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Objective:To prepare the whole bladder acellular matrix (BAM) using the self-designed perfusion decellularization system, and evaluate the feasibility of constructing the tissue engineering bladder with the adipose-derived stem cells (ADSCs).Methods:This study was conducted from October 2020 to April 2021. The self-designed perfusion decellularization system was used, and four different decellularization protocols (group A, group B, group C and group D) were formulated, according to the flow direction of the perfusate and the action time of different decellularization solutions. Among them, the urethral orifice of the bladder tissue was used as the outflow tract of the perfusion fluid in groups A and B. The top of the bladder was cut off and used as the outflow tract of the perfusion fluid in groups C and D. In groups A and C, 1% Triton X-100 was treated for 6 h, and 1% sodium dodecyl sulfate (SDS) was treated for 2 h. In groups B and D, 1% Triton X-100 was treated for 7 h, and 1% sodium dodecyl sulfate (SDS) was treated for 1 h. In addition, the tissue in the normal bladder group was directly obtained from the natural bladder tissue, which did not require perfusion, cryopreservation and thawing. The fast and efficient decellularization protocol was screened out through HE, DAPI, Masson trichrome and Alcian Blue staining and quantitative analyses to prepare the whole bladder scaffold. The prepared BAM was used as the scaffold material, and the ADSCs were used as the seeding cells to construct the tissue engineering bladder. HE and DAPI staining were used to observe the distribution of ADSCs on the BAM.Results:HE and DAPI staining showed that there was no obvious nuclear residue in the group C. Masson trichrome and Alcian Blue staining showed that the collagen structure and glycosaminoglycan were well preserved in the group C. There was no significant difference in bladder wall thickness between the group C and the normal bladder group [(975.44±158.62)μm vs.(1 064.49±168.52)μm, P > 0.05]. The DNA content in the group C [(43.59 ±4.59) ng/mg] was lower than that in the normal bladder group, group A, group B and group D [(532.50±26.69), (135.17±6.99), (182.49±13.69) and(84.00±4.38)ng/mg], and the difference was statistically significant ( P<0.05). The collagen content [(10.98 ± 0.29)μg/mg] and glycosaminoglycan content [(2.30±0.18)μg/mg] in group C were not significantly different with those in the normal bladder group [(11.69±0.49) and (2.36±0.09)μg/mg, P>0.05]. Scanning electron microscopy showed that a large number of pore structures could be observed on the surface of the prepared BAM in groups A-D and were facilitated to cell adhesion. The isolated and cultured ADSCs were identified by flow cytometry to confirm the positive expression of CD90 and CD29, and the negative expression of CD45 and CD106. Live/dead staining and CCK-8 detection confirmed that the prepared BAM in the group C had no cytotoxicity. HE and DAPI staining showed that a large number of ADSCs were distributed on the surface and inside of the tissue engineering bladder. Conclusions:The whole bladder shape BAM prepared by the self-designed perfusion decellularization system could be used as the scaffold material for bladder tissue engineering, and the constructed tissue engineering bladder could be used for bladder repair and reconstruction.
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Objective:To investigate the effect of tissue engineered bladder patch constructed by double-layer silk scaffold and adipose-derived stem cells (ADSCs) in the repair and reconstruction of bladder.Methods:This study was conducted from May 2020 to March 2021. The silk fibroin (SF) aqueous solution was obtained from silkworm cocoons, and a double-layer silk scaffold composed of silk fibroin film and silk fibroin sponge was further prepared. The rat ADSCs were isolated, cultured, and the ADSCs surface markers (CD29, CD90, CD45, CD106) were identified by flow cytometry. The ADSCs were planted on a double-layer silk scaffold to construct a tissue-engineered bladder patch. Thirty-six male SD rats were randomly divided into three groups: tissue engineered bladder patch group (SF-ADSCs group, n=15), double-layer silk scaffold group (SF group, n=15), control group ( n=6). The tissue engineered bladder patch (SF-ADSCs group) and double-layer silk scaffold (SF group) were wrapped on the omentum to promote vascularization. The vascularization was evaluated by HE and immunofluorescence staining. The wrapped tissue engineered bladder patch and double-layer silk scaffold were used to repair the defective bladder. In the control group (six rats), the incision was closed immediately after the bladder tissue fully exposed. At 4 weeks and 12 weeks after operation, the general morphology of bladder tissue and cystography were performed to evaluate the recovery of bladder morphology. After the graft was harvested, HE and Masson's trichrome staining and immunofluorescence staining were used to observe the regeneration of bladder wall tissue. Urodynamics was used to assess the recovery of bladder function at 12 weeks after operation. Results:The flow cytometry results confirmed that the isolated cells positively expressed CD29 and CD90, and there was no significant expression of CD45 and CD106. Gross observation and scanning electron microscope confirmed that the preparation of double-layer silk scaffold not only had a pore structure that was conducive to cell planting, but also had good toughness and was facilitated to surgical suture. The number (43.50±2.66) and area (0.73±0.03)% of vascular-like structures in the SF-ADSCs group after the omentum encapsulation was significantly higher than that in the SF group [(24.50±3.51), (0.55±0.05)%], and the difference was statistically significant ( P<0.05). At 4 weeks after bladder repair, the histological staining of the grafts in the SF-ADSCs and SF groups showed a large number of degraded fragments of double-layer silk scaffold. At 12 weeks, the morphology of the graft in the SF-ADSCs group showed uniform bladder morphology, which was similar to that of normal bladder tissue. Immunofluorescence staining showed that the continuous urothelial layer, abundant smooth muscle tissue, vascular structure and regenerated neurons could be observed in the SF-ADSCs group. Urodynamic test showed that the bladder maximum volume (0.74±0.03)ml and compliance (16.68±0.44)μl/cm H 2O in the SF-ADSCs group, which were better than that in the SF group [(0.47±0.05)ml, (14.89±0.37)μl/cm H 2O], but lower than that in the control group [(1.12±0.08)ml, (19.34±0.45)μl/cm H 2O], and the difference was statistically significant ( P<0.05). Conclusions:The tissue engineered bladder patch constructed with double-layer silk scaffolds and ADSCs could promote the morphological repair of bladder tissue, the regeneration of bladder wall structure and the recovery of bladder physiological function.
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For the damage and loss of tissues and organs caused by urinary system diseases, the current clinical treatment methods have limitations. Tissue engineering provides a therapeutic method that can replace or regenerate damaged tissues and organs through the research of cells, biological scaffolds and biologically related molecules. As an emerging manufacturing technology, three-dimensional (3D) bioprinting technology can accurately control the biological materials carrying cells, which further promotes the development of tissue engineering. This article reviews the research progress and application of 3D bioprinting technology in tissue engineering of kidney, ureter, bladder, and urethra. Finally, the main current challenges and future prospects are discussed.
Subject(s)
Bioprinting , Regeneration , Technology , Tissue Engineering/methodsABSTRACT
Objective:To investigate the effect of ureteral decellularized matrix (UDM) coating on the differentiation of adipose stem cells.Methods:From January 2018 to October 2019, UDM was prepared by perfusion method. H&E staining, DAPI staining, and DNA quantification were used to assessment the residues of cellular component in UDM and normal ureter; Masson′s trichrome staining and collagen quantification evaluated the collagen retention in UDM and normal ureter; the distribution and content of glycosaminoglycan in UDM and normal ureter were analyzed through Alcian Blue staining and glycosaminoglycan quantification. Canine adipose mesenchymal stem cells (cADMSCs) were isolated and cultured and identified by flow cytometry. The UDM was digested by pepsin enzyme to prepare the decellularized matrix coating as the experimental group. Type Ⅰ rat tail collagen coating was used as a control group. The cADMSCs were seeded on different coatings, and the differentiation of the cADMSCs was detected by immunofluorescence staining and Western Blot.Results:H&E and DAPI staining showed that the nuclear residue was not observed in the UDM. The DNA quantification demonstrated that the DNA content in UDM [(38.87±3.40) ng/mg] was significantly lower than that in the normal ureter group [(1 694.63±169.83) ng/mg, P<0.05]. Masson′s trichrome staining and collagen quantification confirmed that the collagen content in UDM [(265.89 ± 16.40) μg/mg] was no significantly different from the normal ureter group [(288.73 ± 16.32) μg/mg, P>0.05]. Alcian Blue staining showed the distribution of glycosaminoglycan in the UDM, and glycosaminoglycan quantification suggested that the content of glycosaminoglycan in the UDM [(1.57 ± 0.19) μg/mg] was lower than that in the normal ureter group [(3.43 ± 0.12) μg/mg] ( P<0.05). Immunofluorescence staining and Western Blot confirmed that the expression of Alpha-smooth muscle Actin (α-SMA) in the experimental group (2.51 ± 0.27, 3.68 ± 0.33, 4.91 ± 0.45) was higher than that in the control group (0.97±0.09, 1.02 ± 0.10, 1.00 ± 0.11) at 3 d, 7 d, 10 d ( P<0.05). The expression of α-SMA in the experimental group increased gradually with culture time ( P<0.05). While no changing of α-SMA expression in the control group was recordered. Conclusions:The prepared UDM removed the cellular components, and retained the collagen structure and bioactive components well; the ureter decellularized matrix coating could promote the differentiation of cADMSCs to smooth muscle cells.