We would like to thank the following institutions for supporting our AV loop research: Staedtler Stiftung, Dr. Fritz Erler Fonds, Else Kr&#246;ner Fesenius Stiftung, Baxter Healthcare GmbH, DFG, IZKF/ELAN/EFI/Office for Gender and Diversity, the Forschungsstiftung Medizin, Friedrich-Alexander University of Erlangen-N&#252;rnberg (FAU), AO Foundation, Manfred Roth Stiftung, Xue Hong, Hans Georg Geis Foundation, Deutscher Akademischer Austauschdienst (DAAD), Germany, and the Ministry of Higher Education and Scientific Research, Iraq. We would like to thank Stefan Fleischer, Marina Milde, Katrin K&#246;hn and Ilse Arnold-Herberth for their excellent technical support.

The Animal Care Committee of the Friedrich-Alexander University of Erlangen-N&#252;rnberg (FAU) and the Government of Middle Franconia, Germany, approved all the experiments. For the experiments, male Lewis rats with a body weight of 300 - 350 g were used.
1. The Arteriovenous Loop Model in the Rat
<ol>
	<li>Implantation Procedure (Figure 2)
	<ol>
		<li>For anesthesia use a special plastic box that is connected via tube to the isoflurane vaporizer and closed by a lid. Turn on supply gas and flow meter between 0.8 - 1.5 L/min.</li>
		<li>Place the rat in the induction plastic box and seal the top. Turn on isoflurane vaporizer to 5%.</li>
		<li>Carefully observe the rat during induction of anesthesia. After 3 - 4 min, the rat will be anesthetized.</li>
		<li>Confirm proper anesthesia: loss of righting reflex, loss of palpebral reflex, withdrawal reflex, positive corneal reflex.</li>
		<li>Remove the rat from the box and check its weight for calculation of drugs. Apply eye ointment to prevent dryness while under anesthesia.</li>
		<li>Administer pain medication and antibiotics (e.g., 7.5 mg/kg enrofloxacin subcutaneously (s. c.), 12.5 mg/kg tramadol and 100 mg/kg metamizole both intravenously (i. v.)). Administer weight-adapted crystalloids during operation (e.g., 30 ml/kg s. c.).</li>
		<li>Place the rat on its back on a warming plate at 37 &#176;C under anesthesia with 1 - 2% isoflurane inhalation administered via mask.</li>
		<li>Monitor anesthesia properly and increase isoflurane if anesthesia level is too low (movement of the rat, response to pain, jaw tone, no loss of reflexes (see 1.1.4.), heart rate increasing). Be careful not to overdose isoflurane (loss of corneal reflexes, high heart rate, decrease in oxygen saturation). Use a special pulse oximetry for small animals for checking the oxygen saturation (95 - 100%) and heart rate (250 - 450/min) of the rat. During the operation monitor temperature of the rat (36 - 40 &#176;C) and adjust temperature of the warming plate if necessary.</li>
		<li>Shave the inner sides of the hind limbs with an electric razor and disinfect the area with antiseptics. Spread the hind limbs and fix them with adhesive tape.</li>
		<li>Lay the rat under a surgical microscope and cover the rat with sterile draping. Ensure that the whole operation procedure is performed under sterile conditions.</li>
		<li>Open the skin in the middle of the left thigh with a longitudinal incision from the upper knee to the groin using a scalpel (No. 10).</li>
		<li>Cut the subcutaneous tissue and fascia in layers of approximately 3 cm in length using dissecting scissors and microforceps until the femoral vascular bundle is exposed from the pelvic artery in the groin to the bifurcation of the femoral artery in the knee.</li>
		<li>Separate the vessels and remove the adventitia using adventitia scissors and microforceps.</li>
		<li>Coagulate the side branches using electric coagulation. Cover the operation field with a damp compress.</li>
		<li>Open the skin on the right side as described for the left side, 1.1.11 - 1.1.14.</li>
		<li>For harvesting the venous graft, ligate the right femoral vein by electric coagulation on the proximal and distal ends at a distance of 1 - 1.5 cm.</li>
		<li>Remove the venous graft with microforceps and flush the venous graft with a heparin solution (50 IU/ml in 0.9% sodium chloride solution) using an irrigation cannula and transfer it to the left thigh. Cover the operation field at the right side with a damp compress.</li>
		<li>Ligate the femoral vein on the left side proximally at the inguinal region with a microvessel clamp. Coagulate the femoral vein distally by electric coagulation, at the upper knee before branching, at a distance of about 2 cm.</li>
		<li>For anastomosis connect the proximal end of the venous graft with the proximal end of the vein by end-to-end anastomosis with an 11-0 suture. Use about 8 interrupted sutures. Begin with placing the first two sutures at the 12 o&#8217;clock and 6 o&#8217;clock positions. Then, put in 2 to 3 more sutures between these points on the front side and then put 2 to 3 more sutures into the back side.</li>
		<li>Ligate the femoral artery in the same way described for the femoral vein (1.1.18.). Make sure the loop vessels are not twisted. Anastomose the distal end of the venous graft with the proximal end of the artery as described for the femoral vein (steps 1.1.19.).</li>
		<li>Administer 25 IU heparin intravenously. Make again sure the loop vessels are not twisted, Remove the clamps and check for leakage and patency of the loop for about 5 min.</li>
		<li>Dribble papaverine (e.g., 4 mg/ml) on the vessels to prevent vascular spasms. If there is patency, the loop expands and the pulse of the artery can be observed.</li>
		<li>Prefill the implantation chamber with the first half (about 500 &#181;l) of the matrix (e.g., a hydrogel or a bone matrix with or without cells). Embed the loop into the implantation chamber.</li>
		<li>Fill the implantation chamber with the second half of the matrix to a total volume of 1,000 &#181;l. Seal the chamber with the chamber lid.</li>
		<li>Fix the implantation chamber on the thigh with a non-absorbable 6-0 suture. Fix the chamber lid with a non-absorbable 6-0 suture. Stop possible bleeding with electric coagulation.</li>
		<li>Close the skin with absorbable 4-0 sutures. Cover the wound with aluminum spray.</li>
		<li>Administer antibiotics and analgesics (e.g., 7.5 mg/kg enrofloxacin s. c., tramadol 12.5 mg/kg per os (p. o.) (through drinking water) for 3 - 5 days and afterwards depending on the behavior of the rat).</li>
		<li>Turn the vaporizer off and allow the rat to breathe supply gas until it begins to awaken.</li>
		<li>Place the rat in a box with thermal support and observe it carefully until fully recovered.</li>
		<li>Do not leave the rat unattended until it has regained sufficient consciousness to maintain sternal recumbency.</li>
		<li>Do not return the rat to the company of other animals until fully recovered.</li>
	</ol>
	</li>
	<li>Explantation Procedure
	<ol>
		<li>Perform the explantation after a short-time or long-time implantation (according to the study design, for examples please see References 27-32). Anesthetize the rat according to steps 1.1.1.-1.1.9.</li>
		<li>Open the skin of the abdomen with a scalpel (No. 10) and move the intestines aside with a compress. Expose the abdominal aorta and the vena cava caudalis using cotton swabs.</li>
		<li>Cannulate the abdominal aorta (e.g., 24 gauge plastic cannula) and cut the vena cava caudalis with dissecting scissors.</li>
		<li>Flush the abdominal aorta with 0.9% sodium chloride solution containing 100 IU/ml heparin until the leaking fluid is clear. Perfuse the aorta with 30 ml perfusion solution (e.g., contrast agent or India ink).</li>
		<li>Euthanize the rat by injection of a lethal dose of embutramide, mebezonium iodide, tetracaine hydrochloride injectable solution (0.1 - 0.2 ml/kg) in deep anesthesia.</li>
		<li>Ligate the abdominal aorta and vena cava caudalis with a non-absorbable 4-0 suture.</li>
		<li>Close the open wound area with 1 - 2 clamps. Store the rat for 24 hr at 4 &#176;C (curing of the perfusion solution).</li>
		<li>Place the rat on its back. Spread the hind limbs and fix them with adhesive tape.</li>
		<li>Open the skin above the chamber with a longitudinal incision with a scalpel (No. 10).</li>
		<li>Remove the connective tissue from the chamber and the loop pedicle using dissecting scissors and microforceps. Cut the loop pedicle at a distance of 1 cm from the chamber opening with dissecting scissors and remove the chamber.</li>
		<li>Open the lid of the chamber and carefully remove the construct from the chamber with forceps and fix it in 4% buffered formalin solution for 24 hr at room temperature. Afterwards use the construct for 3D micro-computed tomography or paraffin-embedded for histological analysis30.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

For over a decade, we have successfully used the arteriovenous (AV) loop for tissue engineering purposes and studying angiogenesis in vivo in the small animal model. We could demonstrate that this microsurgical model is very well suited for engineering different tissues and that it can also be used for angiogenesis or antiangiogenesis studies.
Significance of the Technique with Respect to Existing/Alternative Methods 
Engineered tissues or organs require a functi...

Most tissues and organs in the human body are dependent on a functional blood vessel network that supplies nutrients, exchanges gases and removes waste products. Malfunction of this system caused by local or systemic vascular problems can lead to a multitude of severe diseases. Moreover, in research areas such as tissue engineering or regenerative medicine, a functional blood vessel network within artificially generated tissues or transplanted organs is indispensable for successful clinical application.
For decades researchers have been investigating the exact mechanisms involved in the growing vasculature to gain deeper insight into pathological situations in order to find novel therapeutic interventions and provide better prevention of vascular disorders. In the first step, basic processes such as cell-cell interactions or the effect of molecules on cells of the vascular system are usually investigated by in vitro 2D or 3D experiments. Traditional 2D models are easy to perform, are well established and have contributed greatly to a better understanding of these processes. For the first time in 1980, Folkman et al. reported in vitro angiogenesis seeding of capillary endothelial cells on gelatin coated plates1. This immediately gave way to publication of a multitude of further 2D angiogenesis experiments on endothelial cell tube formation assay2, migration assay3 and the co-culturing of different cell types4, as well as others. These assays are still used today and accepted as standard in vitro methods.
However, this experimental setup is not always appropriate for the study of in vivo cell behavior since most cell types require a 3D environment to form relevant physiological tissue structures5. It could be shown that the architecture of the 3D matrix is decisive for capillary morphogenesis6 and that cell-extra cellular matrix (ECM) interactions and 3D culture conditions regulate important factors involved in tumor angiogenesis7. The 3D matrix provides complex mechanical inputs, can bind effector proteins and establish tissue-scale solute concentration gradients. Moreover, it is considered necessary in order to imitate in vivo morphogenetic and remodeling steps in complex tissues5. In these systems, both angiogenesis and vasculogenesis can be studied. While angiogenesis describes the sprouting of capillaries from preexisting blood vessels8, vasculogenesis refers to the de novo formation of blood vessels through endothelial cells or their progenitors9,10. Maturation of vessels is described in a process called &#39;arteriogenesis&#39; via recruitment of smooth muscle cells11. A typical angiogenic in vitro model is the sprouting of endothelial cells from existing monolayers seeded as a monolayer on gel surfaces, on surface of microspheres embedded within a gel or by building endothelial cell spheroids12. In vasculogenic models single endothelial cells are entrapped in a 3D gel. They interact with adjacent endothelial cells to form vascular structures and networks de novo, typically in combination with supportive cells12.
However, even complex 3D in vitro models cannot mimic in vivo settings completely given the multitude of cell-cell and cell-ECM interactions13. Substances with high in vitro activity do not automatically show the same effects in vivo and vice versa14. For a comprehensive analysis of vascularization processes there is an urgent need to develop in vivo models that better simulate the situation in the body. A large range of in vivo angiogenesis assays are described in the literature, including the chick chorioallantoic membrane assay (CAM), the zebrafish model, the corneal angiogenesis assay, the dorsal air sac model, the dorsal skinfold chamber, the subcutaneous tumor models14. However, these assays are often associated with limitations, such as rapid morphological changes, problems in distinguishing new capillaries from already existing ones in the CAM assay, or the limited space in the corneal angiogenesis assay15. Furthermore, non-mammalian systems are used (e.g., the zebrafish model16), which leads to problems in xenotransplantation17. In the subcutaneous tumor model, angiogenesis originating only from the tumor itself cannot be analyzed since the adjacent tissue greatly contributes to the vascularization process. Moreover, the surrounding tissue can have a decisive role in shaping the tumor microenvironment18.
Not only for studying angiogenesis or vasculogenesis is there a strong need for a standardized and well-characterized in vivo model but also for studying different vascularization strategies in tissue engineering and regenerative medicine. Today, the generation of complex artificial organs or tissues is possible both in vitro and in vivo. 3D bioprinting provides an on-demand fabrication technique for generating complex 3D functional living tissues19. Furthermore, bioreactors can be used for generating tissues20 or even the own body can be used as bioreactor21. However, the main barrier to successful application of artificially generated tissues is the lack of vascularization within the engineered constructs. Immediate connection to the host&#39;s vasculature after transplantation is a major prerequisite for survival, especially in the case of large-scale artificial tissues or organs.
Different in vitro or in vivo prevascularization strategies were developed to establish a functional microvasculature in constructs prior to implantation22. The implantation of a scaffold with in vitro preformed engineered capillaries onto the dorsal skin of mice led to rapid anastomosis of the mice vasculature within a day23. In contrast, a spheroid co-culture consisting of human mesenchymal stem cells and human umbilical vein endothelial cells assembled into a three-dimensional prevascular network developed further after in vivo implantation. However, anastomosis with the host vasculature was limited24. Above all, in poorly vascularized defects, such as necrotic or irradiated areas, this so-called extrinsic vascularization &#8212; the ingrowth of vessels from the surrounding area into the scaffold &#8212; often fails. Intrinsic vascularization, on the other hand, is based on a vascular axis as a source of new capillaries sprouting into the scaffold25. Using the axial vascularization approach, the engineered tissue can be transplanted with its vascular axis and connected to local vessels at the recipient site. Immediately after transplantation, the tissue is adequately supported by oxygen and nutrients, which creates the right conditions for optimal integration.
Due to the limited availability of models for investigating in vivo angiogenesis and in recognition of the growing importance of generating axially vascularized tissue, we developed the microsurgical approach of Erol and Spira further to generate an arteriovenous (AV) loop in the animal model26. The use of a completely closed implantation chamber makes this method very well suited to study blood vessel formation under &#34;controlled&#34;, well characterized in vivo conditions (Figure 1). This model is not only useful for the investigation of angiogenesis but is also optimally suited for the axial vascularization of scaffolds for tissue engineering purposes.

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			<td height="21" style="height:21px;width:355px;">0.9% sodium chloride</td>
			<td style="width:251px;">Berlin-Chemie AG</td>
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			<td height="21" style="height:21px;width:355px;">Lewis rat, male</td>
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			<td height="21" style="height:21px;width:355px;">Metamizol-Natrium&#160;</td>
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			<td height="20" style="height:21px;width:355px;">papaverine / Paveron N</td>
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the arteriovenous (av) loop in a small animal model to study angiogenesis and vascularized tissue engineering

A functional blood vessel network is a prerequisite for the survival and growth of almost all tissues and organs in the human body. Moreover, in pathological situations such as cancer, vascularization plays a leading role in disease progression. Consequently, there is a strong need for a standardized and well-characterized in vivo model in order to elucidate the mechanisms of neovascularization and develop different vascularization approaches for tissue engineering and regenerative medicine.
We describe a microsurgical approach for a small animal model for induction of a vascular axis consisting of a vein and artery that are anastomosed to an arteriovenous (AV) loop. The AV loop is transferred to an enclosed implantation chamber to create an isolated microenvironment in vivo, which is connected to the living organism only by means of the vascular axis. Using 3D imaging (MRI, micro-CT) and immunohistology, the growing vasculature can be visualized over time. By implanting different cells, growth factors and matrices, their function in blood vessel network formation can be analyzed without any disturbing influences from the surroundings in a well controllable environment. 
In addition to angiogenesis and antiangiogenesis studies, the AV loop model is also perfectly suited for engineering vascularized tissues. After a certain prevascularization time, the generated tissues can be transplanted into the defect site and microsurgically connected to the local vessels, thereby ensuring immediate blood supply and integration of the engineered tissue. By varying the matrices, cells, growth factors and chamber architecture, it is possible to generate various tissues, which can then be tailored to the individual patient's needs.

Tissue Engineering 
For bone tissue engineering purposes, a number of different bone substitutes were implanted in the small animal rat AV loop model27,28,33,34. Vascularization could perfectly be demonstrated by 3D micro-computed tomography (micro-CT) (Figure 3A). Vascularization of a processed bovine cancellous bone (PBCB) matrix was significantly higher in the loop group compared to the group without vascularization. A constantly growi...

Watch this Scientific Journal Video about The Arteriovenous AV Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering at JoVE.com

The Arteriovenous AV Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

We describe a microsurgical approach for the generation of an arteriovenous (AV) loop as a model for analyzing vascularization in vivo in an isolated and well-characterized environment. This model is not only useful for the investigation of angiogenesis, but is also optimally suited for engineering axially vascularized and transplantable tissues.

The Arteriovenous (AV) Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

A functional blood vessel network is a prerequisite for the survival and growth of almost all tissues and organs in the human body. Moreover, in ...

the-arteriovenous-av-loop-small-animal-model-to-study-angiogenesis

Research

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Bioengineering

12.4K Views.  University Hospital of Erlangen, Friedrich-Alexander University of Erlangen-Nürnberg (FAU). We describe a microsurgical approach for the generation of an arteriovenous (AV) loop as a model for analyzing vascularization in vivo in an isolated and well-characterized environment. This model is not only useful for the investigation of angiogenesis, but is also optimally suited for engineering axially vascularized and transplantable tissues.

Video: The Arteriovenous AV Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Tissue Engineering of the Intestine in a Murine Model

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to preoperative weights within 40 days.1 In humans, massive small bowel resection can result in short bowel syndrome, a functional malabsorptive state that confers significant morbidity, mortality, and healthcare costs including parenteral nutrition dependence, liver failure and cirrhosis, and the need for multivisceral organ transplantation.2 In this paper, we describe and document our protocol for creating tissue-engineered intestine in a mouse model with a multicellular organoid units-on-scaffold approach. Organoid units are multicellular aggregates derived from the intestine that contain both mucosal and mesenchymal elements,3 the relationship between which preserves the intestinal stem cell niche.4 In ongoing and future research, the transition of our technique into the mouse will allow for investigation of the processes involved during TESI formation by utilizing the transgenic tools available in this species.5The availability of immunocompromised mouse strains will also permit us to apply the technique to human intestinal tissue and optimize the formation of human TESI as a mouse xenograft before its transition into humans. Our method employs good manufacturing practice (GMP) reagents and materials that have already been approved for use in human patients, and therefore offers a significant advantage over approaches that rely upon decellularized animal tissues. The ultimate goal of this method is its translation to humans as a regenerative medicine therapeutic strategy for short bowel syndrome.

This article and the accompanying video present our protocol for generating tissue-engineered intestine in the mouse, using an organoid units-on-scaffold approach.

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to ...

Design of a Biaxial Mechanical Loading Bioreactor for Tissue Engineering

We designed a loading device that is capable of applying uniaxial or biaxial mechanical strain to a tissue engineered biocomposites fabricated for transplantation. While the device primarily functions as a bioreactor that mimics the native mechanical strains, it is also outfitted with a load cell for providing force feedback or mechanical testing of the constructs. The device subjects engineered cartilage constructs to biaxial mechanical loading with great precision of loading dose (amplitude and frequency) and is compact enough to fit inside a standard tissue culture incubator. It loads samples directly in a tissue culture plate, and multiple plate sizes are compatible with the system. The device has been designed using components manufactured for precision-guided laser applications. Bi-axial loading is accomplished by two orthogonal stages. The stages have a 50 mm travel range and are driven independently by stepper motor actuators, controlled by a closed-loop stepper motor driver that features micro-stepping capabilities, enabling step sizes of less than 50 nm. A polysulfone loading platen is coupled to the bi-axial moving platform. Movements of the stages are controlled by Thor-labs Advanced Positioning Technology (APT) software. The stepper motor driver is used with the software to adjust load parameters of frequency and amplitude of both shear and compression independently and simultaneously. Positional feedback is provided by linear optical encoders that have a bidirectional repeatability of 0.1 &mu;m and a resolution of 20 nm, translating to a positional accuracy of less than 3 &mu;m over the full 50 mm of travel. These encoders provide the necessary position feedback to the drive electronics to ensure true nanopositioning capabilities. In order to provide the force feedback to detect contact and evaluate loading responses, a precision miniature load cell is positioned between the loading platen and the moving platform. The load cell has high accuracies of 0.15% to 0.25% full scale.

We designed a novel mechanical loading bioreactor that can apply uniaxial or biaxial mechanical strain to a cartilage biocomposite prior to transplantation into an articular cartilage defect.

We  designed a loading device that is capable of applying uniaxial or biaxial  mechanical strain to a tissue engineered biocomposites fabricated for  ...

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 ...

Tissue Engineering of Tumor Stromal Microenvironment with Application to Cancer Cell Invasion

3D organotypic cultures of epithelial cells on a matrix embedded with mesenchymal cells are widely used to study epithelial cell differentiation and invasion. Rat tail type I collagen and/or matrix derived from Engelbreth-Holm-Swarm mouse sarcoma cells have been traditionally employed as the substrates to model the matrix or stromal microenvironment into which mesenchymal cells (usually fibroblasts) are populated. Although experiments using such matrices are very informative, it can be argued that due to an overriding presence of a single protein (such as in type I Collagen) or a high content of basement membrane components and growth factors (such as in matrix derived from mouse sarcoma cells), these substrates do not best reflect the contribution to matrix composition made by the stromal cells themselves. To study native matrices produced by primary dermal fibroblasts isolated from patients with a tumor prone, genetic blistering disorder (recessive dystrophic epidermolysis bullosa), we have adapted an existing native matrix protocol to study tumor cell invasion. Fibroblasts are induced to produce their own matrix over a prolonged period in culture. This native matrix is then detached from the culture dish and epithelial cells are seeded onto it before the entire coculture is raised to the air-liquid interface. Cellular differentiation and/or invasion can then be assessed over time. This technique provides the ability to assess epithelial-mesenchymal cell interactions in a 3D setting without the need for a synthetic or foreign matrix with the only disadvantage being the prolonged period of time required to produce the native matrix. Here we describe the application of this technique to assess the ability of a single molecule expressed by fibroblasts, type VII collagen, to inhibit tumor cell invasion.

Tissue engineered fibroblast-derived native matrix is an emerging tool to generate a stromal substrate which supports epithelial cell proliferation and differentiation. Here a protocol applying this methodology to assess the impact of different stromal cell types on tumor cell biology is presented.

3D organotypic cultures of epithelial cells on a matrix embedded with mesenchymal cells are widely used to study epithelial cell differentiation and ...

Generation of a Three-dimensional Full Thickness Skin Equivalent and Automated Wounding

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models reflect the human wound situation better than animal models. Although two-dimensional models are widely used to investigate processes such as cellular migration and proliferation, models that are more complex are required to gain a deeper knowledge about wound healing. Besides a suitable model system, the generation of precise and reproducible wounds is crucial to ensure comparable results between different test runs. In this study, the generation of a three-dimensional full thickness skin equivalent to study wound healing is shown. The dermal part of the models is comprised of human dermal fibroblast embedded in a rat-tail collagen type I hydrogel. Following the inoculation with human epidermal keratinocytes and consequent culture at the air-liquid interface, a multilayered epidermis is formed on top of the models. To study the wound healing process, we additionally developed an automated wounding device, which generates standardized wounds in a sterile atmosphere.

The goal of this protocol is to build up a three-dimensional full thickness skin equivalent, which resembles natural skin. With a specifically constructed automated wounding device, precise and reproducible wounds can be generated under maintenance of sterility.

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models ...

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. ...

Use of a Rat Model to Study Ventral Abdominal Hernia Repair

Ventral abdominal hernia is a relatively common clinical condition that sometimes requires herniorraphy (surgical repair). The repair of ventral abdominal hernia typically requires implantation of a material to serve as a mechanical bridge across the defect in the abdominal wall. Biomaterials, such as porcine small intestinal submucosa (SIS), also serve as a lattice for cell growth into the implant and can naturally incorporate into the host tissue. Development of such repair materials benefits from use of animal models in which experimental abdominal wall defects are easily created and are amenable to repair in a reproducible fashion. The method offered here describes surgical creation and repair of ventral abdominal hernia in a rat model. When SIS is used to repair an experimental ventral abdominal hernia in this model, it is rapidly incorporated into host tissue within 28 days of implantation. Histologically, incorporation of their implanted material into host tissue is characterized by a robust fibrovascular response. Future refinements and applications of the rat abdominal hernia model may likely involve diabetic and/or obese animals as a means to more closely mimic common co-morbidities of man.

This manuscript describes methods associated with creation and repair of a ventral abdominal wall defect (hernia). This model can be used to study repair strategies such as those that use implanted materials. In this manuscript, repair of the experimental hernia with porcine small intestinal submucosa is presented as an example.

Ventral abdominal hernia is a relatively common clinical condition that sometimes requires herniorraphy (surgical repair). The repair of ventral abdominal ...

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 ...