The authors thank the Swedish Society for Medical Research, the Promobilia Foundation, the Crown Princess Lovisa Foundation, the Freemason Foundation for Children&#8217;s Welfare, the Swedish Society of Medicine, the Solstickan Foundation, Karolinska Institutet, and the Stockholm City Council for financial support.

All animal protocols were pre-approved by the Stockholm County Committee on Animals and all procedures conformed to the regulations for animal use, as well as relevant federal statutes.
1. Animal Procedures
<ol>
	<li>Preparing the Animal for Surgery
	<ol>
		<li>Prepare the surgical table with all the materials and instruments needed for the operation under sterile conditions. Perform the surgery exclusively under sterile conditions to reduce the risk of infection and optimize the conditions in survival surgery.</li>
		<li>Fast the animal for 12 hr prior to surgery and measure its weight. Administer an intramuscular injection of azaperone (2 mg/kg) for premedication. Anesthetize the animal with intramuscular injections of tiletamine hypochloride (2.5&#8201;mg/kg), zolazepam hypochloride (2.5&#8201;mg/kg), medetomidine (25 &#181;g/kg) and atropine (25 &#181;g/kg).</li>
		<li>Inject phenobarbiturate (15&#8201;mg/kg) prior to endotracheal intubation and continue general anesthesia with 0.8%&#8211;2% isoflurane. Insert a peripheral vein catheter in one ear and infuse glucose (25&#8201;mg/ml) intravenously during the procedure to maintain the well-being of the animal.</li>
		<li>Place monitoring devices on the ear or tail to check temperature, blood pressure, saturation, and peripheral pulse. Control anesthesia by pain stimulation of the ears or hooves. Apply ointment to eyes to prevent dryness while under anesthesia.</li>
	</ol>
	</li>
	<li>Excision of Bladder Biopsy Specimen
	<ol>
		<li>Catheterize the bladder using a 10-French silicone catheter by introducing a speculum to view the urethra and insert the catheter through the urethra and into the urinary bladder to empty the urine under semi-sterile conditions. Fill the bladder with sterile saline solution to a pressure of 20&#8211;25 cm H2O, about 100&#8211;300 ml, and then empty to 20 cm H2O (approx. 8 ml/kg of body weight).</li>
		<li>With the pig carefully turned to a side position, perform a basic preoperative sterilization of the skin with application of chlorhexidine gluconate. Apply a diathermy plate to the shoulder after removing the pelage with shaving shears.</li>
		<li>Turn the pig carefully into a supine position and remove the abdominal pelage using shaving shears and do a basic preoperative sterilization of the abdominal skin with application of chlorhexidine gluconate.
		<ol>
			<li>Embed the extremities with soft clothing to minimize the risk of hyperextension damage to the joints in the extremities. Sterilize the abdominal skin of the animal with successive applications of chlorhexidine gluconate and place sterile draping around the surgical field.</li>
		</ol>
		</li>
		<li>Prior to skin incision, apply an intravenous analgesic consisting of buprenorphine (45 &#181;g/kg), carprofen (3&#8201;mg/kg) and local injection of lidocaine in the midline below the umbilicus. Check for pain reaction by grasping the skin with forceps. Make a lower midline incision through the fascia and peritoneum using diathermy for control of bleeding. Localize the bladder that has a fully intraperitoneal position in the pig and can be freely exposed by mobilizing it through the surgical wound.</li>
		<li>Take hold of the bladder with the forceps and measure it with a sterilized measuring tape and mark an elliptic-shaped biopsy specimen approximately 2 cm longitudinally and 1&#160;cm transversally, or smaller, using a sterilized pen (the pig tolerates a quarter reduction of bladder size very well). Excise the marked area, using a scalpel, and place the bladder biopsy specimen in DMEM under sterile conditions.</li>
		<li>Perform autotransplantation with the biotransplant or close the bladder by suturing it with 5-0 Vicryl in two layers. Close the abdominal fascia carefully with 2-0 or 3-0 running Vicryl. Close the subcutis with 3-0 Vicryl and the skin with 3-0 Ethilon. Place a dressing on the wound and carefully attend to the animal until it has recovered sufficiently from the anesthesia and does not express pain.</li>
		<li>Move the animal to the animal care facility to secure conditions for postoperative care. Put the animal in a single cage with a heating lamp and attend to the animal until full recovery from the anesthesia and then let the animals be stalled in pairs.</li>
		<li>Provide an uneventful recovery regarding pain and well-being and administer buprenorphine (45 &#181;g/kg) intramuscularly for postoperative analgesia and trimetoprim (4 mg/kg) and sulfonamide (20 mg/kg) twice daily for three days and once daily for five days to reduce the risk of postoperative infections.</li>
	</ol>
	</li>
	<li>Excision of Skin Biopsy Specimen
	<ol>
		<li>Prepare the surgical table with all the necessary materials. Anesthetize the animal as previously described in 1.1. Remove the pelage using wax, wash and sterilize the incision area with betadine and 70% alcohol and then place sterile draping around the incision area.</li>
		<li>Use a dermatome to harvest a 0.3 mm partial thickness skin biopsy specimen. Place the skin specimen in DMEM before mincing, as described in 2.2. Cover the wound area with fatty ointment and a dressing.</li>
	</ol>
	</li>
</ol>
2. Minced Tissue Preparation
<ol>
	<li>Bladder Mucosa
	<ol>
		<li>Wash the bladder biopsy twice in DMEM. Place the bladder biopsy specimen onto a sterilized dissecting plate with the mucosa facing upwards and fix one of the sides to the plate using dissection pins.</li>
		<li>Separate the mucosal tissue from the detrusor muscle using fine scissors and forceps (Figure 3) and keep the mucosa moist by dripping saline or DMEM over it.</li>
		<li>Use the mincing device by placing it on the mucosa and then pass the device from one end to the other vertically and horizontally, applying manual pressure to obtain pieces of minced tissue of 0.8 mm x 0.8 mm (0.8 mm is the distance between the rotating cutting blades).</li>
	</ol>
	</li>
	<li>Skin
	<ol>
		<li>If the skin biopsis are thick: place the skin onto a sterile dissecting plate and use surgical scissors to separate the epidermis from subcutaneous fat and dermis. The epidermis is thin and translucent (approx. 0.3 mm) when it is ready for mincing.</li>
		<li>Use the mincing device by placing it on the epidermis. With pressure, pass the device from one end to the other vertically and horizontally to obtain pieces of minced tissue of 0.8 mm x 0.8 mm.</li>
	</ol>
	</li>
</ol>
3. Preparation of Plastic Compressed PCL/Collagen Autografts
<ol>
	<li>Place all ingredients on ice to keep cold. Utensils needed: Falcon tube, 10x DMEM, 1x DMEM, 1 N NaOH and rat tail collagen type 1.
	<ol>
		<li>Mix 2 ml of 10x DMEM (carefully to avoid bubbles) with 12 ml of collagen type 1. Add 1 N NaOH, drop by drop, to bring pH up to 7.4&#8211;8 (color in the medium should indicate the pH by changing from intense yellow to pink). In addition, use a pH strip.</li>
		<li>Carefully add 2 ml of 1x DMEM and mix the solution. Plate approximately 2 ml of collagen in each well of the steel rectangular mold (20x30x10 mm) and incubate at 37 &#176;C in 5% CO2 for 10 min. 
		NOTE: The concentration of collagen should be 2.06 mg/ml in 0.6% acetic acid and the amount of collagen 1 ml/cm2.</li>
		<li>Once the collagen sets into the mold, place the biomaterial (PCL) on top of the collagen gel (20 mm x 30 mm) and pour the remaining collagen (about 6 ml) on top of it. Incubate at 37&#160;&#176;C in 5% CO2 for 20 min.</li>
		<li>Place the minced tissue (for 1:6 expansion) on the top of collagen gel. Press the water out of the construct by mechanical force using plastic compression as follows (Figures 3 and 4).</li>
		<li>Place a thick layer of gauze pads on a sterile surface. Place one stainless steel mesh (400 &#181;m thick) on top of the gauze pads and then a sheet of nylon mesh (110 &#181;m thick). Carefully transfer the collagen gel/minced tissue onto the nylon mesh and carefully remove the rectangular steel mold.</li>
		<li>Place a new layer of nylon mesh on top of the collagen gel/minced tissue. Place a second steel mesh on top of the nylon mesh. Place in position the pressure or loading plate weighing a minimum of 120 g (i.e., a glass plate) for 5 min.</li>
		<li>Remove the weight, nylon, and steel meshes. The autografts are now ready to be sutured to the pig bladder in the full-thickness skin wounds or cultured in vitro.</li>
		<li>For in vitro culturing, cut the thin construct into small pieces fitting 12-well plates. Add 1 ml of keratinocyte medium. Place the plates in the incubator at 37 &#176;C, 5% CO2 and culture up to 6 weeks and change the medium 3 times per week.</li>
	</ol>
	</li>
</ol>
4. Suture of Autografts
<ol>
	<li>Suture of autograft with minced bladder mucosa to the pig bladder
	<ol>
		<li>Keep the autograft moist in DMEM during the waiting time. Suture the autograft with fine running monofilament sutures. Use non-absorbable 5-0 Ethilon for research purposes.</li>
		<li>Check if watertight by filling the bladder with saline through the indwelling urinary catheter. If possible, cover the autograft with a layer of greater omentum. Close the abdominal wall, subcutaneous tissue, and skin as described in 1.2.6. Apply a wound dressing.</li>
	</ol>
	</li>
	<li>Suture of the Autograft with Minced Skin Epidermis to a Full-thickness Wound
	<ol>
		<li>Keep the autograft moist in DMEM during the waiting time.</li>
		<li>Suture the autograft to the bottom of the skin full-thickness wound by interrupted sutures in the corners and in the middle of the autograft to keep the autograft closely attached to the underlying surface.</li>
		<li>Cover the wound with a plastic dressing that keeps the wound moist.</li>
	</ol>
	</li>
</ol>
5. Termination
<ol>
	<li>Sedate the animal with intramuscular injection of zolazepam hypochloride (2.5&#8201;mg/kg) and medetomidine (25 &#181;g/kg) prior to termination and apply monitoring devices to the ear or tail to check for pulse and blood pressure.</li>
	<li>Euthanize the animal by administering a lethal dose of pentobarbital sodium (60-140 mg/kg) intravenously. Check pulse and blood pressure until death has occurred.</li>
</ol>
6. In Vitro Culture
 
NOTE: To evaluate histologically the progression of the minced tissue in the PCL/collagen constructs in vitro, the collagen/PCL/minced patches are cultured in 12-well plates using keratinocyte medium.
<ol>
	<li>Preparation of keratinocyte medium:
	<ol>
		<li>Sterilize a 500 ml glass bottle.</li>
		<li>Mix 400 ml of DMEM with 100 ml of Ham&#8217;s F12 (4:1 mixture). Supplement with 10% fetal bovine serum, 5 &#181;g/ml insulin, 0.4 &#181;g/ml hydrocortisone, 21 &#181;g/ml adenine, 10-10 mol/L cholera toxin, 2 x 10-9 mol/L triiodothyronine, 5 &#181;g/ml transferrin, 10 ng/ml epidermal growth factor, 50 U/ml penicillin and 50 &#181;g/ml streptomycin.</li>
		<li>Sterilize by filtrating through a 0.2 &#956;m filter and collect the filtrate in the sterile 500 ml bottle.</li>
	</ol>
	</li>
</ol>
7. Immunohistochemistry

NOTE: The immunohistochemistry protocol is generally divided into the following steps: (1) fixation and paraffin embedding, (2) micro-sectioning to 5 &#181;m slices, placement on slides, deparaffination, and rehydration, (3) antigen unmasking, staining and mounting. Before starting the last steps in the immunohistochemistry procedure, prepare the washing buffers and the antigen unmasking solution (see separate material details). Prepare the ABC complex solution at least 30 min before use.
<ol>
	<li>Fixation 
	NOTE: At the end of the in vitro culture, fix the patches as follows:
	<ol>
		<li>Prepare Eppendorf tubes with 1 ml of 4% buffered formaldehyde (PFA) (Caution: formaldehyde is toxic. Please read material safety data sheets before working with this chemical. Wear gloves and safety glasses and prepare the solution inside a fume hood).</li>
		<li>Transfer each of the collagen patches to an Eppendorf tube containing 4% PFA. Fix over night at room temperature.</li>
		<li>Place samples in 70% ethanol for long-term storage at 4 &#176;C. Samples are now ready for dehydration and embedding in paraffin blocks before sectioning.</li>
	</ol>
	</li>
	<li>Rehydration
	<ol>
		<li>Place the slides in a staining jar with X-tra solv for 15 min. Repeat by using a new staining jar with X-tra solv. Place the slides in a staining jar with absolute ethanol for 10 min. Repeat by using a new staining jar with absolute ethanol. Place the slides in a staining jar with 95% ethanol for 10 min and thereafter into a staining jar with 70% ethanol for 10 min. Finally wash the slides twice for 5 min with distilled water.</li>
	</ol>
	</li>
	<li>Antigen Unmasking
	<ol>
		<li>Put slides in a Coplin jar with TE-solution and put the jar in a water bath to boil for 20 min. Take the jar out of the water bath carefully. Cool the slides to room temperature for 30 min and wash twice for 5 min in Tris buffer. Place the slides in a staining jar with 3% hydrogen peroxide for 10 min. Wash the slides twice for 5 min in Tris buffer. Draw a circle around the samples using a water repellent marking pen.</li>
		<li>Block nonspecific binding of the antibody using 100&#8211;300 &#181;l of blocking solution. Remove the blocking solution and add 100&#8211;300 &#181;l of primary antibody dissolved at the recommended concentration in Tris buffer. Incubate overnight. Remove the antibody solution and wash sections in Tris buffer twice for 5 min.</li>
		<li>Incubate with the secondary antibody for 1 hr at room temperature. Wash twice for 5 min in Tris buffer. Incubate 30 min using the ABC Elite Kit (follow the manufacturer&#8217;s instructions). Wash twice in Tris buffer.</li>
		<li>Develop antibody reaction by using the Vector VIP Kit, following the manufacturer&#8217;s instructions (1&#8211;7 min incubation generally produces a clear violet intensity). Put the slides in distilled water. Counterstain with Mayer&#8217;s hematoxylin for 30 sec.</li>
		<li>Wash in running water for 5 min. Place the slides in a staining jar with 70% ethanol for 1 min. Repeat by using a new staining jar with 70% ethanol. Place the slides in a staining jar with 95% ethanol for 1 min. Repeat by using a new staining jar with 95% ethanol.</li>
		<li>Place the slides in a staining jar with X-tra solv for 5 min. Remove, one at a time to keep moist. Place a drop of mounting medium on top of each slide and put a cover glass on top (do so carefully to avoid air bubbles). Let the slides dry overnight and view slides under a microscope.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

This study presents an easy-to-use approach to produce bladder wall patches with autologous tissue for transplantation at the surgical table. The patches are formed by the combination of a biodegradable polymer knitting in the middle and collagen with and without minced tissue in the outer surfaces in combination with plastic compression. Plastic compression is a method previously described by other authors and can be defined as a rapid expulsion of fluid from collagen gels12,13. Minced tissue of bladder mucosa or skin is seeded into this scaffold and the formation of a &#160;bladder or skin epithelium could be followed during 6 weeks. Immunohistochemical analyses showed the formation of equivalents characteristic of normal tissue. These results allow not only an in vitro model for studying re-epithelialization and wound healing, but also forms a biomaterial for autologous tissue transplantation. Most importantly for urology purposes, the autografts may be used for in vivo applications of tissue engineering in reconstructive urology as engineered autologous tissue patches without the requirement for in vitro culture before transplantation11.
In respect to cell expansion and in vitro culturing techniques, skin epithelium and uroepithelium share common characteristics. The motive to present the technique for plastic compression and mincing for both epithelial tissues in this study was that harvesting and in vivo studies for skin epithelium is easier than for bladder uroepithelium. By these means, biomaterials and skin epithelium can be studied as a first step to confirm biocompatibility and physical properties before performing more invasive studies in the urogenital organs.
In previous in vivo studies, using the concept of minced tissue with bladder mucosa for tissue regeneration of conduits, findings were that the small transplanted particles required some mechanical support in order to withstand mechanical trauma and to stay in place during the initial take of the transplanted minced particles and during the healing process and tissue regeneration6,7.
In order to overcome these weaknesses, the study aimed at developing a hybrid construct with a biodegradable core of a polymer knitted fabric and plastic-compressed collagen11,14. The collagen provides a favorable surface for cell attachment and growth and allows direct and rapid integration of the minced tissue into the scaffold. However, since the mechanical properties of collagen, even after plastic compression, are still weak and would not sustain the contractions and extensions of the natural bladder movements, a supporting core scaffold for the collagen was added. For more complex structures, transplanted tissue could be expanded around a prefabricated mold to produce a three-dimensional epithelialized structure with a central lumen.
In these studies we used PCL as a stabilizing polymer and the core of the biograft. PCL is widely used in a variety of biomedical devices such as dental fillings, absorbable synthetic surgical meshes and suture materials14. In order to procure maturation of the regenerated tissue, i.e. innervation, smooth muscle tissue and extra cellular matrix, we chose a PCL polymer with a slow degradation rate over several months. However, the degradation rate can be regulated by choice of polymer, thickness and density11,16.
Collagen was chosen as a component of the composite construct due to well-known biocompatibility, safety and good healing characteristics. The collagen promotes ingrowth of granulation tissue, re-modelling and maturation of extra-cellular matrix15. As collagen is degraded, neovascularization and granulation tissue supports the transplanted minced particles that reorganize, migrate and expand5-7,11,16. Besides, collagen is a major natural component of the extracellular matrix both in skin and in the bladder and has been used for creation of scaffolds both in vitro and in vivo including wound healing studies and reconstructions in the urogenital system17,18.
The critical steps within the protocol are easily manageable. First, the biopsy specimens have to be kept under moist and favorable conditions prior to insertion into the transplant or else cell growth will be delayed or absent. A second pitfall may involve getting the right solidity of the collagen. One must make sure that the collagen becomes stiff during the preparation of the autograft by avoiding bubbles inside the collagen gel. Bubbles are avoided by careful pipetting and small bubbles can be ruptured with a needle. Collagen with bubbles should be discarded.
It is also important to get the right size of the minced tissue; before mincing the tissue, make sure that only the thin bladder mucosa is used in cases of bladder expansion or only the epidermis in cases of skin expansion. The last pitfall is when suturing: make sure to move perpendicularly to the transplant with the needle in order to avoid separating the different layers. Edges of the autograft also need to be handled carefully in order not to separate the collagen from the polymer. After suturing the sheets will be harder to separate. These are common problems that may arise when learning the technique.
One limitation of this technique is that the cells in the minced particles rely on diffusion of nutrients and oxygen. Therefore, the thickness of the autograft has to be less than 1 mm and has to be placed in a well-vascularized site. This might be a disadvantage due to risk of leakage of urine through the transplanted part of the bladder. Depending on the plastic compression, different permeability constants can be achieved. In our case the hydraulic permeability (k) value was 0.034 according to previous studies19. In a clinical setting we anticipate that we would need to keep the bladder empty with catheters for approximately 1-2 weeks in order to give time for the urothelial cells to build up a multi-layered continuous urothelium that make the patch un-permeable before active use.
Another limitation is the assumption of a healthy organ for tissue harvest and expansion. In the more severe cases when the bladder is missing or is unfit for expansion, for example, when affected by cancer, the technique might not be suitable.
As for other tissue engineered cell constructs, final conclusions on functional and morphological results can only be answered in long-term in vivo studies. Our next step will be to analyze our results in long-term animal studies in respect to viability and physiological characteristics post-transplantation.
The significance of the presented technique is the possibility to expand tissue in vivo after only a single surgical procedure. Other techniques currently being evaluated are all dependent on expanding the cells in vitro before transplantation. The in vitro procedures have the downside of being associated with high costs, requiring advanced laboratory personnel, and being time-consuming. It may take up to 2 months between the harvesting of tissue or cells and a tissue-engineered autograft; as opposed to less than 1 hr, according to the technique described in this study.
In the future, minced tissue in compressed collagen techniques may be expanded and used in other organs, such as abdominal wall defects, diaphragmatic hernias, and other conditions where a defect&#160;is not easily reconstructed with existing tissue.
In conclusion, an easy-to-perform method is described for transplantation of skin and bladder mucosa that can expand in vivo. The procedure of preparing an autograft is easy to do and can be used under sterile conditions in an ordinary surgical setting. The autograft resists surgical handling and is biodegradable. The core of the autograft can be composed of different biodegradable polymers depending on preferences regarding the rate of degradation and other characteristics, such as elasticity, thickness, and porosity, and according to the patient&#8217;s specific needs. In a clinical setting, the patient would undergo tissue harvest, preparation of the composite autograft and autotransplantation as a single-stage procedure. In addition, all parts of the cell-containing bioconstructs are currently FDA approved.

Most tissue engineering studies on transplantation to the skin and urogenital tract include autologous cell harvests from healthy tissue and cell expansion in specially equipped cell-culturing facilities1,2.
After cell expansion, cells are usually stored for later use when the patient is prepared to receive the autograft. Nitrogen freezers allow long-term storage at low temperatures of -150 &#176;C or lower. The process of freezing must be careful and controlled in order not to lose the cells. One risk of cell death is crystallization of intracellular water during the thawing process, which can lead to rupture of the cell membranes. Cell freezing is usually performed by slow and controlled cooling (-1 &#176;C per min), using a high concentration of cells, fetal bovine serum, and dimethyl sulfoxide. After thawing, the cells need to be processed again by removing the freezing medium and culturing on cell culture plastic or a biomaterial before transplantation back to the patient.
All the above-mentioned steps are time-consuming, laborious, and costly3. In addition, all in vitro processing of cells intended for patient transplantation are highly regulated and requires well-trained and accredited personnel and laboratories4. All in all, to procure a safe and reliable manufacturing process, the technique could only be established in a very small number of technically advanced centers and a wider use in common surgical disorders is doubtful.
In order to overcome the limitations of cell culturing in the laboratory environment, the concept of transplanting minced tissue for cell expansion in vivo is introduced by using the body itself as a bioreactor. For these purposes, the autografts would preferentially be transplanted on a 3D mold according to the shape that is needed for the final reconstruction of the organ of interest5-7.
Originally, the idea of transplanting minced epithelium was presented by Meek in 1958&#160;when he described how epithelium grows from the edges of a wound. He demonstrated that a small piece of skin would increase its margins and thereby its potential for cell expansion by 100% by cutting the piece twice in perpendicular directions (Figure 1)8. The theory has been supported by the use of meshed partial thickness skin grafts for skin transplantation9 and in skin wound healing models10.
<img alt="Figure 1" src="/files/ftp_upload/53061/53061fig1.jpg" width="600" /> 
Figure 1: Meek theory. According to Meek&#8217;s theory, epithelium grows from the edges of a wound. By increasing the area exposed by the mincing technology, minced tissue epithelializes wounds from many spots.
The present study is based on the hypothesis that the same principle could be applied to the subcutaneous tissue by placing minced epithelium around a mold. The epithelial cells would mobilize from minced transplants (reorganize), cover the wound areas (migrate) and divide (expand) in order to form a continuous neoepithelium that covers the wound area and separates the foreign body (the mold) from the inner body (Figure 2).
<img alt="Figure 2" src="/files/ftp_upload/53061/53061fig2.jpg" /> 
Figure 2: Cartoon of a 3D mold with minced epithelium for in vivo intracorporal tissue expansion according to the theory of Meek. By using minced tissue placed on a mold and then transplanted to the subcutaneous tissue, the hypothesis is that the epithelial cells migrate from the edges of the minced tissue, reorganize, and expand so as to form a continuous neoepithelium that covers the wound area and separates the foreign body (the mold) from the inner body.
Although previous in vivo studies show promising results, further improvements could be achieved by reinforcing the autografts so that the regenerated epithelium could withstand mechanical trauma better7. For these purposes, important prerequisites for a successful biomaterial were identified, such as: easy diffusion of nutrients and waste products, possibility to mold in a 3D manner and easiness of surgical handling. Conclusions were made that these needs could be met by adding a composite biomaterial to the minced tissue.
The current study aimed at developing a scaffold composed of minced tissue in plastic-compressed collagen containing a reinforcing core of a biodegradable fabric. By these means, viable cells could migrate from the minced tissue particles and proliferate with morphological features characteristic of the original epithelium (skin or urothelium). Using plastic compression, the scaffold was reduced in size from 1 cm to about 420&#8201;&#181;m as the minced particles were encased in the upper layer collagen. The core fabric could be any polymer but needs to be modified with a hydrophilic surface in order to interlink with the covering collagen layers11.
The method provided an enhanced scaffold integrity by incorporating a knitted mesh consisting of poly(&#949;-caprolactone) (PCL) within two plastic compressed collagen gels using it as a scaffold for culturing minced bladder mucosa or minced skin from pigs. The construct was maintained in cell culture conditions for up to 6 weeks in vitro, demonstrating successful formation of a stratified, multilayered urothelium or squamous skin epithelium on the top of a well-consolidated hybrid construct. The construct was easy to handle and could be sutured in place for bladder augmentation purposes or covering of skin defects. All parts of the tissue scaffold are FDA-approved and the technique could be used for single-stage procedures by tissue harvesting, mincing, plastic compression, and transplanting back to the patient as a single-staged intervention. The procedure could be performed for tissue expansion and reconstruction under sterile conditions in any general surgery unit.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Silicone catheter 10-French</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>DMEM 10X</td>
			<td>Gibco</td>
			<td>31885-023</td>
			<td>Plastic compression section 4</td>
		</tr>
		<tr>
			<td>24 well plates</td>
			<td>Falcon</td>
			<td>08-772-1</td>
			<td>Plastic compression section 4</td>
		</tr>
		<tr>
			<td>3&#39;3,&#39;5-Triiodothyronine</td>
			<td>Sigma-Aldrich</td>
			<td>IRMM469</td>
			<td>&#160; In vitro culture; Section 5</td>
		</tr>
		<tr>
			<td>4% PFA</td>
			<td>Labmed Solutions</td>
			<td>200-001-8</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>70% ethanol</td>
			<td>Histolab</td>
			<td></td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>ABC Elite kit: Biotin -Streptavidin detection kit</td>
			<td>Vector</td>
			<td>PK6102</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Absolute ethanol</td>
			<td>Histolab</td>
			<td>1399.01</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Adenine</td>
			<td>Sigma-Aldrich</td>
			<td>A8626</td>
			<td>&#160; In vitro culture; Section 5</td>
		</tr>
		<tr>
			<td>Atropine 25 &#956;g/kg&#160;</td>
			<td>Temgesic, RB Pharmaceuticals, Great Britain</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Azaperone 2&#8201;mg/kg&#160;</td>
			<td>Stresnil, Janssen-Cilag, Pharma, Austria</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Biosafety Level 2 hood&#160;</td>
			<td></td>
			<td></td>
			<td>Plastic compression; Section 4</td>
		</tr>
		<tr>
			<td>Blocking solution:&#160; Normal serum from the same species as the secondary secondary antibody was generated in.</td>
			<td>Vector</td>
			<td>The blocking solution depends of the&#160; origin of&#160; first antibody</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Buprenorphine 45 &#956;g/kg</td>
			<td>Atropin, Mylan Inc, Canonsburg, PA</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Carprofen 3&#8201;mg/kg&#160;&#160;&#160;</td>
			<td>Rimadyl, Orion Pharma, Sweden</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Chlorhexidine gluconate&#160;</td>
			<td>Hibiscrub 40&#8201;mg/mL, Regent Medical, England</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Cholera toxin&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>C8052</td>
			<td>&#160; In vitro culture; Section 5</td>
		</tr>
		<tr>
			<td>Coplin jar: staining jar for boiling</td>
			<td>Histolab</td>
			<td>6150</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Stainless mold&#160; (33x22x10 mm) custom made</td>
			<td></td>
			<td></td>
			<td>Plastic compression; Section 4</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Gibco</td>
			<td>3188-5023</td>
			<td>Plastic compression section 4. Keep on ice&#160; when using it in plastic compression</td>
		</tr>
		<tr>
			<td>Epidermal growth factor</td>
			<td>Sigma-Aldrich</td>
			<td>E9644</td>
			<td>&#160; In vitro culture; Section 5</td>
		</tr>
		<tr>
			<td>Ethilon (non-absorbable monofilament for skin sutures)</td>
			<td>Ethicon</td>
			<td></td>
			<td>Surgery, Section 1</td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Gibco</td>
			<td>10437-036</td>
			<td>Plastic compression section 4</td>
		</tr>
		<tr>
			<td>Forceps (Adison with tooth)</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Gauze (Gazin Mullkompresse)&#160;</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Ham&#180;s F12</td>
			<td>Gibco</td>
			<td>31765-027</td>
			<td>Plastic compression section 4</td>
		</tr>
		<tr>
			<td>Hematoxylin</td>
			<td>Histolab</td>
			<td>1820</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Humidity chamber</td>
			<td>DALAB</td>
			<td></td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Hydrocortisone</td>
			<td>Sigma-Aldrich</td>
			<td>H0888</td>
			<td>&#160; In vitro culture; Section 5</td>
		</tr>
		<tr>
			<td>Hydrogen peroxide Solution 30%</td>
			<td>Sigma-Aldrich</td>
			<td>H1009</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>Sigma-Aldrich</td>
			<td>I3536</td>
			<td>&#160; In vitro culture; Section 5</td>
		</tr>
		<tr>
			<td>Isoflurane&#160;</td>
			<td>Isoflurane, Baxter, Deerfield, IL</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Lidocaine 5 mg/ml</td>
			<td>Xylocaine, AstraZeneca, Sweden</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Lucose 25&#8201;mg/mL&#160;</td>
			<td>Baxter, Deerfield, IL</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Marker pen pap pen</td>
			<td>Sigma-Aldrich</td>
			<td>Z377821-1EA</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Medetomidine 25 &#956;g/kg&#160;</td>
			<td>Domitor, Orion Pharma, Sweden</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Mincing device</td>
			<td>Applied Tissue Technologies LLC</td>
			<td></td>
			<td>&#160;Minced tissue preparation, section 2</td>
		</tr>
		<tr>
			<td>Monocryl (absorbable monofilament)</td>
			<td>Ethicon</td>
			<td></td>
			<td>Surgery, Section 1</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>NaOH 1N</td>
			<td>Merck Millipore</td>
			<td>106462</td>
			<td>Plastic compression section 4 and cell culture</td>
		</tr>
		<tr>
			<td>Nylon mesh, 110 uM thick pore size 0.04 sqmm</td>
			<td></td>
			<td></td>
			<td>Plastic compression; Section 4</td>
		</tr>
		<tr>
			<td>Oculentum simplex APL: ointment for eye protection</td>
			<td>APL</td>
			<td>Vnr 336164</td>
			<td>Surgery, Section 1</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Gibco</td>
			<td>14190-094</td>
			<td>Plastic compression section 4</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td>Plastic compression section 4</td>
		</tr>
		<tr>
			<td>Phenobarbiturate 15&#8201;mg/kg&#160;</td>
			<td>Pentobarbital, APL, Sweden</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>PLGA Knitted fabric</td>
			<td></td>
			<td></td>
			<td>Plastic compression; Section 4</td>
		</tr>
		<tr>
			<td>Rat-tail collagen</td>
			<td>First LINK, Ltd, UK</td>
			<td>60-30-810</td>
			<td>Plastic compression section 4, keep on ice</td>
		</tr>
		<tr>
			<td>Scalpel blade - 15</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Shaving shears</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Stainless stell mesh, 400 uM thick pore size&#160;</td>
			<td></td>
			<td></td>
			<td>Plastic compression; Section 4</td>
		</tr>
		<tr>
			<td>Steril gloves</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Sterile gowns</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Sterile drapes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sterilium</td>
			<td>Bode Chemie HAMBURG</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Suture Thread Ethilon</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>TE-solution (antigen unmasking solution) consist of 10 mM Tris and 1 mM EDTA, pH 9.0</td>
			<td></td>
			<td></td>
			<td>10 mM Tris/1 mM EDTA,&#160; adjust pH to&#160; 9.0</td>
		</tr>
		<tr>
			<td>Tiletamine hypochloride 2,5&#8201;mg/kg</td>
			<td></td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Transferrin&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>T8158</td>
			<td>&#160; In vitro culture; Section 5</td>
		</tr>
		<tr>
			<td>Trizma Base, H2NC&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>T6066</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Vector VIP kit: Enzyme&#160; peroxidase substrate&#160; kit</td>
			<td>Vector</td>
			<td>SK4600</td>
			<td>Immunocytochemistry; Section 6</td>
		</tr>
		<tr>
			<td>Vicryl (absorbable braded)</td>
			<td>Ethicon</td>
			<td></td>
			<td>Surgery, Section 1</td>
		</tr>
		<tr>
			<td>Tris buffer pH 7.6 (washing buffer)</td>
			<td></td>
			<td></td>
			<td>TE solution: Make 10X&#160; (0,5M Tris, 1,5M NaCl) by mixing: 60,6 g Tris (Trizma Base, H2NC(CH2OH)3, M=121.14 g/mol), add 800 ml&#160; distilled water adjust the pH till 7.6, add 87,7 g NaCl and fill to 1000 ml with&#160; distilled water. Dilute to 1X with distilled water.</td>
		</tr>
		<tr>
			<td>X-tra solv (solvent)</td>
			<td>DALAB</td>
			<td>41-5213-810</td>
			<td>Immunocytochemistry; Section 6. Use under fume hood</td>
		</tr>
		<tr>
			<td>Zolazepam hypochloride</td>
			<td>Zoletil, Virbac, France</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Depilatory wax strips</td>
			<td>Veet</td>
			<td></td>
			<td>Preparing the animal for surgery , Section 1</td>
		</tr>
		<tr>
			<td>Pentobarbital sodium</td>
			<td>Lundbeck</td>
			<td></td>
			<td>Termination, Section 3</td>
		</tr>
	</tbody>
</table>

minced tissue in compressed collagen: a cell-containing biotransplant for single-staged reconstructive repair

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped human cell culture facilities. These methods include isolation of cells in single cell suspension and several laborious and time-consuming events before transplantation back to the patient. Previous studies suggest that the body itself could be used as a bioreactor for cell expansion and regeneration of tissue in order to minimize ex vivo manipulations of tissues and cells before transplanting to the patient. The aim of this study was to demonstrate a method for tissue harvesting, isolation of continuous epithelium, mincing of the epithelium into small pieces and incorporating them into a three-layered biomaterial. The three-layered biomaterial then served as a delivery vehicle, to allow surgical handling, exchange of nutrition across the transplant, and a controlled degradation. The biomaterial consisted of two outer layers of collagen and a core of a mechanically stable and slowly degradable polymer. The minced epithelium was incorporated into one of the collagen layers before transplantation. By mincing the epithelial tissue into small pieces, the pieces could be spread and thereby the propagation of cells was stimulated. After the initial take of the transplants, cell expansion and reorganization would take place and extracellular matrix mature to allow ingrowth of capillaries and nerves and further maturation of the extracellular matrix. The technique minimizes ex vivo manipulations and allow cell harvesting, preparation of autograft, and transplantation to the patient as a simple one-stage intervention. In the future, tissue expansion could be initiated around a 3D mold inside the body itself, according to the specific needs of the patient. Additionally, the technique could be performed in an ordinary surgical setting without the need for sophisticated cell culturing facilities.

This study presents a method that shows how to produce a biomaterial for transplantation using plastic compression of collagen and minced tissue.
Bladder mucosa and skin can be harvested and then mechanically minced into small particles (Figure 3). By plastic compression, the minced particles are incorporated within the composite scaffold composed of a centrally placed biodegradable polymer that is mechanically strong within outer layers of a collagen gel (Figure 4). Minced tissue particles can be separated to allow a 1:6 expansion rate. By pressing out the water content of the collagen, a biotransplant for autologous reconstructive surgery is completed.
Biotransplants can be used for autotransplantation to the animal for in vivo research studies or cultured in an ordinary cell culture environment for further in vitro studies.
The composite autograft allows grasping and suturing and sustains surgical handling (Figure 5). The process from tissue harvest and preparation of the cell-containing autograft for reconstructive surgery takes approximately 20 min and is aimed to be performed as a single-staged procedure.
In in vitro studies cells migrate, expand and reorganize to the surface of the autograft into a single-cell continuous layer in two weeks. After four weeks, the continuous epithelium is approximately 4 cell layers thick (Figure 6). Immunohistochemistry at different points in time reveals that cells proliferate, migrate, and reorganize into a continuous epithelium with a microarchitecture typical for the cell phenotype. The same results apply for autografts with minced skin epithelium as for minced bladder mucosa.

<img alt="Figure 3" src="/files/ftp_upload/53061/53061fig3.jpg" /> 
Figure 3: Minced tissue preparation and plastic compression. The preparation of bladder mucosa (A) for mincing (C) and plastic compression (D&#8211;F), using the mincing device in (B) and the mold in (D) to produce a 420-&#956;m-thick transplant (H). Note that collagen itself is a good extracellular matrix but is difficult to handle mechanically (G); by placing a biodegradable fabric as a reinforcing core, the transplant becomes easy to grasp (H).

<img alt="Figure 4" src="/files/ftp_upload/53061/53061fig4.jpg" /> 
Figure 4: Plastic compression. An animated cartoon showing the process of plastic compression with minced particles.
<img alt="Figure 5" src="/files/ftp_upload/53061/53061fig5.jpg" /> 
Figure 5: Surgical handling. Full-thickness skin wound model in a rat demonstrating sutured plastic-compressed biomaterial with minced skin for autotransplantation marked as T (transplanted autograft) Sutured biomaterial without minced skin marked as S (sham). In this case, minced skin epithelium was transplanted for a 1:3 expansion rate.
<img alt="Figure 6" src="/files/ftp_upload/53061/53061fig6.jpg" /> 
Figure 6: Immunohistochemical staining to visualize the progression of the 3D culture in the collagen-biomaterial minced skin and bladder mucosa. (A&#8211;D) Hematoxylin/eosin staining of (A) native skin, (B) native bladder, (C) minced skin after 5 weeks in culture and (D) minced bladder after 6 weeks in culture. The expression of epithelial and proliferation markers with MNF116 (E) and Ki 67 antibodies (F), respectively, after 2 weeks in culture (minced bladder).

Watch this Scientific Journal Video about Minced Tissue in Compressed Collagen: A Cell-containing Biotransplant for Single-staged Reconstructive Repair at JoVE.com

Minced Tissue in Compressed Collagen: A Cell-containing Biotransplant for Single-staged Reconstructive Repair

Tissue engineering often includes in vitro expansion in order to create autografts for tissue regeneration. In this study a method for tissue expansion, regeneration, and reconstruction in vivo was developed in order to minimize the processing of cells and biological materials outside the body.

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped ...

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9.9K Views.  Karolinska Institutet. Tissue engineering often includes in vitro expansion in order to create autografts for tissue regeneration. In this study a method for tissue expansion, regeneration, and reconstruction in vivo was developed in order to minimize the processing of cells and biological materials outside the body.

Video: Minced Tissue in Compressed Collagen: A Cell-containing Biotransplant for Single-staged Reconstructive Repair

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

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

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

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

Fabrication and Application of Rose Bengal-chitosan Films in Laser Tissue Repair

Photochemical tissue bonding (PTB) is a sutureless technique for tissue repair, which is achieved by applying a solution of rose bengal (RB) between two tissue edges1,2. These are then irradiated by a laser that is selectively absorbed by the RB. The resulting photochemical reactions supposedly crosslink the collagen fibers in the tissue with minimal heat production3. In this report, RB has been incorporated in thin chitosan films to fabricate a novel tissue adhesive that is laser-activated. Adhesive films, based on chitosan and containing ~0.1 wt% RB, are fabricated and bonded to calf intestine and rat tibial nerves by a solid state laser (&lambda;=532 nm, Fluence~110 J/cm2, spot size~0.5 cm). A single-column tensiometer, interfaced with a personal computer, is used to test the bonding strength. The RB-chitosan adhesive bonds firmly to the intestine with a strength of 15 &plusmn; 6 kPa, (n=30). The adhesion strength drops to 2 &plusmn; 2 kPa (n=30) when the laser is not applied to the adhesive. The anastomosis of tibial nerves can be also completed without the use of sutures. A novel chitosan adhesive has been fabricated that bonds photochemically to tissue and does not require sutures.

Sutures are usually needed to repair tissue during surgical procedures. However, their application can be problematic as they are invasive and may damage tissue. The fabrication and application methods of a novel tissue adhesive are here reported. This adhesive film is laser-activated and does not require the use of sutures.

Photochemical tissue bonding (PTB) is a sutureless technique for tissue repair, which is achieved by applying a solution of rose bengal (RB) between two ...

An Improved Method for the Preparation of Type I Collagen From Skin

Soluble type 1 collagen (COL1) is used extensively as an adhesive substrate for cell cultures and as a cellular scaffold for regenerative applications. Clinically, this protein is widely used for cosmetic surgery, dermal injections, bone grafting, and reconstructive surgery. The sources of COL1 for these procedures are commonly nonhuman, which increases the potential for inflammation and rejection as well as xenobiotic disease transmission. In view of this, a method to efficiently and quickly purify COL1 from limited quantities of autologously-derived tissues would circumvent many of these issues; however, standard isolation protocols are lengthy and often require large quantities of collagenous tissues. Here, we demonstrate an efficient COL1 extraction method that reduces the time needed to isolate and purify this protein from about 10 days to less than 3 hr. We chose the dermis as our tissue source because of its availability during many surgical procedures. This method uses traditional extraction buffers combined with forceful agitation and centrifugal filtration to obtain highly-pure, soluble COL1 from small amounts of corium. Briefly, dermal biopsies are washed thoroughly in ice-cold dH2O after removing fat, connective tissue, and hair. The skin samples are stripped of noncollagenous proteins and polysaccharides using 0.5 M sodium acetate and a high speed bench-top homogenizer. Collagen from residual solids is subsequently extracted with a 0.075 M sodium citrate buffer using the homogenizer. These extracts are purified using 100,000 MW cut-off centrifugal filters that yield COL1 preparations of comparable or superior quality to commercial products or those obtained using traditional procedures. We anticipate this method will facilitate the utilization of autologously-derived COL1 for a multitude of research and clinical applications.

Traditional procedures for the isolation of soluble type 1 collagen (COL1) require about 10 days from start to finish because of lengthy buffer incubations and laborious resuspensions of fibrils. Here, we describe a means to purify COL1 from small dermal biopsies in less than 3 hr.

Soluble type 1 collagen (COL1) is used extensively as an adhesive substrate for cell cultures and as a cellular scaffold for regenerative applications. ...

Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides

Collagen is a major structural component of the extracellular matrix that supports tissue formation and maintenance. Although collagen remodeling is an integral part of normal tissue renewal, excessive amount of remodeling activity is involved in tumors, arthritis, and many other pathological conditions. During collagen remodeling, the triple helical structure of collagen molecules is disrupted by proteases in the extracellular environment. In addition, collagens present in many histological tissue samples are partially denatured by the fixation and preservation processes. Therefore, these denatured collagen strands can serve as effective targets for biological imaging. We previously developed a caged collagen mimetic peptide (CMP) that can be photo-triggered to hybridize with denatured collagen strands by forming triple helical structure, which is unique to collagens. The overall goals of this procedure are i)&#160;to image denatured collagen strands resulting from normal remodeling activities in vivo, and ii) to visualize collagens in ex vivo tissue sections using the photo-triggered caged CMPs. To achieve effective hybridization and successful in vivo and ex vivo imaging, fluorescently labeled caged CMPs are either photo-activated immediately before intravenous injection, or are directly activated on tissue sections. Normal skeletal collagen remolding in nude mice and collagens in prefixed mouse cornea tissue sections are imaged in this procedure. The imaging method based on the CMP-collagen hybridization technology presented here could lead to deeper understanding of the tissue remodeling process, as well as allow development of new diagnostics for diseases associated with high collagen remodeling activity.

Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides

This procedure demonstrates in vivo near IR fluorescence imaging of collagen remodeling activities in mice as well as ex vivo staining of collagens in tissue sections using caged collagen mimetic peptides that can be photo-triggered to hybridize with denatured collagen strands.

Collagen is a major structural component of the extracellular matrix that supports tissue formation and maintenance. Although collagen remodeling is an ...

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. ...

Engineered 3D Silk-collagen-based Model of Polarized Neural Tissue

Despite huge efforts to decipher the anatomy, composition and function of the brain, it remains the least understood organ of the human body. To gain a deeper comprehension of the neural system scientists aim to simplistically reconstruct the tissue by assembling it in vitro from basic building blocks using a tissue engineering approach. Our group developed a tissue-engineered silk and collagen-based 3D brain-like model resembling the white and gray matter of the cortex. The model consists of silk porous sponge, which is pre-seeded with rat brain-derived neurons, immersed in soft collagen matrix. Polarized neuronal outgrowth and network formation is observed with separate axonal and cell body localization. This compartmental architecture allows for the unique development of niches mimicking native neural tissue, thus enabling research on neuronal network assembly, axonal guidance, cell-cell and cell-matrix interactions and electrical functions.

Insight into the complex actions of the brain requires advanced research tools. Here we demonstrate a novel silk-collagen-based 3D engineered model of neural tissue resembling brain-like architecture. The model can be used to study neuronal network assembly, axonal guidance, cell-cell interactions and electrical activity.

Despite huge efforts to decipher the anatomy, composition and function of the brain, it remains the least understood organ of the human body. To gain a ...

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D culture of cells in collagen type I gels helps to stabilize cell morphology and function, which is necessary for creating microfluidic tissue models in microdevices. Translating traditional 3D culture techniques for tissue culture plates to microfluidic devices is often difficult because of the limited channel dimensions. In this method, we describe a technique for modifying native type I collagen to generate polycationic and polyanionic collagen solutions that can be used with layer-by-layer deposition to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These thin collagen layers stabilize cell morphology and function, as shown using primary hepatocytes as an example cell, allowing for the long term culture of microtissues in microfluidic devices.

The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D ...

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from traumatic injury, disease and various congenital, genetic and acquired conditions are quite common. Tissue engineering and regenerative medicine technologies have enormous potential to provide a therapeutic solution. However, utilization of biologically relevant animal models in combination with longitudinal assessments of pertinent functional measures are critical to the development of improved regenerative therapeutics for treatment of VML-like injuries. In that regard, a commercial muscle lever system can be used to measure length, tension, force and velocity parameters in skeletal muscle. We used this system, in conjunction with a high power, bi-phase stimulator, to measure in vivo force production in response to activation of the anterior crural compartment of the rat hindlimb. We have previously used this equipment to assess the functional impact of VML injury on the tibialis anterior (TA) muscle, as well as the extent of functional recovery following treatment of the injured TA muscle with our tissue engineered muscle repair (TEMR) technology. For such studies, the left foot of an anaesthetized rat is securely anchored to a footplate linked to a servomotor, and the common peroneal nerve is stimulated by two percutaneous needle electrodes to elicit muscle contraction and dorsiflexion of the foot. The peroneal nerve stimulation-induced muscle contraction is measured over a range of stimulation frequencies (1-200 Hz), to ensure an eventual plateau in force production that allows for an accurate determination of peak tetanic force. In addition to evaluation of the extent of VML injury as well as the degree of functional recovery following treatment, this methodology can be easily applied to study diverse aspects of muscle physiology and pathophysiology. Such an approach should assist with the more rational development of improved therapeutics for muscle repair and regeneration.

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

We describe an in vivo protocol to measure dorsiflexion of the foot following stimulation of the peroneal nerve and contraction of the anterior crural compartment of the rat hindlimb. Such measurements are an indispensable translational tool for evaluating skeletal muscle pathology and tissue engineering approaches to muscle repair and regeneration.

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from ...

Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model

Tissue engineering is a promising field, focused on developing solutions for the increasing demand on tissues and organs regarding transplantation purposes. The process to generate such tissues is complex, and includes an appropriate combination of specific cell types, scaffolds, and physical or biochemical stimuli to guide cell growth and differentiation. Microcarriers represent an appealing tool to expand cells in a three-dimensional (3D) microenvironment, since they provide higher surface-to volume ratios and mimic more closely the in vivo situation compared to traditional two-dimensional methods. The vascular system, supplying oxygen and nutrients to the cells and ensuring waste removal, constitutes an important building block when generating engineered tissues. In fact, most constructs fail after being implanted due to lacking vascular support. In this study, we present a protocol for endothelial cell expansion on recombinant collagen-based microcarriers under dynamic conditions in spinner flask and bioreactors, and we explain how to determine in this setting cell viability and functionality. In addition, we propose a method for cell delivery for vascularization purposes without additional detachment steps necessary. Furthermore, we provide a strategy to evaluate the cell vascularization potential in a perfusion bioreactor on a decellularized&#160;biological matrix. We believe that the use of the presented methods could lead to the development of new cell-based therapies for a large range of tissue engineering applications in the clinical practice.

We propose a cell expansion protocol on macroporous microcarriers and their use as delivery system in a perfusion bioreactor to seed a decellularized tissue matrix. We also include different techniques to determine cell proliferation and viability of cells cultured on microcarriers. Furthermore, we demonstrate functionality of cells after bioreactor cultures.