or proteins. Compared to synthetic polymers, proteinaceous building blocks allow for precise control over emergent supramolecular structures6 on the genetic level. The primary amino acid (aa) sequence of the individual protein building blocks intrinsically encodes the information for their assembly potential from the molecular up to the macroscopic level as well as the three dimensional shape and physical properties of the final supramolecular structure7.
Reported methods for the assembly of different supramolecular structures often involve amphiphilic proteins such as temperature sensitive elastin-like proteins (ELP)5,8,9, recombinant oleosin10 and artificial protein amphiphiles11. Temperature triggered methods have led to the assembly of micelles4,10,12, fibers13, sheets14 and vesicles9,15,16. Methods involving organic solvents have been applied for the formation of dynamic protein based vesicles8,11,14. So far, applied protocols for vesicle formation often lack assembly control over micrometer sized assemblies16,17 or have limited assembly yield5. In addition, some reported ELP based vesicles have impaired encapsulation potential12 or limited stability over time9. Addressing these drawbacks, the presented protocols enable the self-assembly of micrometer and sub micrometer sized supramolecular structures with distinct physiochemical properties, good encapsulation potential and long-time stability. Tailored amphiphilic ELPs assemble into supramolecular structures, spanning the range from spherical coacervates and highly ordered twisted fiber bundles to unilamellar vesicles depending on the applied protocol and associated environmental conditions. Large vesicular Protein Membrane-Based Compartments (PMBC) reveal all main phenotypes such as membrane fusion and phase separation behaviour analogous to liposomes. PMBCs efficiently encapsulate chemically diverse fluorescent cargo molecules which can be monitored using simple epifluorescence microscopy. The repetitive ELP domains used in this study are attractive building blocks for protein based supramolecular architectures18. The ELP pentapeptide repeat unit (VPGVG) is known to tolerate different aa besides proline at the fourth position (valine, V), while preserving its structural and functional properties19. The design of amphiphilic ELPs containing distinctive hydrophilic and hydrophobic domains was realized by inserting aa guest residues (X) in the VPGXG repeat with distinct hydrophobicity, polarity, and charge20. Amphiphilic ELP domains where equipped with hydrophobic phenylalanine (F) or isoleucine (I) while the hydrophilic domain contained charged glutamic acid (E) or arginine (R) as guest residues. A list of eligible amphiphilic ELP constructs and corresponding aa sequences can be found in the supplementary information and references8,21. All building blocks where equipped either with small fluorescent dyes or fluorescent proteins for visualization via fluorescence microscopy. mEGFP and other fluorescent proteins were N-terminally fused to the hydrophilic domains of the ELP amphiphiles . Organic dyes were conjugated via copper-free strain promoted alkyne&#8211;azide cycloaddition (SPAAC) to a co-translationally introduced unnatural amino acid (UAA). The co-translational incorporation of the UAA para-azidophenylalanine (pAzF)22 permits the N-terminal modification of the hydrophilic ELP domain. In this way the green fluorescent dye BDP-FL-PEG4-DBCO (BDP) or any small fluorescent molecule with a strained cyclooctyne can be used as fluorescent probe. Successful incorporation of the UAA pAzF and cycloaddition of the dye via SPAAC can be easily confirmed via LC-MS/MS due to efficient ionization of the corresponding tryptic peptides8. This small organic dye was applied to broaden the solvent choice for assembly protocols, since fluorescent proteins are incompatible with most organic solvents. The two most efficient assembly protocols for supramolecular structures developed in our lab are described below. The THF swelling method is only compatible with organic dye modified amphiphilic ELP. In contrast, the 1-butanol (BuOH) extrusion method is compatible with many proteins as fluorescent probe e.g. mEGFP, since the described method fully preserves the fluorescence of these fusion proteins. In addition, the encapsulation of small molecules and vesicular fusion behavior works best by employing the BuOH extrusion method.

<table><tbody><tr><td>1 &amp;micro;m and 0.2 &amp;micro;m Steril Filter</td><td>VWR</td><td></td><td></td></tr><tr><td>1,4-Dithiothreitol</td><td>Merck</td><td></td><td></td></tr><tr><td>1-butanol. &gt;99.5% p.a.</td><td>Roth</td><td></td><td></td></tr><tr><td>2log DNA ladder</td><td>NEB</td><td></td><td></td></tr><tr><td>2-Mercaptoethanol</td><td>Roth</td><td></td><td></td></tr><tr><td>50 mL Falcon tubes</td><td>VWR</td><td></td><td></td></tr><tr><td>79249 Alkyne Mega Stokes dye</td><td>Sigma Aldrich</td><td></td><td></td></tr><tr><td>Acetic acid glacial</td><td>VWR</td><td></td><td></td></tr><tr><td>Acetonitrile, anhydrous, 99.8%</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Ampicillin sodium-salt, 99%</td><td>Roth</td><td></td><td></td></tr><tr><td>BDP-FL-PEG4-DBCO</td><td>Jena Bioscience</td><td></td><td></td></tr><tr><td>Biofuge</td><td>Heraeus</td><td></td><td></td></tr><tr><td>Bottle Top Filter with PES membrane (45 &amp;micro;m, 22 &amp;micro;m)</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Brillant Blue G250 (Coomassie)</td><td>Roth</td><td></td><td></td></tr><tr><td>BspQI</td><td>NEB</td><td></td><td></td></tr><tr><td>Camera DS Qi1</td><td>Nikon</td><td></td><td></td></tr><tr><td>Centrifuge 5417r</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>Centrifuge 5810r</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>CF-400-Cu square mesh copper grid</td><td>EMS</td><td></td><td></td></tr><tr><td>Chloramphenicol</td><td>Roth</td><td></td><td></td></tr><tr><td>CompactStar CS 4</td><td>VWR</td><td></td><td></td></tr><tr><td>Dextran, Texas Red, 3000 MW, neutral</td><td>Life Technologies</td><td></td><td></td></tr><tr><td>Digital sonifier</td><td>Branson</td><td></td><td></td></tr><tr><td>Dimethylsulfoxide (DMSO)</td><td>Applichem</td><td></td><td></td></tr><tr><td>Dnase I</td><td>Applichem</td><td></td><td></td></tr><tr><td>EarI</td><td>NEB</td><td></td><td></td></tr><tr><td>EcoRI-HF</td><td>NEB</td><td></td><td></td></tr><tr><td>Environmental shaker incubator ES-20</td><td>Biosan</td><td></td><td></td></tr><tr><td>Ethanol absolute</td><td>Roth</td><td></td><td></td></tr><tr><td>Ethidium bromide solution</td><td>Roth</td><td></td><td></td></tr><tr><td>Filter supports</td><td>Avanti</td><td></td><td></td></tr><tr><td>Glass plates</td><td>Bio-Rad</td><td></td><td></td></tr><tr><td>Glycerol Proteomics Grade</td><td>Amresco</td><td></td><td></td></tr><tr><td>Glycin</td><td>Applichem</td><td></td><td></td></tr><tr><td>H4-Azido-Phe-OH</td><td>Bachhem</td><td></td><td></td></tr><tr><td>Heat plate MR HeiTec</td><td>Heidolph</td><td></td><td></td></tr><tr><td>HindIII</td><td>NEB</td><td></td><td></td></tr><tr><td>HisTrap FF crude column</td><td>GE Life Sciences</td><td></td><td>Nickel column</td></tr><tr><td>Hydrochloride acid fuming, 37%, p.a.</td><td>Merck</td><td></td><td></td></tr><tr><td>Illuminator ix 20</td><td>INTAS</td><td></td><td></td></tr><tr><td>Illuminator LAS-4000</td><td>Fujifilm</td><td></td><td></td></tr><tr><td>Imidazole</td><td>Merck</td><td></td><td></td></tr><tr><td>Immersions oil for microscopy</td><td>Merck</td><td></td><td></td></tr><tr><td>Incubators shakers Unimax 1010</td><td>Heidolph</td><td></td><td></td></tr><tr><td>Inkubator 1000</td><td>Heidolph</td><td></td><td></td></tr><tr><td>IPTG, &gt;99%</td><td>Roth</td><td></td><td></td></tr><tr><td>Kanamycinsulfate</td><td>Roth</td><td></td><td></td></tr><tr><td>L(+)-Arabinose</td><td>Roth</td><td></td><td></td></tr><tr><td>Laboratory scales Extend ed2202s/224s-OCE</td><td>Sartorius</td><td></td><td></td></tr><tr><td>LB-Medium</td><td>Roth</td><td></td><td></td></tr><tr><td>Lyophilizer Alpha 2-4 LSC</td><td>Christ</td><td></td><td></td></tr><tr><td>Lysozyme, 20000 U/mg</td><td>Roth</td><td></td><td></td></tr><tr><td>Microscope CM 100</td><td>Philips</td><td></td><td></td></tr><tr><td>Microscope Eclipse TS 100</td><td>Nikon</td><td></td><td></td></tr><tr><td>Microscopy cover glasses (15 x 15 mm)</td><td>VWR</td><td></td><td></td></tr><tr><td>Microscopy slides</td><td>VWR</td><td></td><td></td></tr><tr><td>Microwave</td><td>Studio</td><td></td><td></td></tr><tr><td>Mini-Extruder Set</td><td>Avanti Polar Lipids</td><td></td><td></td></tr><tr><td>NaCl, &gt;99.5%, p.a.</td><td>Roth</td><td></td><td></td></tr><tr><td>Natriumhydroxid pellets</td><td>Roth</td><td></td><td></td></tr><tr><td>Ni-NTA Agarose, PerfectPro</td><td>5 Prime</td><td></td><td></td></tr><tr><td>Nucleopore Track-Etch Membrane</td><td>Avanti</td><td></td><td></td></tr><tr><td>PH meter 766 calimatic</td><td>Knick</td><td></td><td></td></tr><tr><td>Phenylmethylsulfonylflourid (PMSF)</td><td>Roth</td><td></td><td></td></tr><tr><td>Polypropylene Columns (1 mL)</td><td>Qiagen</td><td></td><td></td></tr><tr><td>PowerPac basic</td><td>BioRad</td><td></td><td></td></tr><tr><td>Propanol-2-ol</td><td>Emplura</td><td></td><td></td></tr><tr><td>Protein ladder 10-250 kDa</td><td>NEB</td><td></td><td></td></tr><tr><td>Recirculating cooler F12</td><td>Julabo</td><td></td><td></td></tr><tr><td>Reinforcement rings</td><td>Herma</td><td></td><td></td></tr><tr><td>SacI HF</td><td>NEB</td><td></td><td></td></tr><tr><td>SDS Pellets</td><td>Roth</td><td></td><td></td></tr><tr><td>Sodiumdihydrogen phosphate dihydrate, NaH2PO4</td><td>VWR</td><td></td><td></td></tr><tr><td>Sterile syringe filter 0.2 mm Cellulose Acetate</td><td>VWR</td><td></td><td></td></tr><tr><td>T4 DNA Ligase</td><td>NEB</td><td></td><td></td></tr><tr><td>TEMED</td><td>Roth</td><td></td><td></td></tr><tr><td>TexasRed Dextran-Conjugate</td><td>MolecularProbes</td><td></td><td></td></tr><tr><td>Thermomix comfort</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>THF, &gt;99.5% p.a.</td><td>Acros</td><td></td><td></td></tr><tr><td>Triton X 100</td><td>Roth</td><td></td><td></td></tr><tr><td>Trypton/Pepton from Casein</td><td>Roth</td><td></td><td></td></tr><tr><td>Ultrasonic cleaner</td><td>VWR</td><td></td><td></td></tr><tr><td>Urea p.a.</td><td>Roth</td><td></td><td></td></tr><tr><td>Vacuum pump 2.5</td><td>Vacuubrand</td><td></td><td></td></tr><tr><td>XbaI</td><td>NEB</td><td></td><td></td></tr><tr><td>XhoI</td><td>NEB</td><td></td><td></td></tr><tr><td>ZelluTrans regenerated cellulose tubular membrane (12.0 S/ 3.5 S/ 1.0 V)</td><td>Roth</td><td></td><td></td></tr></tbody></table>

directed assembly of elastin-like proteins into defined supramolecular structures and cargo encapsulation in vitro

Tailored proteinaceous building blocks are versatile candidates for the assembly of supramolecular structures such as minimal cells, drug delivery vehicles and enzyme scaffolds. Due to their biocompatibility and tunability on the genetic level, Elastin-like proteins (ELP) are ideal building blocks for biotechnological and biomedical applications. Nevertheless, the assembly of protein based supramolecular structures with distinct physiochemical properties and good encapsulation potential remains challenging.
Here we provide two efficient protocols for guided self-assembly of amphiphilic ELPs into supramolecular protein architectures such as spherical coacervates, fibers and stable vesicles. The presented assembly protocols generate Protein Membrane-Based Compartments (PMBCs) based on ELPs with adaptable physicochemical properties. PMBCs demonstrate phase separation behavior and reveal method dependent membrane fusion and are able to encapsulate chemically diverse fluorescent cargo molecules. The resulting PMBCs have a high application potential as a drug formulation and delivery platform, artificial cell, and compartmentalized reaction space.

Protocol development for vesicle production 
Figure 1 outlines the two different vesicle preparation methods. The THF swelling method on the left side is composed of three successive steps and results in different supramolecular assemblies of the ELP depending on the temperature. In Figure 1A epifluorescence microscopy images show vesicles assembled from BDP-R20F20 and fibrillary structures assembled from BDP-R40F20. The BuOH method, illustrated on the right side exclusively leads to the formation of ELP vesicles, yielding about two orders of magnitude more vesicles compared to the THF swelling method. The schematic illustration shows the preparation process of BuOH vesicles. For vesicle preparation in Figure 1B BDP-R40F20 was mixed with 10-15% (v/v) BuOH and vesicles were prepared via extrusion of the mixture.
Guiding supramolecular self-assembly into different structures 
Figure 2 shows a schematic illustration and epifluorescence images of different supramolecular structures assembled from BDP-R40F20 via the THF swelling protocol. In this case lyophilized BDP-R40F20 was used for the different assembly protocols. The pH of the buffer and the temperature of the assembly process were&#160;adjusted to form either coacervates, fibrils or vesicles. The coacervates depicted in Figure 2A are 1&#8211;2&#160;&#181;m in diameter and were assembled from BDP-R40F20 at 20 &#176;C and pH 13. Adjustment of the assembly temperature to 90 &#176;C results in the formation of nanofiber bundles (Figure 2B) at pH 4&#8211;13 tested with BDP-R40F20. Stable vesicles could be assembled from the ELP at a temperature of 50 &#176;C and pH 7 (Figure 2C). Small mistakes at one of the crucial steps in the assembly protocol can lead to the formation of aggregates depicted in Figure 2D.
Encapsulation of different cargo 
Figure 3 shows the encapsulation of different cargo into the vesicle lumen of vesicles assembled from F20R20-mEGFP via the BuOH extrusion method. For the encapsulation of the positively charged dye Atto Rho13 in Figure 3A, the dye was mixed with the aqueous ELP solution before addition of (15% v/v) BuOH and syringe extrusion of the mixture. The confocal microscopy images show the vesicles formed from F20R20-mEGFP in the green channel, the red dye AttoRho13 in the red channel and the resulting merged channel shows the successful encapsulation inside the vesicle lumen.
The polysaccharide Dextran Red 3000 was successfully encapsulated using the BuOH extrusion method as described above. Images recorded in green channel depict the vesicles formed from F20R20-mEGFP while red channel shows the polysaccharide cargo. Merged green and red image in Figure 3B confirm the successful Dextran Red 3000 encapsulation in to the vesicle lumen.
Membrane component compatibility and phase separation of mixed BuOH vesicles before/after extrusion 
Figure 4 shows the phase separation and fusion behavior of ELP amphiphiles upon mixing of single PMBC building blocks versus assembled PMBC populations. Mixing amphiphilic ELP building blocks (F20R20-mEGFP and F20R20-mCherry) prior to PMBC assembly leads to homogenously distributed molecules within the assembled PMBC membrane. The homogenous distribution of the fluorophores and associated ELP amphiphiles is evident upon merging the red and green channel of the respective fluorescence images. By mixing vesicle populations assembled from either F20R20-mEGFP or F20R20-mCherry clearly visible membrane patches of red or green fluorescence are evident&#160;immediately after mixing. This indicates that PMBC fusion events of differently labeled PMBCs occur and that these fusing membranes and their constituents stay phase separated for at least 20 min. A similar phase behavior is known from lipid rafts, within lipid membranes23.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60935/60935fig1b.jpg" /> 
Figure 1: Illustration of the THF swelling method and the BuOH extrusion method for the guided self-assembly of amphiphilic ELPs into supramolecular structures such as vesicles or fibers. Schematic workflow and representative epifluorescence images of (A) the THF swelling method with BDP-R20F20 and BDP-R40F20 resulting in different supramolecular structures depending on temperature and pH and (B) the BuOH extrusion method exclusively yielding vesicles from BDP-R40F20&#160;(scale bar 2 &#181;m). This figure has been modified from Schreiber et al. 20198. <a href="https://www.jove.com/files/ftp_upload/60935/60935fig1blarge.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/60935/60935fig2b.jpg" /> 
Figure 2: By applying the THF swelling method BDP-R40F20 self-assembles into different supramolecular structures. The environmental conditions applied during the assembly protocol (e.g. temperature or pH) determine the predominate supramolecular structure formed. Representative supramolecular structures at the respective conditions during the assembly were monitored via epifluorescence microscopy and range from (A) coacervates and (B) fibrils to (C) stable vesicles. (D) Failure in the assembly of defined structures during the THF swelling protocol leads to the formation of unspecific aggregates (scale bar 2 &#181;m). This figure has been modified from8. <a href="https://www.jove.com/files/ftp_upload/60935/60935fig2blarge.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/60935/60935fig3b.jpg" /> 
Figure 3: Different cargos can be encapsulated within ELP vesicles using the BuOH extrusion method. (A) shows representative confocal images of F20R20-mEGFP vesicles with encapsulated positively charged dye AttoRho13 and (B) the encapsulation of the polysaccharide dextran red (scale bar 5 &#181;m). <a href="https://www.jove.com/files/ftp_upload/60935/60935fig3blarge.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/60935/60935fig4b.jpg" /> 
Figure 4: Membrane component compatibility and fusion behavior of vesicle membranes assembled from F20R20 via BuOH extrusion method. (A) Mixing of fluorescent F20R20-mEGFP and F20R20-mCherry protein solution prior to syringe-extrusion leads to PMBC membranes with homogenously distributed amphiphilic proteins visible in green channel (left image), red channel (middle image), and merged channel (right image). (B) PMBCs assembled from either F20R20-mEGFP or F20R20-mCherry and mixed subsequently via syringe extrusion lead to separated ELP amphiphile patches within the PMBC membranes. The separated ELP amphiphiles within the membrane are visible after PMBC fusion for at least 20 min in green channel (left image), red channel (middle image), and the merged channel (right image). Scale bars correspond to 5 &#181;m. This figure has been modified from Schreiber et al. 20198. <a href="https://www.jove.com/files/ftp_upload/60935/60935fig4blarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Directed Assembly of Elastin-like Proteins into defined Supramolecular Structures and Cargo Encapsulation In Vitro at JoVE.com

Directed Assembly of Elastin-like Proteins into defined Supramolecular Structures and Cargo Encapsulation In Vitro

At the interface of organic and aqueous solvents, tailored amphiphilic elastin-like proteins assemble into complex supramolecular structures such as vesicles, fibers and coacervates triggered by environmental parameters. The described assembly protocols yield Protein Membrane-Based Compartments (PMBCs) with tunable properties, enabling the encapsulation of various cargo.

Tailored proteinaceous building blocks are versatile candidates for the assembly of supramolecular structures such as minimal cells, drug delivery ...

directed-assembly-elastin-like-proteins-into-defined-supramolecular

Nanyang Technological University

Research

JoVE Journal

Chemistry

5.8K Views.  University of Freiburg. De efficiënte assemblage van supramoleculaire structuren voor minimaal celontwerp en medicijnlevering blijft een uitdaging. Hier demonstreren we efficiënte protocollen die supramoleculaire structuren opleveren op basis van amphifiele elastine-achtige eiwitten. De gepresenteerde assemblageprotocollen genereren veelzijdige supramoleculaire structuren met aanpasbare fysisch-chemische eigenschappen, zoals vesicles, vezels en coacervates.Eiwitmembraan-gebaseerde compartimenten tonen fases...

Video: Directed Assembly of Elastin-like Proteins into defined Supramolecular Structures and Cargo Encapsulation In Vitro

Formation of Ordered Biomolecular Structures by the Self-assembly of Short Peptides

In nature, complex functional structures are formed by the self-assembly of biomolecules under mild conditions. Understanding the forces that control self-assembly and mimicking this process in vitro will bring about major advances in the areas of materials science and nanotechnology. Among the available biological building blocks, peptides have several advantages as they present substantial diversity, their synthesis in large scale is straightforward, and they can easily be modified with biological and chemical entities1,2. Several classes of designed peptides such as cyclic peptides, amphiphile peptides and peptide-conjugates self-assemble into ordered structures in solution. Homoaromatic dipeptides, are a class of short self-assembled peptides that contain all the molecular information needed to form ordered structures such as nanotubes, spheres and fibrils3-8. A large variety of these peptides is commercially available.
This paper presents a procedure that leads to the formation of ordered structures by the self-assembly of homoaromatic peptides. The protocol requires only commercial reagents and basic laboratory equipment. In addition, the paper describes some of the methods available for the characterization of peptide-based assemblies. These methods include electron and atomic force microscopy and Fourier-Transform Infrared Spectroscopy (FT-IR). Moreover, the manuscript demonstrates the blending of peptides (coassembly) and the formation of a &#34;beads on a string&#34;-like structure by this process.9 The protocols presented here can be adapted to other classes of peptides or biological building blocks and can potentially lead to the discovery of new peptide-based structures and to better control of their assembly.

This paper describes the formation of highly ordered peptide-based structures by the spontaneous process of self-assembly. The method utilizes commercially available peptides and common lab equipment. This technique can be applied to a large variety of peptides and may lead to the discovery of new peptide-based assemblies.

In nature, complex functional structures are formed by the self-assembly of biomolecules under mild conditions. Understanding the forces that control ...

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Biochemistry

In Vitro Polymerization of F-actin on Early Endosomes

Many early endosome functions, particularly cargo protein sorting and membrane deformation, depend on patches of short F-actin filaments nucleated onto the endosomal membrane. We have established a microscopy-based in vitro assay that reconstitutes the nucleation and polymerization of F-actin on early endosomal membranes in test tubes, thus rendering this complex series of reactions amenable to genetic and biochemical manipulations. Endosomal fractions are prepared by floatation in sucrose gradients from cells expressing the early endosomal protein GFP-RAB5. Cytosolic fractions are prepared from separate batches of cells. Both endosomal and cytosolic fractions can be stored frozen in liquid nitrogen, if needed. In the assay, the endosomal and cytosolic fractions are mixed, and the mixture is incubated at 37 &#176;C under appropriate conditions (e.g., ionic strength, reducing environment). At the desired time, the reaction mixture is fixed, and the F-actin is revealed with phalloidin. Actin nucleation and polymerization are then analyzed by fluorescence microscopy. Here, we report that this assay can be used to investigate the role of factors that are involved either in actin nucleation on the membrane, or in the subsequent elongation, branching, or crosslinking of F-actin filaments.

In Vitro Polymerization of F-actin on Early Endosomes

Early endosome functions depend on F-actin polymerization. Here, we describe a microscopy-based in vitro assay that reconstitutes the nucleation and polymerization of F-actin on early endosomal membranes in test tubes, thus rendering this complex series of reactions amenable to biochemical and genetic manipulations.

Many early endosome functions, particularly cargo protein sorting and membrane deformation, depend on patches of short F-actin filaments nucleated onto ...

ECM Protein Nanofibers and Nanostructures Engineered Using Surface-initiated Assembly

The extracellular matrix (ECM) in tissues is synthesized and assembled by cells to form a 3D fibrillar, protein network with tightly regulated fiber diameter, composition and organization. In addition to providing structural support, the physical and chemical properties of the ECM play an important role in multiple cellular processes including adhesion, differentiation, and apoptosis. In vivo, the ECM is assembled by exposing cryptic self-assembly (fibrillogenesis) sites within proteins. This process varies for different proteins, but fibronectin (FN) fibrillogenesis is well-characterized and serves as a model system for cell-mediated ECM assembly. Specifically, cells use integrin receptors on the cell membrane to bind FN dimers and actomyosin-generated contractile forces to unfold and expose binding sites for assembly into insoluble fibers. This receptor-mediated process enables cells to assemble and organize the ECM from the cellular to tissue scales. Here, we present a method termed surface-initiated assembly (SIA), which recapitulates cell-mediated matrix assembly using protein-surface interactions to unfold ECM proteins and assemble them into insoluble fibers. First, ECM proteins are adsorbed onto a hydrophobic polydimethylsiloxane (PDMS) surface where they partially denature (unfold) and expose cryptic binding domains. The unfolded proteins are then transferred in well-defined micro- and nanopatterns through microcontact printing onto a thermally responsive poly(N-isopropylacrylamide) (PIPAAm) surface. Thermally-triggered dissolution of the PIPAAm leads to final assembly and release of insoluble ECM protein nanofibers and nanostructures with well-defined geometries. Complex architectures are possible by engineering defined patterns on the PDMS stamps used for microcontact printing. In addition to FN, the SIA process can be used with laminin, fibrinogen and collagens type I and IV to create multi-component ECM nanostructures. Thus, SIA can be used to engineer ECM protein-based materials with precise control over the protein composition, fiber geometry and scaffold architecture in order to recapitulate the structure and composition of the ECM in vivo.

A method to obtain nanofibers and complex nanostructures from single or multiple extracellular matrix proteins is described. This method uses protein-surface interactions to create free-standing protein-based materials with tunable composition and architecture for use in a variety of tissue engineering and biotechnology applications.

The extracellular matrix (ECM) in tissues is synthesized and assembled by cells to form a 3D fibrillar, protein network with tightly regulated fiber ...

Bioengineering

OaAEP1-Mediated Enzymatic Synthesis and Immobilization of Polymerized Protein for Single-Molecule Force Spectroscopy

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled protein polymerization process is always a challenge. Here, we have developed an enzymatic methodology for constructing polymerized protein step by step in a rationally-controlled sequence. In this method, the C-terminus of a protein monomer is NGL for protein conjugation using OaAEP1 (Oldenlandia affinis asparaginyl endopeptidases) 1) while the N-terminus was a cleavable TEV (tobacco etch virus) cleavage site plus an L (ENLYFQ/GL) for temporary N-terminal protecting. Consequently, OaAEP1 was able to add only one protein monomer at a time, and then the TEV protease cleaved the N-terminus between Q and G to expose the NH2-Gly-Leu. Then the unit is ready for next OaAEP1 ligation. The engineered polyprotein is examined by unfolding individual protein domain using atomic force microscopy-based single-molecule force spectroscopy (AFM-SMFS). Therefore, this study provides a useful strategy for polyprotein engineering and immobilization.

Here, we present a protocol to conjugate protein monomer by enzymes forming protein polymer with a controlled sequence and immobilize it on the surface for single-molecule force spectroscopy studies.

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled ...

Design and Construction of Artificial Extracellular Matrix (aECM) Proteins from Escherichia coli for Skin Tissue Engineering

Recombinant technology is a versatile platform to create novel artificial proteins with tunable properties. For the last decade, many artificial proteins that have incorporated functional domains derived from nature (or created de novo) have been reported. In particular, artificial extracellular matrix (aECM) proteins have been developed; these aECM proteins consist of biological domains taken from fibronectin, laminins and collagens and are combined with structural domains including elastin-like repeats, silk and collagen repeats. To date, aECM proteins have been widely investigated for applications in tissue engineering and wound repair. Recently, Tjin and coworkers developed integrin-specific aECM proteins designed for promoting human skin keratinocyte attachment and propagation. In their work, the aECM proteins incorporate cell binding domains taken from fibronectin, laminin-5 and collagen IV, as well as flanking elastin-like repeats. They demonstrated that the aECM proteins developed in their work were promising candidates for use as substrates in artificial skin. Here, we outline the design and construction of such aECM proteins as well as their purification process using the thermo-responsive characteristics of elastin.

Design and Construction of Artificial Extracellular Matrix aECM Proteins from Escherichia coli for Skin Tissue Engineering

Recombinant technologies have enabled material designers to create novel artificial proteins with customized functionalities for tissue engineering applications. For example, artificial extracellular matrix proteins can be designed to incorporate structural and biological domains derived from native ECMs. Here, we describe the construction and purification of aECM proteins containing elastin-like repeats.

Recombinant technology is a versatile platform to create novel artificial proteins with tunable properties. For the last decade, many artificial proteins ...

Production of Elastin-like Protein Hydrogels for Encapsulation and Immunostaining of Cells in 3D

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue culture systems lack a three-dimensional (3D) matrix, resulting in a significant disconnect between results collected in vitro and in vivo. To address this limitation, researchers have engineered 3D hydrogel tissue culture platforms that can mimic the biochemical and biophysical properties of the in vivo cell microenvironment. This research has motivated the need to develop material platforms that support 3D cell encapsulation and downstream biochemical assays. Recombinant protein engineering offers a unique toolset for 3D hydrogel material design and development by allowing for the specific control of protein sequence and therefore, by extension, the potential mechanical and biochemical properties of the resultant matrix. Here, we present a protocol for the expression of recombinantly-derived elastin-like protein (ELP), which can be used to form hydrogels with independently tunable mechanical properties and cell-adhesive ligand concentration. We further present a methodology for cell encapsulation within ELP hydrogels and subsequent immunofluorescent staining of embedded cells for downstream analysis and quantification.

Recombinant protein-engineered hydrogels are advantageous for 3D cell culture as they allow for complete tunability of the polymer backbone and therefore, the cell microenvironment. Here, we describe the process of recombinant elastin-like protein purification and its application in 3D hydrogel cell encapsulation.

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue ...

In Vesiculo Synthesis of Peptide Membrane Precursors for Autonomous Vesicle Growth

Compartmentalization of biochemical reactions is a central aspect of synthetic cells. For this purpose, peptide-based reaction compartments serve as an attractive alternative to liposomes or fatty acid-based vesicles. Externally or within the vesicles, peptides can be easily expressed and simplify the synthesis of membrane precursors. Provided here is a protocol for the creation of vesicles with diameters of ~200 nm based on the amphiphilic elastin-like polypeptides (ELP) utilizing dehydration-rehydration from glass beads. Also presented are protocols for bacterial ELP expression and purification via inverse temperature cycling, as well as their covalent functionalization with fluorescent dyes. Furthermore, this report describes a protocol to enable the transcription of RNA aptamer dBroccoli inside ELP vesicles as a less complex example for a biochemical reaction. Finally, a protocol is provided, which allows in vesiculo expression of fluorescent proteins and the membrane peptide, whereas synthesis of the latter results in vesicle growth.

Presented here are protocols for the creation of peptide-based small unilamellar vesicles capable of growth. To facilitate in vesiculo production of the membrane peptide, these vesicles are equipped with a transcription-translation system and the peptide-encoding plasmid.

Compartmentalization of biochemical reactions is a central aspect of synthetic cells. For this purpose, peptide-based reaction compartments serve as an ...

Rapid Encapsulation of Reconstituted Cytoskeleton Inside Giant Unilamellar Vesicles

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular processes in vitro. In recent years, encapsulation within GUVs has proven to be a helpful approach for reconstitution experiments in cell biology and related fields. It better mimics confinement conditions inside living cells, as opposed to conventional biochemical reconstitution. Methods for encapsulation inside GUVs are often not easy to implement, and success rates can differ significantly from lab to lab. One technique that has proven to be successful for encapsulating more complex protein systems is called continuous droplet interface crossing encapsulation (cDICE). Here, a cDICE-based method is presented for rapidly encapsulating cytoskeletal proteins in GUVs with high encapsulation efficiency. In this method, first, lipid-monolayer droplets are generated by emulsifying a protein solution of interest in a lipid/oil mixture. After being added into a rotating 3D-printed chamber, these lipid-monolayered droplets then pass through a second lipid monolayer at a water/oil interface inside the chamber to form GUVs that contain the protein system. This method simplifies the overall procedure of encapsulation within GUVs and speeds up the process, and thus allows us to confine and observe the dynamic evolution of network assembly inside lipid bilayer vesicles. This platform is handy for studying the mechanics of cytoskeleton-membrane interactions in confinement.

This article introduces a simple method for expeditious production of giant unilamellar vesicles with encapsulated cytoskeletal proteins. The method proves to be useful for bottom-up reconstitution of cytoskeletal structures in confinement and cytoskeleton-membrane interactions.

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular ...

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell deformation, division, migration, and adhesion. However, studying the dynamics and structure of the actin network in vivo is complicated by the biochemical and genetic regulation within live cells. To build a minimal model devoid of intracellular biochemical regulation, actin is encapsulated inside giant unilamellar vesicles (GUVs, also called liposomes). The biomimetic liposomes are cell-sized and facilitate a quantitative insight into the mechanical and dynamical properties of the cytoskeleton network, opening a viable route for bottom-up synthetic biology. To generate liposomes for encapsulation, the inverted emulsion method (also referred to as the emulsion transfer method) is utilized, which is one of the most successful techniques for encapsulating complex solutions into liposomes to prepare various cell-mimicking systems. With this method, a mixture of proteins of interest is added to the inner buffer, which is later emulsified in a phospholipid-containing mineral oil solution to form monolayer lipid droplets. The desired liposomes are generated from monolayer lipid droplets crossing a lipid/oil-water interface. This method enables the encapsulation of concentrated actin polymers into the liposomes with desired lipid components, paving the way for in vitro reconstitution of a biomimicking cytoskeleton network.

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

In this manuscript, we demonstrate the experimental techniques to encapsulate the F-actin cytoskeleton into giant unilamellar lipid vesicles (also called liposomes), and the method to form a cortex-biomimicking F-actin layer at the inner leaflet of the liposome membrane.

Directed Assembly of Elastin-like Proteins into defined Supramolecular Structures and Cargo Encapsulation In Vitro

Samenvatting

Meer video's verkennen