Name: a decellularization methodology for the production of a natural acellular intestinal matrix
Uploaded: 2013-10-07T22:45:00
Duration: 8 min 10 s
Description: Watch this Scientific Journal Video about A Decellularization Methodology for the Production of a Natural Acellular Intestinal Matrix at JoVE.com

The authors thank the Wake Forest Institute of Regenerative Medicine for their help with development of this protocol. We acknowledge support by grants from the Great Ormond Street Hospital charity, the Foundation Eugenio Litta (Geneva, Switzerland), the Medical Research Council, the Royal College of Surgeons of England, the Sparks Children's Medical Charity, the British Foreign Office for the UK/USA Stem Cell Collaboration Award and the Mittal Research Fund. We would also like to thank the Royal Society/ Wolfson Foundation for the tissue engineering laboratory refurbishment grant obtained for the Pediatric Surgery Department in the Institute of Child Health. PDC and SE are supported by the Great Ormond Street Hospital Children's Charity.

1. Isolation of Rat Intestine
<ol>
	<li>Prepare a 1 L solution of Phosphate Buffered Saline (Sigma, UK) with 1% antibiotic/antimycotic (Sigma, UK) (PBS/AA). Fit the peristaltic pump (Masterflex L/S variable speed) with two tubes. Run 70% ethanol (EtOH) through the tubes of the peristaltic pump for 15 min at maximum speed, followed by PBS/AA for another 15 min.</li>
	<li>Euthanize adult Sprague-Dawley rats weighing approximately 250-350 g by CO2 inhalation and cervical dislocation in accordance with local animal guidelines and approvals (in the UK, Home Office guidelines under the Animals (Scientific Procedures) Act 1986).</li>
	<li>Autoclave the surgical instruments before use. Spray and wipe the abdomen of the animal using 70% EtOH.</li>
	<li>Perform a xifo-pubic laparotomic incision on the abdomen to ensure a clear surgical field.</li>
	<li>Eviscerate the small intestine on the right side of the animal onto a paper towel sprayed with 70% EtOH. Take care to continuously wet the intestine with PBS/AA so as to avoid dehydration of the tissue and denaturation of the extracellular matrix (ECM).</li>
	<li>Locate the superior mesenteric artery (SMA) arising in a 90&#176; angle from the aorta towards the intestine. Use a 27 G cannula to enter the SMA from the aorta and quickly withdraw the needle to avoid tearing the wall of the vessel. Advance the plastic tube of the cannula into the SMA and secure using sutures (Vicryl 5/0).</li>
	<li>Confirm cannulation by injecting 1 ml of PBS/AA. Use spring scissors to incise the aorta proximal and distal to the SMA so as to release the cannula.</li>
	<li>To avoid leakage of feces when dissecting out the large intestine, place one (Silk 5/0) suture knot on the proximal end of the ileocaecal junction and one at the distal end of the sigmoid colon. Then cut the terminal ileum and colon between the two knots and remove the large intestine.</li>
	<li>Cut the intestine at the jejuno-duodenal junction and free the intestine from any connective tissue connections it might have with the abdomen.</li>
	<li>Transfer the small intestine in a Petri dish with PBS/AA. Cannulate the proximal section of the intestine with a plastic Pasteur pipette (LSL Laboratory Consumables) and secure it in place with sutures. Cut the pipette so it will fit the Luer-Lok of a 60 ml syringe (BD Plastipak). Flush the intestinal lumen 2x with 60 ml of PBS/AA to remove feces and debris.</li>
</ol>
2. Decellularization Methodology
<ol>
	<li>Connect the vascular cannula to a three-way stopcock. This will ease handling, minimize tension on the vascular tree and allow removal of bubbles from the tubing. </li>
	<li>Start running deionized water (resistivity 18.2 M&#937;/cm) through the Masterflex L/S variable speed roller pump at 0.6 ml/hr. Ensure no bubbles are present within tubing prior to connecting with the intestinal lumen stopcock. Some bubbles may be present within the lumen from step 1.9, which may impair the decellularization process. Vary the speed of the pump and handle the tissue with forceps carefully to ensure the bubbles exit from the distal end. Do not connect the vascular cannula until this process is finished since high speeds may damage the vascular tree.</li>
	<li>Transfer apparatus to the cold room, since the following step of decellularization is performed at 4 &#176;C. The low temperature allows removal of cells whilst minimizing tissue decomposition. Place the Petri dish in a container that will collect the overflowing solution. Perfuse both the intestinal lumen and the vascular tree with deionized water (resistivity 18.2 M&#8486;/cm) for 24 hr at 0.6 ml/hr. For the first hour check the apparatus regularly to ensure all connections are secure, leakage is minimal and no bubbles are present within the intestine.</li>
	<li>After 24 hr of treatment with deionized water, move decellularization apparatus outside the cold room as the rest of the steps are carried out in room temperature (RT).</li>
	<li>Weigh sodium deoxycholate (Sigma, UK) to prepare 300 ml of a 4% solution in deionized water. Weigh sodium deoxycholate under a suction hood, as it is an oral and eye irritant. Mix using a vortex until the solution is clear. The solution may be maintained at 4 &#176;C for up to 1 month.</li>
	<li>Change the deionized solution to sodium deoxycholate, check the connections, remove any bubbles and perfuse at 0.6 ml/hr for 4 hr.</li>
	<li>Following the sodium deoxycholate step wash with PBS/AA for 30 min to avoid interaction between the sodium deoxycholate and DNase-I.</li>
	<li>Weigh out deoxyribonuclease-I (DNase-I, Sigma) and prepare 250 ml of a 2,000 kU solution in 1 M sodium chloride, pH 7.4. Mix using a vortex until the solution is clear. This solution must be prepared immediately before use and can&#39;t be stored.</li>
	<li>Perfuse with DNase-I for 3 hr at RT at a speed of 0.6 ml/hr.</li>
	<li>Following the DNase-I step, transfer the scaffold into a tissue culture hood and change plates and instruments. It is suggested to use aseptic technique.</li>
	<li>Wash with PBS/AA for 30 min to ensure all remnants of the decellularization solutions are removed. Place the scaffold in a new Petri dish and transfer to a UV Crosslinker (Spectroline, US). Run two sterilization cycles. Store in 5% PBS/AA and change the solution every 3 days.</li>
</ol>

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No conflicts of interest declared.

The most difficult steps in setting up this experiment involve the cannulation of the SMA and the establishment and maintenance of sterility. Cannulating the SMA in rodents without rupturing the wall can be quite difficult due to the size and position of the vessel. Alternatively, a suture may be placed around the proximal aorta prior to the SMA origin, followed by cannulation of the aorta itself distally, directing the plastic cannula into the SMA. During decellularization a combination of poor cannula placement and high flow rates may result in losing the vascular access. Sterility is a major issue due to the amount of bacterial flora present in the small intestine. Washing with PBS/AA following harvest is very important, and any sign of fecal matter or debris must be removed from the lumen. Placing a portion of the scaffold in a falcon tube with DMEM in the incubator following UV sterilization should be an indicator if sterility has been achieved. In the case of bacterial colonization, the media will change the pH with its color turning from red to yellow. To deal with this, further UV cycles are advised as well as washing with PBS containing a high concentration of antibiotic/antimycotic.
A possible modification to obtain a scaffold with both vascular and venous access would be to cannulate the inferior vena cava (IVC) as well as the SMA. Decellularization from the venous and vascular sides should be attempted intermittently for all three solutions. Continuous decellularization may burst the capillaries by having positive pressure on both sides.
Over the years there have been a number of efforts in intestinal TE using different cell-scaffold combinations in vitro and in vivo6,9,12. The majority of the work has been performed using tubular nonwoven 95% PGA-5% PLGA scaffolds coated with collagen type I6,7,13,22. Following seeding with intestinal epithelial organoid units (OUs) and a period of implantation into the omentum of mice, they form cysts with muscle on the outside and epithelium on the inside which can then be tubularized. However, the porosity and simplicity in the design of these scaffolds does not permit for the generation of large pieces of artificial intestine in an in vitro environment such as that of a bioreactor. Additionally, the lack of an innate vascular network that can be connected to the recipient further limits the use of this scaffold for clinical translation. Besides the experiments with the PGA-PLGA scaffolds, other groups have used collagen or SIS scaffolds, both of which do not replicate the intricacy of the intestinal tract. SIS in particular is the only previous published methodology of intestinal tissue engineering and has been used in more than 150 medical settings, demonstrating the ability of decellularized scaffolds to provide mechanical stability and promote cell growth whilst leading to no immunogenic response. However, the lack of appropriate macro- and micro-architecture, has led to poor results in its&#39; use for intestinal TE purposes9-11,23.
The benefits of the decellularization methodology we have described include the preservation of microscopic characteristics such as the luminal crypt-villus architecture that represent an appropriate environment for the repopulation by the intestinal stem cell niche. In the macroscopic aspect, the presence of a hierarchical vascular network will enable attachment to the recipient, allowing provision of nutrients and oxygen to all layer of the TE-intestine21. What is more, the maintenance of ECM components such as collagen, elastin and glyocosaminoglycans has an important role not only for mechanical characteristics but also directing cellular proliferation and differentiation. Most importantly, the ability of the decellularization methodology to be scaled up into larger tissues whilst maintaining the same characteristics is an important feature for clinical translation of TE.
The development of a natural intestinal matrix with a vascular network allows the creation of larger segments of artificial intestine that can be connected to the host.

Tissue engineering (TE) provides a therapeutic alternative to organ transplantation, bypassing issues of immunosuppression and organ shortage. TE has recently had successful applications in the clinic, with replacement of organs such as the bladder1, urethra2 and trachea, both in adults3,4 and children5.
Building a tissue-engineered organ necessitates the combination of a scaffold with the appropriate cells. A scaffold can be prepared using naturally-derived (e.g. collagen) and synthetic (e.g. poly-L-glycolic acid; PLGA) materials, or be obtained by the decellularization of native organs and tissues. Scaffolds that have been used thus far for intestinal TE have mainly been either decellularized (small intestinal submucosa) or synthetic (poly-L-glycolic acid and poly-lactic acid)6-13. These biomaterials are very simple in both macro- and micro-architecture, which may not be ideal if tissue-engineered intestine is to be clinically translated. An optimal biomaterial for the intestine should have an innate vascular tree that can be connected to the host&#39;s blood supply, a tiered tubular wall with different properties to reflect the layers of the intestinal wall and a villus-crypt axis on the luminal side to aid with repopulation by epithelial stem cells.
Decellularization is a novel methodology that produces scaffolds by removing cells from whole organs whilst maintaining their original architecture14. This is preferable to already existing scaffolds since they do not only replicate the structure of the organ but also contain chemical cues embedded within the extracellular matrix (ECM) that aid cellular proliferation and differentiation. In 2008 a cadaveric trachea was decellularized14, seeded with the patient&#39;s own cells, and transplanted to replace the main left bronchus in a young man3. Since then, a number of groups have reported the production of decellularized scaffolds for the heart15, liver16,17, and lung18-20 in small and large animals.
We have also adapted the same methodology to produce a small intestinal decellularized scaffold21. The goal of the method described herein is to produce decellularized intestinal matrices that maintain the macroscopic characteristics of the original tissue such as the blood supply, as well as the microscopic architecture of the villus-crypt axis in the intestinal lumen. We believe this methodology could ultimately be adopted for other organs to improve the efficiency of decellularization.

<table border="1">
	<tbody>
		<tr>
			<td>Name of Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Ethanol solution, 70% in H20</td>
			<td>Sigma</td>
			<td>02877</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline tablets</td>
			<td>Sigma</td>
			<td>79382</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic Antimycotic Solution (100x)</td>
			<td>Sigma</td>
			<td>A5955</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium deoxycholate</td>
			<td>Sigma</td>
			<td>D6750</td>
			<td>Oral and eye irritant; use protection</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I from bovine pancreas</td>
			<td>Sigma</td>
			<td>D5025</td>
			<td></td>
		</tr>
	</tbody>
</table>

a decellularization methodology for the production of a natural acellular intestinal matrix

Successful tissue engineering involves the combination of scaffolds with appropriate cells in vitro or in vivo. Scaffolds may be synthetic, naturally-derived or derived from tissues/organs. The latter are obtained using a technique called decellularization. Decellularization may involve a combination of physical, chemical, and enzymatic methods. The goal of this technique is to remove all cellular traces whilst maintaining the macro- and micro-architecture of the original tissue.
Intestinal tissue engineering has thus far used relatively simple scaffolds that do not replicate the complex architecture of the native organ. The focus of this paper is to describe an efficient decellularization technique for rat small intestine. The isolation of the small intestine so as to ensure the maintenance of a vascular connection is described. The combination of chemical and enzymatic solutions to remove the cells whilst preserving the villus-crypt axis in the luminal aspect of the scaffold is also set out. Finally, assessment of produced scaffolds for appropriate characteristics is discussed.

Upon successful decellularization (Figure 1), the scaffold should have a macroscopic appearance similar to the one of the native intestine, but with translucent walls (Figure 2A). The preservation of macro-architecture should also be evident by the patency of the vascular tree. When dye is injected through the SMA cannula, it should distribute evenly throughout the scaffold and reach the capillary tree (Figure 2B). The scaffold should be free of nuclear and cytoplasmic material, something that can be assessed using a DNA quantification kit and simple histological analysis. Hematoxylin and eosin staining (H&#38;E) should demonstrate an absence of nuclear and cytoplasmic material whilst preserving the small intestinal layers (Figure 3A). In particular, the maintenance of the villi and crypts on the luminal side should be evident following scanning electron microscopy (Figure 4). A conservation of extracellular matrix components such as collagen, elastin and glycosaminoglycans should also be expected. For example, Masson&#39;s Trichrome staining displays intact connective tissue in the scaffold (Figure 3B).
<img alt="Figure 1" fo:content-width="6.5in" src="/files/ftp_upload/50658/50658fig1.jpg" width="600px" /> 
Figure 1. Experimental plan. Timeline of decellularization procedure; PBS/AA: phosphate buffered saline with antibiotic-antimycotic, NaDeoxy: Sodium deoxycholate, RT: room temperature, DNAse-I: deoxyribonuclease-I. <a href="https://www.jove.com/files/ftp_upload/50658/50658fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="6.5in" src="/files/ftp_upload/50658/50658fig2.jpg" width="600px" /> 
Figure 2. Scaffold macroscopic characteristics. Macroscopic appearance of the rat small intestine following successful decellularization (A). Injection of Rosso Ponceau dye through the SMA demonstrates an intact vascular network (B) ; SMA: superior mesenteric artery <a href="https://www.jove.com/files/ftp_upload/50658/50658fig2highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 3" fo:content-width="6.5in" src="/files/ftp_upload/50658/50658fig3.jpg" width="600px" /> 
Figure 3. Histology. H&#38;E (A) and MT (B) staining indicate an absence of cellular material with preservation of the connective tissue architecture; H&#38;E: hematoxylin and eosin, MT: Masson&#39;s trichrome. Scale bar: 100 &#956;m. <a href="https://www.jove.com/files/ftp_upload/50658/50658fig3highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 4" fo:content-width="6.5in" src="/files/ftp_upload/50658/50658fig4.jpg" width="600px" /> 
Figure 4. Electron Microscopy. Scanning electron microscopy indicates the preserved villus-crypt architecture in the scaffold lumen; Scale bar: 100 &#956;m. <a href="https://www.jove.com/files/ftp_upload/50658/50658fig4highres.jpg" target="_blank">Click here to view larger image</a>.

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A Decellularization Methodology for the Production of a Natural Acellular Intestinal Matrix

The article describes a methodology for the production of an acellular matrix from rat intestine. The derivation of intestinal scaffolds is important for future applications in tissue engineering, stem cell biology and drug testing.

Successful tissue engineering involves the  combination of scaffolds with appropriate cells in vitro or in vivo.  Scaffolds may be synthetic, ...

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Bioengineering

Real Time Monitoring of Intracellular Bile Acid Dynamics Using a Genetically Encoded FRET-based Bile Acid Sensor

F&#246;rster Resonance Energy Transfer (FRET) has become a powerful tool for monitoring protein folding, interaction and localization in single cells. Biosensors relying on the principle of FRET have enabled real-time visualization of subcellular signaling events in live cells with high temporal and spatial resolution. Here, we describe the application of a genetically encoded Bile Acid Sensor (BAS) that consists of two fluorophores fused to the farnesoid X receptor ligand binding domain (FXR-LBD), thereby forming a bile acid sensor that can be activated by a large number of bile acids species and other (synthetic) FXR ligands. This sensor can be targeted to different cellular compartments including the nucleus (NucleoBAS) and cytosol (CytoBAS) to measure bile acid concentrations locally. It allows rapid and simple quantitation of cellular bile acid influx, efflux and subcellular distribution of endogenous bile acids without the need for labeling with fluorescent tags or radionuclei. Furthermore, the BAS FRET sensors can be useful for monitoring FXR ligand binding. Finally, we show that this FRET biosensor can be combined with imaging of other spectrally distinct fluorophores. This allows for combined analysis of intracellular bile acid dynamics and i) localization and/or abundance of proteins of interest, or ii) intracellular signaling in a single cell.

We provide a detailed protocol to study bile acid dynamics in living cells using a genetically encoded BAS FRET sensor. This Bile Acid Sensor represents a unique tool to study (regulation of) bile acid transport and FXR activation in a wide range of cell types.

F&#246;rster Resonance Energy Transfer (FRET) has become a powerful tool for monitoring protein folding, interaction and localization in single cells. ...

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

Decellularization and Recellularization Methodology for Human Saphenous Veins

Vascular conduits used during most vascular surgeries are allogeneic or synthetic grafts that often lead to complications caused by immunosuppression and poor patency. Tissue engineering offers a novel solution to generate personalized grafts with a natural extracellular matrix containing the recipient&#39;s cells using the method of decellularization and recellularization. We show a detailed method for performing decellularization of the human saphenous vein and recellularization by perfusion of peripheral blood. The vein was decellularized by perfusing 1% Triton X-100, 1% tri-n-butyl-phosphate (TnBP) and 2,000 Kunitz units of deoxyribonuclease (DNase). Triton X-100 and TnBP were perfused at 35 mL/min for 4 h while DNase was perfused at 10 mL/min at 37 &#176;C for 4 h. The vein was washed in ultrapure water and PBS and then sterilized in 0.1% peracetic acid. It was washed again in PBS and preconditioned in endothelial medium. The vein was connected to a bioreactor and perfused with endothelial medium containing 50 IU/mL heparin for 1 h. Recellularization was performed by filling the bioreactor with fresh blood, diluted 1:1 in Steen solution, and adding endocrine gland-derived vascular endothelial growth factors (80 ng/mL), basic fibroblast growth factors (4 &#181;L/mL), and acetyl salicylic acid (5 &#181;g/mL). The bioreactor was then moved into an incubator and perfused for 48 h at 2 mL/min while maintaining glucose between 3 - 9 mmol/L. Later, the vein was washed with PBS, filled with endothelial medium and perfused for 96 h in the incubator. Treatment with Triton X-100, TnBP and DNase decellularized the saphenous vein in 5 cycles. The decellularized vein looked white in contrast to normal and recellularized veins (light red). The hematoxylin &#38; eosin (H&#38;E) staining showed the presence of nuclei only in normal but not in decellularized veins. In the recellularized vein, H&#38;E-staining showed the presence of cells on the luminal surface of the vein.

Here, we describe a protocol for the saphenous vein decellularization using detergents and recellularization by perfusion of the peripheral blood and the endothelial medium.

Vascular conduits used during most vascular surgeries are allogeneic or synthetic grafts that often lead to complications caused by immunosuppression and ...

Decellularization of Whole Human Heart Inside a Pressurized Pouch in an Inverted Orientation

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ultimately rejection can occur. Creating a functional, autologous bio-artificial heart could solve these challenges. Biofabrication of a heart comprised of scaffold and cells is one option. A natural scaffold with tissue-specific composition as well as micro- and macro-architecture can be obtained by decellularizing hearts from humans or large animals such as pigs. Decellularization involves washing out cellular debris while preserving 3D extracellular matrix and vasculature and allowing &#34;cellularization&#34; at a later timepoint. Capitalizing on our novel finding that perfusion decellularization of complex organs is possible, we developed a more &#34;physiological&#34; method to decellularize non-transplantable human hearts by placing them inside a pressurized pouch, in an inverted orientation, under controlled pressure. The purpose of using a pressurized pouch is to create pressure gradients across the aortic valve to keep it closed and improve myocardial perfusion. Simultaneous assessment of flow dynamics and cellular debris removal during decellularization allowed us to monitor both fluid inflow and debris outflow, thereby generating a scaffold that can be used either for simple cardiac repair (e.g. as a patch or valve scaffold) or as a whole-organ scaffold.

This method enables decellularization of a complex solid organ using a simple protocol based on osmotic shock and perfusion of ionic detergent with minimal organ matrix disruption. It comprises a novel decellularization technique for human hearts inside a pressurized pouch with real-time monitoring of flow dynamics and cellular debris outflow.

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ...

Isolation and Decellularization of a Whole Porcine Pancreas

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an allogeneic or xenogeneic source with human cells. A demonstration of methods for the efficient dissection, decellularization, and recellularization of porcine pancreas might benefit the field. Akin to human pancreases, porcine pancreases have a special anatomical arrangement with three lobes (splenic, duodenal, and connection) rounded by the duodenum and small intestine. The duodenal lobe of the pancreas connects to the duodenum by several small blood vessels. Tissue engineering of the pancreas is complicated because of its exocrine and endocrine nature. In this paper, we show a detailed protocol to dissect the whole porcine pancreas and decellularize it with detergents while saving its structure and some extracellular matrix components. To achieve complete perfusion, the aorta is chosen as inlet and the portal vein as outlet. The other blood vessels (hepatic artery, splenic vein, splenic artery,&#160;mesenteric artery and vein tree) and bile duct are ligated. To prevent the formation of thrombus, the pig is heparinized and, immediately after dissection, the organ is flushed with cold heparin. To inhibit the action of exocrine enzymes, the pancreas decellularization is set at 4 &#176;C. The decellularization is performed by perfusion of Triton X-100, sodium deoxycholate, and deoxyribonuclease, with an intermittent and final extensive washing. With a successful decellularization, the pancreas appears white, and a histological evaluation with hematoxylin and eosin shows an absence of nuclei with a preserved extracellular matrix structure. Thus, the proposed method can be used to successfully dissect and decellularize whole porcine pancreas.

Tissue engineering of the whole pancreas is a challenge because of its exocrine and endocrine functions. We show a method for the dissection of an intact porcine pancreas and the process of successful decellularization by perfusion of detergents Triton X-100, sodium deoxycholate, and deoxyribonuclease.

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an ...

Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust mechanical support for in vitro cell culture but lack the necessary protein and ligand composition to elicit physiological behavior from cells. This manuscript describes a fabrication method for a kidney cortex ECM-derived hydrogel with proper mechanical robustness and supportive biochemical composition. The hydrogel is fabricated by mechanically homogenizing and solubilizing decellularized human kidney cortex ECM. The matrix preserves native kidney cortex ECM protein ratios while also enabling gelation to physiological mechanical stiffnesses. The hydrogel serves as a substrate upon which kidney cortex-derived cells can be maintained under physiological conditions. Furthermore, the hydrogel composition can be manipulated to model a diseased environment which enables the future study of kidney diseases.

Here we present a protocol to fabricate a kidney cortex extracellular matrix-derived hydrogel to retain the native kidney extracellular matrix (ECM) structural and biochemical composition. The fabrication process and its applications are described. Finally, a perspective on using this hydrogel to support kidney-specific cellular and tissue regeneration and bioengineering is discussed.

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust ...

A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of cellular function. The ECM plays a particularly important role in adipose tissue function in obesity, and alterations in adipose tissue ECM deposition and composition are associated with metabolic disease in mice and humans. Tractable in vitro models that permit dissection of the roles of the ECM and cells in contributing to global tissue phenotype are sparse. We describe a novel 3D in vitro model of human ECM-adipocyte culture that permits study of the specific roles of the ECM and adipocytes in regulating adipose tissue metabolic phenotype. Human adipose tissue is decellularized to isolate ECM, which is subsequently repopulated with preadipocytes that are then differentiated within the ECM into mature adipocytes. This method creates ECM-adipocyte constructs that are metabolically active and retain characteristics of the tissues and patients from which they are derived. We have used this system to demonstrate disease-specific ECM-adipocyte crosstalk in human adipose tissue. This culture model provides a tool for dissecting the roles of the ECM and adipocytes in contributing to global adipose tissue metabolic phenotype and permits study of the role of the ECM in regulating adipose tissue homeostasis.

We describe a 3D human extracellular matrix-adipocyte in vitro culture system that permits dissection of the roles of the matrix and adipocytes in contributing to adipose tissue metabolic phenotype.

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of ...

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes all liposomes of the ensemble to be identical. However, new experimental platforms able to observe liposomes at the single-particle level have made it possible to perform highly sophisticated and quantitative studies on protein-membrane interactions or drug carrier properties on individual liposomes, thus avoiding errors from ensemble averaging. Here we present a protocol for preparing, detecting, and analyzing single liposomes using a fluorescence-based microscopy assay, facilitating such single-particle measurements. The setup allows for imaging individual liposomes in a massive parallel manner and is employed to reveal intra-sample size and compositional inhomogeneities. Additionally, the protocol describes the advantages of studying liposomes at the single liposome level, the limitations of the assay, and the important features to be considered when modifying it to study other research questions.

This protocol describes the fabrication of liposomes and how these can be immobilized on a surface and imaged individually in a massive parallel manner using fluorescence microscopy. This allows for the quantification of the size and compositional inhomogeneity between single liposomes of the population.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes ...

Production of Membrane-Filtered Phase-Shift Decafluorobutane Nanodroplets from Preformed Microbubbles

There are many methods that can be used for the production of vaporizable phase-shift droplets for imaging and therapy. Each method utilizes different techniques and varies in price, materials, and purpose. Many of these fabrication methods result in polydisperse populations with non-uniform activation thresholds. Additionally, controlling the droplet sizes typically requires stable perfluorocarbon liquids with high activation thresholds that are not practical in vivo. Producing uniform droplet sizes using low-boiling point gases would be beneficial for in vivo imaging and therapy experiments. This article describes a simple and economical method for the formation of size-filtered lipid-stabilized phase-shift nanodroplets with low-boiling point decafluorobutane (DFB). A common method of generating lipid microbubbles is described, in addition to a novel method of condensing them with high-pressure extrusion in a single step. This method is designed to save time, maximize efficiency, and generate larger volumes of microbubble and nanodroplet solutions for a wide variety of applications using common laboratory equipment found in many biological laboratories.

This protocol describes a method of generating large volumes of lipid encapsulated decafluorobutane microbubbles using probe-tip sonication and subsequently condensing them into phase-shift nanodroplets using high-pressure extrusion and mechanical filtration.

There are many methods that can be used for the production of vaporizable phase-shift droplets for imaging and therapy. Each method utilizes different ...

Light-Controlled Fermentations for Microbial Chemical and Protein Production

Microbial cell factories offer a sustainable alternative for producing chemicals and recombinant proteins from renewable feedstocks. However, overburdening a microorganism with genetic modifications can reduce host fitness and productivity. This problem can be overcome by using dynamic control: inducible expression of enzymes and pathways, typically using chemical- or nutrient-based additives, to balance cellular growth and production. Optogenetics offers a non-invasive, highly tunable, and reversible method of dynamically regulating gene expression. Here, we describe how to set up light-controlled fermentations of engineered Escherichia coli and Saccharomyces cerevisiae for the production of chemicals or recombinant proteins. We discuss how to apply light at selected times and dosages to decouple microbial growth and production for improved fermentation control and productivity, as well as the key optimization considerations for best results. Additionally, we describe how to implement light controls for lab-scale bioreactor experiments. These protocols facilitate the adoption of optogenetic controls in engineered microorganisms for improved fermentation performance.

Optogenetic control of microbial metabolism offers flexible dynamic control over fermentation processes. The protocol here shows how to set up blue light-regulated fermentations for chemical and protein production at different volumetric scales.