This project was funded by the University of Nebraska Medical Center (Start-up funds, MC-S); NIH-COBRE, 5P20GM103480 (T. Bronich) and the American Chemical Society, PRF# 57434-DNI7(MC-S).

1. General Protocol for Synthesis of PPAs
<ol>
	<li>Calculate the scale of synthesis (usually mmol). This scale is based on the mass of the target PPA amount. Keep in mind that the reaction efficiency of SPPS gradually decreases with an increase of amino acid sequence. Therefore, the exact reaction efficiency is difficult to calculate.</li>
	<li>Calculate the weight of the resin to be used according to the loading of the resin. The loading is found on the container or the analysis protocol of the resin and expressed in mmol/g. The following formula can be used for calculating the weight of the resin:</li>
	<li>Carefully weigh out 2-chlorotrityl chloride resin (in our case the loading is ~0.85 mmol)</li>
	<li>Place the resin in a fritted synthesis vessel with the following specifications: Capacity &#8211; 50 mL, Fritted Disc &#8211; 25 mm, Porosity &#8211; Medium.</li>
	<li>Add 15 mL of dichloromethane (DCM) to the resin.</li>
	<li>Affix the synthesis vessel to a variable speed mechanical shaker (flask shaker/stirrer), and turn the vessels to a 45&#176; angle (or horizontally) to maximize agitation.</li>
	<li>Allow the resin beads to swell for 15 min. Swelling enhances the yield of reaction because it facilitates molecular diffusion and accessibility to the active site, enhancing coupling efficiency.</li>
	<li>Add 8 equivalents (2 mmol for a 0.25 mmol scale) of the desired polyamine (Spermine, Spermidine, Diethyelenetriamine, 1,3-diaminopropane, etc.) to the resin and allow it to react for 5 h. 
	Note: A lesser reaction period would reduce yield.</li>
	<li>Do a Kaiser test (see Table of Materials) to confirm successful coupling of the polyamine to the resin. Successful coupling of the primary amines yields a violet/blue color, which corresponds to the free amine.</li>
	<li>Protect the primary amine group using 4 equivalents of 1-(4,4-dimethyl-2,6-dioxocyclohex-1-ylidene)ethyl (Dde), dissolved in anhydrous methanol (15 mL), by shaking the reaction mixture overnight11. 
	Note: A lesser reaction time reduces yield.</li>
	<li>Perform a Kaiser test. An absence of blue color from the resin bead confirms successful protection. In case of unsuccessful protection of the primary amine, repeat the previous step.</li>
	<li>Drain the DCM and wash the resins twice with a mixture of DCM and DMF (2:1, 15 mL).</li>
	<li>Dissolve 20 equivalents (5 mmol) of di-tert butyl di-carbonate (Boc) in DCM (20 mL) and allow the reaction to proceed for 3 h.</li>
	<li>Do a chloranil test (see Table of Materials) to confirm complete protection of the secondary amine. A positive test (protection) yields a colorless or light yellow color. Free secondary amines give a dark green/blue color, while primary amines give a red color.</li>
	<li>Drain the solvent mixture and wash twice with a mixture of DCM and DMF (2:1, 15 mL).</li>
	<li>Add a 2% solution of hydrazine in DMF (10 mL) and shake for 1 h.</li>
	<li>Do a Kaiser test to confirm successful deprotection of the primary amine.</li>
	<li>Add the desired amino acids in the reverse sequence in which you would write/draw them. Peptides are drawn from the N termini to the C termini but are synthesized in the opposite direction, C to N. For example, for a PPA requiring the following peptide core -GLFD-, add the Aspartic acid (D), followed by Phenylalanine (F), Leucine (L), and finally Glycine (G). 
	NOTE: For most amino acids, the following coupling cocktail is appropriate for attachment to the polyamine loaded resins.
	<ol>
		<li>Mix 4 equivalents (1 mmol) of the Fmoc-protected amino acid, 3.95 equivalents of 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexa&#64258;uorophosphate (HBTU), and 15 equivalents of diisopropylethyl amine (DIPEA).</li>
		<li>Dissolve them in a mixture of DCM and DMF (1:1, 15 mL) and sonicate the cocktail for 2 min (until complete dissolution).</li>
		<li>Wait an additional 3&#8211;5 min to ensure activation of the carboxylic acid on the amino acid.</li>
		<li>Add the reaction mixture to the vessel. Carry out the reaction for 2-4 h at ambient temperature. 
		NOTE: We have found the following as efficient alternatives to HBTU (usually requiring lower amounts of the amino acids and the coupling agent): COMU, PyBOP, HATU, DIC/Oxima.</li>
		<li>Do a Kaiser test to confirm successful coupling.</li>
		<li>De-protect the Fmoc-group from the amino acid by adding a 20% solution of 4-methyl piperidine (10 mL) in DMF. Do it twice, each time shaking the reaction mixture for 15 min. 
		NOTE: An efficient alternative for Fmoc removal is piperazine/DBU12.
		<ol>
			<li>Perform a DCM (15 mL) wash between additions.</li>
		</ol>
		</li>
		<li>Do a Kaiser test to confirm successful de-protection of the amino acid.</li>
		<li>Wash the resin thoroughly to remove all traces of 4-methyl. Wash the resin twice with DMF (10 mL), each wash lasting for 5 min, and finally with DCM (10 mL) for 10 min.</li>
		<li>Add all the remaining amino acids by successively repeating the coupling and de-protection steps.</li>
	</ol>
	</li>
	<li>Finally, after coupling all the required amino acids, conjugate the hydrophobic tail to the last amino acid of the polyamine-peptide construct:
	<ol>
		<li>Add 10 equivalents of the desired carboxylic acid functionality to 9.5 equivalents of HBTU and 12 equivalents of DIPEA dissolved in DCM and DMF mixture (1:1, 20 mL). 
		Note: Bear in mind than some tails may need a different proportion of solvent (usually a larger% of DCM) or the addition or another solvent such as N-methylpyrrolidone (NMP).</li>
		<li>Sonicate the cocktail for 5&#8211;10 min until dissolution (we have seen it takes longer for the tail to completely dissolve) and add to the vessel. 
		Note: For tails containing multiple reactive sites, they will need to be protected before coupling them.</li>
		<li>Carry out this reaction (coupling the hydrophobic tail) for at least 5 h, although it is advisable to carry it out overnight for the highest yield.</li>
	</ol>
	</li>
</ol>
2. PPA Cleavage from Solid Support
The purpose of this step is the cleavage of the PPA from the resin, and to remove the Boc protecting groups from the amino acids and polyamine residues.
<ol>
	<li>Wash the resin with DMF (8 mL for 2 min) and twice with DCM (8 mL, each time for 5 min). Before each addition, drain the solvent from the vessel. Once the last wash is performed, dry the resin under vacuum for 15 min.</li>
	<li>Prepare the cleavage solution using the following mixture ratio: 28:1:1 trifluoroacetic acid (TFA): H2O : Triisopropyl silane (TIPS). To make 15 mL of the cleavage cocktail add 14 mL of TFA, to 0.5 mL of H2O and 0.5 mL of TIPS.</li>
	<li>Add this cleavage solution to the resin and shake for 2&#8211;4 h at room temperature.</li>
	<li>Collect the solution in a 25 or 50 mL round bottom flask.</li>
	<li>Concentrate the TFA in vacuo to 1&#8211;2 mL using a rotary evaporator at reduced pressure while heating the mixture at 40 &#176;C (do not surpass 55 &#176;C to avoid decomposition of the PPA).</li>
	<li>After evaporation, add the obtained TFA solution (dropwise) to a round bottom flask containing anhydrous cold ether (15 mL). This will immediately precipitate the PPA.</li>
	<li>In addition, add anhydrous cold ether (5 mL) to the original flask where the TFA was concentrated.</li>
	<li>Sonicate this flask to recover additional solids and combine with the ether solution from step 2.6. 
	Note: The transferring pipette can be washed with a small volume of DCM. Avoid introducing DMF during the process because it tends to form a gel-like substance.</li>
	<li>Place the flask (covered) inside the refrigerator and let it stand overnight to maximize precipitation.</li>
	<li>Collect the precipitated material by vacuum filtration using a sintered disc filter funnel. The ideal pore sizes are fine or medium.</li>
	<li>Wash the precipitate twice with cold ether (5&#8211;10 mL) to remove any residual organics.</li>
</ol>
3. Identification of the Crude Product Using the MALDI Dried-drop Method
<ol>
	<li>Matrix preparation for analysis in positive mode:
	<ol>
		<li>To start molecular weight analysis using Matrix Assisted Laser Desorption/Ionization (MALDI) mass spectrometry, add 10 to 20 mg of &#945;-cyano-4-hydroxycinnamic acid to a microcentrifuge tube.</li>
		<li>Add a solution of water/acetonitrile (1:1, 1.0 mL) with 0.1% Trifluoroacetic acid (TFA). Mix thoroughly by vortexing. 
		Note: This matrix should be used for positively charged peptides. MALDI negative mode is similar but requires a matrix containing 9-aminoacridine with 0.1% ammonia as solvent additive.</li>
		<li>Add 1&#8211;2 &#181;L of the sample to the stainless steel target plate for MALDI and dry the sample in air. Add 1&#8211;2 &#181;L of the matrix and dry it again. 
		Note: The plates have circular markings gridded along the target plate to help locate the particular spot.</li>
	</ol>
	</li>
	<li>Analyze the samples in MALDI instrument to verify their identity. Electrospray ionization (ESI) is a valid alternative to confirm the identity of the PPAs.</li>
</ol>
4. Purification of PPAs Using Preparative Reverse-phase High-performance Liquid Chromatography (HPLC) Purification
<ol>
	<li>Dissolve the PPA precipitate in a minimum amount of acetonitrile and water.
	<ol>
		<li>As a rule of thumb, dissolve 100 mg of dry crude PPA in less than 5 mL of acetonitrile and water. More hydrophobic PPAs might require a greater percentage of acetonitrile to dissolve and might also require a larger volume of solvent in general, depending on the net charge of the PPAs (Table 1).</li>
		<li>If complete dissolution does not take place, add 1% TFA (ACN or H2O). It is also possible to add trace amounts of other compatible solvents including dimethylsulfoxide, methanol, or isopropanol (limit their content to 5%).</li>
	</ol>
	</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Solvent</td>
			<td colspan="2">Positively Charged PPAs</td>
			<td colspan="2">Negatively Charged PPAs</td>
		</tr>
		<tr>
			<td></td>
			<td colspan="2">0.1% TFA in Water</td>
			<td colspan="2">0.1% NH3 in Water</td>
		</tr>
		<tr>
			<td></td>
			<td colspan="2">0.1% TFA in ACN</td>
			<td colspan="2">0.1% NH3 in ACN</td>
		</tr>
	</tbody>
</table>
Table 1: Solvent Systems. Proposed solvent system for positively and negatively charged PPAs.
<ol start="2">
	<li>Sonicate the PPA using a horn sonicator for 20 min at 10 s pulses with Amp1 of 48% or for 2-3 h in an ultrasonic bath.</li>
	<li>Then, filter using a 0.45 &#956;m syringe filter followed by filtration using a 0.20 &#956;m Polytetrafluoroethylene (PTFE) syringe filter. The PPA solution should be clear and free of all particulate materials.
	<ol>
		<li>If filtration is too difficult, sonicate for an extended period of time. It is strongly advised that the PPA solution is purified immediately after syringe filtration, as prolonged storage might cause it to aggregate or gelate, which will necessitate re-sonication and, perhaps, re-filtration.</li>
	</ol>
	</li>
	<li>During and after purification, use either HPLC grade solvents or ones that are filtered through a 0.25 &#956;m syringe filter/membrane. 
	NOTE: The most common solvent conditions to purify PPAs consist in a gradient of H2O and ACN.</li>
	<li>Use a standard reverse-phase HPLC instrument for this protocol. It should include an elution gradient programmer, a binary solvent delivery system, a UV detector capable of detecting at 220 and 254 nm, and a programmable fraction collector. The maximum flow rate should be 50 mL/min.</li>
	<li>Use a C-18 reversed phase column for separation. Columns of the following dimensions can be used depending on the mass of solid being purified and the net charge of the PPA (Table 2).</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>PPA Charge</td>
			<td>Particle Size</td>
			<td>Column Size</td>
			<td>Mass of crude PPA</td>
		</tr>
		<tr>
			<td>+ ve Charged</td>
			<td>5 &#956;m</td>
			<td>150 x 30 mm</td>
			<td>170 mg</td>
		</tr>
		<tr>
			<td>-&#160; ve Charged</td>
			<td>5 &#956;m</td>
			<td>150 x 30 mm</td>
			<td>170 mg</td>
		</tr>
		<tr>
			<td>+ ve Charged</td>
			<td>5 &#956;m</td>
			<td>150 x 21.2 mm</td>
			<td>90 mg</td>
		</tr>
		<tr>
			<td>-&#160; ve Charged</td>
			<td>5 &#956;m</td>
			<td>150 x 21.2 mm</td>
			<td>90 mg</td>
		</tr>
	</tbody>
</table>
Table 2: Suggested columns: Column dimensions, particle sizes and the maximum load capacity per injection for C18 reverse phase HPLC columns
<ol start="7">
	<li>Inject PPA with a volume determined by both the capacity of the column and by the volume of the injection loop of the HPLC. Run the HPLC gradient method according to the settings seen in Table 3 (elution time of 40 min).</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Time</td>
			<td>Solvent A (Acetonitrile)</td>
			<td>Solvent B (Water)</td>
			<td>Flow Rate (mL/min)</td>
		</tr>
		<tr>
			<td>0</td>
			<td>5%</td>
			<td>95%</td>
			<td rowspan="5">Flow rate depends on the column packing and its size.</td>
		</tr>
		<tr>
			<td>2</td>
			<td>5%</td>
			<td>95%</td>
		</tr>
		<tr>
			<td>35</td>
			<td>95%</td>
			<td>5%</td>
		</tr>
		<tr>
			<td>38</td>
			<td>100%</td>
			<td>0%</td>
		</tr>
		<tr>
			<td>40</td>
			<td>5%</td>
			<td>95%</td>
		</tr>
	</tbody>
</table>
Table 3: Suggested gradient: Suggested reverse phase gradient showing the relative composition of water vs acetonitrile over a period of time. The flow rate will depend on the column specifications.
<ol start="8">
	<li>Analyze the fractions collected on the various test tubes using MALDI and determine which tube (or tubes) contains the PPA. This is assessed by studying the molecular weight found in each tube.
	<ol>
		<li>Check purity of the fractions on an analytical HPLC. Use an HPLC-gradient system equipped with a UV detector set at 220 nm.</li>
		<li>If there are impurities, do an additional cycle of purification by HPLC. The above gradient used for preparative HPLC can be scaled down to use with the analytical HPLC using the column converter online software. Each fraction much be &#8805;95% pure for physical characterization or biological evaluation.</li>
	</ol>
	</li>
	<li>Collect the fractions in a large round-bottomed flask or evaporation flask and remove all the acetonitrile and most of the water. The final PPA solution should be no more than 10 mL.</li>
	<li>Transfer this concentrate to a 50 mL centrifuge tube. Be advised that PPAs are detergent-like substances; thus, bubbles will be generated during evaporation of the solvent. We have found that addition of ethyl alcohol diminishes bubble formation.</li>
	<li>Vacuum concentrate the concentrate. During the vacuum evaporation, a large amount of the PPA might adhere to the walls of the container. Recover this by repeatedly rinsing the flask wall with HPLC water. The combined volume of the concentrated PPA and the rinsate should be no more than 30 mL.</li>
	<li>Place the aqueous solution inside of a 50 mL tube.</li>
	<li>Flash freeze this PPA solution in liquid nitrogen: 
	NOTE: Flash freezing results in smaller ice-crystals which will sublime faster in the lyophilizer. Moreover, slower freezing may allow the formation of self-assembled supramolecular structures. Another alternative (but less desirable) is to gradually freeze in a -80 &#176;C freezer or to freeze using a dry ice + acetone mixture
	<ol>
		<li>Before placing the centrifuge tube in the lyophilizer, perforate or loosen the cap of the tube to allow moisture to escape the sample and get collected in the refrigeration unit. Set the lyophilization unit set to -54 &#176;C and 0.016 mbar. 
		NOTE: Another option is to remove the cap and place a wipe (with a rubber band) on top of the opening. Also, we have found that freezing samples at an angle or breaking the ice into small portions allows for a faster and better lyophilization due to an increase in surface area.</li>
		<li>If the solid starts to melt, this may indicate traces of an organic solvent (usually acetonitrile) are present. Repeat the freezing step and assess the condition of lyophilization.</li>
		<li>If the melting persists, there are two potential solutions:
		<ol>
			<li>Split the sample in two portions (to be placed in tubes), dilute with water, and freeze again. Do not use this solution often because large amounts of organic solvents may damage the equipment.</li>
			<li>Alternatively, place the sample in a flask and further evaporate the organic solvent under vacuum. Lyophilizing a sample that is in the liquid form usually leads to bumping and loss of the PPA compound.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
5. Storage of PPAs
<ol>
	<li>Store the PPAs in a -20 &#176;C with the caps tightened and sealed with Parafilm on a tube with a proper label.</li>
	<li>Before use, bring the PPA back to ambient temperature in a vacuum chamber to avoid condensation of water vapor on the PPAs.</li>
</ol>

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The authors have no conflict of interest to declare

The protocols described here can be used to synthesize PPAs as wells as PAs and related peptide-based molecules (such as hybrid PA-peptoids). Although the synthesis of peptides using SPPS is a straightforward procedure, the synthesis of peptides containing biological homing molecules can be particularly challenging. Polyamines like spermine, spermidine, diethyelenetriamine, etc., can function as homing molecules for targeting cancer cells13. The PPAs can self-assemble into nanostructures ...

Self-assembling peptide amphiphiles (PAs) are a class of biomaterials typically comprised of the following segments: (a) hydrophilic head, (b) linker region, and (c) hydrophobic tail. Most PAs described in the literature possess a hydrophilic head comprised of charged or polar amino acid residues1,2,3,4. PAs have found a wide range of applications in biomedicine including drug delivery, disease diagnostics, regenerative medicine, etc.5. Based on their amino acid sequence, PAs can form a wide variety of nanostructures including spherical micelles, and nano-filaments. We have recently reported a class of hybrid polyamine-based peptide amphiphiles, termed PPAs6. The morphologies, self-assembling kinetics, and metabolic degradation, of these biomaterials, were found to be related to their solubilizing head group. Moreover, the PPA nanostructures did not show toxicity towards mammalian cells (MiaPaCa2 and HeLa cell lines) at the tested concentrations. PPA-based nanocarriers are attractive drug-delivery vehicles because: (1) polyamine uptake and metabolism has been shown to be increased in cancerous cells, (2) cationic nanostructures can achieve endosomal escape7,8, which leads to higher circulation and residence within a cell, and (3) they should have a distinct metabolic profile when compared with PA; for example, they will be more stable towards proteases found in the human body (although they maybe sensitive to other enzymes, such as amine oxidases)9,10. Also, PPAs have been found to have diverse morphologies, physicochemical properties, nanoparticle stiffness, and assembly kinetics depending on the length and charge of individual PPA molecule6. Herein, we describe a detailed protocol for the synthesis, identification, and purification of PPAs that can also be applied to the preparation of PAs or similar hybrid peptide molecules.
Because polyamines are not commonly commercially available in their protected forms, and because protecting the primary and secondary amines of polyamines is of utmost importance for conjugating them with amino acids and other molecules, we have outlined the synthetic steps to achieve their protection. The overall goal of this protocol is to provide a simple method for conjugating polyamines to amino acids. Polyamines lack a carboxylic group; thus, they cannot be coupled to Rink Amide or Wang resins. Instead, resins such as 2-chlorotrityl chloride are recommended for the synthetic protocol. The main challenge for PPA synthesis is the presence of primary and secondary amine functional groups. For our purposes, we protected all the secondary amines in the polyamine while keeping the primary amino group on the polyamine free to allow the coupling reaction. The reaction was done on a solid support following the principles of solid phase peptide synthesis (SPPS) to facilitate the work-up after each coupling and deprotection step. The following protocol is for both the manual and automated synthesis of PPAs (although the verification of some steps will be challenging in an automated system). The synthesis of these molecules can also be carried out on an automated synthesizer or with the aid of a microwave reactor (automated or semi-automated). The reaction scheme has been summarized in Figure 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57908/57908fig1.jpg" /> 
Figure 1: (A) A general reaction scheme for the synthesis of PPAs. (B) Representative polyamines that can be used to synthesized PPAs described here. <a href="/files/ftp_upload/57908/57908fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table>
	<tbody>
		<tr>
			<td>2-Chlorotrityl chloride resin&#160;</td>
			<td>AappTec</td>
			<td>RTZ001</td>
			<td></td>
		</tr>
		<tr>
			<td>SynthwareTM synthesis vessel&#160;</td>
			<td>Aldrich</td>
			<td>SYNP120050M</td>
			<td></td>
		</tr>
		<tr>
			<td>Dichloromethane</td>
			<td>Acros</td>
			<td>AC406920250</td>
			<td>Fisher Sci. Catalogue #</td>
		</tr>
		<tr>
			<td>Wrist Shaker</td>
			<td>Boekel Scientific</td>
			<td>401000-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Kaiser test kit</td>
			<td>Sigma-Aldrich</td>
			<td>60017</td>
			<td></td>
		</tr>
		<tr>
			<td>2-[(4,4-dimethyl-2,6-dioxocyclohex-1-ylidene)ethyl-amino]-ethanol</td>
			<td>Sigma-Aldrich</td>
			<td>CDS004772</td>
			<td></td>
		</tr>
		<tr>
			<td>Anhydrous Methanol</td>
			<td>Acros</td>
			<td>AC610981000</td>
			<td>Fisher Sci. Catalogue #</td>
		</tr>
		<tr>
			<td>Chloranil test kit</td>
			<td>TCI</td>
			<td>TCC1771-KIT</td>
			<td>VWR Catalogue #</td>
		</tr>
		<tr>
			<td>Di-tert butyl di-carbonate&#160;</td>
			<td>Acros</td>
			<td>AC194670250</td>
			<td>Fisher Sci. Catalogue #</td>
		</tr>
		<tr>
			<td>Dimethylformamide</td>
			<td>Fisher Scientific</td>
			<td>BP1160-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrazine</td>
			<td>Acros</td>
			<td>AC296815000</td>
			<td>FIsher Sci. Catalogue #</td>
		</tr>
		<tr>
			<td>(2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate)</td>
			<td>p3biosystems</td>
			<td>31001</td>
			<td></td>
		</tr>
		<tr>
			<td>4-methyl piperidine&#160;</td>
			<td>Acros</td>
			<td>AC127515000</td>
			<td>FIsher Sci. Catalogue #</td>
		</tr>
		<tr>
			<td>Trifluoroacetic Acid</td>
			<td>AappTec</td>
			<td>CXZ035</td>
			<td></td>
		</tr>
		<tr>
			<td>Triisopropyl Silane</td>
			<td>Sigma-Aldrich</td>
			<td>233781</td>
			<td></td>
		</tr>
		<tr>
			<td>Ether</td>
			<td>Fisher Scientific</td>
			<td>E138-1</td>
			<td></td>
		</tr>
		<tr>
			<td>&#945;-Cyano-4-hydroxycinnamic acid</td>
			<td>Sigma-Aldrich</td>
			<td>C8982</td>
			<td></td>
		</tr>
		<tr>
			<td>9-Aminoacridine</td>
			<td>Sigma-Aldrich</td>
			<td>92817</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand&#160;Syringe Filters: PTFE Membrane</td>
			<td>Fisher Scientific</td>
			<td>09-730-21</td>
			<td></td>
		</tr>
	</tbody>
</table>

facile protocol for the synthesis of self-assembling polyamine-based peptide amphiphiles (ppas) and related biomaterials

Polyamine-based Peptide Amphiphiles (PPAs) are a new class of self-assembling amphiphilic biomaterials-related to the peptide amphiphiles (PAs). Traditional PAs possess charged amino acids as solubilizing groups (lysine, arginine), which are directly connected to a lipid segment or can contain a linker region made of neutral amino acids. Tuning the peptide sequence of PAs can yield diverse morphologies. Similarly, PPAs possess a hydrophobic segment and neutral amino acids, but also contain polyamine molecules as water solubilizing (hydrophilic) groups. As is the case with PAs, PPAs can also self-assemble into diverse morphologies, including small rods, twisted nano-ribbons, and fused nano-sheets, when dissolved in water. However, the presence of both primary and secondary amines on a single polyamine molecule poses a significant challenge when synthesizing PPAs. In this paper, we show a simple protocol, based on literature precedents, to achieve a facile synthesis of PPAs using solid phase peptide synthesis (SPPS). This protocol can be extended to the synthesis of PAs and other similar systems. We also illustrate the steps that are needed for cleavage from the resin, identification, and purification.

After the synthesis and purification and before physicochemical or biological evaluation, it is recommended the masses of the PPAs are re-checked and the purity ascertained using analytical HPLC. For material characterization or biological evaluation, PPAs need to have a purity of &#62;95%. Figure 2 shows the HPLC trace (Top) and MALDI spectra (Bottom) confirming the presence of the product. HPLC analytical systems will integrate the area under the curve (AUC...

Watch this Scientific Journal Video about Facile Protocol for the Synthesis of Self-assembling Polyamine-based Peptide Amphiphiles PPAs and Related Biomaterials at JoVE.com

Facile Protocol for the Synthesis of Self-assembling Polyamine-based Peptide Amphiphiles PPAs and Related Biomaterials

The synthesis of polyamine-based peptide amphiphiles (PPAs) is a significant challenge due to the presence of multiple amine nitrogens, which requires judicious use of protecting groups to mask these reactive functionalities. In this paper, we describe a facile method for the preparation of these new class of self-assembling molecules.

Facile Protocol for the Synthesis of Self-assembling Polyamine-based Peptide Amphiphiles (PPAs) and Related Biomaterials

Polyamine-based Peptide Amphiphiles (PPAs) are a new class of self-assembling amphiphilic biomaterials-related to the peptide amphiphiles (PAs). ...

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8.0K Views.  University of Nebraska Medical Center. The synthesis of polyamine-based peptide amphiphiles (PPAs) is a significant challenge due to the presence of multiple amine nitrogens, which requires judicious use of protecting groups to mask these reactive functionalities. In this paper, we describe a facile method for the preparation of these new class of self-assembling molecules.

Video: Facile Protocol for the Synthesis of Self-assembling Polyamine-based Peptide Amphiphiles PPAs and Related Biomaterials

Production of Disulfide-stabilized Transmembrane Peptide Complexes for Structural Studies

Physical interactions among the lipid-embedded alpha-helical domains of membrane proteins play a crucial role in folding and assembly of membrane protein complexes and in dynamic processes such as transmembrane (TM) signaling and regulation of cell-surface protein levels. Understanding the structural features driving the association of particular sequences requires sophisticated biophysical and biochemical analyses of TM peptide complexes. However, the extreme hydrophobicity of TM domains makes them very difficult to manipulate using standard peptide chemistry techniques, and production of suitable study material often proves prohibitively challenging. Identifying conditions under which peptides can adopt stable helical conformations and form complexes spontaneously adds a further level of difficulty. Here we present a procedure for the production of homo- or hetero-dimeric TM peptide complexes from materials that are expressed in E. coli, thus allowing incorporation of stable isotope labels for nuclear magnetic resonance (NMR) or non-natural amino acids for other applications relatively inexpensively. The key innovation in this method is that TM complexes are produced and purified as covalently associated (disulfide-crosslinked) assemblies that can form stable, stoichiometric and homogeneous structures when reconstituted into detergent, lipid or other membrane-mimetic materials. We also present carefully optimized procedures for expression and purification that are equally applicable whether producing single TM domains or crosslinked complexes and provide advice for adapting these methods to new TM sequences.

Biophysical and biochemical studies of interactions among membrane-embedded protein domains face many technical challenges, the first of which is obtaining appropriate study material. This article describes a protocol for producing and purifying disulfide-stabilized transmembrane peptide complexes that are suitable for structural analysis by solution nuclear magnetic resonance (NMR) and other analytical applications.

Physical  interactions among the lipid-embedded alpha-helical domains of membrane  proteins play a crucial role in folding and assembly of membrane ...

Structure and Coordination Determination of Peptide-metal Complexes Using 1D and 2D 1H NMR

Copper (I) binding by metallochaperone transport proteins prevents copper oxidation and release of the toxic ions that may participate in harmful redox reactions. The Cu (I) complex of the peptide model of a Cu (I) binding metallochaperone protein, which includes the sequence MTCSGCSRPG (underlined is conserved), was determined in solution under inert conditions by NMR spectroscopy.
NMR is a widely accepted technique for the determination of solution structures of proteins and peptides. Due to difficulty in crystallization to provide single crystals suitable for X-ray crystallography, the NMR technique is extremely valuable, especially as it provides information on the solution state rather than the solid state. Herein we describe all steps that are required for full three-dimensional structure determinations by NMR. The protocol includes sample preparation in an NMR tube, 1D and 2D data collection and processing, peak assignment and integration, molecular mechanics calculations, and structure analysis. Importantly, the analysis was first conducted without any preset metal-ligand bonds, to assure a reliable structure determination in an unbiased manner.

Structure and Coordination Determination of Peptide-metal Complexes Using 1D and 2D 1H NMR

The NMR-solution structure of a metallochaperone model peptide with Cu (I) was determined, and a detailed protocol from sample preparation and 1D and 2D data collection to a three-dimensional structure is described.

Copper (I) binding by metallochaperone transport proteins prevents copper oxidation and release of the toxic ions that may participate in harmful redox ...

Split-and-pool Synthesis and Characterization of Peptide Tertiary Amide Library

Peptidomimetics are great sources of protein ligands. The oligomeric nature of these compounds enables us to access large synthetic libraries on solid phase by using combinatorial chemistry. One of the most well studied classes of peptidomimetics is peptoids. Peptoids are easy to synthesize and have been shown to be proteolysis-resistant and cell-permeable. Over the past decade, many useful protein ligands have been identified through screening of peptoid libraries. However, most of the ligands identified from peptoid libraries do not display high affinity, with rare exceptions. This may be due, in part, to the lack of chiral centers and conformational constraints in peptoid molecules. Recently, we described a new synthetic route to access peptide tertiary amides (PTAs). PTAs are a superfamily of peptidomimetics that include but are not limited to peptides, peptoids and N-methylated peptides. With side chains on both &#945;-carbon and main chain nitrogen atoms, the conformation of these molecules are greatly constrained by sterical hindrance and allylic 1,3 strain. (Figure 1) Our study suggests that these PTA molecules are highly structured in solution and can be used to identify protein ligands. We believe that these molecules can be a future source of high-affinity protein ligands. Here we describe the synthetic method combining the power of both split-and-pool and sub-monomer strategies to synthesize a sample one-bead one-compound (OBOC) library of PTAs.

Peptide tertiary amides (PTAs) are a superfamily of peptidomimetics that include but are not limited to peptides, peptoids and N-methylated peptides. Here we describe a synthetic method which combines&#160;both split-and-pool and sub-monomer strategies to synthesize a one-bead one-compound library of PTAs.

Peptidomimetics are great sources of protein ligands. The oligomeric nature of these compounds enables us to access large synthetic libraries on solid ...

A Direct, Early Stage Guanidinylation Protocol for the Synthesis of Complex Aminoguanidine-containing Natural Products

The guanidine functional group, displayed most prominently in the amino acid arginine, one of the fundamental building blocks of life, is an important structural element found in many complex natural products and pharmaceuticals. Owing to the continual discovery of new guanidine-containing natural products and designed small molecules, rapid and efficient guanidinylation methods are of keen interest to synthetic and medicinal organic chemists. Because the nucleophilicity and basicity of guanidines can affect subsequent chemical transformations, traditional, indirect guanidinylation is typically pursued. Indirect methods commonly employ multiple protection steps involving a latent amine precursor, such as an azide, phthalimide, or carbamate. By circumventing these circuitous methods and employing a direct guanidinylation reaction early in the synthetic sequence, it was possible to forge the linear terminal guanidine containing backbone of clavatadine A to realize a short and streamlined synthesis of this potent factor XIa inhibitor. In practice, guanidine hydrochloride is elaborated with a carefully constructed protecting array that is optimized to survive the synthetic steps to come. In the preparation of clavatadine A, direct guanidinylation of a commercially available diamine eliminated two unnecessary steps from its synthesis. Coupled with the wide variety of known guanidine protecting groups, direct guanidinylation evinces a succinct and efficient practicality inherent to methods that find a home in a synthetic chemist&#39;s toolbox.

Here, we present a protocol for direct, early stage guanidinylation that enables rapid total synthesis of aminoguanidine-containing small organic molecules. An advanced synthetic intermediate used in the synthesis of a blood coagulation factor XIa inhibitor was prepared using this protocol.

The guanidine functional group, displayed most prominently in the amino acid arginine, one of the fundamental building blocks of life, is an important ...

Protocol for the Synthesis of Ortho-trifluoromethoxylated Aniline Derivatives

Molecules bearing trifluoromethoxy (OCF3) group often show desired pharmacological and biological properties. However, facile synthesis of trifluoromethoxylated aromatic compounds remains a formidable challenge in organic synthesis. Conventional approaches often suffer from poor substrate scope, or require use of highly toxic, difficult-to-handle, and/or thermally labile reagents. Herein, we report a user-friendly protocol for the synthesis of methyl 4-acetamido-3-(trifluoromethoxy)benzoate using 1-trifluoromethyl-1,2-benziodoxol-3(1H)-one (Togni reagent II). Treating methyl 4-(N-hydroxyacetamido)benzoate (1a) with Togni reagent II in the presence of a catalytic amount of cesium carbonate (Cs2CO3) in chloroform at RT afforded methyl 4-(N-(trifluoromethoxy)acetamido)benzoate (2a). This intermediate was then converted to the final product methyl 4-acetamido-3-(trifluoromethoxy)benzoate (3a) in nitromethane at 120 &#176;C. This procedure is general and can be applied to the synthesis of a broad spectrum of ortho-trifluoromethoxylated aniline derivatives, which could serve as useful synthetic building blocks for the discovery and development of new pharmaceuticals, agrochemicals, and functional materials.

Protocol for the Synthesis of Ortho-trifluoromethoxylated Aniline Derivatives

An operationally simple procedure for the synthesis of ortho-trifluoromethoxylated aniline derivatives via a two-step sequence of O-trifluoromethylation of N-aryl-N-hydroxyacetamide followed by thermally induced intramolecular OCF3-migration is reported.

Molecules bearing trifluoromethoxy (OCF3) group often show desired pharmacological and biological properties. However, facile synthesis of ...

Synthesis of Protein Bioconjugates via Cysteine-maleimide Chemistry

The chemical linking or bioconjugation of proteins to fluorescent dyes, drugs, polymers and other proteins has a broad range of applications, such as the development of antibody drug conjugates (ADCs) and nanomedicine, fluorescent microscopy and systems chemistry. For many of these applications, specificity of the bioconjugation method used is of prime concern. The Michael addition of maleimides with cysteine(s) on the target proteins is highly selective and proceeds rapidly under mild conditions, making it one of the most popular methods for protein bioconjugation.
We demonstrate here the modification of the only surface-accessible cysteine residue on yeast cytochrome c with a ruthenium(II) bisterpyridine maleimide. The protein bioconjugation is verified by gel electrophoresis and purified by aqueous-based fast protein liquid chromatography in 27% yield of isolated protein material. Structural characterization with MALDI-TOF MS and UV-Vis is then used to verify that the bioconjugation is successful. The protocol shown here is easily applicable to other cysteine - maleimide coupling of proteins to other proteins, dyes, drugs or polymers.

Synthesis of Protein Bioconjugates via Cysteine-maleimide Chemistry

This protocol details the important steps required for the bioconjugation of a cysteine containing protein to a maleimide, including reagent purification, reaction conditions, bioconjugate purification and bioconjugate characterization.

The chemical linking or bioconjugation of proteins to fluorescent dyes, drugs, polymers and other proteins has a broad range of applications, such as the ...

Facile Synthesis of Worm-like Micelles by Visible Light Mediated Dispersion Polymerization Using Photoredox Catalyst

Presented herein is a protocol for the facile synthesis of worm-like micelles by visible light mediated dispersion polymerization. This approach begins with the synthesis of a hydrophilic poly(oligo(ethylene glycol) methyl ether methacrylate) (POEGMA) homopolymer using reversible addition-fragmentation chain-transfer (RAFT) polymerization. Under mild visible light irradiation (&#955; = 460 nm, 0.7 mW/cm2), this macro-chain transfer agent (macro-CTA) in the presence of a ruthenium based photoredox catalyst, Ru(bpy)3Cl2 can be chain extended with a second monomer to form a well-defined block copolymer in a process known as Photoinduced Electron Transfer RAFT (PET-RAFT). When PET-RAFT is used to chain extend POEGMA with benzyl methacrylate (BzMA) in ethanol (EtOH), polymeric nanoparticles with different morphologies are formed in situ according to a polymerization-induced self-assembly (PISA) mechanism. Self-assembly into nanoparticles presenting POEGMA chains at the corona and poly(benzyl methacrylate) (PBzMA) chains in the core occurs in situ due to the growing insolubility of the PBzMA block in ethanol. Interestingly, the formation of highly pure worm-like micelles can be readily monitored by observing the onset of a highly viscous gel in situ due to nanoparticle entanglements occurring during the polymerization. This process thereby allows for a more reproducible synthesis of worm-like micelles simply by monitoring the solution viscosity during the course of the polymerization. In addition, the light stimulus can be intermittently applied in an ON/OFF manner demonstrating temporal control over the nanoparticle morphology.

This article describes a process for producing polymeric self-assembled nanoparticles using visible light mediated dispersion polymerization. Using low energy visible light to control the polymerization allows for the reproducible formation of self-assembled worm-like micelles at high solids content.

Presented herein is a protocol for the facile synthesis of worm-like micelles by visible light mediated dispersion polymerization. This approach begins ...

Self-assembling Morphologies Obtained from Helical Polycarbodiimide Copolymers and Their Triazole Derivatives

A facile method for the preparation of polycarbodiimide-based secondary structures (e.g., nano-rings, &#34;craters,&#34; fibers, looped fibers, fibrous networks, ribbons, worm-like aggregates, toroidal structures, and spherical particles) is described. These aggregates are morphologically influenced by extensive hydrophobic side chain-side chain interactions of the singular polycarbodiimide strands, as inferred by atomic force microscopy (AFM) and scanning electron microscopy (SEM) techniques. Polycarbodiimide-g-polystyrene copolymers (PS-PCDs) were prepared by a combination of synthetic methods, including coordination-insertion polymerization, copper(I)-catalyzed azide alkyne cycloaddition (CuAAC) &#34;click&#34; chemistry, and atom transfer radical polymerization (ATRP). PS-PCDs were found to form specific toroidal architectures at low concentrations in CHCl3. To determine the influence of a more polar solvent medium (i.e., THF and THF/EtOH) on polymer aggregation behavior, a number of representative PS-PCD composites have been tested to show discrete concentration-dependent spherical particles. These fundamental studies are of practical interest to the development of experimental procedures for desirable architectures by directed self-assembly in thin film. These architectures may be exploited as drug carriers, whereas other morphological findings represent certain interest in the area of novel functional materials.

Here, we present a protocol to prepare and visualize secondary structures (e.g., fibers, toroidal architectures, and nano-spheres) derived from helical polycarbodiimides. The morphology characterized by both atomic force microscopy (AFM) and scanning electron microscopy (SEM) was shown to depend on molecular structure, concentration, and the solvent of choice.

A facile method for the preparation of polycarbodiimide-based secondary structures (e.g., nano-rings, &#34;craters,&#34; fibers, looped fibers, fibrous ...

Protocol for the Solid-phase Synthesis of Oligomers of RNA Containing a 2'-O-thiophenylmethyl Modification and Characterization via Circular Dichroism

Solid-phase synthesis has been used to obtain canonical and modified polymers of nucleic acids, specifically of DNA or RNA, which has made it a popular methodology for applications in various fields and for different research purposes. The procedure described herein focuses on the synthesis, purification, and characterization of dodecamers of RNA 5&#39;-[CUA CGG AAU CAU]-3&#39; containing zero, one, or two modifications located at the C2&#39;-O-position. The probes are based on 2-thiophenylmethyl groups, incorporated into RNA nucleotides via standard organic synthesis and introduced into the corresponding oligonucleotides via their respective phosphoramidites. This report makes use of phosphoramidite chemistry via the four canonical nucleobases (Uridine (U), Cytosine (C), Guanosine (G), Adenosine (A)), as well as 2-thiophenylmethyl functionalized nucleotides modified at the 2&#39;-O-position; however, the methodology is amenable for a large variety of modifications that have been developed over the years. The oligonucleotides were synthesized on a controlled-pore glass (CPG) support followed by cleavage from the resin and deprotection under standard conditions, i.e., a mixture of ammonia and methylamine (AMA) followed by hydrogen fluoride/triethylamine/N-methylpyrrolidinone. The corresponding oligonucleotides were purified via polyacrylamide electrophoresis (20% denaturing) followed by elution, desalting, and isolation via reversed-phase chromatography (Sep-pak, C18-column). Quantification and structural parameters were assessed via ultraviolet-visible (UV-vis) and circular dichroism (CD) photometric analysis, respectively. This report aims to serve as a resource and guide for beginner and expert researchers interested in embarking in this field. It is expected to serve as a work-in-progress as new technologies and methodologies are developed. The description of the methodologies and techniques within this document correspond to a DNA/RNA synthesizer (refurbished and purchased in 2013) that uses phosphoramidite chemistry.

Protocol for the Solid-phase Synthesis of Oligomers of RNA Containing a 2'-O-thiophenylmethyl Modification and Characterization via Circular Dichroism

This article provides a detailed procedure on the solid-phase synthesis, purification, and characterization of dodecamers of RNA modified at the C2&#39;-O-position. UV-vis and circular dichroism photometric analyses are used to quantify and characterize structural aspects, i.e., single-strands or double-strands.

Solid-phase synthesis has been used to obtain canonical and modified polymers of nucleic acids, specifically of DNA or RNA, which has made it a popular ...

Synthesis and Mass Spectrometry Analysis of Oligo-peptoids

Peptoids are sequence-controlled peptide-mimicking oligomers consisting of N-alkylated glycine units. Among many potential applications, peptoids have been thought of as a type of molecular information storage. Mass spectrometry analysis has been considered the method of choice for sequencing peptoids. Peptoids can be synthesized via solid phase chemistry using a repeating two-step reaction cycle. Here we present a method to manually synthesize oligo-peptoids and to analyze the sequence of the peptoids using tandem mass spectrometry (MS/MS) techniques. The sample peptoid is a nonamer consisting of alternating N-(2-methyloxyethyl)glycine (Nme) and N-(2-phenylethyl)glycine (Npe), as well as an N-(2-aminoethyl)glycine (Nae) at the N-terminus. The sequence formula of the peptoid is Ac-Nae-(Npe-Nme)4-NH2, where Ac is the acetyl group. The synthesis takes place in a commercially available solid-phase reaction vessel. The rink amide resin is used as the solid support to yield the peptoid with an amide group at the C-terminus. The resulting peptoid product is subjected to sequence analysis using a triple-quadrupole mass spectrometer coupled to an electrospray ionization source. The MS/MS measurement produces a spectrum of fragment ions resulting from the dissociation of charged peptoid. The fragment ions are sorted out based on the values of their mass-to-charge ratio (m/z). The m/z values of the fragment ions are compared against the nominal masses of theoretically predicted fragment ions, according to the scheme of peptoid fragmentation. The analysis generates a fragmentation pattern of the charged peptoid. The fragmentation pattern is correlated to the monomer sequence of the neutral peptoid. In this regard, MS analysis reads out the sequence information of the peptoids.

A protocol is described for the manual synthesis of oligo-peptoids followed by sequence analysis by mass spectrometry.

Peptoids are sequence-controlled peptide-mimicking oligomers consisting of N-alkylated glycine units. Among many potential applications, peptoids have ...