Biomaterials 35 (2014) 6822e6828
Contents lists available at ScienceDirect
Biomaterials journal homepage: www.elsevier.com/locate/biomaterials
Decellularized kidney scaffold-mediated renal regeneration Y.L. Yu a, b, Y.K. Shao c, Y.Q. Ding d, K.Z. Lin b, B. Chen b, e, H.Z. Zhang f, L.N. Zhao a, b, Z.B. Wang b, J.S. Zhang a, b, M.L. Tang a, b, J. Mei a, b, d, * a
Anatomy Department, Wenzhou Medical University, Wenzhou 325035, China Institute of Bioscaffold Transplantation and Immunology, Wenzhou Medical University, Wenzhou 325035, China c School of the First Clinical Medical Sciences, Wenzhou Medical University, Wenzhou 325035, China d Institute of Neuroscience, Wenzhou Medical University, Wenzhou 325035, China e Department of Radiology, The First Affiliated Hospital of Wenzhou Medical University, Wenzhou 32000, China f Department of Nuclear Medicine, The Second Affiliated Hospital of Wenzhou Medical University, Wenzhou 32000, China b
a r t i c l e i n f o
a b s t r a c t
Article history: Received 8 April 2014 Accepted 22 April 2014 Available online 21 May 2014
Renal regeneration approaches offer great potential for the treatment of chronic kidney disease, but their availability remains limited by the clinical challenges they pose. In the present study, we used continuous detergent perfusion to generate decellularized (DC) rat kidney scaffolds. The scaffolds retained intact vascular trees and overall architecture, along with significant concentrations of various cytokines, but lost all cellular components. To evaluate its potential in renal function recovery, DC scaffold tissue was grafted onto partially nephrectomized rat kidneys. An increase of renal size was found, and regenerated renal parenchyma cells were observed in the repair area containing the grafted scaffold. In addition, the number of nestin-positive renal progenitor cells was markedly higher in scaffold-grafted kidneys compared to controls. Moreover, radionuclide scan analysis showed significant recovery of renal functions at 6 weeks post-implantation. Our results provide further evidence to show that DC kidney scaffolds could be used to promote renal recovery in the treatment of chronic kidney disease. Ó 2014 Elsevier Ltd. All rights reserved.
Keywords: Renal regeneration Scaffold Partial nephrectomy Stem/progenitor cells
1. Introduction Chronic kidney disease is a leading cause of mortality and morbidity worldwide, affecting between 8% and 16% of the global adult population [1]. While the incidence of chronic kidney disease is on the rise, available therapeutic options remain limited. Several recent studies have demonstrated that the postnatal mammalian kidney can undergo a certain degree of regenerative repair after partial resection. This regeneration occurs mainly through the proliferation of surviving mature cells or stem cells present in the kidney, such as glomerular parietal epithelial cells (GPECs; a type of renal progenitor cell) [2e8]. Following injury, these cells migrate to the injured area and proliferate, and then redifferentiate into new somatic cells [3,9,10]. These findings suggest that regenerative solutions offer great potential for treating chronic kidney disease. However, achieving clinically significant regeneration has proven to be extremely challenging, due to the complexity of this organ. In
* Corresponding author. Anatomy Department, Wenzhou Medical University, Wenzhou 325035, China. Tel.: þ86 13758481967. E-mail address: [email protected] (J. Mei). http://dx.doi.org/10.1016/j.biomaterials.2014.04.074 0142-9612/Ó 2014 Elsevier Ltd. All rights reserved.
addition, regenerative repair is incomplete and cannot fully restore renal function on its own. Recent advances in tissue engineering, stem cell research, and regenerative medicine have offered new hope for the treatment of kidney diseases [11e16]. In particular, the use of decellularized (DC) extracellular matrix (ECM) scaffolds to help repair damaged or injured tissue has emerged as a promising approach in the field of renal regeneration [17e21]. DC kidney scaffolds are able to act as inductive template for functional organ recovery, allowing the injured area to recellularize with autologous stem cells or differentiated cells [19,22,23]. The particular physiochemical properties of different ECM scaffolds create specific cellular niches within body tissues; thus, the ECM plays a critical biochemical and physical role in initiating and sustaining various cellular functions. In light of this, it is feasible that ECM scaffolds could be used to promote renal regeneration by enabling macroscopic regrowth into a damaged kidney. The goal of the present study was to engineer DC kidney scaffolds capable of inducing renal regeneration after injury. We hypothesized that, by providing a neutral environment that mimics normal physiological conditions, the scaffolds would allow recellularization of the damaged area by cells present in the residual
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
kidney. Our DC kidney scaffolds were created by continuous detergent perfusion of dissected rat whole kidneys. To test the scaffolds in vivo, we grafted some scaffold tissue onto partially nephrectomized rat kidneys and assessed their recovery at various post-surgical time points. 2. Materials and methods
6823
2.5. B-scan ultrasonography B-scan ultrasonography images of both kidneys were obtained on weeks 1, 2 and 4 to assess the size, shape and location of the organ, as well as blood flow to the area in anesthetized rats. A 15 MHz probe (ML6-15; General Electric Company, USA) was placed in the left lower intercostal space at the midaxillary line. The liver was used as an “acoustic window” to aim the probe slightly posteriorly (toward the kidney). The probe was then rocked gently (up and down or side to side) to scan the entire kidney.
2.1. Preparation of DC kidney scaffolds In this study, we developed a new protocol to generate rat DC kidney scaffolds using continuous detergent perfusion. A 24-gauge cannula (Puyi, China) was inserted into the infrarenal abdominal aorta before bilateral kidneys were isolated, and connected with a peristaltic pump (YX1515X-A; Baoding Longer Precision Pump Co., China) to allow continuous rinsing with various detergents. Solutions were perfused at an approximate rate of 8 ml/min in the following order: 50 U/ml heparin in 0.01 M phosphate buffered saline (PBS, pH 7.4) for 30 min, 0.1% tritonX-100 for 3 h, deionized water for 30 min, 0.8% (v/v) sodium lauryl sulfate (SDS) for 3 h, and deionized-water containing 100 U/ml penicillin and 100 mg/ml streptomycin (Therma Scientific, USA) for 24 h.Once the perfusion was complete, DC kidney scaffolds were kept in 50 ml of deionized-water containing the penicillin and streptomycin at 4 C for less than 7 d. 2.2. Vascular corrosion casting and ultrastructural observation of DC kidney scaffolds To determine the integrity of microvasculature in the DC kidney scaffolds, we performed the vascular corrosion casting. Catheterization of the inferior vena cava and abdominal aorta was performed, and 1e2 ml of acetone was injected into the native kidney or DC kidney scaffolds through the inferior vena cava. Then 5 ml of 10% Acrylonitrile Butadiene Styrene (ABS) Sudan solvent mixture was poured through abdominal aorta, and meanwhile 10 ml of 10% ABS blue pigments mixture was perfused via the inferior vena. The samples were cooled in running water and corroded in 50% hydrochloric acid for 1-3 d. The morphology and distribution of vasculature was observed under stereomicroscope and images were recorded with Olympus soft image viewer (Japan). Transmission electron microscope was used to examine the extracellular matrix in the DC kidney scaffolds. DC kidney scaffolds were fixed with 2.5% (v/v) glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) overnight at 4 C and post-fixed with 1% osmium tetroxide for 1 h at 37 C. The samples were then dehydrated with a series of acetone solutions with increasing concentrations, infiltrated with epon resin and baked overnight at 65 C. Ultrathin sections (80 nm) were prepared, stained with 2% uranyl acetate and lead citrate, and observed under a Hitachi electron microscope (H7500; Japan) at 70 kV. We recorded images with a gatan 830 high resolution CCD digital camera. 2.3. Assessing SDS residue levels in DC kidney scaffolds SDS residue levels were assessed by ultraviolet-visible spectrophotometer (UVVIS). The water inside the DC kidney scaffolds (n ¼ 5) was sponged up, and the scaffolds were digested with proteinase K (100 mg/mg; Biomiga, USA) in the protein lysis buffer at 50 C overnight, and centrifuged (14,000 g) for 5minto remove the precipitate. Light absorption of the supernatant at 499 nm was measured, and content of SDS in the supernatant was calculated according to standard concentrationeabsorption curve, which was generated by measuring the absorbance at 499 nm of eight solutions with known SDS concentrations, as reported previously. 2.4. Rat model of renal regeneration with DC kidney scaffolds All procedures involving animal use, housing, and surgery were approved by the administration of Wenzhou Medical University. Male 2-month old Sprague Dawley rats (average body weight: 250 g) were used, and were divided into two treatment groups. Control rats were partially nephrectomized without any scaffold repair, whereas treatment rats underwent partial nephrectomy repaired with DC kidney scaffolds (n ¼ 40 in each group). Animals were anesthetized with an intraperitoneal injection of chloral hydrate (6.1/kg body weight), and an abdominal paramedian incision was made from the pubis to the xyphoid to expose the left kidney. In the beginning of the operation, 1 ml of heparin (50 U/ ml) was injected through the inferior vena cava. After separating the left renal capsule, renal artery and vein were clipped with micro-ligation haemostatic clip (W40130; Chen-He Microsurgical Instruments Factory, China), at which point the timer was started to make sure renal ischemia was kept under 10 min. The left kidney was transected slightly below the renal pelvis (removing about 1/3 of the renal parenchyma). For animals in the treatment group, the wound was grafted with lower 1/3 of DC scaffold by suturing the external capsules between the excised kidney and DC kidney scaffold. Control kidneys were sutured directly after excision. After reperfusion, the left kidney was monitored if there was a leakage of blood for 20 min before closing the abdominal wall. After surgery, all animals were given unlimited access to rat chow and water containing penicillin and streptomycin.
2.6. Singer photon emission computed tomography (SPECT) imaging Radionuclide scans were performed on weeks 4, 6 and 8 on a dual-head EPIC Vertex SPECT equipped with an LEGP collimator (V-60; ADAC, USA). Animals were deprived of food overnight and anesthetized with chloral hydrate (6.1 g/kg body weight) 10 min prior to imaging. The rats were injected intravenously with100 mCi of 99m Tc-DTPA (Jiangsu Institute of Nuclear Medicine, China), and then subjected to a 30 min dynamic SPECT immediately after the injection. For PET quantification, the whole right and left kidneys were included and all images were analyzed by the Gates method [24]. 2.7. Histology and immunofluorescence analysis Samples were prepared for histological and immunofluorescence analyses by following standard protocols for paraffin embedding. Histological analysis, mounted kidney sections were stained by hematoxylin and eosin (H&E) to visualize nuclei (blue), and both cytoplasm and connective tissue (pink). For immunofluorescence, following deparaffinization, rehydration and antigen retrieval, the sections were blocked with 5% normal bovine serum in PBS for 30 min. Sections were then incubated with primary antibodies overnight at 4 C. The primary antibodies used were as follows: mouse anti-nestin (1:200; Abcam,UK) and rabbitanti-AQP1 (1:200; Abcam). After being washed in PBS, sections were then incubated in 488- or 594-conjugated species-specific secondary antibody (Chemicon, USA) for 2 h. Slides were observed with an Olympus fluorescent microscope and images were captured using Olympus soft image viewer. The intensity of immunoreactivity was quantified using the Image-Pro Plus 6.0 software (Media Cybernetics, USA). 2.8. Enzyme-linked immunosorbentassay (ELISA) for quantitative analysis of cytokines in DC kidney scaffolds Total protein in DC kidney scaffolds or intact kidney was extracted using an ELISA kit (R&D Systems, USA). The concentrations of various cytokines including VEGF, TGF, HGF, IL-8, CTGF, FGF and PDGF were assayed with a microplate reader at 450 nm [25]. 2.9. Statistical analysis Quantitative results were reported as mean standard deviation. Independent samples t-test was used to reveal differences in the level of cytokines and intensity of immunofluorescence among different groups. The significance level was set at 0.05 and SPSS 17.0 (SPSS Inc., Chicago, USA) was used for the analyses.
3. Results 3.1. Characterization of DC kidney scaffolds After decellularization with continuous detergent perfusion, DC kidney scaffolds had a somewhat transparent appearance (Fig. 1A). Vascular corrosion casting showed that the vascular tree in the DC kidney scaffolds was well maintained compared with intact kidney (Fig. 1B,C). H&E staining revealed that blue-stained nuclei were not observed, but pink-stained components were present in the DC kidney scaffolds compared with control (Fig. 1D,E). As pink-stained components include both cytoplasm and extracellular matrix, and this may reflect the morphological difference in H&E staining between intact kidney and DC kidney scaffolds (Fig. 1D,E). In addition, PAS and Masson’s staining were used to examine the ECM (e.g. polysaccharides and collagen), and it showed that positivelystained structures were present in DC kidney scaffolds without interruption (Fig. 1F,G). Finally, electron microscopy observation indicated that nuclear structure was not found, but the integrity of extracellular matrix was not disrupted in the DC kidney scaffolds as shown by the presence of continuous membrane of Bowman’s capsule, the basement membrane of the glomerular capillaries and mesangial matrix (Fig. 1H). Taken together, the DC kidney scaffolds
6824
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
Fig. 1. Characterization of the DC kidney scaffolds. (A) Gross appearance of harvested DC kidney scaffolds. (B) Electron microscopy observation shows intact extracellular matrix in DC kidney scaffold. Blue arrows indicate the membrane of Bowman’s capsule, a green arrow indicates the basement membrane of the glomerular capillaries, and a red star points to the mesangial matrix. (C, D) Vascular corrosion casting shows a normal vascular tree of DC kidney scaffold (D) compared with intact kidney (C). (E, F) H&E staining shows the existence of blue-stained nuclei in intact kidney (E) but not DC kidney scaffold (F). (G) PAS staining shows the presence of the ECM (e.g. basement membranes) in DC kidney scaffold. Noted that the capillary loops of the glomeruli are clearly displayed. (H) Masson’s staining shows that green-stained collagenous fibers in DC kidney scaffold. (I) Quantitative assay of cytokines in DC kidney scaffolds. The level of PDGF CTGF, TGF-b and VEGF in the DC kidney is not different from that of native kidney, while that of IL-8, FGF and HGF is reduced. Scar bars ¼ 5 mm (A, C D), 100 mm (EeH) and 5 mm (B).
prepared in our study lose renal cells but keep normal vascular tree and continuous extracellular matrix. Considering that SDS is toxic to cells, UV-VIS was used to assess SDS residue levels in the scaffolds. A standard concentration curve for SDS was generated by measuring the absorbance of eight different known concentrations of SDS. According to this standard curve, the mean residual SDS concentration in the DC kidney scaffolds was 50.0 1.7 mg/g, which is below the safe level of 133.3 mg/g [26]. To assess the potential of DC kidney scaffolds for renal regeneration, we performed an ELISA assay to quantify levels of various cytokines. As shown in Fig. 1I, the levels of cytokines HGF, CTGF, TGF-b and VEGF in DC kidney scaffolds were similar to that of the intact kidney, although some of them (IL-8, FGF and PDGF) were lower than the intact kidney. Thus, although the rat kidneys had been completely decellularized, some cytokines still remained within the scaffolds in concentrations that might be sufficient to contribute to renal regeneration after injury. 3.2. Renal degeneration in DC kidney scaffolds The regenerative potential of the DC kidney scaffolds was tested by grafting them onto partially nephrectomized rat kidneys (Fig. 2A). The lower 1/3 of kidney was removed, and similar sizedDC kidney scaffolds (left lower panel) were grafted onto the cutend. Rats with cut-end-sutured kidney were used as control. As shown by non-invasive B-scan ultrasonography performed at weeks 1, 2 and 4 post-surgery, kidneys treated with DC scaffolds exhibited some macroscopic regrowth, whereas control kidneys did not (Fig. 2B,C). In addition, obvious bloodstream signals were observed in the RDS area starting2 weeks after the RDS transplantation (Fig. 2B,C).
To examine the regeneration in more detail, some rats were sacrificed on 1, 2, 4 and 8 weeks post-surgery and their kidneys were removed for analysis. As shown in Fig. 3AeH, a macroscopic regrowth of the renal tissue into the grafting DC kidney scaffolds occurred in the experimental group with the scaffold graft developing first into granulation tissue, then into a scar by week 8. This regrowth was also reflected by a downward moving of a boundary between the regenerated tissue and the scaffolds overtime (lines in Fig. 3AeH). The size of regenerated kidney was measured 4 weeks post-transplantation, and it showed a significant increase compared with controls (Fig. 2C; control, 1.35 0.05 cm 0.75 0.05 cm; scaffolds-grafted, 1.50 0.1 cm 0.76 0.08 cm; n ¼ 10 for each, P < 0.05). Microscopic examination showed that tissue obtained from the regions above the boundary between the regenerated tissue and the scaffolds (yellow marked area in Fig. 3AeD) contained the renal parenchyma cells on 1 and 2 weeks, and they became to tubular and glomerular morphology on 4 and 8 weeks post-grafting (Fig. 3IeL). In this area, numerous dark hematoxylin-stained cells were also observed, and these small cells gradually reduced in number overtime; they are probably inflammatory cells (Fig. 3IeL). By contrast, the tissue from the region below the boundary contained eosin-stained ECM of the scaffolds, and hematoxylin-stained cells that were also reduced overtime (Fig. 3MeP). Taken together, above data demonstrate an apparent regeneration of the renal tissue in the DC kidney scaffolds of partially-excised kidney. Having found the morphological regeneration in the DC scaffolds-grafted kidney, we were promoted to see if there is any functional recovery. Radionuclide renal scans were performed on 6 and 8 weeks post-grafting. Kidneys grafted with the DC scaffold tissue did not show improved glomerular filtration rate until 6 weeks, compared with controls (Fig. 4B). Six weeks after surgery,
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
6825
Fig. 2. The procedures of grafting and the regeneration of renal tissue in DC kidney scaffolds shown by B-scan ultrasonography and gross appearance of explanted kidneys. (A) Showing the procedures of grafting DC kidney scaffolds. Approximately 1/3 of the left kidney was transversely excised from each rat, and similar sized-scaffolds (arrow in the left lower panel) were grafted onto the cut-end by suturing the external capsules. Cut-end in control group was sutured directly. (B) B-scan ultrasonography images of the DC kidney scaffold-grafted obtained on weeks 1, 2 and 4 to assess the size, shape, and blood flow to the kidneys. Yellow circles represent the left kidneys, the white triangles represent implanted scaffolds area, the red signals represent artery blood flow, and blue ones represent vein flow. (C) Comparison of the long axis of kidney measured by B-scan ultrasonography. N ¼ 5for each, P < 0.05. (D) Gross appearance of explanted kidney scaffolds on weeks 8. D1, D2, D3 show right kidney and DC kidney scaffold-grafted group and control group respectively.
both experimental and control kidneys displayed a very mild delay in renal perfusion, as well as some degree of delay in excretion (Fig. 4C,D). In an attempt to characterize the regenerative cells found in the scaffold-grafted kidneys, we began by looking at the distribution of cells immune positive for nestin, a marker of renal stem/progenitor cells. In intact kidney, nestin immunoreactivity was only distributed in the glomeruli (Fig. 5A). In the scaffoldgrafted kidneys, we examined nestin expression in the tissue located above the boundary between the newly degenerated tissue and DC kidney scaffold at different post-surgery time points. On 1 week, in addition to their typical glomerular distribution, many nestin-positive cells were also present in the
renal tubules and in the interstitium near the incisal border (Fig. 5B, C1, D1), but they were noticeably reduced by 2 weeks (Fig. 5C2, D2) and had become undetectable by week 8 (Fig. 5C3, C4, D3, D4). At this time point, nestin-positive cells were only present in the glomeruli (Fig. 5C4, D4). In control kidney, nestin immunoreactivity was observed in the glomeruli and some scattered cell near the boundary on week 1, but they were not present at later time points (Fig. 5E). In addition, expression of aquaporin-1 (AQP-1), a channel protein typically located in mature proximal tubules [27], was examined. AQP-1 was not observed in the first two weeks post-surgery, but many AQP-1positive cells were observed in scaffold-grafted kidneys on week 3 and 4 (Fig. 5D).
Fig. 3. Experimental kidney recovery over 8 weeks. (AeH) Macroscopic images show longitudinal cross-sections of whole experimental kidneys (gross observation: AeD; observation under stereoscopic microscope: EeH) on weeks 1, 2, 4 and 8 post-surgery. The black line represents the border between the renal parenchyma and the grafted-DC scaffold. Yellow area and blue area were removed for light microscopic examination shown in IeL and MP, respectively. (IeP) H&E staining shows renal parenchyma (IeL; yellow squares in AeD) and DC scaffold (MeP; blue ellipses in AeD) at the four time points. On week 1, the renal parenchyma was seriously damaged, exhibiting extensive glomerular and tubular atrophy and widespread infiltration of inflammatory cells (I). By week 2, tissue atrophy was reduced in the injured renal parenchyma (J), and mild edema was observed on week 4 (K). By week 8, the renal parenchyma had recovered to a near-normal state (L). In the DC scaffold, inflammatory cells were seen infiltrating along the margin on week 1 (M). By week 2, the entire scaffold graft had been infiltrated (N). Granulation tissue had become to form by week 4 (O), and had developed into scar tissue by week 8 (P). For H&E staining in intact kidney, please see Fig. 1D. Scale bars ¼ 100 mm (IeP).
Fig. 4. Radionuclide scan analysis demonstrating a remarkable improvement in GFR compared with the control. (A) Normal renal function in healthy intact rats was used as a baseline comparator. Left panel is renal dynamic imaging. Right lower panel is the renogram showing the renal filtration and drainage, and the appearance of isotope in the bladder, which are also expressed by CTS per second. (B) Statistical analysis revealed significant differences in renal functions between the experimental and control groups at weeks 6 and 8 (n ¼ 10 for each; *P < 0.05). Vertical axis is glomerular filtration rate of the injured left kidney (GFRLI), glomerular filtration rate of the healthy left kidney (GFRLN), the indicator of GFR. (C, D) Radionuclide scan analysis of experimental (C) and control (D) kidneys on 6 weeks post-surgery. White arrows indicate the left kidney.
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
6827
Fig. 5. Immunofluorescent analysis showing that more nestin-positive cells (green) are present in experimental kidneys compared with controls. Tissues used for immunostaining were taken from the region located above the border between newly regenerated tissue and the scaffold at different time points. (A) Immunoreactivity of nestin is only present in the glomeruli of intact kidney. (B) Quantitative analysis showing significantly higher nestin immunoreactivity in experimental kidneys compared to controls, with a pronounced peak at week 1 post-surgery. (C) Nestin-positive cells are more abundant and widely distributed in experimental kidneys. In addition to the glomeruli, nestin protein is also detected in cortical tubules near the incision border, especially on week 1 (C1, C5). By week 8, nestin expression is restricted to the gomeruli (C4, C8). (D) Double immunostaining of nestin (green) and aquaporin (red). Aquaporin-positive cells are hardly observed in scaffold-grafted kidneys on week 1 or 2 (D1, D2), but increased greatly on week 4 and 8 (D3, D4). (E) In control kidneys, weak nestin expression is only seen on week 1.Scale bars ¼ 10 mm (A, D1eD4, E1eE4) and 25 mm (C1eC4).
4. Discussion Tissue scaffolds containing intact vascular and ECM architecture hold great therapeutic promise for the field of organ recovery, particularly in the case of highly vascular organs [18,22,28,29]. Recently, Harald C. Ott and his team were able to recellularize rat kidney scaffold grafts [22], and Giuseppe Orlando and colleagues successfully implanted porcine DC kidney scaffolds into experimental pigs [30]. In these two studies, some degree of both functional and physical renal recovery was attained, but further research is required in order to turn these observations into clinically relevant achievements. The findings presented herein provide additional support to the hypothesis that orthotopic DC scaffold grafts can induce macroscopic growth and regeneration of damaged kidneys, thus promoting in vivo renal function recovery. Our rat kidney scaffolds, which were decellularized by continuous detergent perfusion, retained vascular trees that were intact at all hierarchical levels. Residual detergent levels in the scaffolds were found to be negligible, well below the level deemed safe by the Food and Drug Administration. Overall renal ECM was also maintained, and the levels of various cytokines were maintained.
The ECM represents a substrate to which cells can adhere, but it also provides support and inductive cues for cellular proliferation, differentiation and migration. To date, attempts at recreating a synthetic environment mimicking that of the ECM have proven mostly unsuccessful [31]. In particular, the specific topography or geometry of the ECM has been shown to play a role in promoting osteogenic differentiation [32]. In addition, an artificial ECM containing the angiogenic laminin-1-derived IKVAV sequence was shown to cause a significant increase in the number and area of vascular branches in the chick chorioallantoic membrane [33]. In our model, rats were partially nephrectomized (by removing approximately 30% of the kidney parenchyma from the lower pole), and DC kidney scaffold tissue was grafted onto the resected kidney before suturing shut. Microscopic imaging revealed that progenitor cells were migrating from the injured kidney into the grafted scaffold tissue, suggesting that the scaffold’s intact ECM retains sufficient chemical and/or physical cues to induce the proliferation and differentiation of renal progenitor cells present nearby. Immunohistochemical analysis of post-surgery kidney sections revealed a significant number of nestin-positive cells within the repair zone of scaffold-grafted kidneys. By the interaction of injured kidney with the RDS, many nestin-positive renal progenitor cells were observed, especially in the second week. These observations
6828
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
suggest that several distinct mechanisms may be involved in DC scaffold-mediated regenerative kidney repair. First, the presence of the scaffold graft could potentially contribute to hemostasis by physically protecting/repairing the wound surface. In addition, the intact scaffold ECM might harbor binding sites for proteins that stimulate the proliferation of renal progenitor cells, thus contributing to kidney repair and regeneration. Finally, the grafted tissue may provide physical support as well as biochemical cues that promote the migration of renal cells from the injured kidney into the acellular scaffold. In the present study, we developed an experimental rat model of DC scaffold-induced renal recovery, and focused mainly on the more macroscopic aspects of kidney repair. In the future, we aim to also explore the molecular mechanisms involved in this process, and to expand our model to other test species, such as mice and pigs. 5. Conclusion In summary, we developed a decellularized (DC) rat kidney scaffold with overall architecture and significant concentrations of various cytokines that improves recovery of the kidney after partial nephrectomy, independently from renal endogenous cells and other factors. The main mechanisms responsible for this effect include the mechanical support of the scaffold, facilitation of cell migration and regeneration. Further investigations on therapeutic factors and/or cells that can be seeded within the decellularized scaffold can usher in a new clinical therapy for renal recovery in the treatment of chronic kidney disease. Acknowledgments The authors appreciate the input of Miaozhong Li and Zhiheng Rao in these studies. The authors are grateful to Jiawei Li for help in imaging. These studies were carried out with the support of the National Natural Science Foundation of China (81071576) and the Natural Science Foundation of Zhejiang Province (LY12H15002, LY14H050005). Appendix A. Supplementary data Supplementary video related to this article can be found at http://dx.doi.org/10.1016/j.biomaterials.2014.04.074. References [1] Jha V, Garcia-Garcia G, Iseki K, Li Z, Naicker S, Plattner B, et al. Chronic kidney disease: global dimension and perspectives. Lancet 2013;382:260e72. [2] Franquesa M, Flaquer M, Cruzado JM, Grinyo JM. Kidney regeneration and repair after transplantation. Curr Opin Organ Transplant 2013;18:191e6. [3] Wen D, Ni L, You L, Zhang L, Gu Y, Hao C-M, et al. Upregulation of nestin in proximal tubules may participate in cell migration during renal repair. Am J Physiol Renal Physiol 2012;303:1534e44. [4] Swetha G, Chandra V, Phadnis S, Bhonde R. Glomerular parietal epithelial cells of adult murine kidney undergo EMT to generate cells with traits of renal progenitors. J Cell Mol Med 2011;15:396e413. [5] Reule S, Gupta S. Kidney regeneration and resident stem cells. Organogenesis 2011;7:135e9. [6] Vogetseder A, Picard N, Gaspert A, Walch M, Kaissling B, Hir1 ML. Proliferation capacity of the renal proximal tubule involves the bulk of differentiated epithelial cells. Am J Physiol Cell Physiol 2008;294:22e8.
[7] Humphreys BD, Valerius MT, Kobayashi A, Mugford JW, Soeung S, Duffield JS, et al. Intrinsic epithelial cells repair the kidney after injury. Cell Stem Cell 2008;2:284e91. [8] Lin F, Moran A, Igarashi P. Intrarenal cells, not bone marrow-derived cells, are the major source for regeneration in postischemic kidney. J Clin Invest 2005;115:1756e64. [9] Yamanaka S, Blau HM. Nuclear reprogramming to a pluripotent state by three approaches. Nature 2010;465:704e12. [10] Hanna JH, Saha K, Jaenisch R. Pluripotency and cellular reprogramming: facts, hypotheses, unresolved issues. Cell 2010;143:508e25. [11] Li B, Cohen A, Hudson TE, Motlagh D, Amrani DL, Duffield JS. Mobilized human hematopoietic stem/progenitor cells promote kidney repair after ischemia/ reperfusion injury. Circulation 2010;121:2211e20. [12] Ross EA, Williams MJ, Hamazaki T, Terada N, Clapp WL, Adin C, et al. Embryonic stem cells proliferate and differentiate when seeded into kidney scaffolds. J Am Soc Nephrol 2009;20:2338e47. [13] Krause D, Cantley LG. Bone marrow plasticity revisited: protection or differentiation in the kidney tubule? J Clin Invest 2005;115:1705e8. [14] Morigi M. Mesenchymal stem cells are Renotropic, helping to repair the kidney and improve function in Acute renal Failure. J Am Soc Nephrol 2004;15:1794e804. [15] Bonventre JV. Dedifferentiation and proliferation of surviving epithelial cells in Acute renal Failure. J Am Soc Nephrol 2003;14:55Se61S. [16] Schuldiner M, Yanuka O, Itskovitz-Eldor J, Melton DA, Benvenist N. Effects of eight growth factors on the differentiation of cells derived from human embryonic stem cells. Proc Natl Acad Sci U S A 2000;97:11307e12. [17] Badylak SF, Taylor D, Uygun K. Whole-organ tissue engineering: decellularization and recellularization of three-dimensional matrix scaffolds. Annu Rev Biomed Eng 2011;13:27e53. [18] Orlando G, Booth C, Wang Z, Totonelli G, Ros CL, Moran E, et al. Discarded human kidneys as a source of ECM scaffold for kidney regeneration technologies. Biomaterials 2013;34:5915e25. [19] Ross EA, Abrahamson DR, St John P, Clapp WL, Williams MJ, Terada N, et al. Mouse stem cells seeded into decellularized rat kidney scaffolds endothelialize and remodel basement membranes. Organogenesis 2012;8:49e55. [20] Park KM, Woo HM. Porcine bioengineered scaffolds as new frontiers in regenerative medicine. Transplant Proc 2012;44:1146e50. [21] Song JJ, Ott HC. Organ engineering based on decellularized matrix scaffolds. Trends Mol Med 2011;17:424e32. [22] Song JJ, Guyette JP, Gilpin SE, Gonzalez G, Vacanti JP, Ott HC. Regeneration and experimental orthotopic transplantation of a bioengineered kidney. Nat Med 2013;19:646e51. [23] Nakayama KH, Batchelder CA, Lee CI, Tarantal AF. Decellularized rhesus monkey kidney as a three-dimensional scaffold for renal tissue engineering. Tissue Eng Part A 2010;16:2207e16. [24] Melis M, de Swart J, de Visser M, Berndsen SC, Koelewijn S, Valkema R, et al. Dynamic and static small-animal SPECT in rats for monitoring renal function after 177Lu-labeled Tyr3-octreotate radionuclide therapy. J Nucl Med 2010;51:1962e8. [25] Banks J, Muriel A, Smith JP. Attrition and health in ageing studies: evidence from ELSA and HRS. Longit Life Course Stud 2011;2. [26] Nam CW, Kang SJ, Kang YK, Kwak MK. Cell growth inhibition and apoptosis by SDS-solubilized single-walled carbon nanotubes in normal rat kidney epithelial cells. Arch Pharm Res 2011;34:661e9. [27] Hara-Chikuma M, Verkman AS. Aquaporin-1 facilitates epithelial cell migration in kidney proximal tubule. J Am Soc Nephrol 2006;17:39e45. [28] Uygun BE, Soto-Gutierrez A, Yagi H, Izamis ML, Guzzardi MA, Shulman C, et al. Organ reengineering through development of a transplantable recellularized liver graft using decellularized liver matrix. Nat Med 2010;16:814e20. [29] Serpooshan V, Zhao M, Metzler SA, Wei K, Shah PB, Wang A, et al. The effect of bioengineered acellular collagen patch on cardiac remodeling and ventricular function post myocardial infarction. Biomaterials 2013;34:9048e55. [30] Orlando G, Farney AC, Iskandar SS, Mirmalek-Sani SH, Sullivan DC, Moran E, et al. Production and implantation of renal extracellular matrix scaffolds from porcine kidneys as a platform for renal bioengineering investigations. Ann Surg 2012;256:363e70. [31] Chen XD. Extracellular matrix provides an optimal niche for the maintenance and propagation of mesenchymal stem cells. Birth Defects Res C Embryo Today 2010;90:45e54. [32] Seo CH, Furukawa K, Suzuki Y, Kasagi N, Ichiki T, Ushida T. A topographically optimized substrate with well-ordered lattice micropatterns for enhancing the osteogenic differentiation of murine mesenchymal stem cells. Macromol Biosci 2011;11:938e45. [33] Nakamura M, Yamaguchi K, Mie M, Nakamura M, Akita K, Kobatake E. Promotion of angiogenesis by an artificial extracellular matrix protein containing the laminin-1-derived IKVAV sequence. Bioconjug Chem 2009;20:1759e64.
Contents lists available at ScienceDirect
Biomaterials journal homepage: www.elsevier.com/locate/biomaterials
Decellularized kidney scaffold-mediated renal regeneration Y.L. Yu a, b, Y.K. Shao c, Y.Q. Ding d, K.Z. Lin b, B. Chen b, e, H.Z. Zhang f, L.N. Zhao a, b, Z.B. Wang b, J.S. Zhang a, b, M.L. Tang a, b, J. Mei a, b, d, * a
Anatomy Department, Wenzhou Medical University, Wenzhou 325035, China Institute of Bioscaffold Transplantation and Immunology, Wenzhou Medical University, Wenzhou 325035, China c School of the First Clinical Medical Sciences, Wenzhou Medical University, Wenzhou 325035, China d Institute of Neuroscience, Wenzhou Medical University, Wenzhou 325035, China e Department of Radiology, The First Affiliated Hospital of Wenzhou Medical University, Wenzhou 32000, China f Department of Nuclear Medicine, The Second Affiliated Hospital of Wenzhou Medical University, Wenzhou 32000, China b
a r t i c l e i n f o
a b s t r a c t
Article history: Received 8 April 2014 Accepted 22 April 2014 Available online 21 May 2014
Renal regeneration approaches offer great potential for the treatment of chronic kidney disease, but their availability remains limited by the clinical challenges they pose. In the present study, we used continuous detergent perfusion to generate decellularized (DC) rat kidney scaffolds. The scaffolds retained intact vascular trees and overall architecture, along with significant concentrations of various cytokines, but lost all cellular components. To evaluate its potential in renal function recovery, DC scaffold tissue was grafted onto partially nephrectomized rat kidneys. An increase of renal size was found, and regenerated renal parenchyma cells were observed in the repair area containing the grafted scaffold. In addition, the number of nestin-positive renal progenitor cells was markedly higher in scaffold-grafted kidneys compared to controls. Moreover, radionuclide scan analysis showed significant recovery of renal functions at 6 weeks post-implantation. Our results provide further evidence to show that DC kidney scaffolds could be used to promote renal recovery in the treatment of chronic kidney disease. Ó 2014 Elsevier Ltd. All rights reserved.
Keywords: Renal regeneration Scaffold Partial nephrectomy Stem/progenitor cells
1. Introduction Chronic kidney disease is a leading cause of mortality and morbidity worldwide, affecting between 8% and 16% of the global adult population [1]. While the incidence of chronic kidney disease is on the rise, available therapeutic options remain limited. Several recent studies have demonstrated that the postnatal mammalian kidney can undergo a certain degree of regenerative repair after partial resection. This regeneration occurs mainly through the proliferation of surviving mature cells or stem cells present in the kidney, such as glomerular parietal epithelial cells (GPECs; a type of renal progenitor cell) [2e8]. Following injury, these cells migrate to the injured area and proliferate, and then redifferentiate into new somatic cells [3,9,10]. These findings suggest that regenerative solutions offer great potential for treating chronic kidney disease. However, achieving clinically significant regeneration has proven to be extremely challenging, due to the complexity of this organ. In
* Corresponding author. Anatomy Department, Wenzhou Medical University, Wenzhou 325035, China. Tel.: þ86 13758481967. E-mail address: [email protected] (J. Mei). http://dx.doi.org/10.1016/j.biomaterials.2014.04.074 0142-9612/Ó 2014 Elsevier Ltd. All rights reserved.
addition, regenerative repair is incomplete and cannot fully restore renal function on its own. Recent advances in tissue engineering, stem cell research, and regenerative medicine have offered new hope for the treatment of kidney diseases [11e16]. In particular, the use of decellularized (DC) extracellular matrix (ECM) scaffolds to help repair damaged or injured tissue has emerged as a promising approach in the field of renal regeneration [17e21]. DC kidney scaffolds are able to act as inductive template for functional organ recovery, allowing the injured area to recellularize with autologous stem cells or differentiated cells [19,22,23]. The particular physiochemical properties of different ECM scaffolds create specific cellular niches within body tissues; thus, the ECM plays a critical biochemical and physical role in initiating and sustaining various cellular functions. In light of this, it is feasible that ECM scaffolds could be used to promote renal regeneration by enabling macroscopic regrowth into a damaged kidney. The goal of the present study was to engineer DC kidney scaffolds capable of inducing renal regeneration after injury. We hypothesized that, by providing a neutral environment that mimics normal physiological conditions, the scaffolds would allow recellularization of the damaged area by cells present in the residual
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
kidney. Our DC kidney scaffolds were created by continuous detergent perfusion of dissected rat whole kidneys. To test the scaffolds in vivo, we grafted some scaffold tissue onto partially nephrectomized rat kidneys and assessed their recovery at various post-surgical time points. 2. Materials and methods
6823
2.5. B-scan ultrasonography B-scan ultrasonography images of both kidneys were obtained on weeks 1, 2 and 4 to assess the size, shape and location of the organ, as well as blood flow to the area in anesthetized rats. A 15 MHz probe (ML6-15; General Electric Company, USA) was placed in the left lower intercostal space at the midaxillary line. The liver was used as an “acoustic window” to aim the probe slightly posteriorly (toward the kidney). The probe was then rocked gently (up and down or side to side) to scan the entire kidney.
2.1. Preparation of DC kidney scaffolds In this study, we developed a new protocol to generate rat DC kidney scaffolds using continuous detergent perfusion. A 24-gauge cannula (Puyi, China) was inserted into the infrarenal abdominal aorta before bilateral kidneys were isolated, and connected with a peristaltic pump (YX1515X-A; Baoding Longer Precision Pump Co., China) to allow continuous rinsing with various detergents. Solutions were perfused at an approximate rate of 8 ml/min in the following order: 50 U/ml heparin in 0.01 M phosphate buffered saline (PBS, pH 7.4) for 30 min, 0.1% tritonX-100 for 3 h, deionized water for 30 min, 0.8% (v/v) sodium lauryl sulfate (SDS) for 3 h, and deionized-water containing 100 U/ml penicillin and 100 mg/ml streptomycin (Therma Scientific, USA) for 24 h.Once the perfusion was complete, DC kidney scaffolds were kept in 50 ml of deionized-water containing the penicillin and streptomycin at 4 C for less than 7 d. 2.2. Vascular corrosion casting and ultrastructural observation of DC kidney scaffolds To determine the integrity of microvasculature in the DC kidney scaffolds, we performed the vascular corrosion casting. Catheterization of the inferior vena cava and abdominal aorta was performed, and 1e2 ml of acetone was injected into the native kidney or DC kidney scaffolds through the inferior vena cava. Then 5 ml of 10% Acrylonitrile Butadiene Styrene (ABS) Sudan solvent mixture was poured through abdominal aorta, and meanwhile 10 ml of 10% ABS blue pigments mixture was perfused via the inferior vena. The samples were cooled in running water and corroded in 50% hydrochloric acid for 1-3 d. The morphology and distribution of vasculature was observed under stereomicroscope and images were recorded with Olympus soft image viewer (Japan). Transmission electron microscope was used to examine the extracellular matrix in the DC kidney scaffolds. DC kidney scaffolds were fixed with 2.5% (v/v) glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) overnight at 4 C and post-fixed with 1% osmium tetroxide for 1 h at 37 C. The samples were then dehydrated with a series of acetone solutions with increasing concentrations, infiltrated with epon resin and baked overnight at 65 C. Ultrathin sections (80 nm) were prepared, stained with 2% uranyl acetate and lead citrate, and observed under a Hitachi electron microscope (H7500; Japan) at 70 kV. We recorded images with a gatan 830 high resolution CCD digital camera. 2.3. Assessing SDS residue levels in DC kidney scaffolds SDS residue levels were assessed by ultraviolet-visible spectrophotometer (UVVIS). The water inside the DC kidney scaffolds (n ¼ 5) was sponged up, and the scaffolds were digested with proteinase K (100 mg/mg; Biomiga, USA) in the protein lysis buffer at 50 C overnight, and centrifuged (14,000 g) for 5minto remove the precipitate. Light absorption of the supernatant at 499 nm was measured, and content of SDS in the supernatant was calculated according to standard concentrationeabsorption curve, which was generated by measuring the absorbance at 499 nm of eight solutions with known SDS concentrations, as reported previously. 2.4. Rat model of renal regeneration with DC kidney scaffolds All procedures involving animal use, housing, and surgery were approved by the administration of Wenzhou Medical University. Male 2-month old Sprague Dawley rats (average body weight: 250 g) were used, and were divided into two treatment groups. Control rats were partially nephrectomized without any scaffold repair, whereas treatment rats underwent partial nephrectomy repaired with DC kidney scaffolds (n ¼ 40 in each group). Animals were anesthetized with an intraperitoneal injection of chloral hydrate (6.1/kg body weight), and an abdominal paramedian incision was made from the pubis to the xyphoid to expose the left kidney. In the beginning of the operation, 1 ml of heparin (50 U/ ml) was injected through the inferior vena cava. After separating the left renal capsule, renal artery and vein were clipped with micro-ligation haemostatic clip (W40130; Chen-He Microsurgical Instruments Factory, China), at which point the timer was started to make sure renal ischemia was kept under 10 min. The left kidney was transected slightly below the renal pelvis (removing about 1/3 of the renal parenchyma). For animals in the treatment group, the wound was grafted with lower 1/3 of DC scaffold by suturing the external capsules between the excised kidney and DC kidney scaffold. Control kidneys were sutured directly after excision. After reperfusion, the left kidney was monitored if there was a leakage of blood for 20 min before closing the abdominal wall. After surgery, all animals were given unlimited access to rat chow and water containing penicillin and streptomycin.
2.6. Singer photon emission computed tomography (SPECT) imaging Radionuclide scans were performed on weeks 4, 6 and 8 on a dual-head EPIC Vertex SPECT equipped with an LEGP collimator (V-60; ADAC, USA). Animals were deprived of food overnight and anesthetized with chloral hydrate (6.1 g/kg body weight) 10 min prior to imaging. The rats were injected intravenously with100 mCi of 99m Tc-DTPA (Jiangsu Institute of Nuclear Medicine, China), and then subjected to a 30 min dynamic SPECT immediately after the injection. For PET quantification, the whole right and left kidneys were included and all images were analyzed by the Gates method [24]. 2.7. Histology and immunofluorescence analysis Samples were prepared for histological and immunofluorescence analyses by following standard protocols for paraffin embedding. Histological analysis, mounted kidney sections were stained by hematoxylin and eosin (H&E) to visualize nuclei (blue), and both cytoplasm and connective tissue (pink). For immunofluorescence, following deparaffinization, rehydration and antigen retrieval, the sections were blocked with 5% normal bovine serum in PBS for 30 min. Sections were then incubated with primary antibodies overnight at 4 C. The primary antibodies used were as follows: mouse anti-nestin (1:200; Abcam,UK) and rabbitanti-AQP1 (1:200; Abcam). After being washed in PBS, sections were then incubated in 488- or 594-conjugated species-specific secondary antibody (Chemicon, USA) for 2 h. Slides were observed with an Olympus fluorescent microscope and images were captured using Olympus soft image viewer. The intensity of immunoreactivity was quantified using the Image-Pro Plus 6.0 software (Media Cybernetics, USA). 2.8. Enzyme-linked immunosorbentassay (ELISA) for quantitative analysis of cytokines in DC kidney scaffolds Total protein in DC kidney scaffolds or intact kidney was extracted using an ELISA kit (R&D Systems, USA). The concentrations of various cytokines including VEGF, TGF, HGF, IL-8, CTGF, FGF and PDGF were assayed with a microplate reader at 450 nm [25]. 2.9. Statistical analysis Quantitative results were reported as mean standard deviation. Independent samples t-test was used to reveal differences in the level of cytokines and intensity of immunofluorescence among different groups. The significance level was set at 0.05 and SPSS 17.0 (SPSS Inc., Chicago, USA) was used for the analyses.
3. Results 3.1. Characterization of DC kidney scaffolds After decellularization with continuous detergent perfusion, DC kidney scaffolds had a somewhat transparent appearance (Fig. 1A). Vascular corrosion casting showed that the vascular tree in the DC kidney scaffolds was well maintained compared with intact kidney (Fig. 1B,C). H&E staining revealed that blue-stained nuclei were not observed, but pink-stained components were present in the DC kidney scaffolds compared with control (Fig. 1D,E). As pink-stained components include both cytoplasm and extracellular matrix, and this may reflect the morphological difference in H&E staining between intact kidney and DC kidney scaffolds (Fig. 1D,E). In addition, PAS and Masson’s staining were used to examine the ECM (e.g. polysaccharides and collagen), and it showed that positivelystained structures were present in DC kidney scaffolds without interruption (Fig. 1F,G). Finally, electron microscopy observation indicated that nuclear structure was not found, but the integrity of extracellular matrix was not disrupted in the DC kidney scaffolds as shown by the presence of continuous membrane of Bowman’s capsule, the basement membrane of the glomerular capillaries and mesangial matrix (Fig. 1H). Taken together, the DC kidney scaffolds
6824
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
Fig. 1. Characterization of the DC kidney scaffolds. (A) Gross appearance of harvested DC kidney scaffolds. (B) Electron microscopy observation shows intact extracellular matrix in DC kidney scaffold. Blue arrows indicate the membrane of Bowman’s capsule, a green arrow indicates the basement membrane of the glomerular capillaries, and a red star points to the mesangial matrix. (C, D) Vascular corrosion casting shows a normal vascular tree of DC kidney scaffold (D) compared with intact kidney (C). (E, F) H&E staining shows the existence of blue-stained nuclei in intact kidney (E) but not DC kidney scaffold (F). (G) PAS staining shows the presence of the ECM (e.g. basement membranes) in DC kidney scaffold. Noted that the capillary loops of the glomeruli are clearly displayed. (H) Masson’s staining shows that green-stained collagenous fibers in DC kidney scaffold. (I) Quantitative assay of cytokines in DC kidney scaffolds. The level of PDGF CTGF, TGF-b and VEGF in the DC kidney is not different from that of native kidney, while that of IL-8, FGF and HGF is reduced. Scar bars ¼ 5 mm (A, C D), 100 mm (EeH) and 5 mm (B).
prepared in our study lose renal cells but keep normal vascular tree and continuous extracellular matrix. Considering that SDS is toxic to cells, UV-VIS was used to assess SDS residue levels in the scaffolds. A standard concentration curve for SDS was generated by measuring the absorbance of eight different known concentrations of SDS. According to this standard curve, the mean residual SDS concentration in the DC kidney scaffolds was 50.0 1.7 mg/g, which is below the safe level of 133.3 mg/g [26]. To assess the potential of DC kidney scaffolds for renal regeneration, we performed an ELISA assay to quantify levels of various cytokines. As shown in Fig. 1I, the levels of cytokines HGF, CTGF, TGF-b and VEGF in DC kidney scaffolds were similar to that of the intact kidney, although some of them (IL-8, FGF and PDGF) were lower than the intact kidney. Thus, although the rat kidneys had been completely decellularized, some cytokines still remained within the scaffolds in concentrations that might be sufficient to contribute to renal regeneration after injury. 3.2. Renal degeneration in DC kidney scaffolds The regenerative potential of the DC kidney scaffolds was tested by grafting them onto partially nephrectomized rat kidneys (Fig. 2A). The lower 1/3 of kidney was removed, and similar sizedDC kidney scaffolds (left lower panel) were grafted onto the cutend. Rats with cut-end-sutured kidney were used as control. As shown by non-invasive B-scan ultrasonography performed at weeks 1, 2 and 4 post-surgery, kidneys treated with DC scaffolds exhibited some macroscopic regrowth, whereas control kidneys did not (Fig. 2B,C). In addition, obvious bloodstream signals were observed in the RDS area starting2 weeks after the RDS transplantation (Fig. 2B,C).
To examine the regeneration in more detail, some rats were sacrificed on 1, 2, 4 and 8 weeks post-surgery and their kidneys were removed for analysis. As shown in Fig. 3AeH, a macroscopic regrowth of the renal tissue into the grafting DC kidney scaffolds occurred in the experimental group with the scaffold graft developing first into granulation tissue, then into a scar by week 8. This regrowth was also reflected by a downward moving of a boundary between the regenerated tissue and the scaffolds overtime (lines in Fig. 3AeH). The size of regenerated kidney was measured 4 weeks post-transplantation, and it showed a significant increase compared with controls (Fig. 2C; control, 1.35 0.05 cm 0.75 0.05 cm; scaffolds-grafted, 1.50 0.1 cm 0.76 0.08 cm; n ¼ 10 for each, P < 0.05). Microscopic examination showed that tissue obtained from the regions above the boundary between the regenerated tissue and the scaffolds (yellow marked area in Fig. 3AeD) contained the renal parenchyma cells on 1 and 2 weeks, and they became to tubular and glomerular morphology on 4 and 8 weeks post-grafting (Fig. 3IeL). In this area, numerous dark hematoxylin-stained cells were also observed, and these small cells gradually reduced in number overtime; they are probably inflammatory cells (Fig. 3IeL). By contrast, the tissue from the region below the boundary contained eosin-stained ECM of the scaffolds, and hematoxylin-stained cells that were also reduced overtime (Fig. 3MeP). Taken together, above data demonstrate an apparent regeneration of the renal tissue in the DC kidney scaffolds of partially-excised kidney. Having found the morphological regeneration in the DC scaffolds-grafted kidney, we were promoted to see if there is any functional recovery. Radionuclide renal scans were performed on 6 and 8 weeks post-grafting. Kidneys grafted with the DC scaffold tissue did not show improved glomerular filtration rate until 6 weeks, compared with controls (Fig. 4B). Six weeks after surgery,
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
6825
Fig. 2. The procedures of grafting and the regeneration of renal tissue in DC kidney scaffolds shown by B-scan ultrasonography and gross appearance of explanted kidneys. (A) Showing the procedures of grafting DC kidney scaffolds. Approximately 1/3 of the left kidney was transversely excised from each rat, and similar sized-scaffolds (arrow in the left lower panel) were grafted onto the cut-end by suturing the external capsules. Cut-end in control group was sutured directly. (B) B-scan ultrasonography images of the DC kidney scaffold-grafted obtained on weeks 1, 2 and 4 to assess the size, shape, and blood flow to the kidneys. Yellow circles represent the left kidneys, the white triangles represent implanted scaffolds area, the red signals represent artery blood flow, and blue ones represent vein flow. (C) Comparison of the long axis of kidney measured by B-scan ultrasonography. N ¼ 5for each, P < 0.05. (D) Gross appearance of explanted kidney scaffolds on weeks 8. D1, D2, D3 show right kidney and DC kidney scaffold-grafted group and control group respectively.
both experimental and control kidneys displayed a very mild delay in renal perfusion, as well as some degree of delay in excretion (Fig. 4C,D). In an attempt to characterize the regenerative cells found in the scaffold-grafted kidneys, we began by looking at the distribution of cells immune positive for nestin, a marker of renal stem/progenitor cells. In intact kidney, nestin immunoreactivity was only distributed in the glomeruli (Fig. 5A). In the scaffoldgrafted kidneys, we examined nestin expression in the tissue located above the boundary between the newly degenerated tissue and DC kidney scaffold at different post-surgery time points. On 1 week, in addition to their typical glomerular distribution, many nestin-positive cells were also present in the
renal tubules and in the interstitium near the incisal border (Fig. 5B, C1, D1), but they were noticeably reduced by 2 weeks (Fig. 5C2, D2) and had become undetectable by week 8 (Fig. 5C3, C4, D3, D4). At this time point, nestin-positive cells were only present in the glomeruli (Fig. 5C4, D4). In control kidney, nestin immunoreactivity was observed in the glomeruli and some scattered cell near the boundary on week 1, but they were not present at later time points (Fig. 5E). In addition, expression of aquaporin-1 (AQP-1), a channel protein typically located in mature proximal tubules [27], was examined. AQP-1 was not observed in the first two weeks post-surgery, but many AQP-1positive cells were observed in scaffold-grafted kidneys on week 3 and 4 (Fig. 5D).
Fig. 3. Experimental kidney recovery over 8 weeks. (AeH) Macroscopic images show longitudinal cross-sections of whole experimental kidneys (gross observation: AeD; observation under stereoscopic microscope: EeH) on weeks 1, 2, 4 and 8 post-surgery. The black line represents the border between the renal parenchyma and the grafted-DC scaffold. Yellow area and blue area were removed for light microscopic examination shown in IeL and MP, respectively. (IeP) H&E staining shows renal parenchyma (IeL; yellow squares in AeD) and DC scaffold (MeP; blue ellipses in AeD) at the four time points. On week 1, the renal parenchyma was seriously damaged, exhibiting extensive glomerular and tubular atrophy and widespread infiltration of inflammatory cells (I). By week 2, tissue atrophy was reduced in the injured renal parenchyma (J), and mild edema was observed on week 4 (K). By week 8, the renal parenchyma had recovered to a near-normal state (L). In the DC scaffold, inflammatory cells were seen infiltrating along the margin on week 1 (M). By week 2, the entire scaffold graft had been infiltrated (N). Granulation tissue had become to form by week 4 (O), and had developed into scar tissue by week 8 (P). For H&E staining in intact kidney, please see Fig. 1D. Scale bars ¼ 100 mm (IeP).
Fig. 4. Radionuclide scan analysis demonstrating a remarkable improvement in GFR compared with the control. (A) Normal renal function in healthy intact rats was used as a baseline comparator. Left panel is renal dynamic imaging. Right lower panel is the renogram showing the renal filtration and drainage, and the appearance of isotope in the bladder, which are also expressed by CTS per second. (B) Statistical analysis revealed significant differences in renal functions between the experimental and control groups at weeks 6 and 8 (n ¼ 10 for each; *P < 0.05). Vertical axis is glomerular filtration rate of the injured left kidney (GFRLI), glomerular filtration rate of the healthy left kidney (GFRLN), the indicator of GFR. (C, D) Radionuclide scan analysis of experimental (C) and control (D) kidneys on 6 weeks post-surgery. White arrows indicate the left kidney.
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
6827
Fig. 5. Immunofluorescent analysis showing that more nestin-positive cells (green) are present in experimental kidneys compared with controls. Tissues used for immunostaining were taken from the region located above the border between newly regenerated tissue and the scaffold at different time points. (A) Immunoreactivity of nestin is only present in the glomeruli of intact kidney. (B) Quantitative analysis showing significantly higher nestin immunoreactivity in experimental kidneys compared to controls, with a pronounced peak at week 1 post-surgery. (C) Nestin-positive cells are more abundant and widely distributed in experimental kidneys. In addition to the glomeruli, nestin protein is also detected in cortical tubules near the incision border, especially on week 1 (C1, C5). By week 8, nestin expression is restricted to the gomeruli (C4, C8). (D) Double immunostaining of nestin (green) and aquaporin (red). Aquaporin-positive cells are hardly observed in scaffold-grafted kidneys on week 1 or 2 (D1, D2), but increased greatly on week 4 and 8 (D3, D4). (E) In control kidneys, weak nestin expression is only seen on week 1.Scale bars ¼ 10 mm (A, D1eD4, E1eE4) and 25 mm (C1eC4).
4. Discussion Tissue scaffolds containing intact vascular and ECM architecture hold great therapeutic promise for the field of organ recovery, particularly in the case of highly vascular organs [18,22,28,29]. Recently, Harald C. Ott and his team were able to recellularize rat kidney scaffold grafts [22], and Giuseppe Orlando and colleagues successfully implanted porcine DC kidney scaffolds into experimental pigs [30]. In these two studies, some degree of both functional and physical renal recovery was attained, but further research is required in order to turn these observations into clinically relevant achievements. The findings presented herein provide additional support to the hypothesis that orthotopic DC scaffold grafts can induce macroscopic growth and regeneration of damaged kidneys, thus promoting in vivo renal function recovery. Our rat kidney scaffolds, which were decellularized by continuous detergent perfusion, retained vascular trees that were intact at all hierarchical levels. Residual detergent levels in the scaffolds were found to be negligible, well below the level deemed safe by the Food and Drug Administration. Overall renal ECM was also maintained, and the levels of various cytokines were maintained.
The ECM represents a substrate to which cells can adhere, but it also provides support and inductive cues for cellular proliferation, differentiation and migration. To date, attempts at recreating a synthetic environment mimicking that of the ECM have proven mostly unsuccessful [31]. In particular, the specific topography or geometry of the ECM has been shown to play a role in promoting osteogenic differentiation [32]. In addition, an artificial ECM containing the angiogenic laminin-1-derived IKVAV sequence was shown to cause a significant increase in the number and area of vascular branches in the chick chorioallantoic membrane [33]. In our model, rats were partially nephrectomized (by removing approximately 30% of the kidney parenchyma from the lower pole), and DC kidney scaffold tissue was grafted onto the resected kidney before suturing shut. Microscopic imaging revealed that progenitor cells were migrating from the injured kidney into the grafted scaffold tissue, suggesting that the scaffold’s intact ECM retains sufficient chemical and/or physical cues to induce the proliferation and differentiation of renal progenitor cells present nearby. Immunohistochemical analysis of post-surgery kidney sections revealed a significant number of nestin-positive cells within the repair zone of scaffold-grafted kidneys. By the interaction of injured kidney with the RDS, many nestin-positive renal progenitor cells were observed, especially in the second week. These observations
6828
Y.L. Yu et al. / Biomaterials 35 (2014) 6822e6828
suggest that several distinct mechanisms may be involved in DC scaffold-mediated regenerative kidney repair. First, the presence of the scaffold graft could potentially contribute to hemostasis by physically protecting/repairing the wound surface. In addition, the intact scaffold ECM might harbor binding sites for proteins that stimulate the proliferation of renal progenitor cells, thus contributing to kidney repair and regeneration. Finally, the grafted tissue may provide physical support as well as biochemical cues that promote the migration of renal cells from the injured kidney into the acellular scaffold. In the present study, we developed an experimental rat model of DC scaffold-induced renal recovery, and focused mainly on the more macroscopic aspects of kidney repair. In the future, we aim to also explore the molecular mechanisms involved in this process, and to expand our model to other test species, such as mice and pigs. 5. Conclusion In summary, we developed a decellularized (DC) rat kidney scaffold with overall architecture and significant concentrations of various cytokines that improves recovery of the kidney after partial nephrectomy, independently from renal endogenous cells and other factors. The main mechanisms responsible for this effect include the mechanical support of the scaffold, facilitation of cell migration and regeneration. Further investigations on therapeutic factors and/or cells that can be seeded within the decellularized scaffold can usher in a new clinical therapy for renal recovery in the treatment of chronic kidney disease. Acknowledgments The authors appreciate the input of Miaozhong Li and Zhiheng Rao in these studies. The authors are grateful to Jiawei Li for help in imaging. These studies were carried out with the support of the National Natural Science Foundation of China (81071576) and the Natural Science Foundation of Zhejiang Province (LY12H15002, LY14H050005). Appendix A. Supplementary data Supplementary video related to this article can be found at http://dx.doi.org/10.1016/j.biomaterials.2014.04.074. References [1] Jha V, Garcia-Garcia G, Iseki K, Li Z, Naicker S, Plattner B, et al. Chronic kidney disease: global dimension and perspectives. Lancet 2013;382:260e72. [2] Franquesa M, Flaquer M, Cruzado JM, Grinyo JM. Kidney regeneration and repair after transplantation. Curr Opin Organ Transplant 2013;18:191e6. [3] Wen D, Ni L, You L, Zhang L, Gu Y, Hao C-M, et al. Upregulation of nestin in proximal tubules may participate in cell migration during renal repair. Am J Physiol Renal Physiol 2012;303:1534e44. [4] Swetha G, Chandra V, Phadnis S, Bhonde R. Glomerular parietal epithelial cells of adult murine kidney undergo EMT to generate cells with traits of renal progenitors. J Cell Mol Med 2011;15:396e413. [5] Reule S, Gupta S. Kidney regeneration and resident stem cells. Organogenesis 2011;7:135e9. [6] Vogetseder A, Picard N, Gaspert A, Walch M, Kaissling B, Hir1 ML. Proliferation capacity of the renal proximal tubule involves the bulk of differentiated epithelial cells. Am J Physiol Cell Physiol 2008;294:22e8.
[7] Humphreys BD, Valerius MT, Kobayashi A, Mugford JW, Soeung S, Duffield JS, et al. Intrinsic epithelial cells repair the kidney after injury. Cell Stem Cell 2008;2:284e91. [8] Lin F, Moran A, Igarashi P. Intrarenal cells, not bone marrow-derived cells, are the major source for regeneration in postischemic kidney. J Clin Invest 2005;115:1756e64. [9] Yamanaka S, Blau HM. Nuclear reprogramming to a pluripotent state by three approaches. Nature 2010;465:704e12. [10] Hanna JH, Saha K, Jaenisch R. Pluripotency and cellular reprogramming: facts, hypotheses, unresolved issues. Cell 2010;143:508e25. [11] Li B, Cohen A, Hudson TE, Motlagh D, Amrani DL, Duffield JS. Mobilized human hematopoietic stem/progenitor cells promote kidney repair after ischemia/ reperfusion injury. Circulation 2010;121:2211e20. [12] Ross EA, Williams MJ, Hamazaki T, Terada N, Clapp WL, Adin C, et al. Embryonic stem cells proliferate and differentiate when seeded into kidney scaffolds. J Am Soc Nephrol 2009;20:2338e47. [13] Krause D, Cantley LG. Bone marrow plasticity revisited: protection or differentiation in the kidney tubule? J Clin Invest 2005;115:1705e8. [14] Morigi M. Mesenchymal stem cells are Renotropic, helping to repair the kidney and improve function in Acute renal Failure. J Am Soc Nephrol 2004;15:1794e804. [15] Bonventre JV. Dedifferentiation and proliferation of surviving epithelial cells in Acute renal Failure. J Am Soc Nephrol 2003;14:55Se61S. [16] Schuldiner M, Yanuka O, Itskovitz-Eldor J, Melton DA, Benvenist N. Effects of eight growth factors on the differentiation of cells derived from human embryonic stem cells. Proc Natl Acad Sci U S A 2000;97:11307e12. [17] Badylak SF, Taylor D, Uygun K. Whole-organ tissue engineering: decellularization and recellularization of three-dimensional matrix scaffolds. Annu Rev Biomed Eng 2011;13:27e53. [18] Orlando G, Booth C, Wang Z, Totonelli G, Ros CL, Moran E, et al. Discarded human kidneys as a source of ECM scaffold for kidney regeneration technologies. Biomaterials 2013;34:5915e25. [19] Ross EA, Abrahamson DR, St John P, Clapp WL, Williams MJ, Terada N, et al. Mouse stem cells seeded into decellularized rat kidney scaffolds endothelialize and remodel basement membranes. Organogenesis 2012;8:49e55. [20] Park KM, Woo HM. Porcine bioengineered scaffolds as new frontiers in regenerative medicine. Transplant Proc 2012;44:1146e50. [21] Song JJ, Ott HC. Organ engineering based on decellularized matrix scaffolds. Trends Mol Med 2011;17:424e32. [22] Song JJ, Guyette JP, Gilpin SE, Gonzalez G, Vacanti JP, Ott HC. Regeneration and experimental orthotopic transplantation of a bioengineered kidney. Nat Med 2013;19:646e51. [23] Nakayama KH, Batchelder CA, Lee CI, Tarantal AF. Decellularized rhesus monkey kidney as a three-dimensional scaffold for renal tissue engineering. Tissue Eng Part A 2010;16:2207e16. [24] Melis M, de Swart J, de Visser M, Berndsen SC, Koelewijn S, Valkema R, et al. Dynamic and static small-animal SPECT in rats for monitoring renal function after 177Lu-labeled Tyr3-octreotate radionuclide therapy. J Nucl Med 2010;51:1962e8. [25] Banks J, Muriel A, Smith JP. Attrition and health in ageing studies: evidence from ELSA and HRS. Longit Life Course Stud 2011;2. [26] Nam CW, Kang SJ, Kang YK, Kwak MK. Cell growth inhibition and apoptosis by SDS-solubilized single-walled carbon nanotubes in normal rat kidney epithelial cells. Arch Pharm Res 2011;34:661e9. [27] Hara-Chikuma M, Verkman AS. Aquaporin-1 facilitates epithelial cell migration in kidney proximal tubule. J Am Soc Nephrol 2006;17:39e45. [28] Uygun BE, Soto-Gutierrez A, Yagi H, Izamis ML, Guzzardi MA, Shulman C, et al. Organ reengineering through development of a transplantable recellularized liver graft using decellularized liver matrix. Nat Med 2010;16:814e20. [29] Serpooshan V, Zhao M, Metzler SA, Wei K, Shah PB, Wang A, et al. The effect of bioengineered acellular collagen patch on cardiac remodeling and ventricular function post myocardial infarction. Biomaterials 2013;34:9048e55. [30] Orlando G, Farney AC, Iskandar SS, Mirmalek-Sani SH, Sullivan DC, Moran E, et al. Production and implantation of renal extracellular matrix scaffolds from porcine kidneys as a platform for renal bioengineering investigations. Ann Surg 2012;256:363e70. [31] Chen XD. Extracellular matrix provides an optimal niche for the maintenance and propagation of mesenchymal stem cells. Birth Defects Res C Embryo Today 2010;90:45e54. [32] Seo CH, Furukawa K, Suzuki Y, Kasagi N, Ichiki T, Ushida T. A topographically optimized substrate with well-ordered lattice micropatterns for enhancing the osteogenic differentiation of murine mesenchymal stem cells. Macromol Biosci 2011;11:938e45. [33] Nakamura M, Yamaguchi K, Mie M, Nakamura M, Akita K, Kobatake E. Promotion of angiogenesis by an artificial extracellular matrix protein containing the laminin-1-derived IKVAV sequence. Bioconjug Chem 2009;20:1759e64.
