Engineered 3D Kidney Glomerular Microtissues to Model Podocyte-Centric Diseases for the Validation of New Drug Targets

Abstract

Podocytopathies are a diverse group of kidney diseases characterized by podocyte injury, leading to proteinuria and reduced kidney function. This injury often disrupts cytoskeletal dynamics and cellular adhesion, causing glomerular dysfunction. Current in vitro models fail to accurately mimic the three-dimensional (3D) organization and mechanics of kidney tissue, hindering the understanding of podocyte pathophysiology and therapeutic development. In this study, 3D microtissues are developed that replicate the structure and mechanics of the glomerular filtration barrier, enabling the modeling of chemically and genetically induced podocyte injuries for drug target validation. These microtissues simulate the glomerulus's three-layer structure and hemodynamic mechanical stretch, providing a platform to evaluate relevant mechanobiological signaling pathways and podocyte dynamics. Collective cellular forces are measured to assess podocyte resilience against genetic or chemical injuries. As a proof-of-concept, podocyte injury is modeled through transient receptor potantial canonical 6 (TRPC6) overexpression, a validated target in podocytopathies, and evaluated by the TRPC6 inhibitor SAR7334. The results demonstrated a loss of podocyte contractile forces upon TRPC6 overexpression, with recovery following treatment. This highlights the potential of glomerular microtissues to model podocyte mechano-pathophysiology and serves as a robust platform for screening new therapies.

Keywords: 3D cell models; Glomerular filtration barrier; kidney diseases; podocytopathies; target validation.

Figure 1

Schematic representation of glomerular microtissue development strategy. A) Depiction of the nephron structure, glomerulus, and GFB. The GFB comprises fenestrated GECs, the GBM, and podocytes. Microtissues mimicking the GFB are formed in PDMS devices. B) Methodology for generating microtissues. GECs embedded in collagen‐I allow the initial formation of the GEC microtissues, which is further matured through the incorporation of podocytes, which provide coverage and major force generation. C) Illustration of the tissue generating force and deflecting the pillars. Created in BioRender. Yasa, I. (2025) https://BioRender.com/brlp530.

Figure 2

Engineered glomerular microtissues. A) Brightfield images of microtissues generated by different endothelial cell sources, HUVECs, and GECs. Calcein Green labeled ciPODs were visualized to attach to the surface of microtissues. B) Microtissue force measurement is taken on day 4 of culture, 3 days after ciPODs addition. Glomerular microtissues exhibit significantly higher force compared to endothelial microtissues. Data are reported as mean ± SD, n≥ 9 microtissues and ^** P < 0.01, ^**** P < 0.0001 when compared by one‐way ANOVA with Tukey multiple comparison test. C) Representative two‐dimensional (2D) projected and 3D reconstructed fluorescent confocal images of GEC (i) and glomerular (GEC + ciPOD) microtissues (ii), respectively. GECs were distinguished by their VE‐Cadherin expression and ciPODs by their actin fibers. D) Second harmonic imaging of collagen‐I in both GEC microtissues (i) and glomerular microtissues (ii), showing the organization of the matrix and the barrier between podocytes (yellow arrows) and GECs (red arrows) in the glomerular microtissue (ii).

Figure 3

Expression and synthesis of GBM components within 3D microtissues. A) Analysis of gene expression levels for COL4A1 and LAMA1, indicative of immature GBM formation. B) Profiling of gene expression for COL4A3 and LAMA5, associated with the maturation of GBM. Immunofluorescence assessment reveals robust Col‐IV presence in both C) GEC and D) glomerular microtissues. The podocyte layer with high F‐actin intensity is evident in glomerular microtissues. Data are reported as the mean ± SD. n = 2 independent repeats for GEC microtissues, and n ≥ 3 for other groups; ^* P = 0.005, by one‐way ANOVA.

Figure 4

Cyclic stretching of glomerular microtissues. A‐i) Blood filtration exerts mechanical stress in capillary walls and podocytes are exposed to tangential shear stress and circumferential stress (stretch). ii) Illustration showing the strategy of applying cyclic stretch to glomerular microtissues with a frequency of 0.5 Hz to mimic the mechanical stress which is a critical component of glomeruli physiology. B) Microtissues demonstrated significantly lower forces under mechanical stretch with high linear strain (12%). Data are reported as the mean ± SD. n ≥ 9 microtissues; ^* P < 0.001 when compared to static condition by one‐way ANOVA with Tukey test. C) Confocal imaging revealed the detrimental effect of mechanical stretching with a linear strain of 12 even after 4 h whereas no pronounced effect on cytoskeletal organization and coverage of microtissues was observed when the stretching strain was 2.5%. The scale bar is 500 µm for all images. D) Plot showing the significantly lower coverage after mechanical stretch based on fluorescent image analysis. Data are reported as the mean ± SD. n ≥ 9 microtissues; ^* P = 0.001, ^** P < 0.0001 when compared to static condition by one‐way ANOVA with Tukey test. E) Finite element models assist in understanding the effects of tissue deformation under stretching. Total deformation at the average height of the microtissue when there is no strain, during 2.5% strain stretching and 12% strain stretching. F) Multi‐view representations of the Von Mises equivalent stress contour showing the stress distribution where the red color indicates areas of higher stress. During stretching, stress is predominantly concentrated around the region where the microtissue contacts the inner part of the pillars, suggesting a focal point of mechanical tension.

Figure 5

TNFα driven podocyte injury. A) Overview of the strategy for exposing microtissues to 200 ng ml⁻¹ TNFα for 7 days. B) Plot of static microtissue force measured at day 9 normalized to pre‐TNFα treatment. The fold shows the TNFα driven reduction in the forces. C) Representative immunofluorescence images of nuclei, actin, and vinculin at day 9, with or without TNFα treatment. Induced elongated morphology in podocytes (left panels). Vinculin staining also demonstrated less intensity compared to control microtissues when the microtissues were treated with TNFα. D) Plot of adhesion area (%) calculated based on vinculin staining shows a decrease in the TNFα‐induced damage group. Data reported as mean ± SD. n ≥ 7 microtissues/repeat; repeated at least 3 times. P value calculated by t‐test, ***P < 0.001.

Figure 6

Recapitulation of TRPC6‐driven genetically induced podocyte injury. A‐i) Overview of the strategy and timeline for inducing TRPC6‐driven injury. ii) Confocal images comparing the microtissues formed in the absence or presence of Dox to induce TRPC6 overexpression, which had no effect in the morphology of the microtissues and coverage by podocytes. Dox induction resulted in a strong expression of TRPC6 (red) compared to no induction group as illustrated by representative fluorescent images. iii) Plot showing the distribution of contractile force (normalized to 1‐day post podocyte seeding) and the significant decrease when TRPC6 was induced and overexpressed. P value calculated by two‐tailed t‐test, n≥ 20 microtissues, ^**** P < 0.001. B‐i) Overview of inhibitory compound treatment strategy for preventative and rescue scheme, ii) plot showing the distribution of contractile forces for 1µM and 10 µM SAR7334‐treated and no treatment microtissues. SAR7334 treatment inhibited the force reduction induced by TRPC6 overexpression. P value calculated by one‐way ANOVA, n ≥ 6 microtissues per repeat, ^*** P < 0.001. SAR treatment experiments are repeated 2 times.