We would like to thank John Wirth of the Olbrich Gardens for graciously supplying the specimens used in this project. This work is supported in part by the National Heart, Lung, and Blood Institute (R01HL115282 to G.R.G.) National Science Foundation (DGE1144804 to J.R.G and G.R.G.), and the University of Wisconsin Department of Surgery and Alumni Fund (H.D.L.). This work was also supported in part by the Environmental Protection Agency (STAR grant no. 83573701), the National Institutes of Health (R01HL093282-01A1 and UH3TR000506), and the National Science Foundation (IGERT DGE1144804).

1. Decellularization of Plant Tissue Using the Detergent-based Approach
<ol>
	<li>Use fresh or frozen F. hispida, leaf samples. Freeze unused fresh samples in a -20 &#176;C freezer and store for future use (up to a year). 
	NOTE: Use stem or leaf tissue of nearly any desired plant. Extended storage times can cause damage to the tissues.
	<ol>
		<li>Determine the size and shape of samples to be processed on the basis of the sample&#8217;s intended use (i.e. samples cut into strips are well suited for mechanical testing applications, meanwhile 8 mm disc samples are useful in multi-well culturing applications). Cut the leaf into 8 mm discs with a sharp, clean biopsy punch while submerged under room temperature (20-25 &#176;C) deionized H2O. 
		Note: This protocol can be used on whole leaves and stems. However, smaller samples will decellularize faster.</li>
		<li>Incubate the samples for 5-10 min at room temperature (20-25 &#176;C) deionized H2O on a shake plate set to a low speed setting to wash and/or thaw them. Use enough deionized H2O to ensure all samples are thoroughly wetted.</li>
	</ol>
	</li>
	<li>Prepare a solution of 10% (w/v) sodium dodecyl sulfate (SDS) in deionized H2O. Place the samples in a suitable container (a glass or plastic dish is ideal) and add SDS solution to completely cover the samples. Incubate samples for 5 days at room temperature (20-25 &#176;C) on a shake plate set to a low speed to prevent damage to the samples. 
	NOTE: Do not overcrowd the container as this can slow down the decellularization process and lead to uneven treatment by the SDS. During this step, samples should acquire a brown hue.
	<ol>
		<li>After 5 days, replace the SDS solution with deionized H2O. Incubate the samples for an additional 10-15 min on the shake plate to thoroughly rinse off any residual SDS solution.</li>
	</ol>
	</li>
	<li>Prepare a 1% (v/v) non-ionic surfactant in 10% (v/v) bleach solution. To make a 500 mL solution, mix 5 mL of the non-ionic surfactant with 50 mL of bleach then add 445 mL of deionized H2O. Submerge samples in the freshly prepared solution. 
	NOTE: The non-ionic surfactant/bleach solution does not have a long shelf life, therefore, should be used within 48 h of preparation.</li>
	<li>Replace the non-ionic surfactant/bleach solution every 24 h until the samples are completely cleared (refer to Figure 1A for visual comparison). Then incubate the sample in deionized water on a shake plate for 2 min to rinse off the excess non-ionic surfactant/bleach solution. Set the shake plate to a low setting to prevent damaging the samples. 
	NOTE: The time required to clear the samples used varies, depending on the type and species of the plant. The complete discoloring of the samples is indicative of full decellularization (perform DNA quantification to ensure thorough clearing).
	<ol>
		<li>Lyophilize the samples to store them up to a year (flash freezing using liquid nitrogen is preferred, freezing samples in -80 &#176;C is also acceptable) and store them at room temperature (20-25 &#176;C) in low humidity.</li>
		<li>Reconstitute samples in Tris-HCl (10 mM, pH 8.5) using enough to coat samples. Rinse samples 2-3 times in serum-free media gently using a micropipette before use (i.e. use 150-300 &#181;L per rinse, in a standard 24 well plate). 
		NOTE: Tris-HCl buffer and serum-free media (i.e. DMEM, but any basic cell culture media can be substituted) are more effective in lowering the impact to cell viability of SDS treated samples than those treated with deionized H2O alone.</li>
	</ol>
	</li>
</ol>
2. Preparation of Samples Using the Detergent-free Decellularization Approach
NOTE: The initial steps of this procedure coincide with steps 1.1-1.1.2 (see above).
<ol>
	<li>Prepare a 5% (v/v) bleach (NaClO) and 3% (w/v) sodium bicarbonate (NaHCO3) solution. Warm the solution in a fume hood to 60-70 &#176;C for F. hispida leaf samples cut into 8 mm discs while stirring on a hot plate. 
	NOTE: The sodium bicarbonate can be substituted with sodium carbonate (Na2CO3), or sodium hydroxide (NaOH). The desired temperature varies greatly (from room temperature to 90 &#176;C) and should be adjusted according to the properties of the samples used.</li>
	<li>Once the solution reaches the desired temperature range, submerge the samples and reduce the stirring speed to avoid damaging them. After the samples are visibly cleared (refer to Figure 1B for visual comparison), remove from the bath carefully. Incubate samples once in deionized H2O for 1-2 min to remove excess bleach solution. 
	NOTE: The time required to clear the samples can vary widely. For example, cut parsley can be cleared in 10-15 min, while thicker and/or larger samples such as whole leaves or stems can take hours in high-temperature baths to clear completely.</li>
	<li>Lyophilize the samples and store them at room temperature (20-25 &#176;C) in low humidity.</li>
</ol>

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The authors have nothing to disclose.

Herein, two methods to decellularize plant tissues are described. The results presented here, coupled with the results of prior studies25, suggest that the protocols put forth are likely applicable to a wide spectrum of plant species and can be performed on both stems and leaves. These procedures are simple and do not require specialized equipment, so plant decellularization can be carried out in most laboratories. It is noteworthy that after decellularization, the scaffolds must be functionalized...

Tissue engineering emerged in the 1980s to create living tissue substitutes, and potentially address significant organ and tissue shortages1. One strategy has used scaffolds to stimulate and guide the body to regenerate missing tissues or organs. Although advanced manufacturing approaches such as 3-D printing have produced scaffolds with unique physical properties, the ability to manufacture scaffolds with a diverse range of achievable physical and biological properties remains a challenge2,3. Moreover, due to a lack of a functional vascular network, these techniques have been limited in regenerating 3-dimensional tissues. The use of decellularized animal and human tissues as scaffolds has aided in circumventing this problem4,5,6,7. However, high cost, batch-to-batch variability, and limited availability may limit widespread use of decellularized animal scaffolds8. There are also concerns about potential disease transmission to patients and immunologic reaction to some decellularized mammalian tissues9.
Cellulose, derived from plant and bacterial sources, has been extensively used to generate biomaterials for a wide range of applications in regenerative medicine. Some examples include: bone10,11, cartilage12,13,14 and wound healing15. Scaffolds that are comprised of cellulose have an added benefit in that they are durable and resistant to being broken down by mammalian cells. This is due to the fact that mammalian cells do not produce the enzymes necessary to break down cellulose molecules. In comparison, scaffolds produced using macromolecules from the extracellular matrix, such as collagen, are readily broken down16 and may not be well suited to long-term applications. Collagen scaffolds can be stabilized by chemical cross-linking. However, there is a trade-off due to the inherent toxicity of the cross-linkers that affect the biocompatibility of the scaffolds17. Conversely, cellulose has the potential to remain present at the site of implantation for prolonged periods of time because it is impervious to enzymatic degradation by mammalian cells18,19,20. This can be altered by tuning the rate of degradation through hydrolysis pretreatment and co-delivery of the scaffolds with cellulases21. The biocompatibility of decellularized plant-derived cellulose scaffolds in vivo has also been demonstrated in a study done on mice22.
Through hundreds of millions of years of evolution, plants have refined their structure and composition to increase the efficiency of fluid transport and retention. Plant vascular vessels minimize hydraulic resistance by branching into smaller vessels, similar to the mammalian vasculature according to Murray&#39;s law23. After decellularization, the plant&#39;s complex network of vessels and interconnected pores is maintained. Considering the vast number of distinct plant species readily available, plant-derived scaffolds have the potential to overcome design limitations currently affecting scaffolds in tissue engineering24,25. For instance, Modulevsky et al. demonstrated that angiogenesis and cell migration occurred when decellularized apple tissue was implanted subcutaneously on the back of a mouse22. Similarly, Gershlak et al. showed that endothelial cells could be grown within the vasculature of decellularized leaves24. In a separate experiment, Gershlak et al. were also able to show that cardiomyocytes could be grown on the surface of leaves and were able to contract24.
Plants also include complex organization from the cellular to the macroscopic scale, which is difficult to achieve even with the most advanced manufacturing techniques developed to date. The complex hierarchical design of plant tissues makes them stronger than the sum of their constituents26. Plants possess a plethora of different mechanical properties ranging from rigid and tough components such as stems, to much more flexible and pliable ones such as leaves27. Leaves vary depending on species in terms of size, shape, break strength, the degree of vascularization, and can carry different degrees of hydrophilicity. Overall, these plant properties suggest that decellularized plants can serve as unique and highly functional medical devices, including as tissue engineering scaffolds.
This protocol focuses on two methods to decellularize plant tissues, such as leaves and stems, for use as scaffolds in tissue engineering. The first method is a detergent-based technique that uses a series of baths to remove DNA and cellular matter, which has been adapted from a widely used technique to decellularize mammalian and plant tissues6,22,25,28,29,30. The second method is detergent-free and is adapted from a &#34;skeletonization&#34; protocol generally used to remove the soft tissues of leaves31. Prior work showed that simmering leaves in a bleach and sodium bicarbonate solution facilitated separation of the vasculature from the surrounding soft tissue31. This technique can be cited back to experiments carried out in the 17th and 18th centuries, such as the work of Albertus Seba32 and Edward Parrish33. These experiments centered around leaving plant matter, such as leaves and fruit, submerged in water for extended periods of time (weeks to months) and allowing the softer tissues to decay away naturally. Here the &#34;skeletonization&#34; approach is adapted to use milder conditions, such as longer incubation times at lower temperatures, to remove cellular residues and to avoid significantly disrupting the soft tissue structure. For the experiments detailed herein, three plant types were used: Ficus hispida, Pachira aquatica and a species of Garcinia. Results of DNA quantification, mechanical tests, and impact on cellular metabolic activity from both methods are described.

<table border="0" cellpadding="0" cellspacing="0" style="width:771px;" width="771">
	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:247px;">Sodium dodecyl sulfate</td>
			<td style="width:111px;">Sigma Life Science</td>
			<td style="width:136px;">75746-1KG</td>
			<td style="width:277px;"></td>
		</tr>
		<tr height="85">
			<td height="85" style="height:85px;width:247px;">Triton X-100</td>
			<td style="width:111px;">MP Biomedicals, LLC</td>
			<td style="width:136px;">807426</td>
			<td style="width:277px;">Non-ionic surfactant referenced in paper. Very viscous reagent, can help to cut end of pipette tip when drawing it up.</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:247px;">Concentrated bleach (8.25% sodium hypochlorite)</td>
			<td style="width:111px;">Clorox</td>
			<td style="width:136px;">Item #: 31009</td>
			<td style="width:277px;">Standard concentrated bleach.</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:247px;">Sodium bicarbonate</td>
			<td style="width:111px;">Acros Organics</td>
			<td style="width:136px;">217120010</td>
			<td style="width:277px;">Can be substituted with sodium hydroxide or sodium carbonate.</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:247px;">8 mm Biopunch</td>
			<td style="width:111px;">HealthLink</td>
			<td style="width:136px;">15111-80</td>
			<td style="width:277px;">Cuts samples that fit well in 24 well plate</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:247px;">Belly Dancer-Shake table</td>
			<td style="width:111px;">Stovall Life Sciences</td>
			<td style="width:136px;">BDRAA115S</td>
			<td style="width:277px;">Use low speeds to not damage tissues. Can use any model/brand of shake table.</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:247px;">Isotemp hot/stir plate</td>
			<td style="width:111px;">Fisher Scientific</td>
			<td style="width:136px;"></td>
			<td style="width:277px;">Can use any style/brand of hot/stir plate.</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:247px;">Beaker</td>
			<td style="width:111px;">Any</td>
			<td style="width:136px;"></td>
			<td style="width:277px;">Can use any size beaker as long as it will fit your samples and not overcrowd them.</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:247px;">Tris Hydrochloride</td>
			<td style="width:111px;">Fisher Scientific</td>
			<td style="width:136px;">BP153-500</td>
			<td style="width:277px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">DMEM</td>
			<td style="width:111px;">Corning</td>
			<td style="width:136px;">MT50003PC</td>
			<td style="width:277px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:247px;">Quant-iT Picogreen dsDNA assay</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:136px;">P11496</td>
			<td style="width:277px;">Can use any dsDNA quantification mehtod on hand.</td>
		</tr>
	</tbody>
</table>

two methods for decellularization of plant tissues for tissue engineering applications

The autologous, synthetic, and animal-derived grafts currently used as scaffolds for tissue replacement have limitations due to low availability, poor biocompatibility, and cost. Plant tissues have favorable characteristics that make them uniquely suited for use as scaffolds, such as high surface area, excellent water transport and retention, interconnected porosity, preexisting vascular networks, and a wide range of mechanical properties. Two successful methods of plant decellularization for tissue engineering applications are described here. The first method is based on detergent baths to remove cellular matter, which is similar to previously established methods used to clear mammalian tissues. The second is a detergent-free method adapted from a protocol that isolates leaf vasculature and involves the use of a heated bleach and salt bath to clear the leaves and stems. Both methods yield scaffolds with comparable mechanical properties and low cellular metabolic impact, thus allowing the user to select the protocol which better suits their intended application.

Both methods yielded scaffolds that were suitable for cell culture and tissue engineering applications. Figure 1 shows the general workflow of the decellularization process using an intact leaf for the detergent-based method and cut samples (8 mm diameter) for the detergent-free method. Successful decellularization of Ficus hispida tissues following both methods yielded clear and intact samples (Figure 1A and 1B<...

Watch this Scientific Journal Video about Two Methods for Decellularization of Plant Tissues for Tissue Engineering Applications at JoVE.com

Two Methods for Decellularization of Plant Tissues for Tissue Engineering Applications

Here we present, and contrast two protocols used to decellularize plant tissues: a detergent-based approach and a detergent-free approach. Both methods leave behind the extracellular matrix of the plant tissues used, which can then be utilized as scaffolds for tissue engineering applications.

The autologous, synthetic, and animal-derived grafts currently used as scaffolds for tissue replacement have limitations due to low availability, poor ...

two-methods-for-decellularization-plant-tissues-for-tissue

Research

JoVE Journal

Bioengineering

14.4K Views.  University of Wisconsin-Madison. Here we present, and contrast two protocols used to decellularize plant tissues: a detergent-based approach and a detergent-free approach. Both methods leave behind the extracellular matrix of the plant tissues used, which can then be utilized as scaffolds for tissue engineering applications.

Video: Two Methods for Decellularization of Plant Tissues for Tissue Engineering Applications

Fabrication of Biologically Derived Injectable Materials for Myocardial Tissue Engineering

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications. Briefly, decellularized tissue is lyophilized, milled, enzymatically digested, and then brought to physiological pH. The lyophilization removes all water content from the tissue, resulting in dry ECM that can be ground into a fine powder with a small mill. After milling, the ECM powder is digested with pepsin to form an injectable matrix. After adjustment to pH 7.4, the liquid matrix material can be injected into the myocardium. Results of previous characterization assays have shown that matrix gels produced from decellularized pericardial and myocardial tissue retain native ECM components, including diverse proteins, peptides and glycosaminoglycans. Given the use of this material for tissue engineering, in vivo characterization is especially useful; here, a method for performing an intramural injection into the left ventricular (LV) free wall is presented as a means of analyzing the host response to the matrix gel in a small animal model. Access to the chest cavity is gained through the diaphragm and the injection is made slightly above the apex in the LV free wall. The biologically derived scaffold can be visualized by biotin-labeling before injection and then staining tissue sections with a horse radish peroxidase-conjugated neutravidin and visualizing via diaminobenzidine (DAB) staining. Analysis of the injection region can also be done with histological and immunohistochemical staining. In this way, the previously examined pericardial and myocardial matrix gels were shown to form fibrous, porous networks and promote vessel formation within the injection region.

Methods for preparing an injectable matrix gel from decellularized tissue and injecting it into rat myocardium in vivo are described.

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications.  ...

Decellularization and Recellularization of Whole Livers

The liver is a complex organ which requires constant perfusion for delivery of nutrients and oxygen and removal of waste in order to survive1. Efforts to recreate or mimic the liver microstructure with grounds up approach using tissue engineering and microfabrication techniques have not been successful so far due to this design challenge. In addition, synthetic biomaterials used to create scaffolds for liver tissue engineering applications have been limited in inducing tissue regeneration and repair in large part due to the lack of specific cell binding motifs that would induce the proper cell functions2. Decellularized native tissues such blood vessels3and skin4on the other hand have found many applications in tissue engineering, and have provided a practical solution to some of the challenges. The advantage of decellularized native matrix is that it retains, to an extent, the original composition, and the microstructure, hence enhancing cell attachment and reorganization5.
In this work we describe the methods to perform perfusion-decellularization of the liver, such that an intact liver bioscaffold that retains the structure of major blood vessels is obtained. Further, we describe methods to recellularize these bioscaffolds with adult primary hepatocytes, creating a liver graft that is functional in vitro, and has the vessel access necessary for transplantation in vivo.

Perfusion decellularization is a novel technique to produce whole liver scaffolds that retains the organ's extracellular matrix composition and microarchitecture. Herein, the method of preparing whole organ scaffolds using perfusion decellularization and subsequent repopulation with hepatocytes is described. Functional and transplantable liver grafts can be generated using this technique.

The liver is a complex organ which requires constant perfusion for delivery of nutrients and oxygen and removal of waste in order to survive1. Efforts to ...

Elastomeric PGS Scaffolds in Arterial Tissue Engineering

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and increasing obesity1. Currently, bypass surgery using autologous vessels, allografts, and synthetic grafts are known as a commonly used for arterial substitutes2. However, these grafts have limited applications when an inner diameter of arteries is less than 6 mm due to low availability, thrombotic complications, compliance mismatch, and late intimal hyperplasia3,4. To overcome these limitations, tissue engineering has been successfully applied as a promising alternative to develop small-diameter arterial constructs that are nonthrombogenic, robust, and compliant. Several previous studies have developed small-diameter arterial constructs with tri-lamellar structure, excellent mechanical properties and burst pressure comparable to native arteries5,6. While high tensile strength and burst pressure by increasing collagen production from a rigid material or cell sheet scaffold, these constructs still had low elastin production and compliance, which is a major problem to cause graft failure after implantation. Considering these issues, we hypothesized that an elastometric biomaterial combined with mechanical conditioning would provide elasticity and conduct mechanical signals more efficiently to vascular cells, which increase extracellular matrix production and support cellular orientation.
The objective of this report is to introduce a fabrication technique of porous tubular scaffolds and a dynamic mechanical conditioning for applying them to arterial tissue engineering. We used a biodegradable elastomer, poly (glycerol sebacate) (PGS)7 for fabricating porous tubular scaffolds from the salt fusion method. Adult primary baboon smooth muscle cells (SMCs) were seeded on the lumen of scaffolds, which cultured in our designed pulsatile flow bioreactor for 3 weeks. PGS scaffolds had consistent thickness and randomly distributed macro- and micro-pores. Mechanical conditioning from pulsatile flow bioreactor supported SMC orientation and enhanced ECM production in scaffolds. These results suggest that elastomeric scaffolds and mechanical conditioning of bioreactor culture may be a promising method for arterial tissue engineering.

Elastomeric PGS scaffolds with vascular smooth muscle cells cultured in a pulsatile flow bioreactor may lead to promising small-diameter arterial constructs with native ECM production in a relatively short culture period.

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and ...

Postproduction Processing of Electrospun Fibres for Tissue Engineering

Electrospinning is a commonly used and versatile method to produce scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo undergo biaxial distension to varying extents such as skin, bladder, pelvic floor and even the hard palate as children grow. In producing scaffolds for these purposes there is a need to develop scaffolds of appropriate biomechanical properties (whether achieved without or with cells) and which are sterile for clinical use. The focus of this paper is not how to establish basic electrospinning parameters (as there is extensive literature on electrospinning) but on how to modify spun scaffolds post production to make them fit for tissue engineering purposes - here thickness, mechanical properties and sterilisation (required for clinical use) are considered and we also describe how cells can be cultured on scaffolds and subjected to biaxial strain to condition them for specific applications.
Electrospinning tends to produce thin sheets; as the electrospinning collector becomes coated with insulating fibres it becomes a poor conductor such that fibres no longer deposit on it. Hence we describe approaches to produce thicker structures by heat or vapour annealing increasing the strength of scaffolds but not necessarily the elasticity. Sequential spinning of scaffolds of different polymers to achieve complex scaffolds is also described. Sterilisation methodologies can adversely affect strength and elasticity of scaffolds. We compare three methods for their effects on the biomechanical properties on electrospun scaffolds of poly lactic-co-glycolic acid (PLGA).
Imaging of cells on scaffolds and assessment of production of extracellular matrix (ECM) proteins by cells on scaffolds is described. Culturing cells on scaffolds in vitro can improve scaffold strength and elasticity but the tissue engineering literature shows that cells often fail to produce appropriate ECM when cultured under static conditions. There are few commercial systems available that allow one to culture cells on scaffolds under dynamic conditioning regimes - one example is the Bose Electroforce 3100 which can be used to exert a conditioning programme on cells in scaffolds held using mechanical grips within a media filled chamber.4 An approach to a budget cell culture bioreactor for controlled distortion in 2 dimensions is described. We show that cells can be induced to produce elastin under these conditions. Finally assessment of the biomechanical properties of processed scaffolds cultured with or without cells is described.

Electrospun scaffolds can be processed post production for tissue engineering applications. Here we describe methods for spinning complex scaffolds (by consecutive spinning), for making thicker scaffolds (by multi-layering using heat or vapour annealing), for achieving sterility (aseptic production or sterilisation post production) and for achieving appropriate biomechanical properties.

Electrospinning is a commonly used and versatile method to produce  scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo ...

Engineering Skeletal Muscle Tissues from Murine Myoblast Progenitor Cells and Application of Electrical Stimulation

Engineered muscle tissues can be used for several different purposes, which include the production of tissues for use as a disease model in vitro, e.g. to study pressure ulcers, for regenerative medicine and as a meat alternative 1. The first reported 3D muscle constructs have been made many years ago and pioneers in the field are Vandenburgh and colleagues 2,3. Advances made in muscle tissue engineering are not only the result from the vast gain in knowledge of biochemical factors, stem cells and progenitor cells, but are in particular based on insights gained by researchers that physical factors play essential roles in the control of cell behavior and tissue development. State-of-the-art engineered muscle constructs currently consist of cell-populated hydrogel constructs. In our lab these generally consist of murine myoblast progenitor cells, isolated from murine hind limb muscles or a murine myoblast cell line C2C12, mixed with a mixture of collagen/Matrigel and plated between two anchoring points, mimicking the muscle ligaments. Other cells may be considered as well, e.g. alternative cell lines such as L6 rat myoblasts 4, neonatal muscle derived progenitor cells 5, cells derived from adult muscle tissues from other species such as human 6 or even induced pluripotent stem cells (iPS cells) 7. Cell contractility causes alignment of the cells along the long axis of the construct 8,9 and differentiation of the muscle progenitor cells after approximately one week of culture. Moreover, the application of electrical stimulation can enhance the process of differentiation to some extent 8. Because of its limited size (8 x 2 x 0.5 mm) the complete tissue can be analyzed using confocal microscopy to monitor e.g. viability, differentiation and cell alignment. Depending on the specific application the requirements for the engineered muscle tissue will vary; e.g. use for regenerative medicine requires the up scaling of tissue size and vascularization, while to serve as a meat alternative translation to other species is necessary.

Engineered muscle tissue has great potential in regenerative medicine, as disease model and also as an alternative source for meat. Here we describe the engineering of a muscle construct, in this case from mouse myoblast progenitor cells, and the stimulation by electrical pulses.

Engineered muscle tissues can be used for several different purposes, which include the production of tissues for use as a disease model in vitro, e.g. to ...

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 ...

Surgical Retrieval, Isolation and In vitro Expansion of Human Anterior Cruciate Ligament-derived Cells for Tissue Engineering Applications

Injury to the ACL is a commonly encountered problem in active individuals. Even partial tears of this intra-articular knee ligament lead to biomechanical deficiencies that impair function and stability. Current options for the treatment of partial ACL tears range from nonoperative, conservative management to multiple surgical options, such as: thermal modification, single-bundle repair, complete reconstruction, and reconstruction of the damaged portion of the native ligament. Few studies, if any, have demonstrated any single method for management to be consistently superior, and in many cases patients continue to demonstrate persistent instability and other comorbidities.
The goal of this study is to identify a potential cell source for utilization in the development of a tissue engineered patch that could be implemented in the repair of a partially torn ACL. A novel protocol was developed for the expansion of cells derived from patients undergoing ACL reconstruction. To isolate the cells, minced hACL tissue obtained during ACL reconstruction was digested in a Collagenase solution. Expansion was performed using DMEM/F12 medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (P/S). The cells were then stored at -80 &#186;C or in liquid nitrogen in a freezing medium consisting of DMSO, FBS and the expansion medium. After thawing, the hACL derived cells were then seeded onto a tissue engineered scaffold, PLAGA (Poly lactic-co-glycolic acid) and control Tissue culture polystyrene (TCPS). After 7 days, SEM was performed to compare cellular adhesion to the PLAGA versus the control TCPS. Cellular morphology was evaluated using immunofluorescence staining. SEM (Scanning Electron Microscope) micrographs demonstrated that cells grew and adhered on both PLAGA and TCPS surfaces and were confluent over the entire surfaces by day 7. Immunofluorescence staining showed normal, non-stressed morphological patterns on both surfaces. This technique is promising for applications in ACL regeneration and reconstruction.

Surgical Retrieval, Isolation and In vitro Expansion of Human Anterior Cruciate Ligament-derived Cells for Tissue Engineering Applications

For future applications as a patch to repair partial tears of the Anterior Cruciate Ligament (ACL), human ACL derived cells were isolated from tissue obtained during reconstructive procedures, expanded in vitro and grown on tissue engineered scaffolds. Cellular adhesion and morphology was then performed to confirm biocompatibility on scaffold surface.

Injury to the ACL is a commonly encountered problem in active individuals. Even partial tears of this intra-articular knee ligament lead to biomechanical ...

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. ...

Preparation of Thermoresponsive Nanostructured Surfaces for Tissue Engineering

Thermoresponsive poly(N-isopropylacrylamide) (PIPAAm)-immobilized surfaces for controlling cell adhesion and detachment were fabricated by the Langmuir-Schaefer method. Amphiphilic block copolymers composed of polystyrene and PIPAAm (St-IPAAms) were synthesized by reversible addition-fragmentation chain transfer (RAFT) radical polymerization. A chloroform solution of St-IPAAm molecules was gently dropped into a Langmuir-trough apparatus, and both barriers of the apparatus were moved horizontally to compress the film to regulate its density. Then, the St-IPAAm Langmuir film was horizontally transferred onto a hydrophobically modified glass substrate by a surface-fixed device. Atomic force microscopy images clearly revealed nanoscale sea-island structures on the surface. The strength, rate, and quality of cell adhesion and detachment on the prepared surface were modulated by changes in temperature across the lower critical solution temperature range of PIPAAm molecules. In addition, a two-dimensional cell structure (cell sheet) was successfully recovered on the optimized surfaces. These unique PIPAAm surfaces may be useful for controlling the strength of cell adhesion and detachment.

Nanoscaled sea-island surfaces composed of thermoresponsive block copolymers were fabricated by the Langmuir-Schaefer method for controlling spontaneous cell adhesion and detachment. Both the preparation of the surface and the adhesion and detachment of cells on the surface were visualized.

Thermoresponsive poly(N-isopropylacrylamide) (PIPAAm)-immobilized surfaces for controlling cell adhesion and detachment were fabricated by the ...

Extraction of Plant-based Capsules for Microencapsulation Applications

Microcapsules derived from plant-based spores or pollen provide a robust platform for a diverse range of microencapsulation applications. Sporopollenin exine capsules (SECs) are obtained when spores or pollen are processed so as to remove the internal sporoplasmic contents. The resulting hollow microcapsules exhibit a high degree of micromeritic uniformity and retain intricate microstructural features related to the particular plant species. Herein, we demonstrate a streamlined process for the production of SECs from Lycopodium clavatum spores and for the loading of hydrophilic compounds into these SECs. The current SEC isolation procedure has been recently optimized to significantly reduce the processing requirements which are conventionally used in SEC isolation, and to ensure the production of intact microcapsules. Natural L. clavatum spores are defatted with acetone, treated with phosphoric acid, and extensively washed to remove sporoplasmic contents. After acetone defatting, a single processing step using 85% phosphoric acid has been shown to remove all sporoplasmic contents. By limiting the acid processing time to 30 hr, it is possible to isolate clean SECs and avoid SEC fracturing, which has been shown to occur with prolonged processing time. Extensive washing with water, dilute acids, dilute bases, and solvents ensures that all sporoplasmic material and chemical residues are adequately removed. The vacuum loading technique is utilized to load a model protein (Bovine Serum Albumin) as a representative hydrophilic compound. Vacuum loading provides a simple technique to load various compounds without the need for harsh solvents or undesirable chemicals which are often required in other microencapsulation protocols. Based on these isolation and loading protocols, SECs provide a promising material for use in a diverse range of microencapsulation applications, such as, therapeutics, foods, cosmetics, and personal care products.

This protocol/manuscript describes a streamlined process for the production of sporopollenin exine capsules (SECs) from Lycopodium clavatum spores and for the loading of hydrophilic compounds into these SECs.

Microcapsules derived from plant-based spores or pollen provide a robust platform for a diverse range of microencapsulation applications. Sporopollenin ...