To date, thick tissue defects are typically reconstructed by applying autologous tissue flaps or engineered tissues. In this protocol, we present a new method for engineering vascularized tissue flap bearing an autologous pedicle, to serve as a substitute to autologous flaps.
One of the main factors limiting the thickness of a tissue construct and its consequential viability and applicability in vivo, is the control of oxygen supply to the cell microenvironment, as passive diffusion is limited to a very thin layer. Although various materials have been described to restore the integrity of full-thickness defects of the abdominal wall, no material has yet proved to be optimal, due to low graft vascularization, tissue rejection, infection, or inadequate mechanical properties. This protocol describes a means of engineering a fully vascularized flap, with a thickness relevant for muscle tissue reconstruction. Cell-embedded poly L-lactic acid/poly lactic-co-glycolic acid constructs are implanted around the mouse femoral artery and vein and maintained in vivo for a period of one or two weeks. The vascularized graft is then transferred as a flap towards a full thickness defect made in the abdomen. This technique replaces the need for autologous tissue sacrifications and may enable the use of in vitro engineered vascularized flaps in many surgical applications.
Abdominal wall defects often arise following severe trauma, cancer treatment, burns and removal of infected mesh. These defects often involve significant tissue loss, requiring complicated surgical procedures and presenting a major challenge for plastic reconstruction surgeons 1-4. Tissue engineering researchers seeking new sources for artificial tissues have explored different materials, cell sources and growth factors. Successful restorations of various tissues, such as trachea 5,6, bladder 7, cornea 8, bone 9and skin 10, by implantation of engineered tissues were previously reported. However, fabrication of a thick vascularized engineered tissue, particularly for reconstruction of large defects, remains a significant challenge in tissue engineering.
One of the main factors limiting the thickness of a viable tissue construct is the control of oxygen supply to its constituent cells. When relying on diffusion, construct thickness is limited to that of a very thin layer. The maximum distance between oxygen- and nutrient-supplying capillaries in vivo is approximately 200 µm, which correlates with the diffusion limit of oxygen11,12. Insufficient vascularization can result in tissue ischemia and escalate to tissue resorption or necrosis 13.
In addition, the ideal material used for tissue reconstruction must be biocompatible and non-immunogenic. It must also be capable of promoting further integration of host cells with the biomaterial, and maintaining structural integrity. Various biological 14-16 and synthetic 1,17,18 matrices have been previously explored for tissue reconstruction, however their use remain limited due to lack of effective blood supply, infections or insufficient tissue strength.
In this study, a biocompatible, cell-embedded scaffold comprised of Food and Drug Administration (FDA)-approved poly L-lactic acid (PLLA)/poly lactic-co-glycolic acid (PLGA), was implanted around the femoral artery and vein (AV) vessels of a nude mouse and separated from the surrounding tissue, ensuring vascularization from the AV vessels only. One week post-implantation, the graft was viable, thick and well vascularized. This thick vascularized tissue with the AV vessels, was then transferred as a pedicled flap to an abdominal full-thickness defect in the same mouse. One week post-transfer, the flap was viable, vascularized and well integrated with the surrounding tissue, bearing sufficient strength to support abdominal viscera. Thus, the engineered thick, vascularized tissue flap, bearing an autologous pedicle, presents a novel method for repairing full-thickness abdominal wall defects.
All animal studies were approved by the Committee of the Ethics of Animal Experiments of the Technion. For this procedure, athymic nude mice were used to avoid immunological rejection. If using another type of mouse, the mice should be shaved prior to the surgical procedure and administration of cyclosporine (or another anti-rejection substitute) is recommended.
1. Scaffold Preparation and Cell Embedding
2. Before Starting the Surgical Procedure
3. Flap Construction (Graft Implantation)
4. Flap Transfer
5. Ultrasound Determination of Vascular Perfusion of the Graft
NOTE: Before transfer, vascular perfusion of the graft is measured at one and two weeks after implantation, by ultrasonography.
6. Determination of the Extent of Graft Vascularization
NOTE: The extent of tissue graft vascularization is determined one or two weeks after implantation.
7. Immunohistological and Histological Staining of the Graft
8. Mechanical Properties Assessment
Graft vascularization and perfusion in vivo
The grafts were implanted one or two weeks prior to their transfer as axial flaps. At one and two weeks post-implantation, gross observation of the graft area revealed viable and vascularized tissue grafts. These grafts proved to be highly vascularized, as determined by positive CD31 immunostaining (Figure 1A), and highly perfused, as evidenced by FITC-dextran tail vein injection and ultrasound measurements. Many vessels were already observed at one-week post-implantation, a number that rose significantly after an additional week in the vicinity of the AV vessels. FITC-dextran-based determination of the functional vessel density (FVD) (Figure 1B) showed that the graft was highly vascularized at one-week postimplantation and that no significant changes took place in the additional week in vivo. Vessel patency and perfusion were demonstrated by ultrasound imaging (the FVD was 5.71 mm–1 ± 0.51 mm–1and 10.28 mm–1 ± 2.71 mm–1, one and two weeks post-implantation, respectively). Figure 1C confirmed host femoral vessel patency and integrity after graft implantation. Moreover, ultrasonographic examination revealed perfusion within the graft area, which was slightly higher two weeks post-implantation as compared to one week post-implantation (Figure 1D).
Flap properties
Gross examination of the flaps one week post-transfer revealed a viable, vascularized and well integrated tissue. The flaps underwent firm attachment to their surroundings. When comparing the pre-vascularized, cell-embedded flaps to control acellular, nonvascularized grafts, the former showed superior mechanical properties, as manifested by increased stiffness and strength (Figure 2). We also observed that wound dehiscence and herniation occurred less frequently in mice treated with cell-embedded flaps, when compared to animals treated with control grafts, which can be attributed to the increased mechanical strength of the transplanted tissue.
Figure 1. Graft vascularization. (A) The density of CD31-positive vessels, measured at one and two weeks postimplantation compared to control group. All values are normalized to the graft area (mm2). *= T-test p <0.05. (B) Functional vascular density (FVD) at one and two weeks postimplantation. No significant difference was observed, p = 0.08 (T-test). (C) Patent AV vessels as imaged using ultrasound in the color Doppler mode. Blue and red represent the blood flow in the graft. The yellow quadrant outlines the graft area. (D) Graft perfusion at one and two weeks postimplantation. (E and F) H&E staining of the flaps integrated with the host tissue: black arrow point viable artery; white arrows point abdominal muscle fibers; yellow arrows point scaffold remains. (H and G) FITC dextran distribution as shown by confocal microscopy images. For all determinations, the sample size was n ≥ 3 and all values are represented as mean ± standard error of the mean. Please click here to view a larger version of this figure.
Figure 2. Mechanical properties of the flap one week post-transfer. (A) The stiffness and (B) the ultimate tensile strength (UTS) of the flaps. Control stands an empty graft without cells, cells stands tri-culture grafts. * = T-test p <0.05, n = 3. Values are represented as mean ± standard error of the mean. Please click here to view a larger version of this figure.
The advances in tissue engineering have been met with a growing demand for substitute tissues for reconstruction of various tissue types. A variety of synthetic 1,17,18 and biological 14-16 materials as well as fabrication methods have been assessed for their capacity to address these demands. However, despite the progress in clinical care and in tissue engineering, the restoration of full-thickness abdominal wall defects remains a challenge. A tissue adequate for reconstruction of such massive defects must be (1) thick and (2) vascularized, and demonstrate (3) mechanical integrity, and (4) viability over time 23.
Here, we present a step-by-step detailed protocol for creating a thick, viable, and vascularized tissue flap, which can serve as an alternative to autologous tissue flaps. The fabricated flap was constructed in two steps: (1) A PLLA/PLGA scaffold was implanted around the AV vessels of the mouse hindlimb and separated from the surrounding tissue to allow vascularization by the AV vessels only. (2) The created vascularized, thick tissue flap was then transferred with the AV vessels, which served as its pedicle, into a full-thickness abdominal wall defect.
Proper flap vascularization is essential for its successful integration within the host 17,18,24. Various approaches to create vascularized engineered tissue in order to improve oxygen supply and diffusion in thick tissues, have been discussed in the literature. Among these were methods involving seeding of endothelial cells (ECs) on various scaffold types 13,20,25-30, various techniques to supply angiogenic factors to the implantation site (either by direct administration by injection, use of transformed cells expressing the factors 31-34 or slow release from different scaffolds 35,36), use of bioreactors to ensure engineered tissue perfusion 37 and employment of AV loop chambers 38,39. Here, the tissue graft was vascularized in vivo, by exploitation of autologous vessels. The graft, which proved viable, vascularized and perfused, was then transferred as a thick tissue flap to repair a full-thickness abdominal wall defect. One week post-transfer, the flap was viable and featured sufficient mechanical strength to support the abdominal viscera. Here, we used athymic nude mice, which bear an impaired immune system; when using any other type of mouse or other animal, the possibility of immune reactions should be taken into consideration (especially when seeding the scaffolds with cells). Other limitations of this technique include (1) the transfer of femoral AV vessels, which supply the blood flow to the hindlimb. However, the technique leaves the profunda and the deep femoral vessels untouched and no hindlimb ischemia was observed after flap transfer. Moreover, in larger animals and in humans, the use of peripheral vessels will be advised as they are redundant in the body; (2) the time required to achieve adequate vascularization prior to flap transfer, can be a limiting factor in urgent cases; and (3) flap creation is performed in vivo, which can limit its relevance in humans. In the future, we aim to develop a means of creating this flap ex vivo. The advantage of the presented method lies in the capacity to enhance formation of an autologous tissue (with autologous cells) around the scaffold. The resulting tissue presents a suitable alternative to an autologous tissue flap. Moreover, the scaffold can be seeded with autologous cells prior to implantation, to further improve graft vascularization and integration within the host tissue. Use of autologous cells and a biodegradable and biocompatible scaffold, can circumvent the substantial risk of immuno-rejection reactions and present an alternative to autologous flaps while avoiding harvesting and postoperative scarification.
The described method can translated to experiments in large animals and further expanded to clinical trials in humans. Moreover it can be translated to repair various damaged tissues within the body. In humans and in large animals, the flaps can be generated around peripheral vessels instead of on the femoral AV vessels.
The authors have nothing to disclose.
This research was supported by the FP7 European Research Council Grant 281501, ENGVASC.
small fine straight scissors | Fine Science Tools (FST) | 14090-09 | |
spring scissors | Fine Science Tools (FST) | 15003-08 | |
straight forceps with fine tip | Fine Science Tools (FST) | 11251-20 | |
serrated forceps | Fine Science Tools (FST) | 11050-10 | |
needle holder | Fine Science Tools (FST) | 12500-12 | |
Small vessel cauterizer | Fine Science Tools (FST) | 18000-00 | |
Duratears | Alcon | 5686 | |
Sedaxylan | Euravet | DJ03 | |
Clorketam 1000 | Vetoquinol | 4A0726B | |
Buprenorphine | vetmarket | B15100 | |
4-0 silk sutures | Assut sutures | 647 | |
6-0 polypropylene sutures | Assut sutures | 9351F | |
8-0 silk sutures | Assut sutures | 684568 | |
Insulin syringe (6mm needle) | BD | 324911 | |
Vevo 2100 high-resolution ultrasound system | VisualSonics inc. | ||
MS250 non-linear transducer | VisualSonics inc. | ||
Micromarker non-targeted contrast agent | VisualSonics inc. | VS-11694 | |
tail vein catheter | VisualSonics inc. | VS-11912 | |
Vevo 2100 software | VisualSonics inc. | ||
fluorescein isothiocyanate-conjugated dextran | Sigma | FD500S | |
Matlab | Mathworks, MA, USA | ||
Kimwipes | Kimtech | 34120 | |
antigen unmasking solution | Vector laboratories | H-3300 | |
anti-CD31 antibody | Abcam | ab28364 | |
biotinylated goat anti-rabbit (secondary) antibody | Vector laboratories | BA-1000 | |
streptavidin-peroxidase | Jackson | 016-030-084 | |
Mayer's hamatoxylin solution | Sigma-Aldrich | MHS-16 | |
aminoethylcarbazole (AEC) substrate kit | Life technologies, Invitrogen | 00-2007 | |
Vectamount | Vector laboratories | H-5501 |