Name: solid-phase submonomer synthesis of peptoid polymers and their self-assembly into highly-ordered nanosheets
Uploaded: 2011-11-02T12:00:00
Duration: 13 min 42 s
Description: Watch this Scientific Journal Video about Solid-phase Submonomer Synthesis of Peptoid Polymers and their Self-Assembly into Highly-Ordered Nanosheets at JoVE.com

The authors would like to thank Byoung-Chul Lee, Philip Choi and Samuel Ho for valuable assistance. This work was carried out at the Molecular Foundry at Lawrence Berkeley National Laboratory, which is supported by the Office of Science, Office of Basic Energy Sciences, of the US Department of Energy under Contract No. DE-AC02-05CH11231 and the Defense Threat Reduction Agency under Contract No: IACRO-B0845281.

1. Solid-Phase Submonomer Synthesis of Polypeptoids
Solid-phase synthesis (SPS) is a common technique used to synthesize sequence-specific biopolymers step-wise, directly on an inert solid-support such as a polymeric resin bead. High coupling yields and ease of excess reactant removal are major advantages of SPS. After a coupling reaction to the resin, excess reagents are simply drained and the beads are washed to be ready for the next reaction step. After the final synthesis reaction, the full-length oligomers are cleaved from the resin and the solution-phase material can be further studied. Here, we adapt the SPS procedure to generate sequence-specific peptoid polymers.
<ol>
	<li>Setup: All steps of the manual peptoid synthesis can be carried out in a disposable, polypropylene (PP) fritted cartridge or a fritted glass reaction vessel equipped with a 3-way stopcock. Perform all operations in a fume hood. For incubations in the glass vessel or plastic cartridge, connect one arm to a nitrogen supply to gently bubble the solution for proper mixing. Alternatively, for reaction incubations in the disposable cartridge, seal both ends of the cartridge with caps and place on a rotary shaker. To drain reaction mixtures or washes, connect to house vacuum via a waste trap. The vessel should be fritted with a coarse frit. Siliconize the glass reaction vessel to avoid beads from sticking to the walls of the glass vessel. Prepare a solution of 5% dichlorodimethylsilane in dichloroethane (v/v). Fill a clean and dry reaction vessel to the top with siliconizing solution, let sit for 30 minutes, then drain. Wash the vessel once with DCE and then once with methanol. The siliconizing solution can be reused, so it should be saved. Either air dry or shake off any excess solution and bake the glassware until dry after removing the stopcock. Cool reaction vessel before adding resin.</li>
	<li>Add 100 mg (0.06 mmol) of Rink amide resin to a fritted reaction vessel. Swell the resin by adding 2 mL of dimethylformamide (DMF). Agitate by shaking or bubbling for 10 minutes. Drain the solution by vacuum to isolate the swelled resin.</li>
	<li>Add 1 mL of 20% 4-methylpiperidine in DMF (v/v) to deprotect the Fmoc group. Agitate for 2 minutes and drain. Repeat with a 12 minute incubation.</li>
	<li>Rinse the resin by adding 2 mL of DMF, agitating for 15 seconds, and draining. Repeat 3x.</li>
	<li>Bromoacetylation: Add 1 mL of 0.6 M bromoacetic acid (0.6 mmol) in DMF and 86 &mu;L of N,N'-diisopropylcarbodiimide (0.93 equivalent, 0.56 mmol). Incubate with gentle bubbling for 30 minutes, then drain and rinse with 2 mL of DMF (repeat 4x).</li>
	<li>Displacement: Add 1 mL of 1-2 M amine in N-methylpyrrolidinone. Incubate with bubbling for 30-120 minutes, then drain and rinse with DMF (4x 2 mL).</li>
	<li>Continue to grow the peptoid chain by repeating the submonomer cycle, steps 1.5 (bromoacetylation) and 1.6 (displacement).</li>
</ol>
<ol start="8">
	<li>After the final displacement is done, rinse with 2 mL of DMF (repeat 4x), then 2 mL of dichloromethane (repeat 3x). Cap and store the reaction vessel until cleavage.</li>
	<li>Pause in synthesis (optional): To pause during a peptoid synthesis, finish the displacement reaction and continue to step 1.8. To continue growing the peptoid chain, restart the synthesis by re-swelling the dried resin (step 1.2) and repeating the submonomer cycle (step 1.5 and 1.6). The resin can be dried and stored after any displacement except the 2nd displacement because the resin-peptoid conjugate may form a cyclic diketopiperazine side product.</li>
	<li>For multiple simultaneous syntheses, a solid-phase extraction vacuum manifold is recommended to maximize efficiency. Peptoid synthesis can also be automated by properly programming methods in commercially available peptide synthesizers, such as the Aapptec Apex 396, CEM Liberty microwave synthesizer and Protein Technologies Inc. Prelude synthesizer.</li>
</ol>
2. Cleavage and Side-Chain Deprotection
<ol>
	<li>Transfer all of dried resin to a 20 mL scintillation glass vial.</li>
	<li>Working inside a hood and using proper personal protective equipment, add 4 mL of trifluoroacetic acid (TFA) cleavage cocktail1 (e.g. 95% aq. TFA, see discussion) to the scintillation glass vial and cap tightly. Shake for 10 minutes to 2 hours at room temperature (see discussion).</li>
	<li>Collect the TFA cleavage solution by filtering the resin through a disposable, PP fritted cartridge into a new, pre-weighed 20 mL scintillation glass vial. A disposable, PP pipette is convenient to transfer the cleavage cocktail solutions.</li>
	<li>Add 1 mL of fresh cleavage cocktail to rinse the resin and collect any residual peptoid. Repeat 2x.</li>
	<li>Evaporate TFA by blowing a gentle stream of nitrogen or by using a Biotage V10 evaporator.</li>
	<li>Redissolve the crude oil in 6 mL of acetonitrile/water 1:1 (v/v) for HPLC. Freeze and lyophilize. Repeat.</li>
	<li>Record the weight of the crude product. Store as a dry powder at -20 &deg;C</li>
	<li>Test cleavage (optional): A test cleavage on 0.5% of the resin can be performed to quickly determine the purity and mass of the synthesized peptoid and whether correct cleavage conditions were chosen. Test cleavages are especially useful to monitor the progress of the synthesis.</li>
</ol>
3. Characterization and Purification of the Polypeptoid
<ol>
	<li>Through a combination of analytical HPLC, electrospray LC-MS, and/or MALDI-TOF, determine the purity of the crude product and whether the desired molecular weight is present.</li>
	<li>Prepare a &tilde;5-10 mg/mL solution of the dry peptoid powder in water with minimal acetonitrile as needed for solubility. Filter clear solution of crude peptoid product with 0.45 &mu;m syringe filter to remove dust and particles.</li>
	<li>Analytical HPLC and electrospray LC-MS: Prepare a &tilde; 20 &mu;g/mL crude peptoid solution. Filter 200 &mu;L with a 0.45 &mu;m filter and inject 20 &mu;L.</li>
	<li>MALDI: Mix 1 &mu;L of &tilde;20 mg/mL peptoid with 1 &mu;L matrix. Spot 1 &mu;L on MALDI plate and allow to air dry. Matrix and acquisition mode is dependent on sample (Fig. 5).</li>
	<li>Purify the crude peptoid mixture with reverse-phase prep HPLC. Choose the gradient and column (C4 or C18) based on the hydrophobicity of polypeptoid. Combine purified fractions, freeze, and lyophilize, resulting in a fluffy white powder. Record the weight of the final product.</li>
	<li>Formation of HCl salt (optional): Redissolve the lyophilized powder in 100 mM HCl (aq.) with minimal acetonitrile. Transfer to pre-weighed glass vial. Freeze and re-lyophilize. Repeat 2x. Reweigh to determine mass of peptoid powder.</li>
</ol>
4. Peptoid Nanosheet Formation
This section describes the protocol to form sheets from a single-chain, sequence specific, amphiphilic 36-mer peptoid (Fig. 1). After the peptoid strand is synthesized, purified, and lyophilized as described above, the resulting white powder is dissolved in DMSO to make a 2 mM stock solution.
<ol>
	<li>Prepare 500 &mu;L of 20 &mu;M peptoid solution in sheet formation buffer (10 mM Tris-HCl, 100 mM NaCl, pH 8.0 in water) in a 1 dram glass vial. First, add 445 &mu;L of Milli-Q water, 50 &mu;L of 10x sheet formation buffer, and vortex to mix. Then, add 5 &mu;L of 2 mM peptoid stock solution and gently swirl solution. Cap the glass vial.</li>
	<li>Sheets are formed by the gentle agitation of the dilute aqueous peptoid solution. Slowly tilting the glass vial from the horizontal position to the upright position results in sheets. Gentle shaking also yields sheets; however, the sheets tend to be smaller and with fewer straight edges. A more thorough analysis of the sheet formation mechanism is reported separately.2</li>
	<li>For many high-quality sheets, rotate the glass vials about the horizontal axis slowly (&lt;1 RPM) for one to three days. An Appropriate Technical Resources RKVSD Rotamix tube rotator or a customized rocker can perform this continuously.</li>
	<li>Dialysis of Nanosheets (optional): In certain applications, it may be necessary to remove any free peptoid chains or buffers/salts. Soak a Float-a-Lyzer 100 kD membrane in the desired buffer for 15 minutes. Load 500 &mu;L peptoid sheet solution into the sample chamber. Soak in 500 mL of desired buffer, stirring with a magnetic stir bar at 60 rpm. Allow dialysis of sheets to proceed for 4 hours. Every hour, exchange with a fresh stock of buffer solution.</li>
</ol>
5. Fluorescence Microscopy of Nanosheets
<ol>
	<li>Fluorescence images of the nanosheets were imaged with Nile Red, an environmentally sensitive dye whose fluorescence intensity increases when it is localized in hydrophobic environments (Fig. 2).</li>
	<li>Add 1 &mu;L of 100 &mu;M Nile Red to 100 &mu;L of the nanosheet solution to obtain a final concentration of 1 &mu;M of Nile Red.</li>
	<li>Make a 1% agarose solution in hot water and pour into a plastic Petri dish. Ensure the agarose solution is approximately 1/8 inch thick and allow the solution to cool undisturbed on a flat surface. After the agarose sets, use a spatula to cut and transfer 1 cm x 1 cm squares to a glass slide.</li>
	<li>To collect the sheets in the same plane, spot 1 &mu;L of sheet solution on the piece of agarose. Allow the agarose to absorb the buffer for 2 minutes, leaving the sheets at the surface. Image within 15 minutes, otherwise the agarose will begin to deform due to dehydration.</li>
	<li>To image sheets in solution, load 15 &mu;L inside a 20 mm diameter 0.12 mm gasket on a glass slide. Cover with a coverslip. If the sheets are simply sandwiched between a glass slide and coverslip without a gasket, many sheets will shear and the seemingly minor evaporation will cause the sheets to constantly move.</li>
	<li>Image sheets under epifluorescence illumination (e.g. an Olympus IX81 inverted microscope fitted with an Andor iXonEM+ EMCCD spectra with a Texas red filter).</li>
</ol>
6. Scanning Electron Microscopy (SEM) of Nanosheets
<ol>
	<li>Plasma etch of silicone substrate (optional): The silicon chips are plasma etched to aid in the adsorption of sheets. Place the silicon chips in the vacuum chamber of a plasma cleaner (e.g. Harrick Plasma Cleaner/Sterilizer PDC-32G). Pump down to 200 mTorr and set the RF coil to 18W (high setting for PDC-32G). Etch for 2 minutes.</li>
	<li>Drop 20 &mu;L of peptoid sheet solution on a plasma-treated silicon substrate. Allow to sit for 3 minutes. Remove excess solution with tip of Kim-wipe. Pipette 20 &mu;L of water onto the surface and remove excess solution again to remove buffer and salts. Repeat 4x.</li>
	<li>Alternatively, dialyze the peptoid sheet solutions against water to remove buffer and salt. Drop 20 &mu;L of dialyzed sheet solution on plasma-treated silicon substrates. Air dry the sample.</li>
	<li>Image sheets with SEM (e.g. Zeiss Gemini Ultra-55 Analytical Scanning Electron Microscope) with an in-lens detector and at beam energies between 1 kV and 5 kV (Fig. 3).</li>
</ol>
7. Safety Notes:
<ol>
	<li>Dimethylformamide and Dichloromethane are reasonably suspected carcinogens.</li>
	<li>N,N'-Diisopropylcarbodiimide, 4-methylpiperdine and bromoacetic acid are hazardous to the skin, eyes, and respiratory tract. They should be used in the hood with care. It may be toxic if inhaled or absorbed through the skin, and exposure may result in sensitization. Empty containers retain product residue (liquid/vapor) and should be thoroughly rinsed before removing them from the hood.</li>
	<li>TFA is a strong acid, and is extremely destructive to the upper respiratory tract, eyes, and skin. TFA is also volatile-keep concentrated solutions of TFA in the hood at all times to avoid respiratory damage. Use proper PPE, and caution when handling solutions of TFA. Change gloves promptly if they come in contact with TFA, and immediately clean up any spills.</li>
</ol>
8. Representative Results:
This section describes the synthesis, characterization, and purification of a sequence-specific 36-mer peptoid chain that folds into a highly ordered nanosheet3 (Fig. 1).
The block-charge peptoid H-[Nae-Npe]9-[Nce-Npe]9-NH2 was synthesized on 100 mg of Rink amide resin. A 2 M amine solution was used for all displacement reactions, which were carried out for 60 minutes for residue 1-18 and 120 minutes for residue 19-36. t-Butyl beta-alanine HCl was converted to the free base (see discussion) whereas phenethylamine and boc-ethylenediamine were both used directly. The resin was cleaved with 95% TFA, 2.5% triispropylsilane, 2.5% water for 2 hours. TFA was evaporated and the resulting viscous oil (~180 mg) was re-dissolved in 6 mL acetonitrile:water 1:1 (v/v). Product purity (Fig. 4) and presence of the product mass was confirmed by from analytical RP-HPLC (30-80% acetonitrile in water gradient, both containing 0.1% (v/v) TFA, at 1 mL/min over 30 minutes at 60 &deg;C with a C18, 5 &mu;m, 50 X 2 mm column) and MALDI (Fig. 5).
Purification with reverse phase HPLC on a Vydac C18 column (10 &mu;m, 22 mm x 250 mm) proceeded, using a gradient of 30-60% acetonitrile in water with 0.1% TFA over 60 minutes at 10 mL/min. The column was loaded with 60 mg of crude product for each chromatographic run. The purified fractions were combined based on purity from analytical RP-HPLC (Fig. 4) and lyophilized to yield &tilde;80 mg of a fluffy white powder.
Purified block-charge peptoid molecular weight was confirmed by MALDI. 1 &mu;L of 100 &mu;M purified peptoid in acetonitrile:water 1:1 (v/v) was mixed with 1 &mu;L of matrix (5 mg/mL &alpha;-cyano-4-hydroxycinnamic acid in acetonitrile:water 1:1 v/v and 0.1% TFA) and 1 &mu;L was spotted on the MALDI plate. After the sample air-dried, it was placed in the Applied Biosystem/MDS SCIEX 4800 MALDI TOF/TOF Analyzer. The acquisition and processing modes were linear low mass. The calculated weight was entered in the targeted mass to automatically adjust for the delay time. The laser intensity was set to 3400. The observed mass, 4981.2, matches closely to the calculated mass of 4981.74.
The lyophilized purified powder was dissolved in DMSO to make a 2 mM stock solution, which can be stored at 4 &deg;C. Sheets were prepared by aforementioned protocol and imaged with fluorescence optical microscopy and SEM (Fig. 2 and 3). A variety of shapes with feature sizes ranging up to 300 &mu;m are observed, and notably, straight edges are prominent.
<img alt="Figure 1" src="/files/ftp_upload/3373/3373fig1.jpg" /> 
Figure 1. Sequence of the block-charge peptoid H-[Nae-Npe]9-[Nce-Npe]9-NH2. A single-chain, block charge, amphiphilic polypeptoid 36-mer self-assembles into highly-ordered, two-dimensional nanosheets3. The calculated molecular weight is 4981.74.
<img alt="Figure 2" src="/files/ftp_upload/3373/3373fig2.jpg" /> 
Figure 2. Fluorescence microscopy images of peptoid nanosheets. Sheets were formed from a 20 &mu;M peptoid solution in 10 mM Tris, 100 mM NaCl, pH 8.0. The sheets were imaged on agarose with 1&mu;M Nile Red. Scale bars are 100 &mu;m.
<img alt="Figure 3" src="/files/ftp_upload/3373/3373fig3.jpg" /> 
Figure 3. Scanning electron microscopy images of peptoid nanosheets. Sheets were formed from a 20 &mu;M peptoid solution in 10 mM Tris, 100 mM NaCl, pH 8.0. Scale bars are 5 &mu;m.
<img alt="Figure 4" src="/files/ftp_upload/3373/3373fig4.jpg" /> 
Figure 4. Analytical reverse phase HPLC trace of H-[Nae-Npe]9-[Nce-Npe]9-NH2. The crude and purified analytical HPLC trace (30-80% gradient at 1 mL/min over 30 minutes at 60&deg;C with a C18, 5 &mu;m, 50 x 2 mm column) of the crude and purified block-charge peptoid H-[Nae-Npe]9-[Nce-Npe]9-NH2 is shown.
<img alt="Figure 5" src="/files/ftp_upload/3373/3373fig5.jpg" /> 
Figure 5. MALDI-TOF mass spectroscopy trace of H-[Nae-Npe]9-[Nce-Npe]9-NH2. The observed mass, 4981.2, is in close agreement to the calculated mass, 4981.74.

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We have nothing to disclose.

Applications and Significance
This protocol describes a simple and efficient method of peptoid synthesis and the aqueous self-assembly of the peptoids into nanosheets. Most laboratories are easily capable of synthesizing peptoids because inexpensive materials, basic expertise and straightforward techniques are utilized4. Likewise, the self-assembly of ultra-thin, highly-ordered nanosheets merely requires repeated tilting a vial of a dilute aqueous peptoid solution2. Peptoids are promising materials for biomedical and nanoscience research because they are robust and synthetically flexible yet sequence-specific and highly tunable5. Peptoids have demonstrated biological activity (therapeutics6,7, diagnostics8, intracellular delivery9-10) and folding into hierarchical nanostructures3, 11-14. Because of their modular synthesis, combinatorial peptoid libraries15-19 can be readily synthesized and screened for a broad series of activities or properties. In particular, the nanosheets serve as a potential platform for two-dimensional display scaffolds, membrane mimetics, biological sensors, protein mimetics and device fabrication. With the practically inexhaustible different sequences possible, the realm of peptoid research is quickly expanding.
 
Variables in solid-phase submonomer synthesis of polypeptoids
Because of the ability to choose from an incredibly large and diverse alphabet of monomers20, the submonomer method needs occasional modifications for cases where increasing the coupling efficiency of each step will improve the overall product yield. Incorporation of unprotected heterocyclic side chains requires the use of chloroacetic acid instead of bromoacetic acid21. Extended displacement times and higher amine concentrations are usually employed after about 20 couplings for long peptoid sequences or less nucleophilic amines. Heating the reaction vessel to 35 &deg;C, by using a water-jacketed reaction vessel, helps to drive the reaction. For highly-volatile amines such as isopropylamine, care must be taken to avoid evaporation.
Amines in the form of an HCl salt, such as t-butyl beta-alanine HCl, need to be free-based before being introduced in the displacement reaction. This can be achieved by dissolving or suspending the amine in DCM (&tilde;5g amine/25 mL DCM), and neutralizing with an equimolar solution of aqueous sodium hydroxide in a separatory funnel. The DCM layer is collected and the aqueous layer is washed with additional DCM. The combined DCM layers are dried over sodium sulfate and filtered into a pre-weighed round bottom flask. Remove solvent by rotary evaporation to yield an oil, and record the product weight.
During the cleavage step, TFA cleavage cocktail and cleavage time is dependent on the number and variety of protecting groups used. Guidelines for cleavage cocktails are similar to traditional peptide deprotection cleavages1. Generally, 10 minute incubations are required for sequences without protecting groups or sequences with few highly acid labile protecting groups (e.g. BOC, trityl). Two hour incubations are recommended for sequences with more difficult protecting groups (e.g. t-butyl ester, Mtr, Pbf) or sequences with many protecting groups to ensure full deprotection of each chain. Crude peptoid products will generally dissolve in acetonitrile:water 1:1 (v/v), but higher acetonitrile proportions are common with side chains with a high overall hydrophobicity.

<table border="1">
<tbody>
<tr>
<td>Name of the reagent</td>
<td>Company</td>
<td>Catalogue number</td>
<td>Comments</td>
</tr>
<tr>
<td>Dimethylformamide</td>
<td>EMD</td>
<td>EM-DX1726P-1 </td>
<td>99+%</td>
</tr>
<tr>
<td>N-methylpyrrolidinone</td>
<td>BDH</td>
<td>BDH1141-4LP</td>
<td>99%</td>
</tr>
<tr>
<td>Bromoacetic Acid</td>
<td>Acros Organics</td>
<td>200000-106</td>
<td>99%</td>
</tr>
<tr>
<td>4-Methylpiperidine</td>
<td>Sigma Aldrich</td>
<td>M73206</td>
<td>96%</td>
</tr>
<tr>
<td>N,N&rsquo;-diisopropylcarbodiimide</td>
<td>Chem-Impex</td>
<td>001100</td>
<td>99.5%</td>
</tr>
<tr>
<td>Dichloromethane</td>
<td>EMD</td>
<td>EMD-DX0835</td>
<td>ACS grade</td>
</tr>
<tr>
<td>Acetonitrile</td>
<td>EMD</td>
<td>EM-AX0145P-1</td>
<td>99.8%</td>
</tr>
<tr>
<td>Trifluoroacetic acid</td>
<td>Sigma Aldrich</td>
<td>T6508</td>
<td>99%</td>
</tr>
<tr>
<td>Triisopropylsilane</td>
<td>Sigma Aldrich</td>
<td>233781-10G </td>
<td>For TFA cleavage</td>
</tr>
<tr>
<td>1,2-Dichloroethane</td>
<td>JT Baker</td>
<td>JTH076-33</td>
<td>For siliconization of glass reaction vessels</td>
</tr>
<tr>
<td>Phenethylamine</td>
<td>Sigma Aldrich</td>
<td>407267-100ML </td>
<td>&gt;99.5% Hydrophobic side-chain amine</td>
</tr>
<tr>
<td>Boc-ethylenediamine</td>
<td>CNH Technologies</td>
<td>C-1112</td>
<td>Cationic side-chain amine</td>
</tr>
<tr>
<td>t-Butyl beta-alanine HCl</td>
<td>Chem-Impex International</td>
<td>04407 </td>
<td>Anionic side-chain amine</td>
</tr>
<tr>
<td>&alpha;-Cyano-4-hydroxycinnamic acid</td>
<td>Sigma Aldrich</td>
<td>C8982-10X10MG </td>
<td>For MALDI matrix</td>
</tr>
<tr>
<td>Nile Red</td>
<td>Sigma Aldrich</td>
<td>19123-10MG </td>
<td>For fluorescence Imaging</td>
</tr>
<tr>
<td>Dichlorodimethylsilane</td>
<td>Sigma Aldrich</td>
<td>80430-500G-F</td>
<td>For siliconization of glass reaction vessels</td>
</tr>
<tr>
<td>Disposable PP fritted cartridge</td>
<td>Applied Separations</td>
<td>2416</td>
<td>6 mL polypropylene cartridge with 20 mm PE frit</td>
</tr>
<tr>
<td>Disposable 3 way luer adapter</td>
<td>Cole Parmer</td>
<td>31200-80</td>
<td>Stopcock for disposable manual synthesis reaction vessel</td>
</tr>
<tr>
<td>Luer Lock ring</td>
<td>Cole Parmer</td>
<td>45503-19 </td>
<td>&frac14;&rdquo; fitting for disposable manual synthesis reaction vessel</td>
</tr>
<tr>
<td>Fittings Luer</td>
<td>Cole Parmer</td>
<td>45500-20</td>
<td>&frac14;&rdquo; fitting for disposable manual synthesis reaction vessel</td>
</tr>
<tr>
<td>Disposable PP pipets</td>
<td>VWR</td>
<td>16001-194</td>
<td>For TFA transfers</td>
</tr>
<tr>
<td>Luer lock plastic syringe</td>
<td>National Scientific</td>
<td>S7515-5</td>
<td>6 mL syringes</td>
</tr>
<tr>
<td>1 dram glass vial</td>
<td>VWR</td>
<td>66011-041</td>
<td>With phenolic molded screw cap with polyvinyl-faced pulp liner</td>
</tr>
<tr>
<td>20 mL scintillation vial</td>
<td>VWR</td>
<td>66022-060 </td>
<td>With attached PP cap and pulp foil liner</td>
</tr>
<tr>
<td>Secure-Seal adhesive spacer</td>
<td>Invitrogen</td>
<td>S-24736 </td>
<td>For fluorescence imaging</td>
</tr>
<tr>
<td>Glass slides</td>
<td>Electron Microscopy Sciences</td>
<td>63411</td>
<td>For fluorescence imaging</td>
</tr>
<tr>
<td>Cover slip</td>
<td>VWR</td>
<td>48366-067</td>
<td>For fluorescence imaging</td>
</tr>
<tr>
<td>4&rdquo; Silicon wafer</td>
<td>Ted Pella</td>
<td>16007</td>
<td>Pre-dice in 5x7 mm chips</td>
</tr>
<tr>
<td>0.45 filter</td>
<td>VWR, Acrodisc</td>
<td>28143-924</td>
<td>For HPLC. 
PTFE membrane</td>
</tr>
<tr>
<td>Agarose</td>
<td>BD</td>
<td>212272</td>
<td>For fluorescence imaging</td>
</tr>
<tr>
<td>SPE Vacuum Manifold</td>
<td>Sigma Aldrich</td>
<td>57044</td>
<td>Example of SPE vacuum manifold</td>
</tr>
<tr>
<td>Fritted glass vessel</td>
<td>Ace glass</td>
<td>6402-12</td>
<td>Porosity C frit</td>
</tr>
<tr>
<td>Plasma Cleaner/Sterilizer</td>
<td>Harrick Plasma</td>
<td>PDC-32G</td>
<td>Example of plasma cleaner to prepare silicon chips for SEM</td>
</tr>
</tbody>
</table>

solid-phase submonomer synthesis of peptoid polymers and their self-assembly into highly-ordered nanosheets

Peptoids are a novel class of biomimetic, non-natural, sequence-specific heteropolymers that resist proteolysis, exhibit potent biological activity, and fold into higher order nanostructures. Structurally similar to peptides, peptoids are poly N-substituted glycines, where the side chains are attached to the nitrogen rather than the alpha-carbon. Their ease of synthesis and structural diversity allows testing of basic design principles to drive de novo design and engineering of new biologically-active and nanostructured materials.
Here, a simple manual peptoid synthesis protocol is presented that allows the synthesis of long chain polypeptoids ( up to 50mers) in excellent yields. Only basic equipment, simple techniques (e.g. liquid transfer, filtration), and commercially available reagents are required, making peptoids an accessible addition to many researchers' toolkits. The peptoid backbone is grown one monomer at a time via the submonomer method which consists of a two-step monomer addition cycle: acylation and displacement. First, bromoacetic acid activated in situ with N,N'-diisopropylcarbodiimide acylates a resin-bound secondary amine. Second, nucleophilic displacement of the bromide by a primary amine follows to introduce the side chain. The two-step cycle is iterated until the desired chain length is reached. The coupling efficiency of this two-step cycle routinely exceeds 98% and enables the synthesis of peptoids as long as 50 residues. Highly tunable, precise and chemically diverse sequences are achievable with the submonomer method as hundreds of readily available primary amines can be directly incorporated.
 
Peptoids are emerging as a versatile biomimetic material for nanobioscience research because of their synthetic flexibility, robustness, and ordering at the atomic level. The folding of a single-chain, amphiphilic, information-rich polypeptoid into a highly-ordered nanosheet was recently demonstrated. This peptoid is a 36-mer that consists of only three different commercially available monomers: hydrophobic, cationic and anionic. The hydrophobic phenylethyl side chains are buried in the nanosheet core whereas the ionic amine and carboxyl side chains align on the hydrophilic faces. The peptoid nanosheets serve as a potential platform for membrane mimetics, protein mimetics, device fabrication, and sensors. Methods for peptoid synthesis, sheet formation, and microscopy imaging are described and provide a simple method to enable future peptoid nanosheet designs.

Watch this Scientific Journal Video about Solid-phase Submonomer Synthesis of Peptoid Polymers and their Self-Assembly into Highly-Ordered Nanosheets at JoVE.com

Solid-phase Submonomer Synthesis of Peptoid Polymers and their Self-Assembly into Highly-Ordered Nanosheets

A simple and general manual peptoid synthesis method involving basic equipment and commercially available reagents is outlined, enabling peptoids to be easily synthesized in most laboratories. The synthesis, purification and characterization of an amphiphilic peptoid 36mer is described, as well as its self-assembly into highly-ordered nanosheets.

Peptoids are a novel class of biomimetic, non-natural, sequence-specific heteropolymers that resist proteolysis, exhibit potent biological activity, and ...

solid-phase-submonomer-synthesis-peptoid-polymers-their-self-assembly

Research

JoVE Journal

Bioengineering

Shape Memory Polymers for Active Cell Culture

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent shape upon the application of a stimulus such as heat1-5. In a typical shape memory cycle, the SMP is first deformed at an elevated temperature that is higher than its transition temperature, Ttrans [either the melting temperature (Tm) or the glass transition temperature (Tg)]. The deformation is elastic in nature and mainly leads to a reduction in conformational entropy of the constituent network chains (following the rubber elasticity theory). The deformed SMP is then cooled to a temperature below its Ttrans while maintaining the external strain or stress constant. During cooling, the material transitions to a more rigid state (semi-crystalline or glassy), which kinetically traps or "freezes" the material in this low-entropy state leading to macroscopic shape fixing. Shape recovery is triggered by continuously heating the material through Ttrans under a stress-free (unconstrained) condition. By allowing the network chains (with regained mobility) to relax to their thermodynamically favored, maximal-entropy state, the material changes from the temporary shape to the permanent shape.
Cells are capable of surveying the mechanical properties of their surrounding environment6. The mechanisms through which mechanical interactions between cells and their physical environment control cell behavior are areas of active research. Substrates of defined topography have emerged as powerful tools in the investigation of these mechanisms. Mesoscale, microscale, and nanoscale patterns of substrate topography have been shown to direct cell alignment, cell adhesion, and cell traction forces7-14. These findings have underscored the potential for substrate topography to control and assay the mechanical interactions between cells and their physical environment during cell culture, but the substrates used to date have generally been passive and could not be programmed to change significantly during culture. This physical stasis has limited the potential of topographic substrates to control cells in culture.
Here, active cell culture (ACC) SMP substrates are introduced that employ surface shape memory to provide programmed control of substrate topography and deformation. These substrates demonstrate the ability to transition from a temporary grooved topography to a second, nearly flat memorized topography. This change in topography can be used to control cell behavior under standard cell culture conditions.

A method for developing cell culture substrates with the ability to change topography during culture is described. The method makes use of smart materials known as shape memory polymers that have the ability to memorize a permanent shape. This concept is adaptable to a wide range of materials and applications.

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent ...

Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do not have suitable autologous donor vessels due to disease progression or previous harvest. The aim of vascular tissue engineering is to develop a suitable alternative source for these bypass grafts. In addition, engineered vascular tissue may prove valuable as living vascular models to study cardiovascular diseases. Several promising approaches to engineering blood vessels have been explored, with many recent studies focusing on development and analysis of cell-based methods.2-5 Herein, we present a method to rapidly self-assemble cells into 3D tissue rings that can be used in vitro to model vascular tissues.
To do this, suspensions of smooth muscle cells are seeded into round-bottomed annular agarose wells. The non-adhesive properties of the agarose allow the cells to settle, aggregate and contract around a post at the center of the well to form a cohesive tissue ring.6,7 These rings can be cultured for several days prior to harvesting for mechanical, physiological, biochemical, or histological analysis. We have shown that these cell-derived tissue rings yield at 100-500 kPa ultimate tensile strength8 which exceeds the value reported for other tissue engineered vascular constructs cultured for similar durations (&lt;30 kPa).9,10 Our results demonstrate that robust cell-derived vascular tissue ring generation can be achieved within a short time period, and offers the opportunity for direct and quantitative assessment of the contributions of cells and cell-derived matrix (CDM) to vascular tissue structure and function.

This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do ...

High-throughput Synthesis of Carbohydrates and Functionalization of Polyanhydride Nanoparticles

Transdisciplinary approaches involving areas such as material design, nanotechnology, chemistry, and immunology have to be utilized to rationally design efficacious vaccines carriers. Nanoparticle-based platforms can prolong the persistence of vaccine antigens, which could improve vaccine immunogenicity1. Several biodegradable polymers have been studied as vaccine delivery vehicles1; in particular, polyanhydride particles have demonstrated the ability to provide sustained release of stable protein antigens and to activate antigen presenting cells and modulate immune responses2-12.
The molecular design of these vaccine carriers needs to integrate the rational selection of polymer properties as well as the incorporation of appropriate targeting agents. High throughput automated fabrication of targeting ligands and functionalized particles is a powerful tool that will enhance the ability to study a wide range of properties and will lead to the design of reproducible vaccine delivery devices.
The addition of targeting ligands capable of being recognized by specific receptors on immune cells has been shown to modulate and tailor immune responses10,11,13 C-type lectin receptors (CLRs) are pattern recognition receptors (PRRs) that recognize carbohydrates present on the surface of pathogens. The stimulation of immune cells via CLRs allows for enhanced internalization of antigen and subsequent presentation for further T cell activation14,15. Therefore, carbohydrate molecules play an important role in the study of immune responses; however, the use of these biomolecules often suffers from the lack of availability of structurally well-defined and pure carbohydrates. An automation platform based on iterative solution-phase reactions can enable rapid and controlled synthesis of these synthetically challenging molecules using significantly lower building block quantities than traditional solid-phase methods16,17.
Herein we report a protocol for the automated solution-phase synthesis of oligosaccharides such as mannose-based targeting ligands with fluorous solid-phase extraction for intermediate purification. After development of automated methods to make the carbohydrate-based targeting agent, we describe methods for their attachment on the surface of polyanhydride nanoparticles employing an automated robotic set up operated by LabVIEW as previously described10. Surface functionalization with carbohydrates has shown efficacy in targeting CLRs10,11 and increasing the throughput of the fabrication method to unearth the complexities associated with a multi-parametric system will be of great value (Figure 1a).

In this article, a high throughput method is presented for the synthesis of oligosaccharides and their attachment to the surface of polyanhydride nanoparticles for further use in targeting specific receptors on antigen presenting cells.

Transdisciplinary approaches involving areas such as material design, nanotechnology, chemistry, and immunology have to be utilized to rationally design ...

Optical Control of Living Cells Electrical Activity by Conjugated Polymers

Hybrid interfaces between organic semiconductors and living tissues represent a new tool for in-vitro and in-vivo applications. In particular, conjugated polymers display several optimal properties as substrates for biological systems, such as good biocompatibility, excellent mechanical properties, cheap and easy processing technology, and possibility of deposition on light, thin and flexible substrates. These materials have been employed for cellular interfaces like neural probes, transistors for excitation and recording of neural activity, biosensors and actuators for drug release. Recent experiments have also demonstrated the possibility to use conjugated polymers for all-optical modulation of the electrical activity of cells. Several in-vitro study cases have been reported, including primary neuronal networks, astrocytes and secondary line cells. Moreover, signal photo-transduction mediated by organic polymers has been shown to restore light sensitivity in degenerated retinas, suggesting that these devices may be used for artificial retinal prosthesis in the future. All in all, light sensitive conjugated polymers represent a new approach for optical modulation of cellular activity.
In this work, all the steps required to fabricate a bio-polymer interface for optical excitation of living cells are described. The function of the active interface is to transduce the light stimulus into a modulation of the cell membrane potential. As a study case, useful for in-vitro studies, a polythiophene thin film is used as the functional, light absorbing layer, and Human Embryonic Kidney (HEK-293) cells are employed as the biological component of the interface. Practical examples of successful control of the cell membrane potential upon stimulation with light pulses of different duration are provided. In particular, it is shown that both depolarizing and hyperpolarizing effects on the cell membrane can be achieved depending on the duration of the light stimulus. The reported protocol is of general validity and can be straightforwardly extended to other biological preparations.

A method for stimulation of in-vitro cell cultures electrical activity with visible light, based on the use of organic semiconducting polymers is described.

Hybrid interfaces between organic semiconductors and living tissues represent a new tool for in-vitro and in-vivo applications. In particular, conjugated ...

Fabrication of a Dipole-assisted Solid Phase Extraction Microchip for Trace Metal Analysis in Water Samples

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water samples. A brief overview of the evolution of chip-based SPE techniques is provided. This is followed by an introduction to specific polymeric materials and their role in SPE. To develop an innovative dipole-assisted SPE technique, a chlorine (Cl)-containing SPE functionality was implanted into a poly(methyl methacrylate) (PMMA) microchip. Herein, diverse analytical techniques including contact angle analysis, Raman spectroscopic analysis, and laser ablation-inductively coupled plasma-mass spectrometry (LA-ICP-MS) analysis were employed to validate the utility of the implantation protocol of the C-Cl moieties on the PMMA. The analytical results of the X-ray absorption near-edge structure (XANES) analysis also demonstrated the feasibility of the Cl-containing PMMA used as an extraction medium by virtue of the dipole-ion interactions between the highly electronegative C-Cl moieties and the positively charged metal ions.

The fabrication protocol of a dipole-assisted solid phase extraction microchip for the trace metal analysis is presented.

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water ...

Synthesis of Gold Nanoparticle Integrated Photo-responsive Liposomes and Measurement of Their Microbubble Cavitation upon Pulse Laser Excitation

Photo-responsive nanoparticles (NPs) have received considerable attention because of their potential in providing spatial, temporal, and dosage control over the drug release. However, most of the relevant technologies are still in the development process and are unprocurable by clinics. Here, we describe a facile fabrication of these photo-responsive NPs with commercially available gold NPs and thermo-responsive liposomes. Calcein is used as a model drug to evaluate the encapsulation efficiency and the release kinetic profile upon heat/light stimulation. Finally, we show that this photo-triggered release is due to the membrane disruption caused by microbubble cavitation, which can be measured with hydrophone.

This protocol describes a simple preparation method for gold nanoparticle integrated photo-responsive liposomes with the commercially available materials. It also shows how to measure the microbubble cavitation process of the synthesized liposomes upon the treatment of pulsed laser.

Photo-responsive nanoparticles (NPs) have received considerable attention because of their potential in providing spatial, temporal, and dosage control ...

High-throughput Identification of Bacteria Repellent Polymers for Medical Devices

Medical devices are often associated with hospital-acquired infections, which place enormous strain on patients and the healthcare system as well as contributing to antimicrobial resistance. One possible avenue for the reduction of device-associated infections is the identification of bacteria-repellent polymer coatings for these devices, which would prevent bacterial binding at the initial attachment step. A method for the identification of such repellent polymers, based on the parallel screening of hundreds of polymers using a microarray, is described here. This high-throughput method resulted in the identification of a range of promising polymers that resisted binding of various clinically relevant bacterial species individually and also as multi-species communities. One polymer, PA13 (poly(methylmethacrylate-co-dimethylacrylamide)), demonstrated significant reduction in attachment of a number of hospital isolates when coated onto two commercially available central venous catheters. The method described could be applied to identify polymers for a wide range of applications in which modification of bacterial attachment is important.

A high-throughput microarray method for the identification of polymers which reduce bacterial surface binding on medical devices is described.

Medical devices are often associated with hospital-acquired infections, which place enormous strain on patients and the healthcare system as well as ...

Fabricating Degradable Thermoresponsive Hydrogels on Multiple Length Scales via Reactive Extrusion, Microfluidics, Self-assembly, and Electrospinning

While various smart materials have been explored for a variety of biomedical applications (e.g., drug delivery, tissue engineering, bioimaging, etc.), their ultimate clinical use has been hampered by the lack of biologically-relevant degradation observed for most smart materials. This is particularly true for temperature-responsive hydrogels, which are almost uniformly based on polymers that are functionally non-degradable (e.g., poly(N-isopropylacrylamide) (PNIPAM) or poly(oligoethylene glycol methacrylate) (POEGMA)). As such, to effectively translate the potential of thermoresponsive hydrogels to the challenges of remote-controlled or metabolism-regulated drug delivery, cell scaffolds with tunable cell-material interactions, theranostic materials with the potential for both imaging and drug delivery, and other such applications, a method is required to render the hydrogels (if not fully degradable) at least capable of renal clearance following the required lifetime of the material. To that end, this protocol describes the preparation of hydrolytically-degradable hydrazone-crosslinked hydrogels on multiple length scales based on the reaction between hydrazide and aldehyde-functionalized PNIPAM or POEGMA oligomers with molecular weights below the renal filtration limit. Specifically, methods to fabricate degradable thermoresponsive bulk hydrogels (using a double barrel syringe technique), hydrogel particles (on both the microscale through the use of a microfluidics platform facilitating simultaneous mixing and emulsification of the precursor polymers and the nanoscale through the use of a thermally-driven self-assembly and cross-linking method), and hydrogel nanofibers (using a reactive electrospinning strategy) are described. In each case, hydrogels with temperature-responsive properties similar to those achieved via conventional free radical cross-linking processes can be achieved, but the hydrazone cross-linked network can be degraded over time to re-form the oligomeric precursor polymers and enable clearance. As such, we anticipate these methods (which may be generically applied to any synthetic water-soluble polymer, not just smart materials) will enable easier translation of synthetic smart materials to clinical applications.

Protocols are described for the fabrication of degradable thermoresponsive hydrogels based on hydrazone cross-linking of polymeric oligomers on the bulk scale, microscale, and nanoscale, the latter for preparation of both gel nanoparticles and nanofibers.

While various smart materials have been explored for a variety of biomedical applications (e.g., drug delivery, tissue engineering, bioimaging, etc.), ...

Synthesis of Cationized Magnetoferritin for Ultra-fast Magnetization of Cells

Many important biomedical applications, such as cell imaging and remote manipulation, can be achieved by labeling cells with superparamagnetic iron oxide nanoparticles (SPIONs). Achieving sufficient cellular uptake of SPIONs is a challenge that has traditionally been met by exposing cells to elevated concentrations of SPIONs or by prolonging exposure times (up to 72 hr). However, these strategies are likely to mediate toxicity. Here, we present the synthesis of the protein-based SPION magnetoferritin as well as a facile surface functionalization protocol that enables rapid cell magnetization using low exposure concentrations. The SPION core of magnetoferritin consists of cobalt-doped iron oxide with an average particle diameter of 8.2 nm mineralized inside the cavity of horse spleen apo-ferritin. Chemical cationization of magnetoferritin produced a novel, highly membrane-active SPION that magnetized human mesenchymal stem cells (hMSCs) using incubation times as short as one minute and iron concentrations as lows as 0.2 mM.

A protocol for the synthesis and cationization of cobalt-doped magnetoferritin is presented, as well as a method to rapidly magnetize stem cells with cationized magnetoferritin.

Many important biomedical applications, such as cell imaging and remote manipulation, can be achieved by labeling cells with superparamagnetic iron oxide ...

Engineering Molecular Recognition with Bio-mimetic Polymers on Single Walled Carbon Nanotubes

Semiconducting single-wall carbon nanotubes (SWNTs) are a class of optically active nanomaterial that fluoresce in the near infrared, coinciding with the optical window where biological samples are most transparent. Here, we outline techniques to adsorb amphiphilic polymers and polynucleic acids onto the surface of SWNTs to engineer their corona phases and create novel molecular sensors for small molecules and proteins. These functionalized SWNT sensors are both biocompatible and stable. Polymers are adsorbed onto the nanotube surface either by direct sonication of SWNTs and polymer or by suspending SWNTs using a surfactant followed by dialysis with polymer. The fluorescence emission, stability, and response of these sensors to target analytes are confirmed using absorbance and near-infrared fluorescence spectroscopy. Furthermore, we demonstrate surface immobilization of the sensors onto glass slides to enable single-molecule fluorescence microscopy to characterize polymer adsorption and analyte binding kinetics.

We present a protocol for engineering the corona phase of near infrared fluorescent single walled carbon nanotubes (SWNTs) using amphiphilic polymers and DNA to develop sensors for molecular targets without known recognition elements.

Semiconducting single-wall carbon nanotubes (SWNTs) are a class of optically active nanomaterial that fluoresce in the near infrared, coinciding with the ...