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Reconstruction of renal glomerular tissue using collagen vitrigel scaffold
JOURNAL OF BIOSCIENCE AND BIOENGINEERING Vol. 99, No. 6, 529–540. 2005 DOI: 10.1263/jbb.99.529
© 2005, The Society for Biotechnology, Japan
Reconstruction of Renal Glomerular Tissue Using Collagen Vitrigel Scaffold PI-CHAO WANG1* AND TOSHIAKI TAKEZAWA2 Institute of Applied Biochemistry, University of Tsukuba, 1-1-1 Tennodai, Tsukuba, Ibaraki 305-8572, Japan1 and Laboratory of Animal Cell Biology, National Institute of Agrobiological Sciences, Ikenodai 2, Tsukuba, Ibaraki 305-0901, Japan2 Received 19 April 2005/Accepted 25 April 2005
The construction of renal glomerular tissue has provided an important tool not only for the understanding of renal physiology and pathology in blood ultrafiltration and cell dysfunction, but also in the application of tissue engineering to glomeruli regeneration and nephritic therapy. In this study, a novel method to reconstruct glomerular tissue combining cultured cells on a collagen vitrigel scaffold is described. The method consists of two newly developed techniques, one to isolate glomerular epithelial and mesangial cells rapidly from kidney, which facilitates the prolongation of cell population doublings and allows a long-term cell culture without losing cellular features, and another to prepare a stable and thin transparent collagen gel membrane termed collagen vitrigel that can facilitate three-dimensional cultures for reconstructing an epithelial-mesenchymal model. By combining the two methods, we cocultured glomerular epithelial and mesangial cells on both surfaces of the collagen vitrigel by the manipulation of two-dimensional cultures, resulting in the successful reconstruction of a three-dimensional glomerular organoid. The coculture results showed that the collagen vitrigel maintains cell growth and cell viability for more than 1 month, and surprisingly, the epithelial layer demonstrated polarity formation, which usually appears in in vivo normal epithelial cells existing at the glomerular basement membrane, but seldom appears in epithelial cells cultured in vitro. Moreover, the coculture results showed that fibronectin, an extracellular matrix component, and integrin b1, a receptor of fibronectin, were detected in high amounts on both cells, suggesting our collagen vitrigel can provide a suitable environment for cell–cell interactions that stabilize the cell structure and may contribute to the polarity formation of epithelial cells. [Key words: collagen vitrigel, tissue reconstruction, glomerular mesangial cells, glomerular epithelial cells, extracellular matrix, three-dimensional culture]
Kidney has an important function in maintaining homeostasis and one of the essential functions of the kidney is the ultrafiltration of blood (1). The human kidney contains about one million filtration units, called nephrons. Each nephron is a self-functioning unit consisting of a renal corpuscle and a tubular system. The renal corpuscle consists of the glomerulus, a tuft of capillaries that is surrounded by Bowman’s capsule. The formation of urine starts with the ultrafiltration of blood in the glomerulus (2). The glomerulus is a ball of capillaries surrounded by Bowman’s capsule (3, 4). The glomerulus contains three cell types and two types of extracellular matrices (ECMs), the mesangial matrix and the glomerular basement membrane (GBM) (Fig. 1A). The three cell types include mesangial cells (MCs), glomerular endothelial cells (GENs), and epithelial cells (podocytes). Mesan-
gial cells and endothelial cells are closely associated at the inner surface of the glomerular basement membrane, whereas visceral epithelial cells cover the outer shell (5). The mesangial cells are multipotential in their functions and capacities and are believed to be critical to the regulation of glomerular hemodynamics (6, 7). Because the glomerular mesangial cells contract in vitro in response to a variety of hormones, it has been proposed that the cells modify the glomerular capillary filtration surface area (8). Moreover, as specialized pericytes, they also participate in the regulation of macromolecular trafficking and clearance, and as both a source and target of numerous hormones and autocrines (9). Much insight into mesangial cell function has been obtained by studying mesangial cells in culture (10, 11). The accumulation of extracellular matrix (ECM) in the mesangium or the overproliferation of mesangial cells causes the irreversible progression of glomerular sclerosis and chronic renal failure, and has been observed in the course of glomeru-
* Corresponding author. e-mail: [email protected] phone: +81-(0)29-853-7098 fax: +81-(0)29-853-4605 529
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FIG. 1. Structure of glomerulus. (A) Cross-sectional structure of glomerulus. Glomerulus is surrounded by Bowman’s capsule with an extensive basement membrane (GBM). Three different types of cells, mesangial cells (MC), endothelial cells, and epithelial cells (podocytes), were located in the glomerulus. Mesangial cells and endothelial cells are closely associated at the inner surface of the glomerular basement membrane, whereas visceral epithelial cells (podocytes) cover the outer shell. Filtration slits are formed through podocyte foot processes. Nephrotic syndrome is indicated on the left hand side, and normal glomerular morphology is shown on the right hand side. (B) Normal podocytes form polarity (arrow). (C) Nephrotic syndrome caused the podocytes to lose their polarity and develop a flat morphological change in cell architecture (arrow).
lar nephritis and diabetic nephropathy (12, 13). Glomerular epithelial cells (podocytes) cover the outer aspect of the glomerular capillary. The podocyte has a complex cellular organization and the cell can be divided into three distinct parts: the cell body, primary major processes, and secondary foot processes. Only the foot processes are attached to the GBM, leaving the rest of the cell hanging freely in the urinary space of Bowman’s capsule. Podocytes have several important functions in ultrafiltration. First, they produce and maintain the GBM. For this purpose, podocytes have polarity (Fig. 1B, arrow) and a slit diaphragm through which urine can pass smoothly while proteins remain within the glomeruli and return to the circulation. In the pathological condition of glomerular nephritis, epithelial cells loose their polarity and develop flat morphological changes in their cell architecture (14) (Fig. 1C, arrow). Flat epithelial cells cause the slit diaphragm to become more narrow and are involved in most diseases affecting the glomerulus, such as Alport syndrome, Nail patella syndrome, congenital nephritic syndrome of the Finnish type, and focal segmental glomerulosclerosis (FSGS). Several methods have been developed to isolate glomeruli and culture the three types of glomerular cells. However, the methods have two obstacles; the difficulty obtaining pure cell populations and the short life span of the cells cultured in vitro. Among the three cell types, isolation of the podocytes is the most difficult. Kriz et al. (14) and Yaoita et al. (15) first reported the isolation of podocytes. However, their explant method was complicated and time-consuming. In addition, the isolated podocytes loose their original mor-
phology quickly and tend to become unviable soon after the first cell passage. In order to solve these problems, we developed a digestion method using collagenase to detach Bowman’s capsule in a short period and isolate glomeruli easily and rapidly. The glomerular epithelial cells that proliferated from such glomeruli showed a significant multiarborizing morphology and survived longer than those reported previously. Using this method following slight modification, mesangial cells can also proliferate well and the cell population doubling level can be prolonged more than twofold compared to cells isolated by the traditional method. The communication circuits between cells or between cells and matrices have been found to be important for the maintenance of normal tissue physiology and for the initiation and persistence of pathophysiologic abnormalities of glomerular inflammation which may lead to glomerular sclerosis. Recently, using a cell culture technique and a coculture system, the interactions between monocytes/macrophages and mesangial cells (16, 17), and between glomerular endothelial cells and mesangial cells (18) were investigated. However, very few cocultures have been performed for glomerular epithelial cells (podocytes) and mesangial cells. Several experiments were conducted to reconstitute apical epithelial cells in vitro, but they seldom succeeded. In a coculture system using a commercially available transwell, rat glomerular epithelial cells were reported to secrete soluble factors that regulate the proliferation of quiescent rat mesangial cells, but the factors have not been characterized yet (19). It has also been reported that rat mesangial cells secrete hepatocyte growth factor (HGF) to maintain
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the renal structure and protect epithelial cells from apoptosis by using the same coculture system (20). However, the transwell system is composed of a membrane well sitting above the bottom well, and its function mainly resides in the separation of different cells from each other instead of providing a scaffold for different cells to adhere thereon. To the contrary, the glomerular basement membrane in vivo plays an important role to provide various cells with a suitable scaffold to which cells can adhere directly and have a closer relationship between cells and ECM. Various three-dimensional culture systems have been developed by devising the structures and components of cellular scaffolds to reconstruct organoids that can be utilized for in vitro normal or pathological models to examine drug effects and/or toxicities, for ex vivo extracorporeal devices to assist defective organs, or for in vivo grafts (21). Culture systems to reorganize a three-dimensional multicellular mass are classified into the following five categories: (i) gel culture systems utilizing collagen gel and matrigel scaffolds (22–24); (ii) spheroid culture systems utilizing thermo-responsive polymer, agarose, and polyurethane foam scaffolds (25–28); (iii) medium circulation culture systems utilizing scaffolds such as hollow fiber and cotton gauze (29, 30); (iv) tissue architecture-incorporated culture systems utilizing scaffolds such as acellular dermis, small intestinal submucosa, and tissue/organ sections for histopathology (TOSHI) (31–34), and (v) premolded biodegradable polymer-incorporated culture systems utilizing a polyglycolic acid (PGA) scaffold (35, 36). These three-dimensional culture systems have contributed not only to basic life science research but also to applied biomedical research. Human umbilical vein endothelial cells (HUVECs) cultured on matrigel form a network of capillary-like structures (22) and human microvasular endothelial cells (HMVECs) also form a network of branching tubular capillary-like structures into an overlaid collagen gel with embedded fibroblasts (23), demonstrating the reconstruction of in vitro angiogenesis models. The hetero-spheroids composed of mesenchymal cells (e.g., human dermal fibroblasts) and epithelial cells such as rat primary hepatocytes (25), human epidermal keratinocytes (26), and human carcinoma cell lines (27) were prepared by the following procedure. One type of epithelial cell was cocultured on a monolayer of mesenchymal cells precultured on a thermoresponsive scaffold. Each epithelial cell-attached mesenchymal cell sheet was detached from the scaffold by decreasing the temperature, and the harvested cell sheet was cultured on a nonadhesive agarose scaffold, resulting in the reconstruction of in vitro liver, skin, and cancer models. An extracorporeal circulation device for a defective kidney was reconstructed by seeding porcine renal tubule cells intraluminally into the hollow fiber cartridge as a scaffold (29). An extracorporeal circulation device for liver failure was reconstructed by seeding porcine hepatocytes into a multicapillary polyurethane foam packed-bed module as a scaffold to form numerous hepatocyte spheroids that were used as a pig ischemic liver (28). Transplantable skin neo-organs were reconstructed by culturing epidermal keratinocytes on a fibroblast-embedded collagen gel scaffold (24) or an acellular dermis scaffold (31, 32). A transplantable urinary bladder
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neo-organ was reconstructed by culturing canine urothelial cells and smooth muscle cells on the luminal surface and exterior surface of a premolded bladder-shaped polymer scaffold containing PGA, respectively (35). However, these three-dimensional culture systems, in comparison with two-dimensional systems, have some drawbacks. For example, the culture process is complicated, the observation of cells by a phase-contrast microscope is difficult or impossible, the reproducibility of the distribution of seeded cells is not always excellent, and aseptic handling for medium changes or the coculture of secondary cells is also difficult. Therefore, our aim was to develop a breakthrough technology that could overcome these problems. Opaque egg white or fish eyeballs that are prepared by boiling can be converted into thin, transparent, and rigid materials like glass by evaporating the moisture. This phenomenon is known as the vitrification of denatured proteins (37). On the other hand, it is well known that a clear collagen sol can become an opaque collagen gel by a gelation process with optimum salt concentration, pH, and temperature (38). The application of vitrification technology to a traditional opaque collagen gel results in its conversion into a rigid material like glass. A novel, stable collagen gel can be formed by dehydrating the glass-like material and has been named collagen vitrigel because it is a stable collagen gel membrane prepared via the three processes of gelation, vitrification, and rehydration. Collagen vitrigel possesses valuable physical properties (i.e., a thin and transparent membrane with enhanced gel strength). A framework-embedded collagen vitrigel scaffold enabled us to reconstruct a three-dimensional organoid in the same way as with an epithelial-mesenchymal model by performing conventional two-dimensional cultures for both surfaces of the scaffold and overcome the above problems concerning conventional three-dimensional cultures (39). In this study, we succeeded in the coculture of renal glomerular epithelial and mesangial cells by utilizing the collagen vitrigel and accomplished tissue reconstruction, especially polarity re-formation in epithelial cells that was seldom observed in individual cultures of epithelial cells on culture dishes. Furthermore, the interaction between mesangial and epithelial cells as well as the interaction between cell–ECM adhesion can be easily detected by using the collagen vitrigel, which may contribute to the investigation and clarification of the mechanisms of cell–cell and cell–ECM adhesion. I. METHODS FOR THE ISOLATION OF GLOMERULI, EPITHELIAL CELLS, AND MESANGIAL CELLS Glomeruli were isolated from male ICR mice using sterile technique and the cortex diced into small pieces which were forced through a stainless steel mesh (pore size, 150 mm). Unencapsulated glomeruli, which pass through this mesh, were then passed through the second and third meshes with pore sizes of 90 mm and 75 mm, respectively. The third mesh was turned upside down, and glomeruli that remained on the third mesh were washed with Hanks solution supplemented with 100 U/ml penicillin and 50 mg/ml streptomycin (hereafter referred to as basal medium). The existing
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FIG. 2. Morphology of glomerular epithelial cells (podocytes) and mesangial cells isolated by our method. (A) Glomerular epithelial cells (podocytes) with aborizing morphology. (B) Mesangial cell with contractile morphology.
method for isolating glomerular epithelial cells (podocytes) is called the explant method, which consists of many fine mince steps at the initial stage and thus the entire epithelial cell body is minced into small pieces and causes the outgrowth of epithelial cells very similar to endothelial cells in morphology (15). Instead of using fine mince steps, we developed a collagenase digestion method to remove Bowman’s capsule completely without harming the epithelial cell bodies. Collagenase at a concentration of 230–300 U/ml was found to be the optimum amount to detach the Bowman’s capsule. After the addition of collagenase to the glomeruli solution, the mixture was harvested immediately at 800 rpm and the supernatant containing single cells and small tubular fragments was aspirated off and the pellet containing glomeruli was washed with basal medium and centrifuged several times to completely remove the collagenase and single cells. The glomeruli were seeded in culture dishes coated with type I collagen. For the isolation of glomerular epithelial cells, culture medium consisting of basal medium supplemented with 10% FBS was used. Glomeruli were seeded on dishes and examined after 1 week of undisturbed outgrowth at 37°C in a 5% CO2 incubator. After the cell outgrowth had reached 50%, each dish was gently treated with 0.25% trypsin to remove fibroblasts and other types of cells and then DHSF medium (l–1: DMEM, 5 g; Ham’s F12, 5.3 g; NaHCO3, 1.9 g; ITS, 6.25 g; EGF, 1 mg; supplemented with 10% FBS) was added. Such treatment was conducted once every week until the homologous cells reached confluency. Figure 2A shows epithelial cells isolated by our method using collagenase digestion. The morphology of the epithelial cells was arborizing and could be maintained for more than 1 month without losing its specific morphology. However, after 1 month of culture, the tips of the epithelial cells gradually detached from the dish until finally they were suspended in the medium. Interestingly, after gentle trypsinization, the detached tips of the glomerular epithelial cells recovered and proliferated again. This has never been observed using the previous method because epithelial cells (podocytes) were reported to be fully differentiated cells without proliferation capability. To isolate glomerular mesangial cells, after the outgrowth of mesangial cells appeared in mesangial basal medium (MsBM; Cambrex Bioscience, Walkersville, MD, USA), trypsin was added to remove the outgrowing epithelial cells. Mesangial cells were passaged twice to make sure no other types of glomerular cells coexisted in the cell culture. The
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third passage was carefully performed after the contractile morphology had disappeared and all of the cells had turned into a small spindle shape. At this stage, the cells reached superconfluency. The superconfluent mesangial cells could be maintained for many more passages using the traditional method. Figure 2B shows the morphology of mesangial cells isolated by our method. Since mesangial cells play an important role in regulating blood pressure in glomeruli, they exhibit dynamic cell migration and specific contractile morphology. The appropriate time for mesangial cell passage has been suggested to be when cell proliferation reaches 70–80% confluency, which is usually 3–4 d after seeding (40, 41). However, such a method cannot maintain mesangial cells over 30 population doubling levels (PDLs) because the cells detach from the dish and eventually tend to become unviable. Our method in which culturing the primary cells continues until superconfluency, which is far beyond 80% confluency, provides an alternative way to prolong cell passage to 70 PDLs. Mesangial cells reaching superconfluency exhibited a spindle shape rather than a contractile flat morphology, but the contractile flat morphology could be restored after passaging the superconfluent cells. Identification was conducted on cells whose morphology was restored by immunofluorescence microscopic analysis using FITC-labeling monoclonal anti-Thy-1 antibody, a specific marker of mesangial cells (42–45). Figure 3A shows that the morphology-restoring mesangial cells were stained by anti-Thy-1 antibody, a marker for mesangial cells, suggesting these cells were mesangial cells although their morphology changed under a superconfluent state. RT-PCR showed the morphology-restoring cells strongly expressed Thy-1 and no nephrin band can be detected, where nephrin (46–48) and b-actin served as negative and positive markers, respectively (Fig. 3B, lane 3). On the other hand, neither Thy-1 nor nephrin was detected in epithelial cells (Fig. 3B, lane 1), while slight bands of both Thy-1 and nephrin were detected in the whole kidney organ sample (Fig. 3B, lane 2). These facts prove that our method for culturing mesangial cells in the long term is possible and that the application of mesangial cells to various cultures in vitro is promising. II. PREPARATION OF COLLAGEN VITRIGEL The collagen vitrigel was prepared via the three processes of gelation, vitrification, and rehydration. An equal volume of a 0.5% solution of acid-solubilized type-I collagen and the culture medium was uniformly mixed at 4°C to prepare a collagen sol and 2.0 ml of the collagen sol was poured into a hydrophilic culture dish with a diameter of 35 mm. The culture dish was incubated at 37°C and 5% CO2 for 2 h to induce complete gelation of the collagen. The collagen gel was then dried in a class 100 clean air chamber at 10°C and 40% humidity for an initial 2 d to remove most of the moisture in the gel. Dehydration was continued to sufficient dryness at room temperature for several additional days to convert the gel into a rigid glass-like material. The collagen vitrigel was prepared by pouring 3 ml of PBS onto the vitrified rigid material in the culture dish, letting it stand for about 10 min, and repeating the process three times to completely
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FIG. 3. Identification of morphology-restoring glomerular mesangial cells by immuno-fluorescene microscopic analysis and RT-PCR. (A) Mesangial cell stained with Thy-1 immunofluorescence. (B) RT-PCR of gene purified from epithelial cell (lane 1), whole kidney (lane 2) and mesangial cell (lane 3). Thy-1, A specific protein marker for mesangial cells. Nephrin, A specific protein marker for slit diaphragm at foot processes of podocyte. b-Actin, Control marker.
rehydrate and balance it with PBS. To prepare the framework-embedded collagen vitrigels, a nylon membrane cut into a ring shape with an inner-outer diameter of 23–33 mm or a pressed slid sheet cut into a circular shape with a diameter of 33 mm was sterilized with 70% ethanol for 10 min, washed with PBS three times to completely remove the ethanol, and inserted into a culture dish before pouring the collagen sol. The silk sheet was prepared by adding moisture to uniformly spread cocoon filaments, and then pressing them at a high temperature (120°C) and agglutinating the sericin. Figure 4 shows a schematic representation of the protocol for the preparation of collagen vitrigels with and without the framework of a nylon membrane ring or a pressed silk sheet. The collagen sol poured into culture dishes already containing a nylon ring membrane formed an opaque and soft collagen gel in which the nylon ring was embedded (Fig. 4A). The gel was converted into a rigid glass-like material with salt precipitations derived from the culture medium by drying sufficiently (Fig. 4B). The vitrified material was regenerated as a stable thin gel membrane by rehydrating with PBS (Fig. 4C) which we termed vitrigel. The vitrigel, washed and balanced with PBS, is absolutely transparent (Fig. 4E). Vitrigel embedded with the frameworks of a nylon ring (Fig. 4E) and a silk sheet (Fig. 4F) showed selfsupporting membrane architectures while the vitrigel without frameworks appeared as a self-twisting membrane structure (Fig. 4D) when they were held with forceps. III. PHYSICAL PROPERTIES OF COLLAGEN VITRIGEL The physical properties of the collagen vitrigel, including the changes in net weight, mechanical strength, and transparency in the vitrification process were evaluated as follows. The net weight of each collagen gel in the vitrification process was calculated by subtracting the premeasured dish weight from the gross weight of the gel and dish. Weights
were measured using a digital scale (AG245; Mettler-Toledo International, Greifensee, Germany). In the vitrification process, the collagen gel outwardly showed a dried state within the first 2 d. The net weight of the collagen gel was reduced to about 14.5% of the initial net weight after a vitrification period of 2 d and reached a plateau level of about 1.5% of the initial net weight after 7 d (Fig. 5). The mechanical strength of both collagen gels and collagen vitrigels prepared from different vitrification periods was measured with a digital force gauge (FGC-0.2B or FGC-2B; Nidec Shimpo, Kyoto) by modifying a previous method (49). Each nylon ring-embedded collagen gel or collagen vitrigel was carefully detached from the culture dish by forceps and then inserted and fastened between two glass cylinders with inner-outer diameters of 20–40 mm. A flat disk with a diameter of 12 mm connected to the force gauge was automatically approached at a speed of 9 mm/min and vertically inserted into the center region of each collagen gel or collagen vitrigel. The elastic force of the collagen gel or collagen vitrigel right before breaking was measured and recorded as the maximum elastic force (g force) indicated by the force gauge. Figure 6 shows that the mechanical strength of the collagen vitrigels increased as the vitrification period increased, which reached 813+ 41 g force (n = 3) after 84 d, which is about 19.65 times that of conventional collagen gels. The transparency of both collagen gels and collagen vitrigels prepared from different vitrification periods was measured with a spectrophotometer (V-550; JASCO, Tokyo). Each nylon ring-embedded collagen gel or collagen vitrigel was carefully detached from the culture dish by forceps and pasted on a flat quartz glass specific for the spectrophotometer. The absorbance of wavelengths from 200 to 900 nm was monitored and the optical density (OD) curve at 400 nm was plotted. Figure 7 shows that the absorbance at 400 nm of conventional opaque collagen gels was about 0.34 while that of collagen vitrigels after 2 d was about 0.2. The OD curve reached a plateau level of about 0.1, representing a
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FIG. 4. Schematic protocol for the preparation of collagen vitrigels with and without the framework of a nylon membrane ring or a pressed silk sheet. A vitrigel is produced via three processes, gelation, vitrification, and rehydration. Gross aspects of a conventional gel state (A), a vitrified state (B), and a rehydrated state (C) in the preparation process of a nylon ring-embedded collagen vitrigel, and of a collagen vitrigel without frameworks (D), a nylon ring-embedded collagen vitrigel (E), and a silk sheet-embedded collagen vitrigel (F) that are held by forceps. Bar represents 10 mm. Reproduced from Ref. 39 with permission of the publisher.
transparent state in collagen vitrigels with a vitrification period of more than 42 d. The OD from 350 to 650 nm was almost the same as that at 400 nm (data not shown). Morphological observations of both collagen gel and collagen vitrigel with a vitrification period of 72 d were per-
formed using a phase-contrast microscope (TE300; Nikon, Tokyo). Figure 8 shows the morphological results for both gels. Fine filament structures surrounded by many small particles were observed in a traditional collagen gel (Fig. 8A), whereas numerous small particles were observed in the col-
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FIG. 5. Changes in net weight of collagen gels in the vitrification process. The net weight of each collagen gel was calculated by subtracting the dish weight from the gross weight of the gel and dish. The value of the vitrification period at day 0 represents the net weight of the collagen gel prepared by incubating a collagen sol at 37°C for 2 h. Averages and standard deviations from five independent samples are presented. Reproduced from Ref. 39.
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FIG. 8. Phase-contrast microphotographs of a conventional collagen gel (A) and a collagen vitrigel (B). The conventional collagen gel and the collagen vitrigel were prepared by incubating a collagen sol at 37°C for 2 h and by rehydrating the collagen gel to a vitrified state for 72 d, respectively. Bars represent 100 mm. Reproduced from Ref. 39.
FIG. 9. Phase-contrast microphotographs of a pressed silk sheet (A) and a silk sheet-embedded collagen vitrigel (B). The silk sheetembedded collagen vitrigel was prepared from a 14-d vitrification period. Bars represent 100 mm. Reproduced from Ref. 39 with permission of the publisher. Reproduced from Ref. 39.
FIG. 6. Changes in the mechanical strength of collagen vitrigels prepared from different vitrification periods. The mechanical strength was recorded as the maximum elastic force indicated by the force gauge. The value of the vitrification period at day 0 represents the mechanical strength of the collagen gel prepared by incubating a collagen sol at 37°C for 2 h. Averages and standard deviations from three independent samples are presented. Reproduced from Ref. 39.
ment was about 15 mm and the filaments formed networks with pore size axes of shorter than 200 mm (Fig. 9A). The basic architectures of the pressed silk sheet were conserved in a silk sheet-embedded collagen vitrigel, although the density of the filaments increased slightly due to the compressed process of vitrification of the collagen gel (Fig. 9B). The mechanical strengths of the silk sheets immersed in PBS for 10 min and the silk sheet-embedded collagen vitrigels with a vitrification period of 14 d were 24,790 ± 3540 (n = 5) and 21,110 ± 2470 g force (n = 5), respectively. The latter mechanical strength was about 195 times stronger than the collagen vitrigels without the silk sheet prepared with the same vitrification period. IV. COCULTURE OF GLOMERULAR MESANGIAL CELLS AND EPITHELIAL CELLS ON COLLAGEN VITRIGEL AND RECONSTRUCTION OF RENAL GLOMERULAR TISSUE
FIG. 7. Changes in the transparency of collagen vitrigels prepared from different vitrification periods. The transparency was recorded as the absorbance of wavelength 400 nm monitored by a spectrophotometer. The value of the vitrification period at day 0 represents the transparency of the collagen gel prepared by incubating a collagen sol at 37°C for 2 h. Averages and standard deviations from three independent samples are presented. Reproduced from Ref. 39.
lagen vitrigel after a vitrification period of 72 d (Fig. 8B). Examination of a pressed silk sheet using a phase-contrast microscope revealed that the diameter of each cocoon fila-
The collagen vitrigel used in this study was immersed in PBS for 30 min and then placed on the inner ring edge of in vitro fertilization dishes (BD Falcon, Franklin Lakes, NJ, USA) so that cells facing the top and bottom sides of the collagen vitrigel could be immersed separately in different media. Since mesangial cells grow more slowly than epithelial cells, 1´104 mesangial cells were first seeded on one side of the collagen vitrigel. After an overnight culture, most of the mesangial cells adhered to the collagen vitrigel and cell proliferation continued for 1 week. The collagen vitrigel was then turned upside-down, and 1´104 epithelial cells were
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FIG. 10. Microscopic observation of glomerular epithelial cells and mesangial cells cocultured on opposite sides of collagen vitrigel. (A, D) Epithelial cells stained with HE, where cells on panel A reached to confluency; (B, E) mesangial cells stained with HE, where cells on panel E reached to confluency; (C, F) both epithelial and mesangial cells stained with HE. Bars represent 10 mm.
FIG. 12. Phase contrast microphotographs of collagen vitrigel section stained with HE. (A) Both sides of glomerular epithelial and mesangial cells; (B) glomerular epithelial cell side; and (C) mesangial cell side. Bars represent 10 mm.
seeded on the opposite side of the collagen vitrigel. Epithelial cells adhered to the gel immediately and proliferated to confluency in 3 d after seeding. MsBM medium was added to the mesangial cell side and DHSF medium was added to the epithelial cell side. Cells cultured in commercial dishes were used as a control. All of the cell cultures were carried out for 1 month by replacing with fresh media every week. After 1 month of cell culture on the collagen vitrigel, cells that had grown on the gel were examined by phase contrast microscopy. In order to distinguish the morphology and viability between mesangial cells and epithelial cells at opposite sides of the collagen vitrigel, cells were stained with hemotoxylin and eosin (HE) according to the standard procedure. The viability of mesangial cells and epithelial cells cultured on the collagen vitrigel was examined by incubating the cells in a LIVE/DEAD viability/cytotoxicity kit (Molecular Probe, Eugene, OR, USA) at room temperature for 30 min. Viable cells selectively incorporate the nonfluorescent calcein-AM, cleave the AM moiety by endogenous esterase, and yield green fluorescent calcein molecules. Dead cells selectively incorporate ethidium homodimer-1 into the
FIG. 13. Phase contrast microphotographs of glomerular epithelial cells on collagen vitrigel section (A) and on commercial dishes coated with collagen (B) stained with HE. Bars represent 2 mm.
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FIG. 11. Identification of cell viability of glomerular mesangial cells and epithelial cells cocultured on collagen vitrigel. (A) Epithelial cells stained with HE; (B) viable (green) mesangial cells stained with LIVE/DEAD kit; (C) dead (red) mesangial cells stained with LIVE/DEAD kit; (D) epithelial cells stained with HE; (E) viable (green) epithelial cells stained with LIVE/DEAD kit; and (F) dead (red) epithelial cells stained with LIVE/DEAD kit. Bars represent 1 mm.
FIG. 14. Immunofluorescent microphotographs of glomerular mesangial and epithelial cells in single cell culture and coculture system of collagen vitrigel. Monoclonal antibodies of a-actin, fibronectin, and integrin b1 were used to identify the cell–ECM interaction. (A, C) Mesangial cells stained with a-actin in individual culture and coculture, respectively; (E, G) mesangial cells stained with fibronectin in individual culture and coculture, respectively; (I, K) mesangial cells stained with integrin b1 in individual culture and coculture, respectively; (B, D) epithelial cells stained with a-actin in individual culture and coculture, respectively; (F, H) epithelial cells stained with fibronectin in individual culture and coculture, respectively; (J, L) epithelial cells stained with integrin b1 in individual culture and coculture, respectively. Panels C and D are microphotographs from the same location of vitrigel where mesangial and epithelial cells were cultured on the opposite sides and stained by a-actin labeled with rhodamin-phalloidin (red), respectively. The same for panels G and H, and panels K and L, except that each pair was stained by anti-fibronectin-antibody labeled with FITC (green) and anti-integrin b1 antibody labeled with FITC (green), respectively. Bars represent 1 mm.
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nuclei, and yield red fluorescent ethidium molecules in the nuclei. The viable and dead cells were examined using a fluorescent microscope (DM RXA; Leica, Wetzlar, Germany). Figure 10 shows that cells on collagen vitrigel can be observed by microscopy without any difficulty. Micrograph focusing on the glomerular epithelial cells and mesangial cells is shown in Fig. 10A and 10D, and 10B and 10E, respectively. Figures 10C and 10F show that both cells can be observed simultaneously on the collagen vitrigel after HE staining. Cell viability was confirmed by staining with the LIVE/DEAD kit. The results indicated that mesangial cells as well as epithelial cells grew to confluency (Fig. 11A, D), and that the green fluorescence produced by viable cells was as high as 99% (Fig. 11B, E) while the red fluorescence produced by dead cells was only 1% for both mesangial and epithelial cells (Fig. 11C, F). Cross sections of cultured specimens were obtained by cutting them around their center at a thickness of 4 mm using a microtome (RM2145; Leica Microsystems, Wetzlar, Germany) and then staining with hematoxylin and eosin according to the standard procedure for optical observation. Figure 12A shows that both cells adhered to the opposite sides of the collagen vitrigel. The sections of glomerular epithelial cells and mesangial cells were labeled as B and C, respectively. The enlarged areas of sections B and C (Fig. 12B, C) showed mesangial cells had a flat morphology while epithelial cells revealed polarity formation. In order to verify that the polarity was only formed on epithelial cells cultured on vitrigel in the coculture system, glomerular epithelial cells cultured on collagen vitrigel and commercial dishes coated with collagen for 2 weeks were further examined. Cross sections of both cellular samples proved that only epithelial cells cocultured with mesangial cells on the collagen vitrigel showed polarity formation (Fig. 13A), whereas those cultured on commercial dishes had a flat morphology (Fig. 13B).
be enhanced on mesangial (Fig. 14G) and epithelial cells (Fig. 14H) in cocultures as compared to those in individual cultures (Fig. 14E, F). Since mesangial cells are known to be responsible for fibronectin synthesis, we suspect that the synthesized fibronectin permeated to the epithelial cell side from the mesangial cell side through collagen vitrigel and stimulated integrin, the receptor of fibronectin, on the epithelial cell surface. In order to clarify this point, integrin b1 for both cells was detected. As expected, the expression of integrin b1 was enhanced in mesangial cells (Fig. 14K) and epithelial cells (Fig. 14L) compared to that in individual cell cultures (Fig. 14I, J). Although it cannot be ruled out that the serum in the medium contains fibronectin, it is easy to determine that fibronectin was consumed in the long-term cultures, especially in the individual cell cultures. It is likely that the fibronectin detected in the epithelial cells of the coculture system was due to its secretion by mesangial cells. This fact suggests that secreted fibronectin plays a role in inducing the expression of integrin, the ECM receptor, on the surface of epithelial cells, and enhances the adhesion of epithelial cells to collagen vitrigel. The stable adhesion of epithelial cells to a scaffold may contribute to stabilization of the cytoskeletal organization and promote polarity formation. In conclusion, the collagen vitrigel system provides an optimum environment mimicking the glomerular basement membrane to reconstitute renal glomerular tissue when coculturing various glomerular cells on it. Using this system, the polarity of epithelial cells can be re-formed and both epithelial and mesangial cells can maintain their viability and morphology in the long term. This collagen vitrigel system also provides an easy detection system with which to investigate the interactions between various cells and between cells to ECM which may contribute to the clarification of the mechanism of tissue regeneration. ACKNOWLEDGMENTS
V. CELL–ECM AND CELL–CELL INTERACTIONS FOR GLOMERULAR MESANGIAL CELLS AND EPITHELIAL CELLS IN COCULTURE AND INDIVIDUAL CULTURE The cell–ECM and cell–cell interactions were investigated since cocultures of glomerular mesangial cells and epithelial cells maintained stable adhesion to the collagen vitrigel for more than 1 month without impairing the cell viability. Stable cell adhesion is known to be linked to cell adhesion molecules, signal transduction, cell-to-cell contact, and extracellular matrix (50–53). For the interaction of cell–ECM adhesion, the expression of fibronectin, an ECM protein expressed by glomerular mesangial cells (54, 55) and its cellular receptor, integrin b1 (56, 57), were investigated. The results are shown in Fig. 14. The cytoskeletons of mesangial cells and epithelial cells stained with a-actin in individual cell cultures as controls are presented in Fig. 14A and 14B, while those of mesangial and epithelial cells in cocultures are shown in Fig. 14C and 14D. All of the cells showed proper cytoskeleton organization implying that the cells adhered to the collagen vitrigel and proliferation proceeded smoothly. Fibronectin expression was found to
This work was supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan (nos. 15360434 and 16656252) and by the 21st century COE program of the Ministry of Education, Culture, Sports, Science and Technology of Japan to one of the authors (PC.W.). The authors are grateful to Mr. Takuya Notani and Mr. Hideyuki Chiba for their help with the experiments.
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Reconstruction of Renal Glomerular Tissue Using Collagen Vitrigel Scaffold PI-CHAO WANG1* AND TOSHIAKI TAKEZAWA2 Institute of Applied Biochemistry, University of Tsukuba, 1-1-1 Tennodai, Tsukuba, Ibaraki 305-8572, Japan1 and Laboratory of Animal Cell Biology, National Institute of Agrobiological Sciences, Ikenodai 2, Tsukuba, Ibaraki 305-0901, Japan2 Received 19 April 2005/Accepted 25 April 2005
The construction of renal glomerular tissue has provided an important tool not only for the understanding of renal physiology and pathology in blood ultrafiltration and cell dysfunction, but also in the application of tissue engineering to glomeruli regeneration and nephritic therapy. In this study, a novel method to reconstruct glomerular tissue combining cultured cells on a collagen vitrigel scaffold is described. The method consists of two newly developed techniques, one to isolate glomerular epithelial and mesangial cells rapidly from kidney, which facilitates the prolongation of cell population doublings and allows a long-term cell culture without losing cellular features, and another to prepare a stable and thin transparent collagen gel membrane termed collagen vitrigel that can facilitate three-dimensional cultures for reconstructing an epithelial-mesenchymal model. By combining the two methods, we cocultured glomerular epithelial and mesangial cells on both surfaces of the collagen vitrigel by the manipulation of two-dimensional cultures, resulting in the successful reconstruction of a three-dimensional glomerular organoid. The coculture results showed that the collagen vitrigel maintains cell growth and cell viability for more than 1 month, and surprisingly, the epithelial layer demonstrated polarity formation, which usually appears in in vivo normal epithelial cells existing at the glomerular basement membrane, but seldom appears in epithelial cells cultured in vitro. Moreover, the coculture results showed that fibronectin, an extracellular matrix component, and integrin b1, a receptor of fibronectin, were detected in high amounts on both cells, suggesting our collagen vitrigel can provide a suitable environment for cell–cell interactions that stabilize the cell structure and may contribute to the polarity formation of epithelial cells. [Key words: collagen vitrigel, tissue reconstruction, glomerular mesangial cells, glomerular epithelial cells, extracellular matrix, three-dimensional culture]
Kidney has an important function in maintaining homeostasis and one of the essential functions of the kidney is the ultrafiltration of blood (1). The human kidney contains about one million filtration units, called nephrons. Each nephron is a self-functioning unit consisting of a renal corpuscle and a tubular system. The renal corpuscle consists of the glomerulus, a tuft of capillaries that is surrounded by Bowman’s capsule. The formation of urine starts with the ultrafiltration of blood in the glomerulus (2). The glomerulus is a ball of capillaries surrounded by Bowman’s capsule (3, 4). The glomerulus contains three cell types and two types of extracellular matrices (ECMs), the mesangial matrix and the glomerular basement membrane (GBM) (Fig. 1A). The three cell types include mesangial cells (MCs), glomerular endothelial cells (GENs), and epithelial cells (podocytes). Mesan-
gial cells and endothelial cells are closely associated at the inner surface of the glomerular basement membrane, whereas visceral epithelial cells cover the outer shell (5). The mesangial cells are multipotential in their functions and capacities and are believed to be critical to the regulation of glomerular hemodynamics (6, 7). Because the glomerular mesangial cells contract in vitro in response to a variety of hormones, it has been proposed that the cells modify the glomerular capillary filtration surface area (8). Moreover, as specialized pericytes, they also participate in the regulation of macromolecular trafficking and clearance, and as both a source and target of numerous hormones and autocrines (9). Much insight into mesangial cell function has been obtained by studying mesangial cells in culture (10, 11). The accumulation of extracellular matrix (ECM) in the mesangium or the overproliferation of mesangial cells causes the irreversible progression of glomerular sclerosis and chronic renal failure, and has been observed in the course of glomeru-
* Corresponding author. e-mail: [email protected] phone: +81-(0)29-853-7098 fax: +81-(0)29-853-4605 529
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FIG. 1. Structure of glomerulus. (A) Cross-sectional structure of glomerulus. Glomerulus is surrounded by Bowman’s capsule with an extensive basement membrane (GBM). Three different types of cells, mesangial cells (MC), endothelial cells, and epithelial cells (podocytes), were located in the glomerulus. Mesangial cells and endothelial cells are closely associated at the inner surface of the glomerular basement membrane, whereas visceral epithelial cells (podocytes) cover the outer shell. Filtration slits are formed through podocyte foot processes. Nephrotic syndrome is indicated on the left hand side, and normal glomerular morphology is shown on the right hand side. (B) Normal podocytes form polarity (arrow). (C) Nephrotic syndrome caused the podocytes to lose their polarity and develop a flat morphological change in cell architecture (arrow).
lar nephritis and diabetic nephropathy (12, 13). Glomerular epithelial cells (podocytes) cover the outer aspect of the glomerular capillary. The podocyte has a complex cellular organization and the cell can be divided into three distinct parts: the cell body, primary major processes, and secondary foot processes. Only the foot processes are attached to the GBM, leaving the rest of the cell hanging freely in the urinary space of Bowman’s capsule. Podocytes have several important functions in ultrafiltration. First, they produce and maintain the GBM. For this purpose, podocytes have polarity (Fig. 1B, arrow) and a slit diaphragm through which urine can pass smoothly while proteins remain within the glomeruli and return to the circulation. In the pathological condition of glomerular nephritis, epithelial cells loose their polarity and develop flat morphological changes in their cell architecture (14) (Fig. 1C, arrow). Flat epithelial cells cause the slit diaphragm to become more narrow and are involved in most diseases affecting the glomerulus, such as Alport syndrome, Nail patella syndrome, congenital nephritic syndrome of the Finnish type, and focal segmental glomerulosclerosis (FSGS). Several methods have been developed to isolate glomeruli and culture the three types of glomerular cells. However, the methods have two obstacles; the difficulty obtaining pure cell populations and the short life span of the cells cultured in vitro. Among the three cell types, isolation of the podocytes is the most difficult. Kriz et al. (14) and Yaoita et al. (15) first reported the isolation of podocytes. However, their explant method was complicated and time-consuming. In addition, the isolated podocytes loose their original mor-
phology quickly and tend to become unviable soon after the first cell passage. In order to solve these problems, we developed a digestion method using collagenase to detach Bowman’s capsule in a short period and isolate glomeruli easily and rapidly. The glomerular epithelial cells that proliferated from such glomeruli showed a significant multiarborizing morphology and survived longer than those reported previously. Using this method following slight modification, mesangial cells can also proliferate well and the cell population doubling level can be prolonged more than twofold compared to cells isolated by the traditional method. The communication circuits between cells or between cells and matrices have been found to be important for the maintenance of normal tissue physiology and for the initiation and persistence of pathophysiologic abnormalities of glomerular inflammation which may lead to glomerular sclerosis. Recently, using a cell culture technique and a coculture system, the interactions between monocytes/macrophages and mesangial cells (16, 17), and between glomerular endothelial cells and mesangial cells (18) were investigated. However, very few cocultures have been performed for glomerular epithelial cells (podocytes) and mesangial cells. Several experiments were conducted to reconstitute apical epithelial cells in vitro, but they seldom succeeded. In a coculture system using a commercially available transwell, rat glomerular epithelial cells were reported to secrete soluble factors that regulate the proliferation of quiescent rat mesangial cells, but the factors have not been characterized yet (19). It has also been reported that rat mesangial cells secrete hepatocyte growth factor (HGF) to maintain
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the renal structure and protect epithelial cells from apoptosis by using the same coculture system (20). However, the transwell system is composed of a membrane well sitting above the bottom well, and its function mainly resides in the separation of different cells from each other instead of providing a scaffold for different cells to adhere thereon. To the contrary, the glomerular basement membrane in vivo plays an important role to provide various cells with a suitable scaffold to which cells can adhere directly and have a closer relationship between cells and ECM. Various three-dimensional culture systems have been developed by devising the structures and components of cellular scaffolds to reconstruct organoids that can be utilized for in vitro normal or pathological models to examine drug effects and/or toxicities, for ex vivo extracorporeal devices to assist defective organs, or for in vivo grafts (21). Culture systems to reorganize a three-dimensional multicellular mass are classified into the following five categories: (i) gel culture systems utilizing collagen gel and matrigel scaffolds (22–24); (ii) spheroid culture systems utilizing thermo-responsive polymer, agarose, and polyurethane foam scaffolds (25–28); (iii) medium circulation culture systems utilizing scaffolds such as hollow fiber and cotton gauze (29, 30); (iv) tissue architecture-incorporated culture systems utilizing scaffolds such as acellular dermis, small intestinal submucosa, and tissue/organ sections for histopathology (TOSHI) (31–34), and (v) premolded biodegradable polymer-incorporated culture systems utilizing a polyglycolic acid (PGA) scaffold (35, 36). These three-dimensional culture systems have contributed not only to basic life science research but also to applied biomedical research. Human umbilical vein endothelial cells (HUVECs) cultured on matrigel form a network of capillary-like structures (22) and human microvasular endothelial cells (HMVECs) also form a network of branching tubular capillary-like structures into an overlaid collagen gel with embedded fibroblasts (23), demonstrating the reconstruction of in vitro angiogenesis models. The hetero-spheroids composed of mesenchymal cells (e.g., human dermal fibroblasts) and epithelial cells such as rat primary hepatocytes (25), human epidermal keratinocytes (26), and human carcinoma cell lines (27) were prepared by the following procedure. One type of epithelial cell was cocultured on a monolayer of mesenchymal cells precultured on a thermoresponsive scaffold. Each epithelial cell-attached mesenchymal cell sheet was detached from the scaffold by decreasing the temperature, and the harvested cell sheet was cultured on a nonadhesive agarose scaffold, resulting in the reconstruction of in vitro liver, skin, and cancer models. An extracorporeal circulation device for a defective kidney was reconstructed by seeding porcine renal tubule cells intraluminally into the hollow fiber cartridge as a scaffold (29). An extracorporeal circulation device for liver failure was reconstructed by seeding porcine hepatocytes into a multicapillary polyurethane foam packed-bed module as a scaffold to form numerous hepatocyte spheroids that were used as a pig ischemic liver (28). Transplantable skin neo-organs were reconstructed by culturing epidermal keratinocytes on a fibroblast-embedded collagen gel scaffold (24) or an acellular dermis scaffold (31, 32). A transplantable urinary bladder
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neo-organ was reconstructed by culturing canine urothelial cells and smooth muscle cells on the luminal surface and exterior surface of a premolded bladder-shaped polymer scaffold containing PGA, respectively (35). However, these three-dimensional culture systems, in comparison with two-dimensional systems, have some drawbacks. For example, the culture process is complicated, the observation of cells by a phase-contrast microscope is difficult or impossible, the reproducibility of the distribution of seeded cells is not always excellent, and aseptic handling for medium changes or the coculture of secondary cells is also difficult. Therefore, our aim was to develop a breakthrough technology that could overcome these problems. Opaque egg white or fish eyeballs that are prepared by boiling can be converted into thin, transparent, and rigid materials like glass by evaporating the moisture. This phenomenon is known as the vitrification of denatured proteins (37). On the other hand, it is well known that a clear collagen sol can become an opaque collagen gel by a gelation process with optimum salt concentration, pH, and temperature (38). The application of vitrification technology to a traditional opaque collagen gel results in its conversion into a rigid material like glass. A novel, stable collagen gel can be formed by dehydrating the glass-like material and has been named collagen vitrigel because it is a stable collagen gel membrane prepared via the three processes of gelation, vitrification, and rehydration. Collagen vitrigel possesses valuable physical properties (i.e., a thin and transparent membrane with enhanced gel strength). A framework-embedded collagen vitrigel scaffold enabled us to reconstruct a three-dimensional organoid in the same way as with an epithelial-mesenchymal model by performing conventional two-dimensional cultures for both surfaces of the scaffold and overcome the above problems concerning conventional three-dimensional cultures (39). In this study, we succeeded in the coculture of renal glomerular epithelial and mesangial cells by utilizing the collagen vitrigel and accomplished tissue reconstruction, especially polarity re-formation in epithelial cells that was seldom observed in individual cultures of epithelial cells on culture dishes. Furthermore, the interaction between mesangial and epithelial cells as well as the interaction between cell–ECM adhesion can be easily detected by using the collagen vitrigel, which may contribute to the investigation and clarification of the mechanisms of cell–cell and cell–ECM adhesion. I. METHODS FOR THE ISOLATION OF GLOMERULI, EPITHELIAL CELLS, AND MESANGIAL CELLS Glomeruli were isolated from male ICR mice using sterile technique and the cortex diced into small pieces which were forced through a stainless steel mesh (pore size, 150 mm). Unencapsulated glomeruli, which pass through this mesh, were then passed through the second and third meshes with pore sizes of 90 mm and 75 mm, respectively. The third mesh was turned upside down, and glomeruli that remained on the third mesh were washed with Hanks solution supplemented with 100 U/ml penicillin and 50 mg/ml streptomycin (hereafter referred to as basal medium). The existing
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FIG. 2. Morphology of glomerular epithelial cells (podocytes) and mesangial cells isolated by our method. (A) Glomerular epithelial cells (podocytes) with aborizing morphology. (B) Mesangial cell with contractile morphology.
method for isolating glomerular epithelial cells (podocytes) is called the explant method, which consists of many fine mince steps at the initial stage and thus the entire epithelial cell body is minced into small pieces and causes the outgrowth of epithelial cells very similar to endothelial cells in morphology (15). Instead of using fine mince steps, we developed a collagenase digestion method to remove Bowman’s capsule completely without harming the epithelial cell bodies. Collagenase at a concentration of 230–300 U/ml was found to be the optimum amount to detach the Bowman’s capsule. After the addition of collagenase to the glomeruli solution, the mixture was harvested immediately at 800 rpm and the supernatant containing single cells and small tubular fragments was aspirated off and the pellet containing glomeruli was washed with basal medium and centrifuged several times to completely remove the collagenase and single cells. The glomeruli were seeded in culture dishes coated with type I collagen. For the isolation of glomerular epithelial cells, culture medium consisting of basal medium supplemented with 10% FBS was used. Glomeruli were seeded on dishes and examined after 1 week of undisturbed outgrowth at 37°C in a 5% CO2 incubator. After the cell outgrowth had reached 50%, each dish was gently treated with 0.25% trypsin to remove fibroblasts and other types of cells and then DHSF medium (l–1: DMEM, 5 g; Ham’s F12, 5.3 g; NaHCO3, 1.9 g; ITS, 6.25 g; EGF, 1 mg; supplemented with 10% FBS) was added. Such treatment was conducted once every week until the homologous cells reached confluency. Figure 2A shows epithelial cells isolated by our method using collagenase digestion. The morphology of the epithelial cells was arborizing and could be maintained for more than 1 month without losing its specific morphology. However, after 1 month of culture, the tips of the epithelial cells gradually detached from the dish until finally they were suspended in the medium. Interestingly, after gentle trypsinization, the detached tips of the glomerular epithelial cells recovered and proliferated again. This has never been observed using the previous method because epithelial cells (podocytes) were reported to be fully differentiated cells without proliferation capability. To isolate glomerular mesangial cells, after the outgrowth of mesangial cells appeared in mesangial basal medium (MsBM; Cambrex Bioscience, Walkersville, MD, USA), trypsin was added to remove the outgrowing epithelial cells. Mesangial cells were passaged twice to make sure no other types of glomerular cells coexisted in the cell culture. The
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third passage was carefully performed after the contractile morphology had disappeared and all of the cells had turned into a small spindle shape. At this stage, the cells reached superconfluency. The superconfluent mesangial cells could be maintained for many more passages using the traditional method. Figure 2B shows the morphology of mesangial cells isolated by our method. Since mesangial cells play an important role in regulating blood pressure in glomeruli, they exhibit dynamic cell migration and specific contractile morphology. The appropriate time for mesangial cell passage has been suggested to be when cell proliferation reaches 70–80% confluency, which is usually 3–4 d after seeding (40, 41). However, such a method cannot maintain mesangial cells over 30 population doubling levels (PDLs) because the cells detach from the dish and eventually tend to become unviable. Our method in which culturing the primary cells continues until superconfluency, which is far beyond 80% confluency, provides an alternative way to prolong cell passage to 70 PDLs. Mesangial cells reaching superconfluency exhibited a spindle shape rather than a contractile flat morphology, but the contractile flat morphology could be restored after passaging the superconfluent cells. Identification was conducted on cells whose morphology was restored by immunofluorescence microscopic analysis using FITC-labeling monoclonal anti-Thy-1 antibody, a specific marker of mesangial cells (42–45). Figure 3A shows that the morphology-restoring mesangial cells were stained by anti-Thy-1 antibody, a marker for mesangial cells, suggesting these cells were mesangial cells although their morphology changed under a superconfluent state. RT-PCR showed the morphology-restoring cells strongly expressed Thy-1 and no nephrin band can be detected, where nephrin (46–48) and b-actin served as negative and positive markers, respectively (Fig. 3B, lane 3). On the other hand, neither Thy-1 nor nephrin was detected in epithelial cells (Fig. 3B, lane 1), while slight bands of both Thy-1 and nephrin were detected in the whole kidney organ sample (Fig. 3B, lane 2). These facts prove that our method for culturing mesangial cells in the long term is possible and that the application of mesangial cells to various cultures in vitro is promising. II. PREPARATION OF COLLAGEN VITRIGEL The collagen vitrigel was prepared via the three processes of gelation, vitrification, and rehydration. An equal volume of a 0.5% solution of acid-solubilized type-I collagen and the culture medium was uniformly mixed at 4°C to prepare a collagen sol and 2.0 ml of the collagen sol was poured into a hydrophilic culture dish with a diameter of 35 mm. The culture dish was incubated at 37°C and 5% CO2 for 2 h to induce complete gelation of the collagen. The collagen gel was then dried in a class 100 clean air chamber at 10°C and 40% humidity for an initial 2 d to remove most of the moisture in the gel. Dehydration was continued to sufficient dryness at room temperature for several additional days to convert the gel into a rigid glass-like material. The collagen vitrigel was prepared by pouring 3 ml of PBS onto the vitrified rigid material in the culture dish, letting it stand for about 10 min, and repeating the process three times to completely
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FIG. 3. Identification of morphology-restoring glomerular mesangial cells by immuno-fluorescene microscopic analysis and RT-PCR. (A) Mesangial cell stained with Thy-1 immunofluorescence. (B) RT-PCR of gene purified from epithelial cell (lane 1), whole kidney (lane 2) and mesangial cell (lane 3). Thy-1, A specific protein marker for mesangial cells. Nephrin, A specific protein marker for slit diaphragm at foot processes of podocyte. b-Actin, Control marker.
rehydrate and balance it with PBS. To prepare the framework-embedded collagen vitrigels, a nylon membrane cut into a ring shape with an inner-outer diameter of 23–33 mm or a pressed slid sheet cut into a circular shape with a diameter of 33 mm was sterilized with 70% ethanol for 10 min, washed with PBS three times to completely remove the ethanol, and inserted into a culture dish before pouring the collagen sol. The silk sheet was prepared by adding moisture to uniformly spread cocoon filaments, and then pressing them at a high temperature (120°C) and agglutinating the sericin. Figure 4 shows a schematic representation of the protocol for the preparation of collagen vitrigels with and without the framework of a nylon membrane ring or a pressed silk sheet. The collagen sol poured into culture dishes already containing a nylon ring membrane formed an opaque and soft collagen gel in which the nylon ring was embedded (Fig. 4A). The gel was converted into a rigid glass-like material with salt precipitations derived from the culture medium by drying sufficiently (Fig. 4B). The vitrified material was regenerated as a stable thin gel membrane by rehydrating with PBS (Fig. 4C) which we termed vitrigel. The vitrigel, washed and balanced with PBS, is absolutely transparent (Fig. 4E). Vitrigel embedded with the frameworks of a nylon ring (Fig. 4E) and a silk sheet (Fig. 4F) showed selfsupporting membrane architectures while the vitrigel without frameworks appeared as a self-twisting membrane structure (Fig. 4D) when they were held with forceps. III. PHYSICAL PROPERTIES OF COLLAGEN VITRIGEL The physical properties of the collagen vitrigel, including the changes in net weight, mechanical strength, and transparency in the vitrification process were evaluated as follows. The net weight of each collagen gel in the vitrification process was calculated by subtracting the premeasured dish weight from the gross weight of the gel and dish. Weights
were measured using a digital scale (AG245; Mettler-Toledo International, Greifensee, Germany). In the vitrification process, the collagen gel outwardly showed a dried state within the first 2 d. The net weight of the collagen gel was reduced to about 14.5% of the initial net weight after a vitrification period of 2 d and reached a plateau level of about 1.5% of the initial net weight after 7 d (Fig. 5). The mechanical strength of both collagen gels and collagen vitrigels prepared from different vitrification periods was measured with a digital force gauge (FGC-0.2B or FGC-2B; Nidec Shimpo, Kyoto) by modifying a previous method (49). Each nylon ring-embedded collagen gel or collagen vitrigel was carefully detached from the culture dish by forceps and then inserted and fastened between two glass cylinders with inner-outer diameters of 20–40 mm. A flat disk with a diameter of 12 mm connected to the force gauge was automatically approached at a speed of 9 mm/min and vertically inserted into the center region of each collagen gel or collagen vitrigel. The elastic force of the collagen gel or collagen vitrigel right before breaking was measured and recorded as the maximum elastic force (g force) indicated by the force gauge. Figure 6 shows that the mechanical strength of the collagen vitrigels increased as the vitrification period increased, which reached 813+ 41 g force (n = 3) after 84 d, which is about 19.65 times that of conventional collagen gels. The transparency of both collagen gels and collagen vitrigels prepared from different vitrification periods was measured with a spectrophotometer (V-550; JASCO, Tokyo). Each nylon ring-embedded collagen gel or collagen vitrigel was carefully detached from the culture dish by forceps and pasted on a flat quartz glass specific for the spectrophotometer. The absorbance of wavelengths from 200 to 900 nm was monitored and the optical density (OD) curve at 400 nm was plotted. Figure 7 shows that the absorbance at 400 nm of conventional opaque collagen gels was about 0.34 while that of collagen vitrigels after 2 d was about 0.2. The OD curve reached a plateau level of about 0.1, representing a
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FIG. 4. Schematic protocol for the preparation of collagen vitrigels with and without the framework of a nylon membrane ring or a pressed silk sheet. A vitrigel is produced via three processes, gelation, vitrification, and rehydration. Gross aspects of a conventional gel state (A), a vitrified state (B), and a rehydrated state (C) in the preparation process of a nylon ring-embedded collagen vitrigel, and of a collagen vitrigel without frameworks (D), a nylon ring-embedded collagen vitrigel (E), and a silk sheet-embedded collagen vitrigel (F) that are held by forceps. Bar represents 10 mm. Reproduced from Ref. 39 with permission of the publisher.
transparent state in collagen vitrigels with a vitrification period of more than 42 d. The OD from 350 to 650 nm was almost the same as that at 400 nm (data not shown). Morphological observations of both collagen gel and collagen vitrigel with a vitrification period of 72 d were per-
formed using a phase-contrast microscope (TE300; Nikon, Tokyo). Figure 8 shows the morphological results for both gels. Fine filament structures surrounded by many small particles were observed in a traditional collagen gel (Fig. 8A), whereas numerous small particles were observed in the col-
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FIG. 5. Changes in net weight of collagen gels in the vitrification process. The net weight of each collagen gel was calculated by subtracting the dish weight from the gross weight of the gel and dish. The value of the vitrification period at day 0 represents the net weight of the collagen gel prepared by incubating a collagen sol at 37°C for 2 h. Averages and standard deviations from five independent samples are presented. Reproduced from Ref. 39.
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FIG. 8. Phase-contrast microphotographs of a conventional collagen gel (A) and a collagen vitrigel (B). The conventional collagen gel and the collagen vitrigel were prepared by incubating a collagen sol at 37°C for 2 h and by rehydrating the collagen gel to a vitrified state for 72 d, respectively. Bars represent 100 mm. Reproduced from Ref. 39.
FIG. 9. Phase-contrast microphotographs of a pressed silk sheet (A) and a silk sheet-embedded collagen vitrigel (B). The silk sheetembedded collagen vitrigel was prepared from a 14-d vitrification period. Bars represent 100 mm. Reproduced from Ref. 39 with permission of the publisher. Reproduced from Ref. 39.
FIG. 6. Changes in the mechanical strength of collagen vitrigels prepared from different vitrification periods. The mechanical strength was recorded as the maximum elastic force indicated by the force gauge. The value of the vitrification period at day 0 represents the mechanical strength of the collagen gel prepared by incubating a collagen sol at 37°C for 2 h. Averages and standard deviations from three independent samples are presented. Reproduced from Ref. 39.
ment was about 15 mm and the filaments formed networks with pore size axes of shorter than 200 mm (Fig. 9A). The basic architectures of the pressed silk sheet were conserved in a silk sheet-embedded collagen vitrigel, although the density of the filaments increased slightly due to the compressed process of vitrification of the collagen gel (Fig. 9B). The mechanical strengths of the silk sheets immersed in PBS for 10 min and the silk sheet-embedded collagen vitrigels with a vitrification period of 14 d were 24,790 ± 3540 (n = 5) and 21,110 ± 2470 g force (n = 5), respectively. The latter mechanical strength was about 195 times stronger than the collagen vitrigels without the silk sheet prepared with the same vitrification period. IV. COCULTURE OF GLOMERULAR MESANGIAL CELLS AND EPITHELIAL CELLS ON COLLAGEN VITRIGEL AND RECONSTRUCTION OF RENAL GLOMERULAR TISSUE
FIG. 7. Changes in the transparency of collagen vitrigels prepared from different vitrification periods. The transparency was recorded as the absorbance of wavelength 400 nm monitored by a spectrophotometer. The value of the vitrification period at day 0 represents the transparency of the collagen gel prepared by incubating a collagen sol at 37°C for 2 h. Averages and standard deviations from three independent samples are presented. Reproduced from Ref. 39.
lagen vitrigel after a vitrification period of 72 d (Fig. 8B). Examination of a pressed silk sheet using a phase-contrast microscope revealed that the diameter of each cocoon fila-
The collagen vitrigel used in this study was immersed in PBS for 30 min and then placed on the inner ring edge of in vitro fertilization dishes (BD Falcon, Franklin Lakes, NJ, USA) so that cells facing the top and bottom sides of the collagen vitrigel could be immersed separately in different media. Since mesangial cells grow more slowly than epithelial cells, 1´104 mesangial cells were first seeded on one side of the collagen vitrigel. After an overnight culture, most of the mesangial cells adhered to the collagen vitrigel and cell proliferation continued for 1 week. The collagen vitrigel was then turned upside-down, and 1´104 epithelial cells were
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FIG. 10. Microscopic observation of glomerular epithelial cells and mesangial cells cocultured on opposite sides of collagen vitrigel. (A, D) Epithelial cells stained with HE, where cells on panel A reached to confluency; (B, E) mesangial cells stained with HE, where cells on panel E reached to confluency; (C, F) both epithelial and mesangial cells stained with HE. Bars represent 10 mm.
FIG. 12. Phase contrast microphotographs of collagen vitrigel section stained with HE. (A) Both sides of glomerular epithelial and mesangial cells; (B) glomerular epithelial cell side; and (C) mesangial cell side. Bars represent 10 mm.
seeded on the opposite side of the collagen vitrigel. Epithelial cells adhered to the gel immediately and proliferated to confluency in 3 d after seeding. MsBM medium was added to the mesangial cell side and DHSF medium was added to the epithelial cell side. Cells cultured in commercial dishes were used as a control. All of the cell cultures were carried out for 1 month by replacing with fresh media every week. After 1 month of cell culture on the collagen vitrigel, cells that had grown on the gel were examined by phase contrast microscopy. In order to distinguish the morphology and viability between mesangial cells and epithelial cells at opposite sides of the collagen vitrigel, cells were stained with hemotoxylin and eosin (HE) according to the standard procedure. The viability of mesangial cells and epithelial cells cultured on the collagen vitrigel was examined by incubating the cells in a LIVE/DEAD viability/cytotoxicity kit (Molecular Probe, Eugene, OR, USA) at room temperature for 30 min. Viable cells selectively incorporate the nonfluorescent calcein-AM, cleave the AM moiety by endogenous esterase, and yield green fluorescent calcein molecules. Dead cells selectively incorporate ethidium homodimer-1 into the
FIG. 13. Phase contrast microphotographs of glomerular epithelial cells on collagen vitrigel section (A) and on commercial dishes coated with collagen (B) stained with HE. Bars represent 2 mm.
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FIG. 11. Identification of cell viability of glomerular mesangial cells and epithelial cells cocultured on collagen vitrigel. (A) Epithelial cells stained with HE; (B) viable (green) mesangial cells stained with LIVE/DEAD kit; (C) dead (red) mesangial cells stained with LIVE/DEAD kit; (D) epithelial cells stained with HE; (E) viable (green) epithelial cells stained with LIVE/DEAD kit; and (F) dead (red) epithelial cells stained with LIVE/DEAD kit. Bars represent 1 mm.
FIG. 14. Immunofluorescent microphotographs of glomerular mesangial and epithelial cells in single cell culture and coculture system of collagen vitrigel. Monoclonal antibodies of a-actin, fibronectin, and integrin b1 were used to identify the cell–ECM interaction. (A, C) Mesangial cells stained with a-actin in individual culture and coculture, respectively; (E, G) mesangial cells stained with fibronectin in individual culture and coculture, respectively; (I, K) mesangial cells stained with integrin b1 in individual culture and coculture, respectively; (B, D) epithelial cells stained with a-actin in individual culture and coculture, respectively; (F, H) epithelial cells stained with fibronectin in individual culture and coculture, respectively; (J, L) epithelial cells stained with integrin b1 in individual culture and coculture, respectively. Panels C and D are microphotographs from the same location of vitrigel where mesangial and epithelial cells were cultured on the opposite sides and stained by a-actin labeled with rhodamin-phalloidin (red), respectively. The same for panels G and H, and panels K and L, except that each pair was stained by anti-fibronectin-antibody labeled with FITC (green) and anti-integrin b1 antibody labeled with FITC (green), respectively. Bars represent 1 mm.
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nuclei, and yield red fluorescent ethidium molecules in the nuclei. The viable and dead cells were examined using a fluorescent microscope (DM RXA; Leica, Wetzlar, Germany). Figure 10 shows that cells on collagen vitrigel can be observed by microscopy without any difficulty. Micrograph focusing on the glomerular epithelial cells and mesangial cells is shown in Fig. 10A and 10D, and 10B and 10E, respectively. Figures 10C and 10F show that both cells can be observed simultaneously on the collagen vitrigel after HE staining. Cell viability was confirmed by staining with the LIVE/DEAD kit. The results indicated that mesangial cells as well as epithelial cells grew to confluency (Fig. 11A, D), and that the green fluorescence produced by viable cells was as high as 99% (Fig. 11B, E) while the red fluorescence produced by dead cells was only 1% for both mesangial and epithelial cells (Fig. 11C, F). Cross sections of cultured specimens were obtained by cutting them around their center at a thickness of 4 mm using a microtome (RM2145; Leica Microsystems, Wetzlar, Germany) and then staining with hematoxylin and eosin according to the standard procedure for optical observation. Figure 12A shows that both cells adhered to the opposite sides of the collagen vitrigel. The sections of glomerular epithelial cells and mesangial cells were labeled as B and C, respectively. The enlarged areas of sections B and C (Fig. 12B, C) showed mesangial cells had a flat morphology while epithelial cells revealed polarity formation. In order to verify that the polarity was only formed on epithelial cells cultured on vitrigel in the coculture system, glomerular epithelial cells cultured on collagen vitrigel and commercial dishes coated with collagen for 2 weeks were further examined. Cross sections of both cellular samples proved that only epithelial cells cocultured with mesangial cells on the collagen vitrigel showed polarity formation (Fig. 13A), whereas those cultured on commercial dishes had a flat morphology (Fig. 13B).
be enhanced on mesangial (Fig. 14G) and epithelial cells (Fig. 14H) in cocultures as compared to those in individual cultures (Fig. 14E, F). Since mesangial cells are known to be responsible for fibronectin synthesis, we suspect that the synthesized fibronectin permeated to the epithelial cell side from the mesangial cell side through collagen vitrigel and stimulated integrin, the receptor of fibronectin, on the epithelial cell surface. In order to clarify this point, integrin b1 for both cells was detected. As expected, the expression of integrin b1 was enhanced in mesangial cells (Fig. 14K) and epithelial cells (Fig. 14L) compared to that in individual cell cultures (Fig. 14I, J). Although it cannot be ruled out that the serum in the medium contains fibronectin, it is easy to determine that fibronectin was consumed in the long-term cultures, especially in the individual cell cultures. It is likely that the fibronectin detected in the epithelial cells of the coculture system was due to its secretion by mesangial cells. This fact suggests that secreted fibronectin plays a role in inducing the expression of integrin, the ECM receptor, on the surface of epithelial cells, and enhances the adhesion of epithelial cells to collagen vitrigel. The stable adhesion of epithelial cells to a scaffold may contribute to stabilization of the cytoskeletal organization and promote polarity formation. In conclusion, the collagen vitrigel system provides an optimum environment mimicking the glomerular basement membrane to reconstitute renal glomerular tissue when coculturing various glomerular cells on it. Using this system, the polarity of epithelial cells can be re-formed and both epithelial and mesangial cells can maintain their viability and morphology in the long term. This collagen vitrigel system also provides an easy detection system with which to investigate the interactions between various cells and between cells to ECM which may contribute to the clarification of the mechanism of tissue regeneration. ACKNOWLEDGMENTS
V. CELL–ECM AND CELL–CELL INTERACTIONS FOR GLOMERULAR MESANGIAL CELLS AND EPITHELIAL CELLS IN COCULTURE AND INDIVIDUAL CULTURE The cell–ECM and cell–cell interactions were investigated since cocultures of glomerular mesangial cells and epithelial cells maintained stable adhesion to the collagen vitrigel for more than 1 month without impairing the cell viability. Stable cell adhesion is known to be linked to cell adhesion molecules, signal transduction, cell-to-cell contact, and extracellular matrix (50–53). For the interaction of cell–ECM adhesion, the expression of fibronectin, an ECM protein expressed by glomerular mesangial cells (54, 55) and its cellular receptor, integrin b1 (56, 57), were investigated. The results are shown in Fig. 14. The cytoskeletons of mesangial cells and epithelial cells stained with a-actin in individual cell cultures as controls are presented in Fig. 14A and 14B, while those of mesangial and epithelial cells in cocultures are shown in Fig. 14C and 14D. All of the cells showed proper cytoskeleton organization implying that the cells adhered to the collagen vitrigel and proliferation proceeded smoothly. Fibronectin expression was found to
This work was supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan (nos. 15360434 and 16656252) and by the 21st century COE program of the Ministry of Education, Culture, Sports, Science and Technology of Japan to one of the authors (PC.W.). The authors are grateful to Mr. Takuya Notani and Mr. Hideyuki Chiba for their help with the experiments.
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