This work was supported in part by National Science Foundation grant 1739308, NSF CAREER grant 1944130 and by the Air Force Office of Science Research grant number FA9550-18-1-0199. &#947;PNA sequences were a generous gift from Dr. Tumul Srivastava of Trucode Gene Repair, Inc. We would like to thank Dr. Erik Winfree and Dr. Rizal Hariadi for their helpful conversations on DNA Design Toolbox MATLAB code. We would also like to thank Joseph Suhan, Mara Sullivan and the Center for Biological Imaging for their assistance in the collection of TEM data.

1. &#947;PNA sequence design
<ol>
	<li>Download the DNA Design Toolbox22 developed by the Winfree Lab at Caltech23 into the folder containing programming scripts for designing sequences.</li>
	<li>Within that sequence design folder, open a fourth-generation programming language compatible with the file extension &#8220;.m&#8221;, and then add the previously downloaded &#8220;DNAdesign&#8221; folder to the path using the following command: 
	&#62;&#62; addpath DNAdesign</li>
	<li>Subsequently run the following script named &#8220;PNA3nanofiber.m&#8221; (see Supplementary Figure 1) using the following command: 
	&#62;&#62; PNA3nanofiber</li>
	<li>Run this script to create variables called &#8220;thisSeqs&#8221; and &#8220;thisScore.&#8221; The variable &#8220;thisSeqs&#8221; contains the sequences of the designed oligomers, and &#8220;thisScore&#8221; is the penalty score.</li>
	<li>Run the script multiple times to obtain the most minimal score. A sample of 20 such runs are shown in Table 1.</li>
	<li>Manually confirm the desired Watson-Crick complementarity of each domain of the generated sequences for the specified SST structural motif. Sequence specifications for the 3-helix nanofiber structure are shown in Table 2.</li>
	<li>Verify the following for generated sequences.
	<ol>
		<li>Avoid using four consecutive C and G bases.</li>
		<li>Manually select specific sequences for N-terminal functionalization with fluorescent dye and biotin molecules to enable fluorescent microscopy studies.</li>
		<li>Include a minimum of 3 mini-PEG gamma-modifications on the backbone to enable pre-organized helical conformation in the single-stranded oligomers as described by Sahu et al.21</li>
	</ol>
	</li>
</ol>
2. Preparation of &#947;PNA stock strands
<ol>
	<li>Obtain &#947;PNA component strands of specified sequences at 50 nmol scale synthesis with high performance liquid chromatography (HPLC)purification from a commercial manufacturer.</li>
	<li>Resuspend each strand in deionized water to 300 &#181;M concentrations. Store the resuspended strands in -20 &#176;C freezer up to several months until needed for experimentation.</li>
</ol>
3. Melting curve studies of &#947;PNA oligomer subsets
<ol>
	<li>Obtain melting temperature ranges for different combinations of complementary 2-oligomer and 3-oligomer subsets by running a melt curve study in aqueous buffers such as 1x phosphate buffered saline (PBS) or preferred polar organic solvent such as Dimethylformamide (DMF) or Dimethyl sulfoxide (DMSO) (see Figure 2). 
	NOTE: Thermal melting curves at &#62;50% of DMF start to lose the upper baseline and show severe disturbances partly because of high absorbance of DMF at the wavelength required for the experiments. This is a noted phenomenon in the literature24. It is however possible to obtain the melting curves at 5 &#181;M per strand concentrations in these solvent conditions.
	<ol>
		<li>Aliquot 16.7 &#181;L from the 300 &#181;M stock of each oligomer and make the final volume to a 1000 &#181;L either in 1x PBS or 100% (v/v) DMSO or 100% (v/v) DMF effectively obtaining 5 &#181;M final concentration per oligomer. Transfer this oligomer subset mixture to a 1 cm optical path, quartz cuvette.</li>
		<li>Perform variable temperature UV-Vis experiments in a spectrophotometer equipped with a programmable temperature block.</li>
		<li>Collect data points for the melting curves over a temperature range of 15&#8210;90 &#176;C for both cooling (annealing) and heating (melting) cycles at a rate of 0.5&#8210;1 &#176;C/min. Keep the samples for 10 min at 90 &#176;C before cooling and at 15 &#176;C before heating. Determine the melting temperature (Tm) from the peak of the first derivative of the heating curve.</li>
		<li>Verify that melting temperature ranges for 2-oligomer subsets are above 35 &#176;C in all solvent cases. Additionally, verify that the corresponding 3-oligomer subsets that contain an additional strand to their equivalent 2-oligomer subset shows a considerable increase in Tm due to increased co-operativity. 
		NOTE: This would verify that a single pot self-assembly of multiple oligomers can co-operatively fold into the desired nanostructure with reasonable thermal stability.</li>
	</ol>
	</li>
</ol>
4. Self-assembly protocol for multiple distinct &#947;PNA oligomers
NOTE: To devise a self-assembly thermal ramp protocol for &#947;PNA nanostructures, slow-ramp annealing is desirable.
<ol>
	<li>In the case of oligomer sequences generated, anneal the samples for 22.5 h in a thermal cycler cooling from 90 to 20 &#176;C. Typically, melting temperature obtained for 2-oligomer and 3-oligomer &#947;PNA subsets lie in ranges of 40&#8210;70 &#176;C for different solvent conditions.</li>
	<li>Program the thermal cycler as follows: hold at 90 &#176;C for 5 min, ramp down from 90 to 70 &#176;C at a constant rate of 0.1 &#176;C/min, ramp down from 70 to 40 &#176;C at a rate of 0.1 &#176;C/3 min, ramp down from 40 to 20 &#176;C at a rate of 0.1 &#176;C/min and hold at 4 &#176;C (see Table 3). Samples can be stored in 4 &#176;C for 12&#8210;24 h before characterization. 
	NOTE: While 2- and 3- oligomer subsets of our nanofiber system can form in 1x PBS, the full micron-scale nanofibers aggregate in 1x PBS. Therefore, solvent conditions should be optimized based on the scale and size of structure being formed as well as the type and density of gamma modifications.</li>
	<li>For micron scale long 3-helix nanofibers, prepare anneal batch samples in 75% DMSO: H2O (v/v), 75% DMF: H2O (v/v), 40% 1,4-Dioxane: H2O (v/v) based on solvent optimization studies15.</li>
	<li>Prepare anneal batches as follows (see Table 4). First, prepare 10 &#181;L sub-stocks at 20 &#181;M concentrations from the 300 &#181;M main stocks for each oligomer by aliquoting 0.67 &#181;L from the main stock and making the volume to 10 &#181;l using deionized water.</li>
	<li>Aliquot 1 &#181;L from the 20 &#181;M sub-stocks for each oligomer and add it to a 200 &#181;L PCR tube. This accounts for a total volume of 9 &#181;L for 9 oligomers.</li>
	<li>Add either 30 &#181;L of anhydrous DMSO/DMF for the 75% DMSO and 75% DMF cases with an additional 1 &#181;L of deionized water to make a final volume of 40 &#181;L with each oligomer at 500 nM final concentrations. Add 16 &#181;L of 1,4-Dioxane and make the volume with deionized water to 40 &#181;L for the 40% Dioxane solvent condition.</li>
	<li>Load the anneal batches on to the thermal cycler and anneal using the protocol mentioned in step 4.2.</li>
</ol>
5. Total internal reflection fluorescence (TIRF) microscopy imaging
<ol>
	<li>Prepare a humidity chamber from an empty pipette tips box. Fill the box with approximately 5 mL of water to prevent drying of the sample flow channels described as follows (see Figure 3).</li>
	<li>Prepare flow chamber with a microscope slide, 2 double-sided tape strips, and nitrocellulose-coated coverslip. To coat the coverslip with nitrocellulose, dip the coverslip in a beaker containing 0.1% collodion in amyl acetate and air-dry.
	<ol>
		<li>Prepare the nitrocellulose solution by making a 20-fold dilution from commercially available 2% collodion in amyl acetate with isoamyl acetate solvent.</li>
	</ol>
	</li>
	<li>Prepare biotinylated bovine serum albumin (Biotin-BSA) solution by weighing 1 mg of Biotin-BSA and dissolving it in 1 mL of 1x PBS. Flow 15 &#956;L of 1 mg/mL Biotin-BSA in 1x PBS buffer by placing the flow channel at an angle. Incubate the flow channel for 2&#8211;4 min at room temperature in a humidified chamber.</li>
	<li>Prepare the wash buffer by dissolving 1 mg of BSA in 1 mL of 1x PBS. Wash the excess Biotin-BSA by flowing 15 &#956;L of the wash buffer. To passivate the surface, incubate the flow channel for 2&#8211;4 mins at room temperature in a humidified chamber.</li>
	<li>Prepare the streptavidin solution by measuring 0.5 mg of streptavidin and 1 mg of BSA and then dissolving using 1 mL of 1x PBS. Flow 15 &#956;L of 0.5 mg/mL streptavidin in 1x PBS containing 1 mg/mL BSA. Incubate the flow channel for 2&#8211;4 min at room temperature in a humidified chamber. Flow 15 &#956;L of the wash buffer to wash away unbound Streptavidin.</li>
	<li>Flow 15 &#956;L of previously annealed batch of &#947;PNA oligomers at 500 nM concentration per strand in either 75% DMSO, 75% DMF or 40% 1,4-Dioxane condition. Incubate the flow channel for 2&#8211;4 mins at room temperature in a humidified chamber.</li>
	<li>Prepare 1 mM Trolox in preferred solvent conditions by diluting 10-fold from a 10 mM stock of Trolox in DMSO (measure 2.5 mg of Trolox and dissolve in 1 mL of DMSO). Wash the unbound nanostructures in the flow channel with 15 &#181;L of 1 mM Trolox having the same solvent composition as the nanostructures. 
	NOTE: The &#62;50% DMSO content in the flow channel would produce minor signal disturbances due to the different index of refraction of the solvent condition during TIRF which is typically calibrated for coverslip-water TIRF angles. This can be rectified by washing with 1 mM Trolox made by diluting the 10 mM Trolox 10-fold using deionized water. This technique provides clearer images but is only useful for assessing whether formation occurred, however, because it produces two-phase micro-bubbles in the flow channel. As a result, nanostructures might be visible at the interface of the two-phases as shown in Figure 4D.</li>
	<li>Transfer the flow channel on to the slide holder and image using a fluorescence microscope equipped with TIRF imaging using a 60x oil-immersion objective and a 1.5x magnifier. Scan the flow channel at either 60x or 90x magnification by monitoring the Cy3 channel (see Figure 4).</li>
</ol>
6. Transmission electron microscopy (TEM) imaging
<ol>
	<li>Weigh 0.5 g of uranyl acetate in 50 mL of distilled water to prepare a 1% aqueous uranyl acetate stain solution. Filter the 1% aqueous uranyl acetate stain solution using a 0.2 &#956;m filter attached to a syringe. 
	NOTE: Alternatively, 2% aqueous uranyl acetate could be purchased from a commercial manufacturer.</li>
	<li>Purchase commercially available formvar support layer coated Copper grids with 300-mesh size. 
	NOTE: It is important to note that the formvar support layer could be dissolved away by solvents like DMSO beyond 2 min. For longer sample incubation, commercially available formvar stabilized with silicon monoxide on copper grids are available which allow the grid to be more hydrophilic than carbon-coated grids and can withstand vigorous sample conditions and electron beam as shown in Figure 5C.</li>
	<li>Pipette 4 &#956;L of sample onto the grid for 15 s. Use a piece of filter paper to wick off the sample by bringing the filter paper in contact with the grid from the side.</li>
	<li>Immediately add 4 &#956;L of the stain solution onto the grid for 5 s.</li>
	<li>Wick off the stain as before and hold the filter paper against the grid for 1&#8210;2 min to make sure the grid is dry. Samples should be typically imaged within 1&#8210;2 h after staining. Alternatively, if the grid is ensured to be completely dry, grids can be stored in a TEM grid storage box for up to 3 days before imaging.</li>
	<li>Transfer the grid to a TEM specimen holder and image using a transmission electron microscope operated at 80 kV with magnification ranging from 10 K to 150K (see Figure 5).</li>
</ol>
7. Different morphologies for &#947;PNA-DNA hybrids based on selective replacement with DNA
<ol>
	<li>Obtain DNA oligomers of specified sequences (see Table 5) from commercial oligonucleotide manufacturers synthesized at 25 nmol scale using standard desalting. Resuspend these DNA sequences using RNase-free deionized water at 20 &#181;M stock concentrations.</li>
	<li>For contiguous &#947;PNA strand replacements with DNA, sequences D3, D5 and D9 can replace strands p3, p5 and p9. Similarly, for crossover &#947;PNA strand replacements with DNA, sequences D1, D4 and D7 can replace strands p1, p4 and p7.</li>
	<li>Aliquot 1 &#181;L from the 20 &#181;M sub-stocks for each &#947;PNA or DNA oligomer and add it to a 200 &#181;L PCR tube as in step 2.7. Add 30 &#181;L of anhydrous DMSO and 1 &#181;L of RNase-free deionized water to make the final volume to 40 &#181;L (see Table 6).</li>
	<li>Load the anneal batches on to the thermal cycler and anneal using the protocol mentioned in step 4.2.</li>
	<li>Characterize &#947;PNA-DNA hybrid nanostructures using the TIRF protocol following steps in section 5 or using TEM imaging protocol mentioned in section 6 (see Figure 6).</li>
</ol>
8. Different morphologies for &#947;PNA nanofibers in varying concentrations of SDS
<ol>
	<li>Prepare a 20% (wt/v) SDS main stock by measuring 20 mg of SDS and dissolving it in 100 &#181;L of deionized water.</li>
	<li>Prepare a 6% (wt/v) SDS sub-stock by aliquoting 3 &#181;L from the 20% SDS stock and making the volume to 10 &#181;L using deionized water.</li>
	<li>Anneal the &#947;PNA oligomers in final concentrations of 5.25 mM and 17.5 mM SDS as follows. Aliquot 1 &#181;L from the 20 &#181;M sub-stocks for each &#947;PNA oligomer and add it to a 200 &#181;L PCR tube as in step 2.7. Add 30 &#181;L of anhydrous DMSO and 1 &#181;L of 6% and 20% SDS to make the final volume to 40 &#181;L to achieve SDS final concentrations of 5.25 mM and 17.5 mM, respectively (see Table 7).</li>
	<li>Load the anneal batches of varying SDS concentrations on to the thermal cycler and anneal using the protocol mentioned in step 4.2.</li>
	<li>Characterize &#947;PNA nanostructures in the presence of SDS using the TIRF protocol following steps in section 5 or using TEM imaging protocol mentioned in section 6 (see Figure 7).</li>
</ol>

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The authors declare no competing financial interests.

This article focuses on adapting and improving existing nucleic acid nanotechnology protocols towards organic solvent mixtures. The methods described here focus on modifications and troubleshooting within a defined experimental space of select polar aprotic organic solvents. There is yet unexplored potential for other established nucleic acid nanotechnology protocols to be adapted within this space. This could improve potential applications through integration in other fields such as polymer and peptide synthesis which t...

Successful construction of numerous complex nanostructures1,2,3,4,5,6,7,8,9,10,11,12 in aqueous or substantially hydrated media made using naturally occurring nucleic acids such as DNA1,2,3,4,5,6,7,8,9,10 and RNA11,12 has been shown in previous works. However, naturally occurring nucleic acids undergo duplex helical conformational changes or have reduced thermal stabilities in organic solvent mixtures13,14.
Previously, our lab has reported a method towards the construction of 3-helix nanofibers using gamma-position modified synthetic nucleic acid mimics called gamma-peptide nucleic acids (&#947;PNA)15 (Figure 1A). The need for such a development and potential applications of the synthetic nucleic acid mimic PNA has been discussed within the field16,17. We have shown, through an adaptation of the single-stranded tile (SST) strategy presented for DNA nanostructures18,19,20, that 9 sequentially distinct &#947;PNA oligomers can be designed to form 3-helix nanofibers in select polar aprotic organic solvent mixtures such as DMSO and DMF. The &#947;PNA oligomers were commercially ordered with modifications of (R)-diethylene glycol (mini-PEG) at three &#947;-positions (1, 4 and 8 base-positions) along each 12-base oligomer based on methods published by Sahu et al.21 These gamma-modifications cause the helical pre-organization that is associated with the higher binding affinity and thermal stability of &#947;PNA relative to unmodified PNA.
This article is an adaptation of our reported work in which we investigate the effects of solvent solution and substitution with DNA on the formation of &#947;PNA-based nanostructures15. The aim of this article is to provide detailed descriptions of the design as well as detailed protocols for solvent-adapted methods that were developed for the self-assembly and characterization of &#947;PNA nanofiber. Thus, we first introduce the modular SST strategy, a general platform for nanostructure design using the synthetic nucleic acid mimic PNA.
The helical pitch for PNA duplexes has been reported to be 18 bases per turn in comparison to DNA duplexes, which undergo one turn per 10.5 bases (Figure 1B). Therefore, the domain-length of the demonstrated &#947;PNA SSTs was set at 6 bases to accommodate one third of a full turn or 120&#176; of rotation to enable interaction between three triangularly arrayed helices. Also, unlike previous SST motifs, each SST contains just 2 domains, effectively creating a 1-dimensional ribbon-like structure that wraps to form a three-helix bundle (Figure 1C). Each 12-base &#947;PNA oligomer is gamma modified at the 1, 4 and 8 positions to ensure uniformly spaced distribution of mini-PEG groups across the overall SST motif. Additionally, within the motif, there are two types of oligomers: &#8220;contiguous&#8221; strands that exist on a single helix and helix-spanning &#8220;crossover&#8221; strands (Figure 1D). In addition, oligomers P8 and P6 are labelled with fluorescent Cy3 (green star) and biotin (yellow oval), respectively (Figure 1D), to enable detection of structure formation using fluorescence microscopy. Altogether, the SST motif is made of 9 total oligomers to enable the formation of 3-helix nanofibers through programmed complementarity of each individual domain to the corresponding domain on a neighboring oligomer (Figure 1E).

<table>
	<tbody>
		<tr>
			<td>&#947;PNA strands/oligomers</td>
			<td>Trucode Gene Repair Inc.</td>
			<td></td>
			<td>Section 2.1</td>
		</tr>
		<tr>
			<td>UV-Vis Spectrophotometer</td>
			<td>Agilent</td>
			<td>Varian Cary 300</td>
			<td>Section 3.1.2</td>
		</tr>
		<tr>
			<td>Quartz cuvettes</td>
			<td>Starna</td>
			<td>29-Q-10</td>
			<td>Section 3.1.1</td>
		</tr>
		<tr>
			<td>Thermal cycler</td>
			<td>Bio Rad</td>
			<td>C1000 touch</td>
			<td>Section 4.1</td>
		</tr>
		<tr>
			<td>0.2 mL PCR tubes</td>
			<td>VWR</td>
			<td>53509-304</td>
			<td>Section 4.5</td>
		</tr>
		<tr>
			<td>Anhydrous DMF</td>
			<td>VWR</td>
			<td>EM-DX1727-6</td>
			<td>Section 4.6</td>
		</tr>
		<tr>
			<td>Anhydrous DMSO</td>
			<td>VWR</td>
			<td>EM-MX1457-6</td>
			<td>Section 4.6</td>
		</tr>
		<tr>
			<td>Anhydrous 1,4-Dioxane</td>
			<td>Fisher Scientific</td>
			<td>AC615121000</td>
			<td>Section 4.6</td>
		</tr>
		<tr>
			<td>10X Phosphate Buffered Saline (PBS)</td>
			<td>VWR</td>
			<td>75800-994</td>
			<td>Section 3.1.1</td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>VWR</td>
			<td>89085-399</td>
			<td>Section 5.2</td>
		</tr>
		<tr>
			<td>Glass cover slips</td>
			<td>VWR</td>
			<td>48382-126</td>
			<td>Section 5.2</td>
		</tr>
		<tr>
			<td>2% Collodion in Amyl Acetate</td>
			<td>Sigma-Aldrich</td>
			<td>9817</td>
			<td>Section 5.2</td>
		</tr>
		<tr>
			<td>Isoamyl Acetate</td>
			<td>VWR</td>
			<td>200001-180</td>
			<td>Section 5.2</td>
		</tr>
		<tr>
			<td>Biotinylated Bovine Serum Albumin (Biotin-BSA)</td>
			<td>Sigma-Aldrich</td>
			<td>A8549</td>
			<td>Section 5.3</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma-Aldrich</td>
			<td>A2153</td>
			<td>Section 5.4</td>
		</tr>
		<tr>
			<td>Streptavidin</td>
			<td>Sigma-Aldrich</td>
			<td>189730</td>
			<td>Section 5.5</td>
		</tr>
		<tr>
			<td>Trolox</td>
			<td>Sigma-Aldrich</td>
			<td>238813</td>
			<td>Section 5.7</td>
		</tr>
		<tr>
			<td>Total Internal Reflection Fluorescence microscope</td>
			<td>Nikon</td>
			<td>Nikon Ti2-E</td>
			<td>Section 5.8</td>
		</tr>
		<tr>
			<td>Transmission Electron Microscope</td>
			<td>Joel</td>
			<td>JEM 1011</td>
			<td>Section 6.6</td>
		</tr>
		<tr>
			<td>Tweezers</td>
			<td>Dumont</td>
			<td>0203-N5AC-PO</td>
			<td>Section 6.3</td>
		</tr>
		<tr>
			<td>Uranyl Acetate</td>
			<td>Electron Microscopy Sciences</td>
			<td>22400</td>
			<td>Section 6.1</td>
		</tr>
		<tr>
			<td>Formvar, 300 mesh, Copper grids</td>
			<td>Ted Pella Inc.</td>
			<td>1701-F</td>
			<td>Section 6.2</td>
		</tr>
		<tr>
			<td>Formvar-Silicon monoxide Type A, 300 mesh, Copper grids</td>
			<td>Ted Pella Inc.</td>
			<td>1829</td>
			<td>Section 6.2</td>
		</tr>
		<tr>
			<td>DNA oligomers/strands</td>
			<td>IDT</td>
			<td></td>
			<td>Section 7.1</td>
		</tr>
		<tr>
			<td>Sodium Dodecyl Sulphate (SDS)</td>
			<td>VWR</td>
			<td>97064-860</td>
			<td>Section 8.1</td>
		</tr>
	</tbody>
</table>

self-assembly of gamma-modified peptide nucleic acids into complex nanostructures in organic solvent mixtures

Current strategies in DNA and RNA nanotechnology enable the self-assembly of a variety of nucleic acid nanostructures in aqueous or substantially hydrated media. In this article, we describe detailed protocols that enable the construction of nanofiber architectures in organic solvent mixtures through the self-assembly of uniquely addressable, single-stranded, gamma-modified peptide nucleic acid (&#947;PNA) tiles. Each single-stranded tile (SST) is a 12-base &#947;PNA oligomer composed of two concatenated modular domains of 6 bases each. Each domain can bind to a mutually complimentary domain present on neighboring strands using programmed complementarity to form nanofibers that can grow to microns in length. The SST motif is made of 9 total oligomers to enable the formation of 3-helix nanofibers. In contrast with analogous DNA nanostructures, which form diameter-monodisperse structures, these &#947;PNA systems form nanofibers that bundle along their widths during self-assembly in organic solvent mixtures. Self-assembly protocols described here therefore also include a conventional surfactant, Sodium Dodecyl Sulfate (SDS), to reduce bundling effects.

The protocols discussed in the sections above describe the design of an adapted SST motif from DNA nanofibers for the robust generation of self-assembled nanofibers structures using multiple, distinct &#947;PNA oligomers. This section describes the interpretation of data obtained from the successful recreation of the protocols described.
Following the protocol described in section 5 for TIRF imaging of samples of &#947;PNA oligomers annealed in 75% DMSO: H2O (v/v) most readily provi...

Watch this Scientific Journal Video about Self-Assembly of Gamma-Modified Peptide Nucleic Acids into Complex Nanostructures in Organic Solvent Mixtures at JoVE.com

Self-Assembly of Gamma-Modified Peptide Nucleic Acids into Complex Nanostructures in Organic Solvent Mixtures

This article provides protocols for the design and self-assembly of nanostructures from gamma-modified peptide nucleic acid oligomers in organic solvent mixtures.

Current strategies in DNA and RNA nanotechnology enable the self-assembly of a variety of nucleic acid nanostructures in aqueous or substantially hydrated ...

self-assembly-gamma-modified-peptide-nucleic-acids-into-complex

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

4.1K Views.  Carnegie Mellon University. This article provides protocols for the design and self-assembly of nanostructures from gamma-modified peptide nucleic acid oligomers in organic solvent mixtures.

Video: Self-Assembly of Gamma-Modified Peptide Nucleic Acids into Complex Nanostructures in Organic Solvent Mixtures

Fabrication of Electrochemical-DNA Biosensors for the Reagentless Detection of Nucleic Acids, Proteins and Small Molecules

As medicine is currently practiced, doctors send specimens to a central laboratory for testing and thus must wait hours or days to 
receive the results. Many patients would be better served by rapid, bedside tests. To this end our laboratory and others have developed a versatile, reagentless 
biosensor platform that supports the quantitative, reagentless, electrochemical detection of nucleic acids (DNA, RNA), proteins (including antibodies) and small 
molecules analytes directly in unprocessed clinical and environmental samples. In this video, we demonstrate the preparation and use of several biosensors in this 
"E-DNA" class. In particular, we fabricate and demonstrate sensors for the detection of a target DNA sequence in a polymerase chain reaction mixture, an HIV-specific antibody and the drug cocaine. The preparation procedure requires only three hours of hands-on effort followed by an overnight incubation, and their use requires only minutes.

"E-DNA" sensors, reagentless, electrochemical biosensors that perform well even when challenged directly in blood and other complex matrices, have been adapted to the detection of a wide range of nucleic acid, protein and small molecule analytes. Here we present a general procedure for the fabrication and use of such sensors.

As medicine is currently practiced, doctors send specimens to a central laboratory for testing and thus must wait hours or days to 
receive the results. ...

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 ...

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.

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 ...

On-chip Isotachophoresis for Separation of Ions and Purification of Nucleic Acids

Electrokinetic techniques are a staple of microscale applications because of their unique ability to perform a variety of fluidic and electrophoretic processes in simple, compact systems with no moving parts. Isotachophoresis (ITP) is a simple and very robust electrokinetic technique that can achieve million-fold preconcentration1,2 and efficient separation and extraction based on ionic mobility.3 For example, we have demonstrated the application of ITP to separation and sensitive detection of unlabeled ionic molecules (e.g. toxins, DNA, rRNA, miRNA) with little or no sample preparation4-8 and to extraction and purification of nucleic acids from complex matrices including cell culture, urine, and blood.9-12
ITP achieves focusing and separation using an applied electric field and two buffers within a fluidic channel system. For anionic analytes, the leading electrolyte (LE) buffer is chosen such that its anions have higher effective electrophoretic mobility than the anions of the trailing electrolyte (TE) buffer (Effective mobility describes the observable drift velocity of an ion and takes into account the ionization state of the ion, as described in detail by Persat et al.13). After establishing an interface between the TE and LE, an electric field is applied such that LE ions move away from the region occupied by TE ions. Sample ions of intermediate effective mobility race ahead of TE ions but cannot overtake LE ions, and so they focus at the LE-TE interface (hereafter called the "ITP interface"). Further, the TE and LE form regions of respectively low and high conductivity, which establish a steep electric field gradient at the ITP interface. This field gradient preconcentrates sample species as they focus. Proper choice of TE and LE results in focusing and purification of target species from other non-focused species and, eventually, separation and segregation of sample species.
We here review the physical principles underlying ITP and discuss two standard modes of operation: "peak" and "plateau" modes. In peak mode, relatively dilute sample ions focus together within overlapping narrow peaks at the ITP interface. In plateau mode, more abundant sample ions reach a steady-state concentration and segregate into adjoining plateau-like zones ordered by their effective mobility. Peak and plateau modes arise out of the same underlying physics, but represent distinct regimes differentiated by the initial analyte concentration and/or the amount of time allotted for sample accumulation.
We first describe in detail a model peak mode experiment and then demonstrate a peak mode assay for the extraction of nucleic acids from E. coli cell culture. We conclude by presenting a plateau mode assay, where we use a non-focusing tracer (NFT) species to visualize the separation and perform quantitation of amino acids.

Isotachophoresis (ITP) is a robust electrokinetic separation and preconcentration technique with applications ranging from toxin detection to sample preparation. We review the physical principles of ITP and the methodology of applying this technique to two specific example applications: separation and detection of small molecules and purification of nucleic acids from cell culture lysate.

Electrokinetic techniques are a staple of microscale applications because of their unique ability to perform a variety of fluidic and electrophoretic ...

Continuously-stirred Anaerobic Digester to Convert Organic Wastes into Biogas: System Setup and Basic Operation

Anaerobic digestion (AD) is a bioprocess that is commonly used to convert complex organic wastes into a useful biogas with methane as the energy carrier 1-3. Increasingly, AD is being used in industrial, agricultural, and municipal waste(water) treatment applications 4,5. The use of AD technology allows plant operators to reduce waste disposal costs and offset energy utility expenses. In addition to treating organic wastes, energy crops are being converted into the energy carrier methane 6,7. As the application of AD technology broadens for the treatment of new substrates and co-substrate mixtures 8, so does the demand for a reliable testing methodology at the pilot- and laboratory-scale.
Anaerobic digestion systems have a variety of configurations, including the continuously stirred tank reactor (CSTR), plug flow (PF), and anaerobic sequencing batch reactor (ASBR) configurations 9. The CSTR is frequently used in research due to its simplicity in design and operation, but also for its advantages in experimentation. Compared to other configurations, the CSTR provides greater uniformity of system parameters, such as temperature, mixing, chemical concentration, and substrate concentration. Ultimately, when designing a full-scale reactor, the optimum reactor configuration will depend on the character of a given substrate among many other nontechnical considerations. However, all configurations share fundamental design features and operating parameters that render the CSTR appropriate for most preliminary assessments. If researchers and engineers use an influent stream with relatively high concentrations of solids, then lab-scale bioreactor configurations cannot be fed continuously due to plugging problems of lab-scale pumps with solids or settling of solids in tubing. For that scenario with continuous mixing requirements, lab-scale bioreactors are fed periodically and we refer to such configurations as continuously stirred anaerobic digesters (CSADs).
This article presents a general methodology for constructing, inoculating, operating, and monitoring a CSAD system for the purpose of testing the suitability of a given organic substrate for long-term anaerobic digestion. The construction section of this article will cover building the lab-scale reactor system. The inoculation section will explain how to create an anaerobic environment suitable for seeding with an active methanogenic inoculum. The operating section will cover operation, maintenance, and troubleshooting. The monitoring section will introduce testing protocols using standard analyses. The use of these measures is necessary for reliable experimental assessments of substrate suitability for AD. This protocol should provide greater protection against a common mistake made in AD studies, which is to conclude that reactor failure was caused by the substrate in use, when really it was improper user operation 10.

Laboratory-scale anaerobic digesters allow scientists to research new ways of optimizing existing applications of anaerobic biotechnology and to evaluate the methane producing potential of various organic wastes. This article introduces a generalized model for the construction, inoculation, operation, and monitoring of a laboratory-scale continuously stirred anaerobic digester.

Anaerobic digestion (AD) is a bioprocess that is commonly used to convert complex organic wastes into a useful biogas with methane as the energy carrier ...

An Injectable and Drug-loaded Supramolecular Hydrogel for Local Catheter Injection into the Pig Heart

Regeneration of lost myocardium is an important goal for future therapies because of the increasing occurrence of chronic ischemic heart failure and the limited access to donor hearts. An example of a treatment to recover the function of the heart consists of the local delivery of drugs and bioactives from a hydrogel. In this paper a method is introduced to formulate and inject a drug-loaded hydrogel non-invasively and side-specific into the pig heart using a long, flexible catheter. The use of 3-D electromechanical mapping and injection via a catheter allows side-specific treatment of the myocardium. To provide a hydrogel compatible with this catheter, a supramolecular hydrogel is used because of the convenient switching from a gel to a solution state using environmental triggers. At basic pH this ureido-pyrimidinone modified poly(ethylene glycol) acts as a Newtonian fluid which can be easily injected, but at physiological pH the solution rapidly switches into a gel. These mild switching conditions allow for the incorporation of bioactive drugs and bioactive species, such as growth factors and exosomes as we present here in both in vitro and in vivo experiments. The in vitro experiments give an on forehand indication of the gel stability and drug release, which allows for tuning of the gel and release properties before the subsequent application in vivo. This combination allows for the optimal tuning of the gel to the used bioactive compounds and species, and the injection system.

Supramolecular hydrogelators based on ureido-pyrimidinones allow full control over the macroscopic gel properties and the sol&#8211;gel switching behavior using pH. Here, we present a protocol for formulating and injecting such a supramolecular hydrogelator via a catheter delivery system for local delivery directly in relevant areas in the pig heart.

Regeneration of lost myocardium is an important goal for future therapies because of the increasing occurrence of chronic ischemic heart failure and the ...

Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer (SALB) Method

In order to mimic cell membranes, the supported lipid bilayer (SLB) is an attractive platform which enables in vitro investigation of membrane-related processes while conferring biocompatibility and biofunctionality to solid substrates. The spontaneous adsorption and rupture of phospholipid vesicles is the most commonly used method to form SLBs. However, under physiological conditions, vesicle fusion (VF) is limited to only a subset of lipid compositions and solid supports. Here, we describe a one-step general procedure called the solvent-assisted lipid bilayer (SALB) formation method in order to form SLBs which does not require vesicles. The SALB method involves the deposition of lipid molecules onto a solid surface in the presence of water-miscible organic solvents (e.g., isopropanol) and subsequent solvent-exchange with aqueous buffer solution in order to trigger SLB formation. The continuous solvent exchange step enables application of the method in a flow-through configuration suitable for monitoring bilayer formation and subsequent alterations using a wide range of surface-sensitive biosensors. The SALB method can be used to fabricate SLBs on a wide range of hydrophilic solid surfaces, including those which are intractable to vesicle fusion. In addition, it enables fabrication of SLBs composed of lipid compositions which cannot be prepared using the vesicle fusion method. Herein, we compare results obtained with the SALB and conventional vesicle fusion methods on two illustrative hydrophilic surfaces, silicon dioxide and gold. To optimize the experimental conditions for preparation of high quality bilayers prepared via the SALB method, the effect of various parameters, including the type of organic solvent in the deposition step, the rate of solvent exchange, and the lipid concentration is discussed along with troubleshooting tips. Formation of supported membranes containing high fractions of cholesterol is also demonstrated with the SALB method, highlighting the technical capabilities of the SALB technique for a wide range of membrane configurations.

Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer SALB Method

We present an experimental protocol to form a supported lipid bilayer on solid substrates without using lipid vesicles. We demonstrate a one-step method to form a lipid bilayer on silicon dioxide and gold as well as supported membranes with cholesterol-enriched domain for various biological applications.

In order to mimic cell membranes, the supported lipid bilayer (SLB) is an attractive platform which enables in vitro investigation of membrane-related ...

Wet Chemistry and Peptide Immobilization on Polytetrafluoroethylene for Improved Cell-adhesion

Endowing materials surface with cell-adhesive properties is a common strategy in biomaterial research and tissue engineering. This is particularly interesting for already approved polymers that have a long standing use in medicine because these materials are well characterized and legal issues associated with the introduction of newly synthesized polymers may be avoided. Polytetrafluoroethylene (PTFE) is one of the most frequently employed materials for the manufacturing of vascular grafts but the polymer lacks cell adhesion promoting features. Endothelialization, i.e., complete coverage of the grafts inner surface with a confluent layer of endothelial cells is regarded key to optimal performance, mainly by reducing thrombogenicity of the artificial interface.
This study investigates the growth of endothelial cells on peptide-modified PTFE and compares these results to those obtained on unmodified substrate. Coupling with the endothelial cell adhesive peptide Arg-Glu-Asp-Val (REDV) is performed via activation of the fluorin-containing polymer using the reagent sodium naphthalenide, followed by subsequent conjugation steps. Cell culture is accomplished using Human Umbilical Vein Endothelial Cells (HUVECs) and excellent cellular growth on peptide-immobilized material is demonstrated over a two-week period.

Cell-adhesiveness is key to many approaches in biomaterial research and tissue engineering. A step-by-step technique is presented using wet-chemistry for the surface modification of the important polymer PTFE with peptides.

Endowing materials surface with cell-adhesive properties is a common strategy in biomaterial research and tissue engineering. This is particularly ...

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 ...