The authors gratefully acknowledge Neil Whitehouse (WPI, Higgins Machine Shop) for his assistance with CNC machining. Additionally, we would like to thank Adriana Hera (WPI Computing and Communications Center) for her help with MATLAB programming, as well as Kate Beverage and Joseph Cotnoir (WPI Academic Technology Center) for assistance with Camtasia. Sophie Burke and Jacleen Becker (WPI Academic Technology Center) provided supplemental video footage. This work was funded by the National Institutes of Health (R15 HL097332), the UMass Medical School-WPI Pilot Research Initiative, the American Heart Association (undergraduate research fellowship to J.Z.H), and Worcester Polytechnic Institute (Summer Undergraduate Research Fellowship to J.Z.H. and institutional start-up funds to M.W.R.).

1. Cell seeding mold fabrication
Begin by milling a 1/2" thick piece of polycarbonate to create 15, round-bottomed, annular wells with a center post diameter of 2 mm. The milled channels are 6 mm deep and 3.75 mm wide. Clean and dry the polycarbonate mold to remove any plastic debris from the milling process.
Mix polydimethylsiloxane (PDMS) at a 10:1 ratio (w/w) of base to curing agent, degas to remove all air bubbles, and pour onto the polycarbonate mold. Degas again to remove any remaining bubbles, and cure in the oven at 60&deg;C for 4 hours.
Once cured, carefully remove the PDMS template by slowly peeling it away from the polycarbonate, wash with soap and water, and autoclave. Also autoclave a solution of two percent agarose (w/v) dissolved in Dulbecco's modified Eagle medium (DMEM).
Place the PDMS template on a level surface and fill with molten agarose by first pipetting agarose into each of the center post molds, and then pipetting into the space around it. Allow the agarose to solidify (approximately 15 minutes), then invert the mold to release the agarose from the PDMS. Cut excess agarose from around each of the wells, and place the agarose wells into 12-well plates.
2. Cell culture and ring seeding
Add cell culture media (DMEM with 10% FBS and 1% penicillin-streptomycin) around the outside of the agarose mold, without covering the top of the molds. Place the plates in the incubator and allow them to equilibrate for approximately 15 minutes while you prepare the cells.
At 90% confluence, trypsinize rat aortic smooth muscle cells (rSMCs) and resuspend at a concentration of 5x106 cells/ml. Pipette 100 &mu;L of cell suspension into each of the annular agarose wells, using a circular motion to apply cells to each well.
Make sure incubator shelves are level, then place the plates in the incubator and allow them to sit undisturbed for 24 hours. Exchange medium every two days thereafter by aspirating the media from around the well, and re-filling each well until the agarose well is completely submerged.
3. Tissue ring harvesting and thickness measurements
At the conclusion of culture, remove the ring from the agarose mold by sliding it over the top of the center post.
Place the ring in a small Petri dish with phosphate buffered saline (PBS), center the sample, and acquire an image using a digital imaging system.
Use an image analysis program for edge detection and measure the thickness of the ring in 4 positions (top, bottom, left and right) around its circumference. Calculate the mean thickness value (t) for each individual ring, and use this value to calculate the cross-sectional area (2&pi;(t/2)2).
4. Mechanical testing of rings
Set up the tensile testing machine (Instron EPS 1000) in the horizontal position. Attach a 1N &plusmn;1mN load cell and custom thin wire grips (created by bending stainless steel wire).
Mount the ring sample on the two thin wire grips. Extend the grips until a 5 mN tare load is applied to the sample. Enter the cross-sectional area (calculated from thickness measurements) and record the gauge length.
Pre-cycle the rings 8 times between the tare load and 50 kPa stress at a rate of 10 mm/min. After the eighth pre-cycle, pull to failure at 10 mm/min. Failure is noted as a decrease in force by 40% from one measurement to the next. For each ring sample, measure the ultimate tensile stress (UTS) and failure strain and calculate the maximum tangent modulus (MTM) from the acquired data.
5. Tissue ring assembly to fabricate tissue tubes
Custom tube holders are machined from round 5 cm polycarbonate disks with rectangular cutouts (2 cm x 3 cm). Drill threaded holes through these disks to allow two holders to be screwed together. Autoclave the custom tube holders, and then transfer to the biosafety cabinet and place in an empty Petri dish.
Use surgical scissors to cut the ends of 1.9 mm diameter silicone tubes at an angle to create beveled edges, and then autoclave the silicone tubes. While the tubes are being autoclaved, fill a Petri dish with media.
Remove the rings from the agarose wells and place them in the Petri dish with media. Place autoclaved silicone tubes in the same Petri dish to wet them before use.
Use forceps to place a beveled end of a silicone tube into the center of a ring and gently slide the ring onto the silicone tubing. Repeat with the desired number of rings. Slide the rings into contact with one another by gently pushing them successively in both directions along the tube. This method can be used for rings harvested after only one day in culture.
Once rings are mounted, align silicone tubes within the polycarbonate holders and screw the two parts of the holder together. Place the holder in a 100 mm Petri dish and add 55 mL media. Exchange media every 3 days for the duration of culture.
To remove the tissue tubes, release the silicone tubes from the polycarbonate holder and use forceps to slide the tissue tubes off the silicone tube and into a Petri dish filled with PBS.
6. Representative Results:
When the protocol is performed correctly, cells aggregate to form tissue rings with an inner diameter equal to the diameter of the corresponding mold post within 24 hours. The ring edges are usually smooth in appearance and (if cultured on a level surface), are uniform in thickness around the entire circumference. The tissue rings are easy to handle and can be removed from their wells for subsequent mechanical and histological analysis (see in Gwyther et al., 20118). Tissue ring morphology, matrix composition, and mechanical properties vary depending on the number and type of cells seeded.
Representative results from tissue rings made from varying cell types, seeding conditions, and culture length are shown in Table 1. Two mm ID rings created from 5x105 rSMCs exhibited a higher UTS than rings created from our previously published 2 mm rings (made from 6.6x105 rSMCs).8 Similar to rat cells, human smooth muscle cells (hSMC) readily aggregated to form tissue rings, which contained a large amount of collagen after only 14 days in culture (data not shown). Human mesenchymal stem cells (hMSC) also aggregated and formed cohesive rings, but lacked mechanical strength and broke during the initial precycling phases of uniaxial tensile testing.
<table border="1">
	<tbody>
		<tr>
			<td>&nbsp;</td>
			<td>n</td>
			<td>Cell number</td>
			<td>Thickness (mm)</td>
			<td>Length of culture (days)</td>
			<td>UTS (kPa)</td>
			<td>MTM (kPa)</td>
			<td>Failure strain (mm/mm)</td>
		</tr>
		<tr>
			<td>rSMC rings</td>
			<td>6</td>
			<td>660,000</td>
			<td>0.94&plusmn;0.12</td>
			<td>14</td>
			<td>97&plusmn;30</td>
			<td>497&plusmn;91</td>
			<td>0.50&plusmn;0.08</td>
		</tr>
		<tr>
			<td>rSMC rings</td>
			<td>4</td>
			<td>500,000</td>
			<td>0.53&plusmn;0.02</td>
			<td>7</td>
			<td>113&plusmn;8</td>
			<td>189&plusmn;15</td>
			<td>0.88&plusmn;0.05</td>
		</tr>
		<tr>
			<td>hSMC rings</td>
			<td>3</td>
			<td>750,000</td>
			<td>0.51&plusmn;0.05</td>
			<td>14</td>
			<td>160&plusmn;30</td>
			<td>270&plusmn;20</td>
			<td>0.92&plusmn;0.08</td>
		</tr>
		<tr>
			<td>hMSC rings</td>
			<td>3</td>
			<td>750,000</td>
			<td>0.40&plusmn;0.07</td>
			<td>14</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
	</tbody>
</table>
Table 1. Table showing the culture parameters and mechanical properties of 2 mm rings generated from different cell types, seeding concentrations, and culture durations.
<img alt="Figure 1" src="/files/ftp_upload/3366/3366fig1.jpg" /> 
Figure 1. (A) Schematic of the tissue ring generation process. (B) Custom polycarbonate mold with milled annular wells. Central post diameters are 2 mm. (C) PDMS template after it was peeled away from the polycarbonate mold. (D) Aggregated tissue ring cultured in an agarose mold with a 2 mm post. (E) Two mm diameter tissue ring in PBS. Scale bars = 6 mm (B, C) and 2 mm (D, E).
<img alt="Figure 2" src="/files/ftp_upload/3366/3366fig2.jpg" /> 
Figure 2. Representative plot of stress-strain curve produced from uniaxial tensile testing.

<ol>
	<li>Roger, V. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heart+Disease+and+Stroke+Statistics--2011+Update:+A+Report+From+the+American+Heart+Association.">Heart Disease and Stroke Statistics--2011 Update: A Report From the American Heart Association.</a> Circulation. 123, e18-e209 (2011).</li><li>Kelm, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+concept+for+scaffold-free+vessel+tissue+engineering:+self-assembly+of+microtissue+building+blocks.">A novel concept for scaffold-free vessel tissue engineering: self-assembly of microtissue building blocks.</a> J. 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No conflicts of interest declared.

Recently, there has been an increased interest in cell-based or "scaffold-less" tissue engineering methods to address some of the limitations of scaffold-based tissue engineering approaches. Given that cell-derived tissues are created from cells and the matrix they produce, they inherently contain much higher cell densities, contain no exogenous materials, and can be made entirely from human cells and proteins. Vascular grafts made from human cells can attain substantial mechanical strength in the absence of exogenous scaffolds (e.g., burst pressures 3400 mmHg compared to 1600 mmHg for human saphenous veins)12. Although cell-based vascular tissues exhibit improved cell density and mechanical strength, most current fabrication methods (such as "sheet-based engineering"3,4,12 or "bioprinting"5,13) require extensive culture periods (&gt; 3 months) or specialized equipment for 3D tissue construction. The cell-derived tissue ring method described here enables rapid cellular self-assembly to form robust 3D tissue constructs within a short period of time and without the use of specialized equipment.
This protocol details the procedure we developed to create 2 mm inner diameter rat smooth muscle cell-derived tissue rings. In the present example, tissue rings were cultured for 7 days (then cultured for an additional 7 days for ring fusion and tube formation). However, 2 mm rat (and human) smooth muscle cell rings can be removed from the wells and are cohesive enough for handling (e.g., transfer onto silicone tubes) as early as one day after cell seeding. In addition, robust tissue rings with varying inner diameters (2, 4, and 6 mm) can be created with this method by simply changing the post diameter of the original polycarbonate mold.8 We also recently modified the polycarbonate mold design to enable five 2 mm seeding wells to be cast in a single multi-well agarose chamber, which uses less PDMS and agarose, and fits in one well of a 6-well plate (data not shown). Changes in the post diameter, the width of the seeding well, the radius of curvature of the rounded bottom, the number of seeding wells, or the depth of the seeding wells can all be modified simply by changing the specifications in the CAD file for CNC machining of the polycarbonate mold. Finally, a single polycarbonate mold can be used to fabricate an unlimited number of PDMS templates, and each PDMS template can be cleaned, autoclaved and reused dozens of times.
In addition to changing the size of the tissue rings, we have made rings from many different cell types, including: primary rat SMCs (Cell Applications, R354-05), primary human coronary artery SMCs (Lonza, CC-2583), primary human dermal fibroblasts11 (generous gift of Dr. George Pins, WPI Dept. of Biomedical Engineering), rat lung fibroblasts (RFL-6, ATCC CCL-192), and mesenchymal stem cells (Lonza, PT-2501). Each of these cell types aggregates and contracts around the center posts to form tissue rings; although the cellular organization, ECM composition, and mechanical properties of the constructs vary for each cell type. The seeding parameters for each cell type must be empirically determined based on the size of the cells and their ability to aggregate. Therefore, while this system of creating cell-derived tissue rings is extremely versatile, the protocol may need slight adjustments for optimal tissue formation with different cell types.
Tissue ring geometry facilitates easy loading and assessment of tissue material properties by uniaxial tensile testing, as described. There is also substantial precedent for using blood vessel ring segments to measure vascular contraction and physiological function. Preliminary studies indicate that cell-derived tissue rings can be mounted on a wire myograph device for measurement of pharmacological responsiveness and contractile force generation (data not shown). Altogether, the ability to rapidly fabricate self-assembled cell rings for histological, mechanical, physiological, and biochemical analysis suggests a powerful new tool that may be useful for modeling vascular tissue structure and function in health and disease.

<table border="1">
<tbody>
<tr>
	<th>Name of the reagent/equipment</th>
	<th>Company</th>
	<th>Catalog #</th>
</tr>
<tr>
	<td>Polydimethylsiloxane (PDMS)</td>
	<td>Dow Corning</td>
	<td>Sylgard 184</td>
</tr>
<tr>
	<td>Agarose</td>
	<td>Lonza</td>
	<td>50000</td>
</tr>
<tr>
	<td>DMEM</td>
	<td>Mediatech</td>
	<td>15-017-CV</td>
</tr>
<tr>
	<td>Fetal Bovine Serum (FBS)</td>
	<td>PAA</td>
	<td>A05-201</td>
</tr>
<tr>
	<td>Penicillin/Streptomycin</td>
	<td>Mediatech</td>
	<td>30-002-CI</td>
</tr>
<tr>
	<td>Digital imaging system</td>
	<td>DVT Corporation</td>
	<td>Model 630</td>
</tr>
<tr>
	<td>Uniaxial testing machine</td>
	<td>Instron</td>
	<td>ElectroPuls E1000</td>
</tr>
<tr>
	<td>Edge Detection Software	</td>
	<td>DVT Corporation</td>
	<td>Framework 2.4.6</td>
</tr>
</tbody>
</table>

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.

Watch this Scientific Journal Video about Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering at JoVE.com

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

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 ...

directed-cellular-self-assembly-to-fabricate-cell-derived-tissue

Research

JoVE Journal

Bioengineering

18.9K Görüntüleme.  Worcester Polytechnic Institute. Bu makalede, hücresel öz-montaj hücre kökenli doku halkaları oluşturmak için çok yönlü bir yöntem özetlemektedir. Düz kas hücreleri içine numaralı seribaşı halka şeklinde agaroz kuyuları agrega ve 7 gün içinde sağlam bir üç boyutlu (3D) dokular oluşturmak için sözleşme. Milimetre ölçekli doku halkaları mekanik test için elverişli ve doku montaj için yapı taşları olarak hizmet.

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

Electric Field-controlled Directed Migration of Neural Progenitor Cells in 2D and 3D Environments

Endogenous electric fields (EFs) occur naturally in vivo and play a critical role during tissue/organ development and regeneration, including that of the central nervous system1,2. These endogenous EFs are generated by cellular regulation of ionic transport combined with the electrical resistance of cells and tissues. It has been reported that applied EF treatment can promote functional repair of spinal cord injuries in animals and humans3,4. In particular, EF-directed cell migration has been demonstrated in a wide variety of cell types5,6, including neural progenitor cells (NPCs)7,8. Application of direct current (DC) EFs is not a commonly available technique in most laboratories. We have described detailed protocols for the application of DC EFs to cell and tissue cultures previously5,11. Here we present a video demonstration of standard methods based on a calculated field strength to set up 2D and 3D environments for NPCs, and to investigate cellular responses to EF stimulation in both single cell growth conditions in 2D, and the organotypic spinal cord slice in 3D. The spinal cordslice is an ideal recipient tissue for studying NPC ex vivo behaviours, post-transplantation, because the cytoarchitectonic tissue organization is well preserved within these cultures9,10. Additionally, this ex vivo model also allows procedures that are not technically feasible to track cells in vivo using time-lapse recording at the single cell level. It is critically essential to evaluate cell behaviours in not only a 2D environment, but also in a 3D organotypic condition which mimicks the in vivo environment. This system will allow high-resolution imaging using cover glass-based dishes in tissue or organ culture with 3D tracking of single cell migration in vitro and ex vivo and can be an intermediate step before moving onto in vivo paradigms.

This protocol demonstrates methods used to establish 2D and 3D environments in custom-designed electrotactic chambers, which can track cells in vivo/ex vivo using time-lapse recording at the single cell level, in order to investigate galvanotaxis/electrotaxis and other cellular responses to direct current (DC) electric fields (EFs).

Endogenous electric fields (EFs) occur naturally in vivo and play a critical role during tissue/organ development and regeneration, including that of the ...

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Capillary Force Lithography for Cardiac Tissue Engineering

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical discoveries with the aims of developing functional tissues for cardiac regeneration as well as in vitro screening assays. However, the ability to create high-fidelity models of heart tissue has proven difficult. The heart&#8217;s extracellular matrix (ECM) is a complex structure consisting of both biochemical and biomechanical signals ranging from the micro- to the nanometer scale2. Local mechanical loading conditions and cell-ECM interactions have recently been recognized as vital components in cardiac tissue engineering3-5.
A large portion of the cardiac ECM is composed of aligned collagen fibers with nano-scale diameters that significantly influences tissue architecture and electromechanical coupling2. Unfortunately, few methods have been able to mimic the organization of ECM fibers down to the nanometer scale. Recent advancements in nanofabrication techniques, however, have enabled the design and fabrication of scalable scaffolds that mimic the in vivo structural and substrate stiffness cues of the ECM in the heart6-9.
Here we present the development of two reproducible, cost-effective, and scalable nanopatterning processes for the functional alignment of cardiac cells using the biocompatible polymer poly(lactide-co-glycolide) (PLGA)8 and a polyurethane (PU) based polymer. These anisotropically nanofabricated substrata (ANFS) mimic the underlying ECM of well-organized, aligned tissues and can be used to investigate the role of nanotopography on cell morphology and function10-14.
Using a nanopatterned (NP) silicon master as a template, a polyurethane acrylate (PUA) mold is fabricated. This PUA mold is then used to pattern the PU or PLGA hydrogel via UV-assisted or solvent-mediated capillary force lithography (CFL), respectively15,16. Briefly, PU or PLGA pre-polymer is drop dispensed onto a glass coverslip and the PUA mold is placed on top. For UV-assisted CFL, the PU is then exposed to UV radiation (&#955; = 250-400 nm) for curing. For solvent-mediated CFL, the PLGA is embossed using heat (120 &#176;C) and pressure (100 kPa). After curing, the PUA mold is peeled off, leaving behind an ANFS for cell culture. Primary cells, such as neonatal rat ventricular myocytes, as well as human pluripotent stem cell-derived cardiomyocytes, can be maintained on the ANFS2.

In this protocol, we demonstrate the fabrication of biomimetic cardiac cell culture substrata made from two distinct polymeric materials using capillary force lithography. The described methods provide a scalable, cost-effective technique to engineer the structure and function of macroscopic cardiac tissues for in vitro and in vivo applications.

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical ...

Evaluation of Polymeric Gene Delivery Nanoparticles by Nanoparticle Tracking Analysis and High-throughput Flow Cytometry

Non-viral gene delivery using polymeric nanoparticles has emerged as an attractive approach for gene therapy to treat genetic diseases1 and as a technology for regenerative medicine2. Unlike viruses, which have significant safety issues, polymeric nanoparticles can be designed to be non-toxic, non-immunogenic, non-mutagenic, easier to synthesize, chemically versatile, capable of carrying larger nucleic acid cargo and biodegradable and/or environmentally responsive. Cationic polymers self-assemble with negatively charged DNA via electrostatic interaction to form complexes on the order of 100 nm that are commonly termed polymeric nanoparticles. Examples of biomaterials used to form nanoscale polycationic gene delivery nanoparticles include polylysine, polyphosphoesters, poly(amidoamines)s and polyethylenimine (PEI), which is a non-degradable off-the-shelf cationic polymer commonly used for nucleic acid delivery1,3 . Poly(beta-amino ester)s (PBAEs) are a newer class of cationic polymers4 that are hydrolytically degradable5,6 and have been shown to be effective at gene delivery to hard-to-transfect cell types such as human retinal endothelial cells (HRECs)7, mouse mammary epithelial cells8, human brain cancer cells9 and macrovascular (human umbilical vein, HUVECs) endothelial cells10.
A new protocol to characterize polymeric nanoparticles utilizing nanoparticle tracking analysis (NTA) is described. In this approach, both the particle size distribution and the distribution of the number of plasmids per particle are obtained11. In addition, a high-throughput 96-well plate transfection assay for rapid screening of the transfection efficacy of polymeric nanoparticles is presented. In this protocol, poly(beta-amino ester)s (PBAEs) are used as model polymers and human retinal endothelial cells (HRECs) are used as model human cells. This protocol can be easily adapted to evaluate any polymeric nanoparticle and any cell type of interest in a multi-well plate format.

A protocol for nanoparticle tracking analysis (NTA) and high-throughput flow cytometry to evaluate polymeric gene delivery nanoparticles is described. NTA is utilized to characterize the nanoparticle particle size distribution and the plasmid per particle distribution. High-throughput flow cytometry enables quantitative transfection efficacy evaluation for a library of gene delivery biomaterials.

Non-viral gene delivery using polymeric nanoparticles has emerged as an attractive approach for gene therapy to treat genetic diseases1 and as a ...

Tissue Engineering of a Human 3D in vitro Tumor Test System

Cancer is one of the leading causes of death worldwide. Current therapeutic strategies are predominantly developed in 2D culture systems, which inadequately reflect physiological conditions in vivo. Biological 3D matrices provide cells an environment in which cells can self-organize, allowing the study of tissue organization and cell differentiation. Such scaffolds can be seeded with a mixture of different cell types to study direct 3D cell-cell-interactions. To mimic the 3D complexity of cancer tumors, our group has developed a 3D in vitro tumor test system.	
	Our 3D tissue test system models the in vivo situation of malignant peripheral nerve sheath tumors (MPNSTs), which we established with our decellularized porcine jejunal segment derived biological vascularized scaffold (BioVaSc). In our model, we reseeded a modified BioVaSc matrix with primary fibroblasts, microvascular endothelial cells (mvECs) and the S462 tumor cell line. For static culture, the vascular structure of the BioVaSc is removed and the remaining scaffold is cut open on one side (Small Intestinal Submucosa SIS-Muc). The resulting matrix is then fixed between two metal rings (cell crowns).	
	Another option is to culture the cell-seeded SIS-Muc in a flow bioreactor system that exposes the cells to shear stress. Here, the bioreactor is connected to a peristaltic pump in a self-constructed incubator. A computer regulates the arterial oxygen and nutrient supply via parameters such as blood pressure, temperature, and flow rate. This setup allows for a dynamic culture with either pressure-regulated pulsatile or constant flow.	
	In this study, we could successfully establish both a static and dynamic 3D culture system for MPNSTs. The ability to model cancer tumors in a more natural 3D environment will enable the discovery, testing, and validation of future pharmaceuticals in a human-like model.

Tissue Engineering of a Human 3D in vitro Tumor Test System

Methods to create human 3D tumor tissues as test systems are described. These technologies are based on a decellularized Biological Vascularized Scaffold (BioVaSc), primary human cells and a tumor cell line, which can be cultured under static as well as under dynamic conditions in a flow bioreactor.

Cancer is one of the leading causes of death worldwide.  Current therapeutic strategies are predominantly developed in 2D culture  systems, which ...

Self-reporting Scaffolds for 3-Dimensional Cell Culture

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting 3D cellular construct can often be more relevant to studying the molecular events and cell-cell interactions than similar experiments studied in 2D. To create effective 3D cultures with high cell viability throughout the scaffold the culture conditions such as oxygen and pH need to be carefully controlled as gradients in analyte concentration can exist throughout the 3D construct. Here we describe the methods of preparing biocompatible pH responsive sol-gel nanosensors and their incorporation into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds along with their subsequent preparation for the culture of mammalian cells. The pH responsive scaffolds can be used as tools to determine microenvironmental pH within a 3D cellular construct. Furthermore, we detail the delivery of pH responsive nanosensors to the intracellular environment of mammalian cells whose growth was supported by electrospun PLGA scaffolds. The cytoplasmic location of the pH responsive nanosensors can be utilized to monitor intracellular pH (pHi) during ongoing experimentation.

Biocompatible pH responsive sol-gel nanosensors can be incorporated into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds. The produced self-reporting scaffolds can be used for in&#160;situ monitoring of microenvironmental conditions whilst culturing cells upon the scaffold. This is beneficial as the 3D cellular construct can be monitored in real-time without disturbing the experiment.

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting ...

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...

Sandwich-like Microenvironments to Harness Cell/Material Interactions

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. However in vivo, most of the cells are completely surrounded by the extracellular matrix (ECM), resulting in a three-dimensional (3D) distribution of receptors. This may trigger differences in the outside-in signaling pathways and thus in cell behavior.
This article shows that stimulating the dorsal receptors of cells already adhered to a 2D substrate by overlaying a film of a new material (a sandwich-like culture) triggers important changes with respect to standard 2D cultures. Furthermore, the simultaneous excitation of ventral and dorsal receptors shifts cell behavior closer to that found in 3D environments. Additionally, due to the nature of the system, a sandwich-like culture is a versatile tool that allows the study of different parameters in cell/material interactions, e.g., topography, stiffness and different protein coatings at both the ventral and dorsal sides. Finally, since sandwich-like cultures are based on 2D substrates, several analysis procedures already developed for standard 2D cultures can be used normally, overcoming more complex procedures needed for 3D systems.

The following protocol describes the procedure to assemble sandwich-like cultures to be used as an intermediate stage between bi-dimensional (2D) and three-dimensional (3D) cellular environments. The engineered systems can have applications in microscopy, biomechanics, biochemistry and cell biology assays.

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. ...

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and trade one defect for another. Tissue engineering holds the promise to address this increasing demand. However, most tissue engineering strategies fail to generate stable and functional tissue substitutes because of poor vascularization. This paper focuses on an in vivo tissue engineering chamber model of intrinsic vascularization where a perfused artery and a vein either as an arteriovenous loop or a flow-through pedicle configuration is directed inside a protected hollow chamber. In this chamber-based system angiogenic sprouting occurs from the arteriovenous vessels and this system attracts ischemic and inflammatory driven endogenous cell migration which gradually fills the chamber space with fibro-vascular tissue. Exogenous cell/matrix implantation at the time of chamber construction enhances cell survival and determines specificity of the engineered tissues which develop. Our studies have shown that this chamber model can successfully generate different tissues such as fat, cardiac muscle, liver and others. However, modifications and refinements are required to ensure target tissue formation is consistent and reproducible. This article describes a standardized protocol for the fabrication of two different vascularized tissue engineering chamber models in vivo.

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

This is a guideline for constructing in vivo vascularized tissue using a microsurgical arteriovenous loop or a flow-through pedicle configuration inside a tissue engineering chamber. The vascularized tissues generated can be employed for organ regeneration and replacement of tissue defects, as well as for drug testing and disease modeling.

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and ...

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 ...