Financial support was provided to J.M.S. from the NIH Biomechanics in Regenerative Medicine T32 Training Program (2T32EB003392), to Q.J. from the Dowd-ICES Fellowship and to A.W.F. from the NIH Director's New Innovator Award (1DP2HL117750).

1. Fabrication of Master Mold Using Photolithography
<ol>
	<li>The ECM protein nanofibers, nanofabrics and nanostructures to be fabricated are first designed using Computer Aided Design (CAD) software. This CAD file is then transferred to a photomask. The type of photomask will depend on the resolution of the features; with a transparency-based photomask adequate for feature sizes down to ~10 &#956;m. Smaller features &#60;10 &#956;m will require a chrome on glass photomask. All of the nanofibers and nanostructures presented here were fabricated using a transparency-based photomask, and thus were nanometer-scale in thickness but not lateral dimensions. 
	Note: It is important to distinguish which regions of the photomask will be dark (prevent UV light to pass through) and which will be transparent (allow UV light to pass through) as this, along with the type of photoresist (positive or negative), will dictate the final topography of the master mold.</li>
	<li>To begin fabrication of the master mold, dehydrate a 4&#34; silicon wafer by placing it on a hotplate set to 150 &#176;C for 15 min.</li>
	<li>Center the wafer on the vacuum chuck of a spin-coater. Pour SU8-2015 negative photoresist onto the middle of the wafer and continue pouring in concentric circles until about two thirds of the wafer is covered. 
	Note: Keep the bottle of SU8 close to the wafer when pouring to minimize the formation of bubbles.</li>
	<li>Program the spincoater as follows:
	<ul>
		<li>Spread cycle: 500 rpm with an acceleration of 100 rpm/sec for 10 sec.</li>
		<li>Spin cycle: 4,000 rpm with an acceleration of 100 rpm/sec for 30 sec.</li>
	</ul>
	Note: This spincoating recipe will form a photoresist layer that is ~10 &#956;m in thickness. By changing the spinning speed or the SU8 formulation, the thickness can be adjusted.</li>
	<li>Soft bake the wafer by placing it on a hotplate set to 95 &#176;C for 3 min.</li>
	<li>Expose the wafer with UV light through the photomask for a total dosage of 140 mJ/cm2. 
	Note: SU8 is a negative photoresist therefore regions where UV light is able to pass through the photomask will remain after developing and become raised features on the master mold.</li>
	<li>Post exposure bake the wafer by placing it on a hotplate set to 95 &#176;C for 4 min.</li>
	<li>Develop the wafer by placing it in SU8 developer for 3 min. After 3 min, rinse the wafer with isopropyl alcohol. If a white film is produced during the rinsing, the wafer is not fully developed and it should be placed back in the developer for another 30 sec. Rinse again with isopropyl alcohol. Repeat this process until a white film does not form during the isopropyl alcohol rinsing.</li>
	<li>Dry the wafer in a stream of nitrogen and place in a 150 mm petri dish to protect from dust.</li>
</ol>
2. Making the PDMS Stamps
<ol>
	<li>Prepare the PDMS prepolymer by combining the elastomer base and curing agent in a 10:1 w/w ratio. Typically 80 g of base and 8 g of curing agent are used to ensure there is sufficient PDMS to cover the master mold in a 1 cm thick layer.</li>
	<li>Mix and degas the PDMS using a centripetal mixer set to the following:
	<ul>
		<li>Mix: 2,000 rpm for 2 min</li>
		<li>Degas: 2,000 rpm for 2 min.</li>
	</ul>
	</li>
	<li>If a mixer is unavailable mix the PDMS by hand for 10 min using a 10 ml serological pipette. Degas the mixture by placing it in a vacuum desiccator for 30 min to remove bubbles.</li>
	<li>Pour enough PDMS prepolymer over the master mold (patterned silicon wafer) to form a 1 cm thick layer. Cure the PDMS by baking at 65 &#176;C for 4 hr&#160;or at room temperature for 48 hr.</li>
	<li>Once cured, The regions containing the patterns can be cut out to form the PDMS stamps. To distinguish the feature side from the backside of the PDMS stamp, cut a notch out of one of the corners on the backside of the stamp.</li>
</ol>
3. Microcontact Printing of ECM Patterns
<ol>
	<li>Clean 25 mm diameter glass coverslips by sonication in 95% ethanol for 1 hr and then dry in a 65 &#176;C oven.</li>
	<li>Prepare the PIPAAm solution by dissolving PIPAAm in 1-butanol at a concentration of 10% (w/v, typically 1 g in 10 ml).</li>
	<li>Center a glass coverslip on the vacuum chuck of the spincoater and pipette 200 &#956;l of the PIPAAm solution so that the entire glass surface is covered.</li>
	<li>Spincoat the coverslip at 6,000 rpm for 1 min.</li>
	<li>Clean the PDMS stamps by sonication in 50% ethanol for 30 min and then dry under a stream of nitrogen. 
	Note: Drying and subsequent steps should be performed in a biosafety cabinet to maintain sterility for applications where the ECM nanostructures will be used with cells.</li>
	<li>Coat the patterned surface of each PDMS stamp with 200 &#956;l of the protein solution, typically 50 &#956;g/ml in sterile distilled water for FN. Incubate for 1 hr at room temperature. 
	Note: This coating volume is for a 1.5 cm2 PDMS stamp and will need to be adjusted depending on the size of the PDMS stamp, the ECM protein used and the concentration of the ECM protein in solution.</li>
	<li>Wash the PDMS stamps in distilled water to remove excess protein and dry thoroughly under a stream of nitrogen. 
	Note: Any water left on the stamp will trigger the premature dissolution of the PIPAAm coating on the coverslip and prevent proper protein transfer.</li>
	<li>For sterile fabrication, place the PIPAAm-coated coverslips inside a closed petri dish and sterilize using UV exposure, 45 min under the UV light in a biosafety cabinet is sufficient. If sterility is not required this step can be omitted.</li>
	<li>Perform microcontact printing by placing the feature side of the PDMS stamp in contact with the PIPAAm-coated coverslip. If required, use forceps to tap lightly on the back of the stamps to remove any air bubbles and ensure uniform contact.</li>
	<li>After 5 min, peel off the PDMS stamp from the coverslip.</li>
	<li>At this stage, additional ECM proteins can be patterned to create more complex and multicomponent structures. Up to 3 printings have been verified to work with this process, and more may be feasible.</li>
</ol>
4. Release of ECM Nanofibers and Nanostructures
<ol>
	<li>Place the patterned PIPAAm coated coverslip in a 35 mm petri dish and inspect the pattern fidelity using phase contrast microscopy. Depending on the pattern, a CCD camera may be necessary to resolve the features of the pattern. Fluorescence microscopy can also be used to inspect the pattern provided the ECM proteins are fluorescently labeled.</li>
	<li>Add 3 ml of 40 &#176;C distilled water to the petri dish and allow the water to gradually cool.</li>
	<li>The dissolution of the PIPAAm layer and the release of the ECM protein patterns can be monitored using phase contrast microscopy. If the application does not permit the use of optical techniques, the release can be monitored by measuring the solution temperature. Typically, the water is cooled to room temperature, well below the LCST of PIPAAm (32 &#176;C), to ensure the ECM protein nanostructures have been released.</li>
	<li>After release, the nanofibers, nanofabrics and other nanostructures are floating in water. To use them for further applications they need to be manipulated. The exact approach will depend on the experimental objective and may include steps such as immobilizing onto another surface, moving with a micromanipulator system or embedding in a hydrogel.</li>
</ol>

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The authors declare that they have no competing financial interests.

The SIA method presented here mimics cell-mediated matrix assembly and enables the engineering of ECM protein nanofibers and nanostructures with tunable size, organization and composition. While not identical to cell-generated ECM, SIA creates ECM composed of nanoscale protein fibrils20 that undergo reversible folding/unfolding during mechanical strain21 and can bind cells20. This provides a unique capability to build ECM protein materials that recapitulate many properties of the ECM foun...

The extracellular matrix (ECM) in tissues is composed of multifunctional proteins involved in physical and chemical regulation of multiple cell processes including adhesion, proliferation, differentiation, and apoptosis1-3. The ECM is synthesized, assembled, and organized by cells and the constituent protein fibrils have unique compositions, fiber size, geometries and interconnected architectures that vary with tissue type and developmental stage. Recent work has demonstrated that the ECM can provide instructive cues to guide cells to form engineered tissues4, suggesting that recapitulating the ECM in terms of composition and structure might enable the development of biomimetic materials for tissue engineering and biotechnology applications.
A number of fabrication methods have been developed to engineer polymeric scaffolds that can mimic aspects of the ECM in tissues. For example, electrospinning and phase separation have both demonstrated the ability to form porous matrices of fibers with diameters ranging from tens of micrometers down to tens of nanometers5-7. Both techniques have also shown that highly porous matrices of nanofibers can support cell adhesion and infiltration into the scaffold8. However, these approaches are limited in the possible fiber geometries, orientations and 3D architectures that can be created. Electrospinning typically produces scaffolds with either randomly oriented or highly aligned fibers whereas phase separation produces scaffolds with randomly oriented fibers. There are also limitations on the materials, with researchers typically using synthetic polymers, such as poly(&#949;-caprolactone)8 and poly(lactic-co-glycolic acid)9, that are subsequently coated with ECM proteins to promote cell adhesion. Natural biopolymers are also used, including collagen type I10, gelatin11, fibrinogen12, chitosan13, and silk14, but represent only a small subset of the proteins found in native tissue. Most tissues contain a larger milieu of ECM proteins and polysaccharides including fibronectin (FN), laminin (LN), collagen type IV and hyaluronic acid that are difficult or impossible to fabricate nanofibers using existing methods.
To address this challenge, we have focused our research efforts on mimicking the way cells synthesize, assemble and organize ECM protein fibrils in their surroundings. While the specific fibrillogenesis process varies for different ECM proteins, typically a conformational change in the ECM protein molecule is triggered by an enzymatic or receptor-mediated interaction, which exposes cryptic self-assembly sites. Here we use FN as a model system to better understand the fibrillogenesis process. Briefly, FN homodimers bind to integrin receptors on the cell surface via the RGD amino acid sequence in the 10th type III repeat unit. Once bound, the integrins move apart via actomyosin contraction and unfold the FN dimers to expose cryptic self-assembly sites. The exposure of these FN-FN binding sites enables the FN dimers to assemble into an insoluble fibril right on the cell surface15. Work in cell-free systems has demonstrated that cryptic FN-FN binding sites can be revealed through unfolding using denaturants16 or surface tension at an air-liquid-solid interface17-19. However, the FN fibers created by these techniques are restricted to specific fiber sizes and geometries and are typically bound to a surface.
Here we describe an approach termed surface-initiated assembly (SIA) 20 that overcomes these limitations by utilizing protein-surface interactions to create free-standing insoluble nanofibers, nanofabrics (2D sheets) and other nanostructures composed of single or multiple ECM proteins (Figure 1). In this process, ECM proteins are adsorbed from a compact, globular conformation in solution and partially denatured (unfolded) onto a patterned, hydrophobic polydimethylsiloxane (PDMS) stamp. The ECM proteins are then transferred in this state onto a thermally responsive poly(N-isopropylacrylamide) (PIPAAm) surface through microcontact printing22. When hydrated with 40 &#176;C water the PIPAAm remains a solid, but when cooled to 32 &#176;C it passes through a lower critical solution temperature (LCST) where it becomes hydrophilic, swells with water and then dissolves, releasing the assembled ECM nanostructures off of the surface. The SIA method provides control over the dimensions with nanometer-scale precision. By controlling key parameters such as composition, fiber geometry, and architecture, it is possible to recapitulate many properties of the ECM found in vivo and to develop advanced scaffolds for tissue engineering and biotechnology applications.

<table border="0" cellpadding="0" cellspacing="0" width="651">
	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Poly(N-isopropylacrylamide) / PIPAAm</td>
			<td style="width:145px;">Polysciences</td>
			<td style="width:123px;">21458-10</td>
			<td style="width:167px;">40,000 Mw</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Sylgard 184 Silicone kit (PDMS)</td>
			<td style="width:145px;">Dow Corning</td>
			<td style="width:123px;"></td>
			<td style="width:167px;">Mix 10 parts base with 1 part curing agent.&#160;</td>
		</tr>
		<tr height="21">
			<td height="42" style="height:42px;width:216px;">Butanol</td>
			<td style="width:145px;"></td>
			<td style="width:123px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Fibronectin</td>
			<td style="width:145px;">BD biosciences</td>
			<td style="width:123px;">354008</td>
			<td style="width:167px;">Human, 1mg</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Laminin</td>
			<td style="width:145px;">BD biosciences</td>
			<td style="width:123px;">354239</td>
			<td style="width:167px;">Ultrapure, mouse, 1mg</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Negative Photoresist</td>
			<td style="width:145px;">Microchem</td>
			<td style="width:123px;">SU8-2015</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">SU8 Developer</td>
			<td style="width:145px;">Microchem</td>
			<td style="width:123px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="55">
			<td height="55" style="height:55px;width:216px;">Sonicator Branson M3510</td>
			<td style="width:145px;">Branson Ultrasonic Corporation</td>
			<td style="width:123px;">CPN-952-318</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="51">
			<td height="51" style="height:51px;width:216px;">Thinky ARE-250 Mixer</td>
			<td style="width:145px;">Thinky Corporation</td>
			<td style="width:123px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Spincoater</td>
			<td style="width:145px;">Specialty Coating Systems</td>
			<td style="width:123px;">G3P-8</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Glass cover 25mm diameter, No 1.5</td>
			<td style="width:145px;">Fisher Scientific</td>
			<td style="width:123px;">12-545-86</td>
			<td style="width:167px;"></td>
		</tr>
	</tbody>
</table>

ecm protein nanofibers and nanostructures engineered using surface-initiated assembly

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

SIA is capable of engineering ECM protein nanofibers with precise control over fiber dimensions. To demonstrate this, arrays of FN nanofibers with planar dimensions of 50 x 20 &#956;m were patterned onto a PIPAAm coated coverslip (Figure 2A). Upon release, the fibers contracted because they were under an inherent pre-stress when patterned on the PIPAAm surface (Figure 2B). Analysis of the FN nanofibers revealed they were monodisperse pre-release with an average length of 50.19 &#177; 0.4...

Watch this Scientific Journal Video about ECM Protein Nanofibers and Nanostructures Engineered Using Surface-initiated Assembly at JoVE.com

ECM Protein Nanofibers and Nanostructures Engineered Using Surface-initiated Assembly

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

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

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Bioengineering

12.4K Views.  Carnegie Mellon University. A method to obtain nanofibers and complex nanostructures from single or multiple extracellular matrix proteins is described. This method uses protein-surface interactions to create free-standing protein-based materials with tunable composition and architecture for use in a variety of tissue engineering and biotechnology applications.

Video: ECM Protein Nanofibers and Nanostructures Engineered Using Surface-initiated Assembly

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On the end of the microscope's cantilever is a nano-scale probe, which traverses image areas ranging from nanometers to micrometers, measuring the elevation of macromolecules resting on the substrate surface at any given point. Electrostatic forces cause proteins, lipids, and nucleic acids to loosely attach to the substrate in random orientations and permit imaging. The generated data resemble a topographical map, where the macromolecules resolve as three-dimensional particles of discrete sizes (Figure 1) 1,2. Tapping mode AFM involves the repeated oscillation of the cantilever, which permits imaging of relatively soft biomaterials such as DNA and proteins. One of the notable benefits of AFM over other nanoscale microscopy techniques is its relative adaptability to visualize individual proteins and macromolecular complexes in aqueous buffers, including near-physiologic buffered conditions, in real-time, and without staining or coating the sample to be imaged. 
The method presented here describes the imaging of DNA and an immunoadsorbed transcription factor (i.e. the glucocorticoid receptor, GR) in buffered solution (Figure 2). Immunoadsorbed proteins and protein complexes can be separated from the immunoadsorbing antibody-bead pellet by competition with the antibody epitope and then imaged (Figure 2A). This allows for biochemical manipulation of the biomolecules of interest prior to imaging. Once purified, DNA and proteins can be mixed and the resultant interacting complex can be imaged as well. Binding of DNA to mica requires a divalent cation 3,such as Ni2+ or Mg2+, which can be added to sample buffers yet maintain protein activity. Using a similar approach, AFM has been utilized to visualize individual enzymes, including RNA polymerase 4 and a repair enzyme 5, bound to individual DNA strands. These experiments provide significant insight into the protein-protein and DNA-protein biophysical interactions taking place at the molecular level. Imaging individual macromolecular particles with AFM can be useful for determining particle homogeneity and for identifying the physical arrangement of constituent components of the imaged particles. While the present method was developed for visualization of GR-chaperone protein complexes 1,2 and DNA strands to which the GR can bind, it can be applied broadly to imaging DNA and protein samples from a variety of sources.

A tapping mode atomic force microscope (AFM) method for the visualization of plasmid DNA, cytoplasmic proteins, and DNA-protein complexes is described. The method includes alternate approaches for preparing samples for AFM imaging following biochemical manipulation. DNA containing specific protein interacting regions are observed in near-physiologic buffer conditions.

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On ...

Utilization of Plasmonic and Photonic Crystal Nanostructures for Enhanced Micro- and Nanoparticle Manipulation

A method to manipulate the position and orientation of submicron particles nondestructively would be an incredibly useful tool for basic biological research. Perhaps the most widely used physical force to achieve noninvasive manipulation of small particles has been dielectrophoresis(DEP).1 However, DEP on its own lacks the versatility and precision that are desired when manipulating cells since it is traditionally done with stationary electrodes. Optical tweezers, which utilize a three dimensional electromagnetic field gradient to exert forces on small particles, achieve this desired versatility and precision.2 However, a major drawback of this approach is the high radiation intensity required to achieve the necessary force to trap a particle which can damage biological samples.3 A solution that allows trapping and sorting with lower optical intensities are optoelectronic tweezers (OET) but OET's have limitations with fine manipulation of small particles; being DEP-based technology also puts constraint on the property of the solution.4,5
This video article will describe two methods that decrease the intensity of the radiation needed for optical manipulation of living cells and also describe a method for orientation control. The first method is plasmonic tweezers which use a random gold nanoparticle (AuNP) array as a substrate for the sample as shown in Figure 1. The AuNP array converts the incident photons into localized surface plasmons (LSP) which consist of resonant dipole moments that radiate and generate a patterned radiation field with a large gradient in the cell solution. Initial work on surface plasmon enhanced trapping by Righini et al and our own modeling have shown the fields generated by the plasmonic substrate reduce the initial intensity required by enhancing the gradient field that traps the particle.6,7,8 The plasmonic approach allows for fine orientation control of ellipsoidal particles and cells with low optical intensities because of more efficient optical energy conversion into mechanical energy and a dipole-dependent radiation field. These fields are shown in figure 2 and the low trapping intensities are detailed in figures 4 and 5. The main problems with plasmonic tweezers are that the LSP's generate a considerable amount of heat and the trapping is only two dimensional. This heat generates convective flows and thermophoresis which can be powerful enough to expel submicron particles from the trap.9,10 The second approach that we will describe is utilizing periodic dielectric nanostructures to scatter incident light very efficiently into diffraction modes, as shown in figure 6.11 Ideally, one would make this structure out of a dielectric material to avoid the same heating problems experienced with the plasmonic tweezers but in our approach an aluminum-coated diffraction grating is used as a one-dimensional periodic dielectric nanostructure. Although it is not a semiconductor, it did not experience significant heating and effectively trapped small particles with low trapping intensities, as shown in figure 7. Alignment of particles with the grating substrate conceptually validates the proposition that a 2-D photonic crystal could allow precise rotation of non-spherical micron sized particles.10 The efficiencies of these optical traps are increased due to the enhanced fields produced by the nanostructures described in this paper.

Plasmonic tweezers and photonic crystal nanostructures are shown to produce useful enhancements in the efficiency and orientation control of optically trapping micro- and nano-particles.

A method to manipulate the position and orientation of submicron particles nondestructively would be an incredibly useful tool for basic biological ...

Synthesis, Assembly, and Characterization of Monolayer Protected Gold Nanoparticle Films for Protein Monolayer Electrochemistry

Colloidal gold nanoparticles protected with alkanethiolate ligands called monolayer protected gold clusters (MPCs) are synthesized and subsequently incorporated into film assemblies that serve as adsorption platforms for protein monolayer electrochemistry (PME). PME is utilized as the model system for studying electrochemical properties of redox proteins by confining them to an adsorption platform at a modified electrode, which also serves as a redox partner for electron transfer (ET) reactions. Studies have shown that gold nanoparticle film assemblies of this nature provide for a more homogeneous protein adsorption environment and promote ET without distance dependence compared to the more traditional systems modified with alkanethiol self-assembled monolayers (SAM).1-3 In this paper, MPCs functionalized with hexanethiolate ligands are synthesized using a modified Brust reaction4 and characterized with ultraviolet visible (UV-Vis) spectroscopy, transmission electron microscopy (TEM), and proton (1H) nuclear magnetic resonance (NMR). MPC films are assembled on SAM modified gold electrode interfaces by using a "dip cycle" method of alternating MPC layers and dithiol linking molecules. Film growth at gold electrode is tracked electrochemically by measuring changes to the double layer charging current of the system. Analogous films assembled on silane modified glass slides allow for optical monitoring of film growth and cross-sectional TEM analysis provides an estimated film thickness. During film assembly, manipulation of the MPC ligand protection as well as the interparticle linkage mechanism allow for networked films, that are readily adaptable, to interface with redox protein having different adsorption mechanism. For example, Pseudomonas aeruginosa azurin (AZ) can be adsorbed hydrophobically to dithiol-linked films of hexanethiolate MPCs and cytochrome c (cyt c) can be immobilized electrostatically at a carboxylic acid modified MPC interfacial layer. In this report, we focus on the film protocol for the AZ system exclusively. Investigations involving the adsorption of proteins on MPC modified synthetic platforms could further the understanding of interactions between biomolecules and man-made materials, and consequently aid the development of biosensor schemes, ET modeling systems, and synthetic biocompatible materials.5-8

Alkanethiolate stabilized gold colloids known as monolayer protected clusters (MPCs) are synthesized, characterized, and assembled into thin films as an adsorption interface for protein monolayer electrochemistry of simple redox protein like Pseudomonas aeruginosa azurin (AZ) and cytochrome c (cyt c).

Colloidal gold nanoparticles protected with alkanethiolate ligands called monolayer protected gold clusters (MPCs) are synthesized and subsequently ...

Engineered Vascularized Muscle Flap

One of the main factors limiting the thickness of a tissue construct and its consequential viability and applicability in vivo, is the control of oxygen supply to the cell microenvironment, as passive diffusion is limited to a very thin layer. Although various materials have been described to restore the integrity of full-thickness defects of the abdominal wall, no material has yet proved to be optimal, due to low graft vascularization, tissue rejection, infection, or inadequate mechanical properties. This protocol describes a means of engineering a fully vascularized flap, with a thickness relevant for muscle tissue reconstruction. Cell-embedded poly L-lactic acid/poly lactic-co-glycolic acid constructs are implanted around the mouse femoral artery and vein and maintained in vivo for a period of one or two weeks. The vascularized graft is then transferred as a flap towards a full thickness defect made in the abdomen. This technique replaces the need for autologous tissue sacrifications and may enable the use of in vitro engineered vascularized flaps in many surgical applications.

To date, thick tissue defects are typically reconstructed by applying autologous tissue flaps or engineered tissues. In this protocol, we present a new method for engineering vascularized tissue flap bearing an autologous pedicle, to serve as a substitute to autologous flaps.

One of the main factors limiting the thickness of a tissue construct and its consequential viability and applicability in vivo, is the control of oxygen ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

Scaling of Engineered Vascular Grafts Using 3D Printed Guides and the Ring Stacking Method

Coronary artery disease remains a leading cause of death, affecting millions of Americans. With the lack of autologous vascular grafts available, engineered grafts offer great potential for patient treatment. However, engineered vascular grafts are generally not easily scalable, requiring manufacture of custom molds or polymer tubes in order to customize to different sizes, constituting a time-consuming and costly practice. Human arteries range in lumen diameter from about 2.0-38 mm and in wall thickness from about 0.5-2.5 mm. We have created a method, termed the "Ring Stacking Method," in which variable size rings of tissue of the desired cell type, demonstrated here with vascular smooth muscle cells (SMCs), can be created using guides of center posts to control lumen diameter and outer shells to dictate vessel wall thickness. These tissue rings are then stacked to create a tubular construct, mimicking the natural form of a blood vessel. The vessel length can be tailored by simply stacking the number of rings required to constitute the length needed. With our technique, tissues of tubular forms, similar to a blood vessel, can be readily manufactured in a variety of dimensions and lengths to meet the needs of the clinic and patient.

Scalable engineered blood vessels would improve clinical applicability. Using easily sizable 3D-printed guides, rings of vascular smooth muscle were created and stacked into a tubular form, forming a vascular graft. Grafts can be sized to meet the range of human coronary artery dimensions by simply changing the 3D-printed guide size.

Coronary artery disease remains a leading cause of death, affecting millions of Americans. With the lack of autologous vascular grafts available, ...

Fabrication of Custom Agarose Wells for Cell Seeding and Tissue Ring Self-assembly Using 3D-Printed Molds

Engineered tissues are being used clinically for tissue repair and replacement, and are being developed as tools for drug screening and human disease modeling. Self-assembled tissues offer advantages over scaffold-based tissue engineering, such as enhanced matrix deposition, strength, and function. However, there are few available methods for fabricating 3D tissues without seeding cells on or within a supporting scaffold. Previously, we developed a system for fabricating self-assembled tissue rings by seeding cells into non-adhesive agarose wells. A polydimethylsiloxane (PDMS) negative was first cast in a machined polycarbonate mold, and then agarose was gelled in the PDMS negative to create ring-shaped cell seeding wells. However, the versatility of this approach was limited by the resolution of the tools available for machining the polycarbonate mold. Here, we demonstrate that 3D-printed plastic can be used as an alternative to machined polycarbonate for fabricating PDMS negatives. The 3D-printed mold and revised mold design is simpler to use, inexpensive to produce, and requires significantly less agarose and PDMS per cell seeding well. We have demonstrated that the resulting agarose wells can be used to create self-assembled tissue rings with customized diameters from a variety of different cell types. Rings can then be used for mechanical, functional, and histological analysis, or for fabricating larger and more complex tubular tissues.

This protocol describes a platform for fabricating self-assembled tissue rings in variable sizes using a customized 3D-printed plastic mold. PDMS negatives are cured in the 3D-printed mold; then agarose is cast in the cured PDMS negatives. Cells are seeded into the resulting agarose wells where they aggregate into tissue rings.

Engineered tissues are being used clinically for tissue repair and replacement, and are being developed as tools for drug screening and human disease ...

Nondestructive Monitoring of Degradable Scaffold-Based Tissue-Engineered Blood Vessel Development Using Optical Coherence Tomography

Engineered vascular grafts with structural and mechanical properties similar to natural blood vessels are expected to meet the growing demand for arterial bypass. Characterization of the growth dynamics and remodeling process of degradable polymer scaffold-based tissue-engineered blood vessels (TEBVs) with pulsatile stimulation is crucial for vascular tissue engineering. Optical imaging techniques stand out as powerful tools for monitoring vascularization of engineered tissue enabling high-resolution imaging in real-time culture. This paper demonstrates a nondestructive and fast real-time imaging strategy to monitor the growth and remodeling of TEBVs in long-term culture by using optical coherence tomography (OCT). Geometric morphology is evaluated, including vascular remodeling process, wall thickness, and comparison of TEBV thickness in different culture time points and presence of pulsatile stimulation. Finally, OCT provides practical possibilities for real-time observation of the degradation of polymer in the reconstructing tissues under pulsatile stimulation or not and in each vessel segment, by compared with the assessment of polymer degradation using scanning electron microscopic(SEM) and polarized microscope.

A step by step protocol for nondestructive and long-period monitoring the process of vascular remodeling and scaffold degradation in real-time culture of biodegradable polymeric scaffold-based tissue-engineered blood vessels with pulsatile stimulation using optical coherence tomography is described here.

Engineered vascular grafts with structural and mechanical properties similar to natural blood vessels are expected to meet the growing demand for arterial ...

Tissue-Engineered Graft for Circumferential Esophageal Reconstruction in Rats

The use of biocompatible materials for circumferential esophageal reconstruction is a technically challenging task in rats and requires an optimal implant technique with nutritional support. Recently, there have been many attempts at esophageal tissue engineering, but the success rate has been limited due to difficulty in early epithelization in the special environment of peristalsis. Here, we developed an artificial esophagus that can improve the regeneration of the esophageal mucosa and muscle layers through a two-layered tubular scaffold, a mesenchymal stem cell-based bioreactor system, and a bypass feeding technique with modified gastrostomy. The scaffold is made of polyurethane (PU) nanofibers in a cylindrical shape with a three-dimensional (3D) printed polycaprolactone strand wrapped around the outer wall. Prior to transplantation, human-derived mesenchymal stem cells were seeded into the lumen of the scaffold, and bioreactor cultivation was performed to enhance cellular reactivity. We improved the graft survival rate by applying surgical anastomosis and covering the implanted prosthesis with a thyroid gland flap, followed by temporary nonoral gastrostomy feeding. These grafts were able to recapitulate the findings of initial epithelialization and muscle regeneration around the implanted sites, as demonstrated by histological analysis. In addition, increased elastin fibers and neovascularization were observed in the periphery of the graft. Therefore, this model presents a potential new technique for circumferential esophageal reconstruction.

Esophageal reconstruction is a challenging procedure, and development of a tissue-engineered esophagus that enables regeneration of esophageal mucosa and muscle and that can be implanted as an artificial graft is necessary. Here, we present our protocol to generate an artificial esophagus, including scaffold manufacturing, bioreactor cultivation, and various surgical techniques.

The use of biocompatible materials for circumferential esophageal reconstruction is a technically challenging task in rats and requires an optimal implant ...

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.

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 ...