Chronic kidney diseases and acute kidney injury are mechanistically distinct kidney diseases. While chronic kidney diseases are associated with podocyte injury, acute kidney injury affects renal tubular epithelial cells. Despite these differences, a cardinal feature of both acute and chronic kidney diseases is dysregulated actin cytoskeleton. We have shown that pharmacological activation of GTPase dynamin ameliorates podocyte injury in murine models of chronic kidney diseases by promoting actin polymerization. Here we establish dynamin’s role in modulating stiffness and polarity of renal tubular epithelial cells by crosslinking actin filaments into branched networks. Activation of dynamin’s crosslinking capability by a small molecule agonist stabilizes the actomyosin cortex of the apical membrane against injury, which in turn preserves renal function in various murine models of acute kidney injury. Notably, a dynamin agonist simultaneously attenuates podocyte and tubular injury in the genetic murine model of Alport syndrome. Our study provides evidence for the feasibility and highlights the benefits of novel holistic nephron-protective therapies.
The leading causes of acute kidney injury (AKI) are ischemia, hypoxia, or nephrotoxicity1. While it can be reversed, AKI represents a significant healthcare problem with high mortality and no definitive treatment. Regardless of its etiology, AKI primarily injures polarized epithelial cells of the renal tubules whose apical microvilli form the tubular brush border that participates in coordinating essential electrolyte and water transport2. An early morphological feature of AKI is loss of the brush border and cell polarity due to the breakdown of the actomyosin cortex at the apical membrane1.
The establishment and maintenance of cell polarity involve signaling cascades, membrane trafficking, and cytoskeletal dynamics, all of which are highly coordinated3. The organization of the apical membrane is largely determined by the architecture of the actomyosin networks4, which establishes cortex stiffness, thus facilitating the clustering of polarity proteins. While myosin II motors are considered the primary generator of cortical stiffness5,6, the architecture of the actomyosin cortex is established by a myriad of actin-binding proteins (ABPs)7.
In addition to known ABPs, the brush border of renal tubules is highly enriched in dynamin8, a GTPase best known for its role in endocytosis9. Dynamin has an intrinsic propensity to assemble into multiple oligomerization states such as dimers, tetramers, rings, and spirals9. We identified for the first time direct dynamin–actin interactions10 and showed that dynamin’s oligomerization regulates actin polymerization in podocytes11,12, specialized cells essential for the selectivity of the kidney filter. Using murine models of CKD, we have shown that activation of dynamin-dependent actin polymerization reverses podocyte injury by restoring their unique structure and function12.
Here we show that in renal tubular epithelial cells dynamin cross-links filamentous actin (F-actin) into branched networks. Dynamin’s cross-linking capability is defined by its oligomerization state and the length of F-actin. Pharmacological activation of dynamin oligomerization counteracts AKI by stabilizing the actin networks and thus cell integrity, which partially protects renal epithelial cells from oxidative stress-induced injury. Our study identifies the actomyosin cortex of the apical membrane of the renal tubular cell as a druggable target in AKI via dynamin as a proxy.
Dynamin oligomerization establishes the stiffness and morphology of the apical membrane
To examine the role of dynamin–actin interactions in polarized renal tubular epithelial cells, we utilized Bis-T-23, an allosteric activator of actin-dependent dynamin oligomerization in a reconstituted system13, in the cells11,13, and in the whole organism12. Cellular phenotypes were assessed in Madin-Darby Canine Kidney (MDCK) cells by following the status of the F-actin and the staining pattern of a tight junction protein zonula occludens-1 (ZO-1), which is considered a biomarker of cell polarity. Cytochalasin D (CytoD) and latrunculin A (LatA), known inhibitors of actin polymerization, decreased F-actin levels, and induced discontinuous ZO-1 staining (Supplementary Fig. 1a). In contrast, Bis-T-23 induced a slight increase in F-actin levels without any effect on ZO-1 staining. Addition of Bis-T-23 prior to but not after LatA, partially preserved F-actin levels and cell polarity. Neither DMSO vehicle nor dynamin inhibitor dynole14 exhibited any effect (Supplementary Fig. 1a).
Scanning electron microscopy (SEM) allowed us to visualize drug-induced alterations of cell morphology focusing on the apical membrane (Fig. 1a). The average MDCK cell height was 11 ± 2 µm, and the average length of the microvilli was 0.63 ± 0.2 µm (Table 1), which is within the range observed in the kidney15. LatA decreased cell height, microvilli length and shifted the uniformly distributed microvilli into clusters, whereas Bis-T-23 induced the opposite effects (Table 1, Fig. 1a). When added prior to LatA, Bis-T-23 partially preserved cell height and microvilli length. Since microvilli exhibit exquisite length control defined by the cortical actin at their base16, these data provide evidence that Bis-T-23 modified the actomyosin cortex at the apical membrane.
To determine the exact effect that Bis-T-23 had on the cortical actin, we visualized the actomyosin cortex within the lamellipodia using platinum replica electron microscopy (PR-EM). LatA decreased the density of the actin networks, and this effect was partially abrogated by the addition of Bis-T-23 prior to LatA (Fig. 1b). As LatA accelerates actin filament depolymerization by sequestering actin monomers, we next examined whether the observed preservation of actomyosin cortex by Bis-T-23 was due to its positive effect on actin polymerization. In contrast to the potent stimulation of actin polymerization observed in podocyte cell extracts10,11, Bis-T-23 only marginally increased actin polymerization in MDCK cell extract (Supplementary Fig. 1b). Similarly, immunodepletion of endogenous dynamin-2 (Dyn2) from the extract or inhibition of its GTPase activity by dynole resulted in marginal impairment of actin polymerization (Supplementary Fig. 1c). While LatA and CytoD significantly impaired actin polymerization, the addition of Bis-T-23 prior to LatA or CytoD was not able to overcome their inhibitory effects (Supplementary Fig. 1d, e). Jasplakinolide, a drug that induces actin polymerization by stimulating actin filament nucleation17, did not significantly increase the overall level of polymerization (Supplementary Fig. 1d), suggesting that MDCK cell lysate exhibit a near maximal level of polymerized actin. Together, these data indicated that the effects of Bis-T-23 on the morphology of the apical membrane in MDCK cells were driven by a mechanism other than actin polymerization.
Given the common knowledge of enriched localization of Dyn2 and F-actin at the brush border of renal epithelial cells8, and dynamin’s role in endocytosis, we next investigated if Bis-T-23 was affecting actin indirectly via alterations in endocytosis. As expected, both Dyn2 and F-actin co-localized at the actomyosin cortex underneath the apical membrane, within the microvilli, and at clathrin-coated pits (CCPs), defined by their distinct shape and size (Supplementary Fig. 1f). We examined the dynamics of CCPs using total internal reflection fluorescence (TIRF) microscopy18,19. Bis-T-23, even at its highest concentration, had no effect on the distribution of the CCPs’ lifetimes, whereas dynole decreased the number of productive CCPs (Supplementary Fig. 1g). This lack of correlation between the level of endocytosis and alterations in cell morphology asserts that Bis-T-23 targets the cortical actin without influencing dynamin’s role in endocytosis.
Since renal cell polarity is maintained by the architecture and sustained contraction of the actomyosin networks, which establishes cell stiffness at the apical membrane20, we next measured cell stiffness using atomic force microscopy (AFM). Nanowizard IV system and JPK analysis software were used to determine changes in Young’s Modulus21 under different experimental settings (Supplementary Fig. 2a). Treatment with LatA resulted in a significant decrease in cell–cell contact stiffness and apical cell stiffness in MDCK cells (Fig. 1c–e). In contrast, Bis-T-23 significantly increased cell stiffness compared to the DMSO vehicle (Fig. 1c–e), consistent with its positive effects on cell height, microvilli number, and the density of actin networks (Table 1 and Fig. 1b)22. The addition of Bis-T-23 prior to LatA strongly reduced the negative effect of LatA on cell stiffness (Fig. 1c–e), in accordance with Bis-T-23’s positive effect on actin networks and apical cell morphology (Table 1, Fig. 1b).
We have also determined cell stiffness using the BioScope II system as an alternative experimental approach for AFM. In this instance, force indentation curves were obtained following the model of Discher and coworkers computed with Matlab software23. Similar trends with regard to cell stiffness were recorded for the interplay between LatA and Bis-T-23 (Supplementary Fig. 2b, 2c). In addition, dynole did not exhibit any effect on cell stiffness, whereas CytoD significantly decreased cell stiffness (Supplementary Fig. 2d, e), in accordance with their actin phenotypes (Supplementary Fig. 1a). Together, these data establish the correlation between the status of the actomyosin cortex, cell stiffness, the morphology of the apical membrane, and cell polarity. These findings also convincingly demonstrate the role of dynamin oligomerization in defining mechanical parameters of epithelial cell polarity via its effect on the actomyosin cortex.
Dynamin cross-links actin filaments into branched networks that underlie cell polarity
In order to elucidate the molecular mechanism by which dynamin oligomerization influences the architecture of the actomyosin cortex, we next examined the effect of dynamin on actin filaments in a reconstituted system. Based on the current hypothesis, the length of actin filaments defines their mode of cross-linking6. As the average length of cortical actin filaments within a network at the leading edge is between 100 and 150 nm24, we examined the effects of dynamin on the organization of shorter filaments generated by capping F-actin with gelsolin (Gsn-actin) (Fig. 2a). The addition of Dyn2 resulted in the formation of large, branched networks (Fig. 2b, c). Based on the sizes and shapes of recombinant Dyn2 (Fig. 2d), the networks were formed predominantly by Dyn2 dimers (Dyn2DIMER) and tetramers (Dyn2TETRA) that interacted with several actin filaments (Fig. 2e): Dyn2DIMER bound up to two filaments, Dyn2TETRA bound up to four filaments, and Dyn2RING bound up to six filaments. Low magnification of the images revealed that dynamin-dependent networks form a pattern of smaller and larger ring-like shapes (Fig. 2c).
To correlate observations from the reconstituted system with dynamin’s role in cells, we next determined the localization of endogenous Dyn2 on cortical actin networks using a monoclonal anti-Dyn2 antibody followed by a gold-conjugated secondary antibody (Supplementary Fig. 3a–c). As seen in the reconstituted system, dynamin associated with a distinct number of F-actin within branched networks (Fig. 2f). Together, these data identify a novel activity of dynamin, that is cross-linking F-actin into branched networks.
To correlate dynamin’s cross-linking capability and the protective effect of Bis-T-23 on the actomyosin cortex and the morphology of the apical membrane, we next examined the effects of Bis-T-23 on dynamin-mediated networks in reconstituted systems (Fig. 3a). Based on contour plots, which provide topographical representations of varying filament densities, Bis-T-23 increased overall network density (Fig. 3a), which could be explained by the increase in the number of F-actin bound to dynamin due to the increase in its oligomerization. In addition, dynamin more potently cross-linked shorter filaments than long F-actin (Fig. 3a–c), suggesting that dynamin’s capability to form branched networks is defined by its oligomerization status and the length of actin filaments. The ability to cross-link actin filaments into networks was shared by two dynamin isoforms, ubiquitously expressed Dyn2 and neuron-specific dynamin-1 (Dyn1) (Fig. 3b, c).
We and others have shown that dynamin binds F-actin via two distinct domains: one situated in the Middle domain10,12,25,26,27, and one part of the C-terminal proline-arginine rich domain (PRD)28. Similar networks were formed by Dyn1 and Dyn1∆PRD (Supplementary Fig. 3d), demonstrating that PRD is dispensable for network formation. Finally, Dyn2 also cross-linked F-actin into tight bundles and hyper-bundles (Supplementary Fig. 4a, 4b), as seen before10,28. In contrast to network formation, dynamin’s ability to form bundles is dependent on PRD-actin interactions28. Bis-T-23 promoted the formation of hyper-bundles (Supplementary Fig. 4c, 4d), suggesting a mechanism by which Bis-T-23 increased the number of microvilli on the apical membrane (Table 1). This hypothesis is supported by the presence of anti-dynamin antibodies within actin bundles that underlie filopodia (Supplementary Fig. 4e). Together, these data show that dynamin’s ability to cross-link filaments into networks and bundles is influenced by its propensity to oligomerize as well as the length of actin filaments (Supplementary Fig. 4f).
Insights generated by the reconstituted system were tested in the cell-based assays. Expression of both dynamin isoforms lacking PRD (Dyn∆PRD) in MDCK cells did not affect the ZO-1 staining pattern when compared to wild-type dynamins, and Dyn2∆PRD also increased F-actin levels (Supplementary Fig. 5a). These data suggested that dynamin’s ability to form tight bundles does not contribute to the overall level of F-actin or the cell polarity in MDCK cells. In contrast, expression of Dyn2K/E, mutant impaired in F-actin binding10, decreased F-actin levels and cell polarity compared to Dyn2 expressing cells (Supplementary Fig. 5a–c), implicating the Middle domain in the formation of actomyosin networks that underlie cell polarity. This hypothesis is further supported by AFM experiments showing that Dyn2K/E also decreased cell stiffness at the apical membrane when compared to Dyn1WT (Supplementary Fig. 5d).
These studies are complementary to experiments using mouse inner medullary collecting duct (mIMCD) cells, another cell line representing polarized renal epithelial cells. In Dyn2 knockdown mIMCD cells, a significant decrease in F-actin levels and a patchy ZO-1-staining pattern were observed (Supplementary Fig. 6a–c). The F-actin phenotype was rescued to normal level by the expression of either wild type or Dyn2∆PRD, but not by Bis-T-23 (Supplementary Fig. 6d–g). Together these data provide evidence for the essential role of dynamin and its oligomerization in the establishment and maintenance of epithelial cell polarity. They also strongly suggest that actin networks that underlie the apical membrane are formed by interactions between F-actin and the Middle domain, and that dynamin’s ability to form bundles and hyper-bundles is not essential for cell polarity.
Pharmacological stabilization of cortical actin via dynamin ameliorates AKI
Loss of renal tubules brush border due to disassembly of actomyosin cortex is a predominant feature of AKI2,29. A widely used anti-cancer drug often leading to AKI is cisplatin30. A pivotal consequence of cisplatin injury in renal cells is an increase in reactive oxygen species (ROS) level31. Moreover, increased levels of ROS lead to a decrease in actin dynamics32, which causes disruption of the mitochondrial membrane; this further exacerbates ROS production33,34. Thus, we examined whether pharmacological stimulation of dynamin’s cross-linking capability is sufficient to counteract cisplatin-induced injury.
As seen before35, cisplatin decreased MDCK cell height, microvilli number, and length, and affected cell polarity without affecting endocytosis (Table 1, Supplementary Fig. 7a–d). TEM analysis suggested that these phenotypes could be attributed to cisplatin-induced loss of F-actin (Fig. 4a). The addition of Bis-T-23 prior to cisplatin abrogated its effects by stabilizing the actomyosin cortex against cisplatin-induced injury (Fig. 4a–c). Similar phenotypes were observed in porcine kidney proximal tubule (LLC-PK1) cells (Supplementary Fig. 8). Together, these data provide evidence that Bis-T-23 preserves the actomyosin cortex from cisplatin-induced disassembly.
In a complementary physiologically relevant approach36, Bis-T-23 counteracted cisplatin’s ability to reduce transepithelial electrical resistance (TER) as measured across the live epithelial monolayer using a transwell culture system (Fig. 4d). Since TER measures the integrity of tight junctions in cell culture, these data further establish the beneficial effect of Bis-T-23 on epithelial cell integrity upon injury.
As altered actin dynamics can further enhance ROS production, we next examined whether stabilization of the actin network by Bis-T-23 can counteract oxidative stress in cisplatin-injured cells. We elected to assess the status of cellular biomolecule carbonyls, a stable biomarker of oxidative damage produced by high levels of ROS, using our recently developed fluorescent sensor TFCH37 (Fig. 4e). Indeed, the addition of Bis-T-23 prior to cisplatin partially diminished cisplatin-induced increase in carbonylation (Fig. 4f). Similar trends regarding carbonylation were observed after treatment with Bis-T-23 and the actin depolymerizer LatA (Supplementary Fig. 9a). Together these data provide evidence that stabilization of the actin networks can counteract the feedback loop between oxidative stress and the status of actin in injured cells.
An ex vivo rat kidney model was next utilized to visualize Bis-T-23-driven preservation of the actomyosin cortex at the brush border. Cisplatin induced a significant loss of F-actin staining at the brush border of proximal tubules (Fig. 4g). A similar loss of F-actin at the brush border upon the onset of severe ischemia-reperfusion injury was recently observed using intraviral imaging38. Pre-treatment with Bis-T-23 partially protected cisplatin-induced loss of F-actin at the brush border of renal tubules (Fig. 4g), further demonstrating the effect of Bis-T-23 on actomyosin cortex at the apical membrane of renal epithelial cells. As seen in the cell culture, Bis-T-23 did not affect endocytosis in the tissue slices (Supplementary Fig. 9b), further demonstrating that Bis-T-23 does not affect dynamin’s role in endocytosis in renal proximal tubules.
Finally, we examined the reno-protective effect of Bis-T-23 in a mouse model of cisplatin-induced AKI39. BL6 mice were injected with either Bis-T-23 or DMSO once per day starting 24 h prior to injection of cisplatin. As seen before40, serum creatinine (SCr) and blood urine nitrogen (BUN) levels significantly increased 3 days post cisplatin injection (Fig. 4h). DMSO exhibited mild reno-protection as seen before41, due to its oxidant scavenging ability42. Daily administration of Bis-T-23 outperformed DMSO with regard to reno-protection (Fig. 4h). As all animals were sacrificed on day 5 due to the systemic toxicity of cisplatin (Supplementary Fig. 9c), we could not examine the long-term benefits of Bis-T-23 on kidney function. Together, these studies persuasively demonstrate that preserving renal cell integrity via stabilization of the actomyosin cortex at the apical membrane counteracts cisplatin-induced nephrotoxicity.
Stabilization of the actin networks counteracts iohexol-induced AKI
While the exact mechanism of contrast dye-induced AKI is not fully deciphered, it remains a prime cause of hospital-acquired AKI43. Physiologically, contrast dye increases the osmotic load, decreases renal blood flow, and induces renal arterial constriction44. At the cellular level, ROS is a critical player in contrast to dye-induced AKI45. Given the role of ROS in regulating the actin cytoskeleton and our hypothesis that stabilization of the actomyosin cortex can counteract oxidative damage, we examined the effects of Bis-T-23 on a contrast dye-, iohexol-induced AKI46.
The cellular effects of iohexol were examined in human kidney proximal tubular (HK-2) cells. As seen for cisplatin, iohexol induced loss of F-actin, which was accompanied by a significant increase in the level of carbonylated molecules, an irreversible consequence of elevated ROS levels (Fig. 5a, b). Both phenotypes were partially counteracted by pre-treatment with Bis-T-23, further establishing the synergy between the status of the actin cytoskeleton and the level of oxidative stress in these cells. Complementary to cell-based assays, daily administration of Bis-T-23 preserved renal function based on the Scr and BUN levels in BL6 mice challenged with contrast dye (Fig. 5c). A much-coveted response of improved survival rate was also observed in the animals that received Bis-T-23 along with the contrast dye (Fig. 5d).
We have recently shown that elevated serum levels of soluble urokinase-type plasminogen activator receptor (suPAR) sensitize the kidney to iohexol-induced AKI47. We next investigated if dynamin-mediated protection of the tubular cells can also render reno-protection in BL6 mice expressing high levels of suPAR from fat tissue (suPAR-Tg)48. As expected47, the suPAR-Tg mice exhibited a decrease in kidney function 24–48 h post iohexol injection (Fig. 5e). In contrast, the animals that received a daily dose of Bis-T-23 exhibited significantly better kidney function despite the contrast dye challenge (Fig. 5e). Notably, a significant decrease in the mortality rate was observed in the mice cohort that received Bis-T-23 (Fig. 5f). As anticipated, the extent of oxidative damage (biomolecule carbonylation) in HK-2 cells concomitantly treated with Bis-T-23 was significantly lower than in cells that received only a combination of suPAR and Iohexol (Supplementary Fig. 9d).
In contrast to ROS-mediated cellular injury caused by cisplatin and iohexol, suPAR promotes tubular cell injury partly by increasing oxygen consumption rate (OCR)47. To link the physiological effect of Bis-T-23 in suPAR transgenic mice to its effect on cell metabolism, we examined whether stabilization of the actin network via dynamin decreases suPAR-driven OCR. OCR was measured in real-time in HK-2 cells under basal conditions and in response to sequential injections of mitochondrial inhibitors using a Seahorse XFe24 extracellular flux analyzer (Fig. 5g). The addition of an anti-suPAR antibody or Bis-T-23 ameliorated suPAR-driven increases in mitochondrial basal respiration, ATP production, maximum rate of respiration, and spare-respiratory capacity (Fig. 5f). These data demonstrate the positive effects of Bis-T-23 on cell metabolism upon injury. Together, these studies further affirm the feasibility of protecting against multiple types of AKI and thereby improving the survival rate in rodent models via dynamin as a proxy.
Simultaneous stabilization of actin in distinct kidney cells attenuates nephron injury
We have previously shown that pharmacological activation of dynamin-driven actin polymerization restored the structure and function of podocytes in diverse murine models of CKD12. In this study, we demonstrated dynamin’s ability to preserve the integrity of tubular cells upon acute injury by cross-linking the actin cytoskeleton. Together, these insights led us to envision the possibility of simultaneously counteracting both, glomerular and tubular injury, by pharmacologically targeting dynamin.
To test this hypothesis, we examined whether Bis-T-23 could delay the loss of kidney function in the mouse model of Alport syndrome (AS). AS is an inherited form of progressive kidney failure due to mutations in the COL4A3, COL4A4, or COL4A5 genes that together encode type IV collagen, a major component of the glomerular basement membrane (GBM)49,50. Although the primary defect in AS is foot process effacement and proteinuria due to alteration in the composition of the GBM, the tubular response is also critical to the pathogenesis of AS and may precede the deterioration of glomerular function51,52,53. Indeed, signaling between tubules and glomerulus influences the response of the podocytes to collagen deficiency54. Currently, there is no specific therapy for AS.
Daily administration of Bis-T-23 slowed down the increase in proteinuria in Col4a3−/− animals55 (Fig. 6a), demonstrating that dynamin-mediated increase in actin polymerization in podocytes can counteract even genetic defects in GBM. Bis-T-23 also slowed down the decrease in kidney function, indicated by a small increase in BUN levels, protected tubules from developing hyaline casts, and extended the lifespan of animals (Fig. 6a–c). These data further attest to the positive impact of increased dynamin cross-linking activity on the preservation of renal tubular cell integrity. Together, our study demonstrates the suitability and the unique advantage of pharmacological targeting of nephron as a unit via dynamin as a proxy (Fig. 6d).
Since the identification of direct dynamin–actin interactions10, a growing body of evidence is establishing this GTPase as one of the major regulators of the actin cytoskeleton in the cell. In contrast to canonical ABPs, the mechanisms by which dynamin influences actin are both, highly versatile and cell-type specific. This is due to the combination of dynamin’s multiple oligomerization states and its ability to bind F-actin by two different binding sites. F-actin interactions with dynamin rings and helices via its C-terminal PRD result in actin bundles and hyper-bundles, which have been implicated in the formation of filopodia56 and membrane protrusions during myoblast fusion28. Interactions between dynamin’s Middle domain and F-actin have been implicated in regulating actin networks in lamellipodia26, postsynaptic cytoskeleton organization, and neuromuscular junction development27, actin bundle rigidity in invadosomes during myoblast fusion25, and formation of podocyte foot processes12. While the formation of foot processes is driven by dynamin-stimulated actin polymerization, this study suggests that the molecular mechanism by which dynamin influences these other actin-driven processes might be in part due to its ability to cross-link F-actin into branched networks. Our current study expands the role of dynamin-dependent network formation to include formation of actomyosin cortex at the apical membrane of polarized epithelial cells and the establishment of cell stiffness.
Despite the mechanistic diversity, dynamin’s capability for cross-linking F-actin or promoting actin polymerization is enhanced by the increase in its oligomerization state (this study and refs. 12,28). Dynamin oligomerization is cooperative and is regulated by its concentration57, the length of actin filaments (this study and ref. 13), and SH3-domain-containing proteins25,58. The combination of all these mechanisms ultimately defines the temporal, spatial, and cell-type specificity of dynamin’s role in modifying and/or establishing diverse actin structures.
Changes in mitochondrial function and cell metabolism have been linked to a multitude of AKI etiologies59. Mitochondria are dynamic organelles that respond to physiological signals and are significant sources of ROS in healthy60 and injured cells61. Cisplatin and iohexol directly damage mitochondria leading to increased production of ROS37,43,62,63,64, while suPAR affects the bioenergetic parameters of renal cells47. As many cytoskeletal proteins are sensitive to ROS32,65, a decrease in actin dynamics in the presence of elevated levels of ROS has been reported32. Meanwhile, the dysregulated actin cytoskeleton further augments ROS production33, suggesting a feedback loop between oxidative stress and the actin cytoskeleton. Here we show, by using a combination of cell-based assays focusing on cell polarity, real-time extracellular flux experiments, and our new carbonylation-specific fluorophore, that stabilization of actomyosin cortex via dynamin partially attenuates the feedback loop between oxidative stress and actin cytoskeleton dynamics regardless of the initial type of injury (cisplatin, LatA, iohexol, or suPAR).
It is well established that CKD and AKI, though mechanistically distinct, are closely interconnected, with AKI being recognized as a risk factor for CKD development and progression66. As of now, AKI remains undruggable67. This is particularly relevant in light of the current coronavirus disease 2019 (COVID-19) pandemic, where hospitalized patients with COVID-19 and an elevated suPAR plasma level develop AKI at alarming rates68. Elevated suPAR levels have been linked to both AKI47 and CKD69, further reinforcing the molecular link between these two distinct kidney diseases. Therefore, it is becoming increasingly essential to develop therapeutics that can protect against an array of renal insults on multiple cell types70. Here we report the reno-protective effect of a dynamin agonist in a genetic model of AS, which exhibits injury to both podocytes and tubular cells. Our study establishes a comprehensive approach for developing novel therapeutics that engage with the actin network in multiple cell types within the nephron and treat myriad kidney diseases regardless of the site of injury.
MDCK (Madin-Darby canine kidney) cells (ATCC CCL-34) and mouse inner medullar collecting duct (mIMCD-3) cells (ATCC CRL-2123) were grown in DMEM/F12 (ThermoFisher Scientific) containing 10% fetal bovine serum (FBS) and 1X Antibiotic-Antimycotic (penicillin, streptomycin, and Amphtericin B; Gibco). LLC-PK1 (Lilly Laboratories Cell-Porcine Kidney 1) cells (ATCC CL-101) were grown in DMEM with 4.5 g/L glucose, L-glutamine, sodium pyruvate (Corning), 10% FBS, and 1X Antibiotic-Antimycotic. MDCK and LLC-PK1 cells were grown for ~24 h and ~72 h, respectively, before initiating experiments. HK-2 (Human Kidney 2) proximal tubular cells (ATCC CRL-2190) were cultured in DMEM/Ham’s F12 medium (Corning) containing 10% FBS, 100 U/ml penicillin, and 100 μg/ml streptomycin when used for bioenergetic experiments. For analyzying the status of actin cytoskeleton and cellular carbonylation, HK-2 cells were grown in DMEM F-12 (ATCC) containing 10% FBS, 1X Antibiotic-Antimycotic, and 1X ITS (insulin, transferrin, and sodium selenite) liquid media supplement (Sigma-Aldrich). To establish dynamin 2 (Dyn2) knock-down cell line (mIMCD-Dyn2KD), cells were infected with lentiviruses carrying shRNA encoding DNM2 gene (5′CGGCCTAGTGGACATGACAATGAACTCGAGTTCATTGTCATGTCCACTAGGTTTTTG3′)10 or scrambled shRNA (Sigma-Aldrich) in complete media containing polybrene (8 µg/ml). After 24 h, the media was replaced every other day with media containing puromycin to select for stable transfectants. The extent of Dyn2 knock-down was assessed by western blot. All cell lines were grown on collagen-coated glass coverslips.
Mouse monoclonal anti-Dyn1/2 (Hudy 1) (Sigma-Aldrich MABT188); mouse monoclonal anti-dynamin 2 (Sigma-Aldrich SAB4200661); rabbit polyclonal anti-Dyn2 (ThermoFisher Scientific PA5-19800); rabbit polyclonal anti-ZO-1 (ThermoFisher Scientific 61-7300); mouse monoclonal anti-actin antibodies (Sigma-Aldrich A4700); human anti-uPAR antibody (R&D Systems MAB807).
Rabbit anti-mouse IgG1, HRP (ThermoFisher Scientific PA1-86329); chicken anti-human actin (beta) antibody (USBiological A0760-43); goat anti-Mouse IgG, HRP (ThermoFisher Scientific A10551); donkey anti-rabbit gold-conjugated (18 nm) antibody (Jackson ImmunoResearch Laboratories 711-215-152); goat anti-mouse IgG gold-conjugated (15 nm) (Ted Pella #15752-EM.GMHL15); rabbit anti-chicken IgG gold-conjugated (10 nm) (Ted Pella #15971-EM.RCHL10); Goat anti-rabbit IgG (H + L) cross-adsorbed secondary antibody, Alexa Fluor 488 (ThermoFisher Scientific# A-11008); Goat anti-mouse IgG (H + L) cross-absorbed secondary antibody, Alexa Fluor 488 (ThermoFisher Scientific# A-11001); Goat anti-mouse IgG (H + L) cross-absorbed secondary antibody, Alexa Fluor 532 (ThermoFisher Scientific# A-11002).
Human Dyn1, rat Dyn2, human Dyn1∆PRD, and rat Dyn2∆PRD were expressed using Bac-to-Bac baculovirus expression systems (ThermoFisher Scientific) in Sf21 insect cells10. Recombinant human gelsolin (Gsn) (gift from Fumihiko Nakamura, Brigham and Women’s Hospital); purified non-labeled G-actin, pyrene labeled G-actin (Cytoskeleton); human uPAR protein (R&D Systems); Rhodamine phalloidin (ThermoFisher Scientific).
Drugs, small molecules, and other reagents
Bis-T-23 (Ryngo 1-23), Dynole 34-2 (Abcam); Latrunculin A (LatA), cytochalasin D (CytoD), jasplakinolide (Jasp), rhodamine-phalloidin, rhodamine-transferrin, rhodamine-albumin, Hoechst 33342, ProLongTM Gold antifade reagent (ThermoFisher Scientific). Glutathione Sepharose 4B (GE Healthcare); protein G PLUS-agarose beads (Santa Cruz Biotechnology); TFCH (Bane laboratory, Binghamton University, SUNY)37; cisplatin, NG-nitro-L-arginine methyl ester (L-NAME), indomethacin (Sigma-Aldrich); iohexol (Omnipaque, 350 mg iodine/ml, TCI America). For cell-based assays, cisplatin and iohexol were dissolved in media before use; other small molecule solutions were prepared in 100% DMSO and frozen until use.
Immunocytochemistry for ZO-1 and F-actin
Bis-T-23 and cisplatin
MDCK cells were treated with DMSO (0.1%) or Bis-T-23 (5 µM; 0.1% DMSO) for 1 h before adding cisplatin (35 µM) for 23 h. LLC-PK1 cells were treated similarly with 10 µM cisplatin and Bis-T-23.
Bis-T-23 and LatA
MDCK cells were incubated in serum-free media (SFM) for 1 h first. Subsequently, in one set, cells were treated with DMSO (0.1%) or Bis-T-23 (30 µM; 0.1% DMSO) for 10 min before adding LatA (0.2 µM; 0.1% DMSO) for 20 min. In another set, cells were treated with identical concentrations of DMSO or LatA for 10 min before adding Bis-T-23 for 10 min. Cells treated with identical concentrations of only DMSO, Bis-T-23, or LatA were used as controls.
Bis-T-23 and iohexol
HK-2 cells were treated with media, DMSO (0.1%), or Bis-T-23 (10 µM; 0.1% DMSO) for 1 h. The media was replaced with only media or media containing identical concentrations of iohexol (250 mg/mL)+DMSO or iohexol+Bis-T-23 for 3 h.
Dynamin mutants overexpression or knockdown
Multiple dynamin mutants were adenovirally expressed in MDCK, mIMCD, and mIMCD-Dyn2KD cells10. Where indicated, cells were treated with 0.1% DMSO or 30 µM Bis-T-23. At the end of all treatments, cells were rinsed with 10 mM sodium phosphate buffer containing 0.9% NaCl, pH 7.4 (PBS), or Dulbecco’s phosphate buffered saline (DPBS, Sigma-Aldrich), fixed with 4% paraformaldehyde (PFA), rinsed with PBS or DPBS again, stained with Rhodamine Phalloidin (1:250) and/or ZO-1 antibody (1:70), and mounted with ProLongTM Gold antifade reagent.
Post drug treatment, media was replaced with SFM containing rhodamine transferrin (50 µg/ml) for 10 min. The cells were then processed for imaging.
Oxidative stress-induced carbonylation assessment using TFCH assay
Bis-T-23 and LatA
MDCK cells were treated with DMSO (0.1%, vehicle for Bis-T-23 or LatA) or Bis-T-23 (10 µM) for 20 min before adding DMSO or LatA (0.2 µM) for 10 min. TFCH (20 µM, 0.5% DMSO) was added for 30 min, and samples were processed for imaging37.
Bis-T-23 and cisplatin
MDCK cells were treated with DMSO (0.1%) or Bis-T-23 (5 µM, 0.1% DMSO) for 1 h before adding media or cisplatin (5 µM) for 22.5 h. TFCH was added for 30 min, and samples were processed for imaging37.
Bis-T-23 and iohexol
HK-2 cells were incubated with only media or media containing DMSO (0.1%) or Bis-T-23 (10 µM, 0.1% DMSO) for 1 h. The media was replaced with only media or media containing identical concentrations of iohexol (100 mg/mL)+DMSO or iohexol+Bis-T-23. After 3 h, the cells were washed with fresh media for 2–3 min, exposed to media containing TFCH, and processed for imaging.
Bis-T-23, suPAR, and iohexol
HK-2 cells were incubated with media containing vehicle (0.2% PBS + 0.1% DMSO), suPAR (10 ng/mL)+vehicle or suPAR+Bis-T-23 (10 µM)+vehicle for 21 h. Subsequently, Iohexol (100 mg/mL) was added to a subset of samples for an additional 3 h before washing and performing the TFCH assay.
MDCK, mIMCD, mIMCD Dyn2KD, HK-2 and LLC-PKI cell imaging, and quantification
All images were captured using a LSM 5 PASCAL or a 600 LSM (Zeiss). For image quantification, all images within an experimental set were acquired using identical optical parameters, thresholded, and analyzed using Fiji (Image J) or Image J (NIH) software. To measure the continuity of tight junctions, a free line was drawn along ZO-1 staining. The graph represents a sum of the length of ZO-1 staining as a percentage of the total length of the cell membrane. A defined region of interest (ROI) drawn with the rectangle tool on the ZO-1-associated cell membrane was quantified to assess ZO-1 intensity. For MDCK cells, the intensity of the F-actin within each cell was quantified. For mIMCD or mIMCD-Dyn2KD cells, the intensity of F-actin in a fixed ROI within each cell was quantified. Cellular carbonylation level was determined by quantifying TFCH fluorescence per cell37. In the scatter dot plots for TFCH assays, each dot represents the average intensity of 1–14 cells. The representative images showing cellular carnbonylation levels were processed identically within a set and assigned a pseudocolor for visual clarity. The staining intensity of treated samples was normalized to the non-treated or vehicle control. An unpaired two-tailed t-test or one-way ANOVA with Tukey’s multiple comparisons test was performed using Prism (GraphPad). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001; whereas P > 0.05 was considered not significant.
Total internal reflection fluorescence microscopy (TIRFM)
Actin polymerization assay using cell extracts
The cytosolic protein extract (100 μg) was treated with dynamin modulators: Bis-T-23 (10 µM), Dynole (10 µM), or DMSO (0.5–1%), for 10 min, or actin modulators: LatA (0.2 µM), CytoD (0.25 µM), or Jasp (1 μM) for 20 m. Actin polymerization was initiated by adding actin polymerization buffer (10 mM Tris-HCl, pH. 7.4, 100 mM KCl, 2 mM MgCl2, 0.5 mM ATP, 0.2 mM CaCl2) and pyrene G-actin (20%, 1 μM). Changes in pyrene actin fluorescence were monitored (excitation: 365 nm; emission: 407 nm) using a cytation5 imaging reader (BioTek).
Immunodepletion of endogenous Dyn2 from MDCK cells
Cytosolic extracts were incubated with anti-Dyn2 antibodies (1 μg/200 μg cytosolic proteins) at 4 ˚C for 2 h, followed by incubation with protein G PLUS-agarose beads at 4 ˚C for 2 h. After centrifugation, the supernatant was used as the ∆Dyn cytosol for the actin polymerization assay. Mock-depletion was performed in the same manner with a non-specific mouse anti-rabbit IgG antibody (1 μg/200 μg cytosolic proteins). The efficacy of the depletion was confirmed to be >90% by Western blot using the same antibody.
Cell polarity assessment using a transwell system
MDCK cells were grown in collagen-coated transwells (Corning) for 48 h before treatment with DMSO (0.1%) or Bis-T-23 (30 µM; 0.1% DMSO) for 1 h and cisplatin (50 µM) or media for another 23 h. A Millicell ERS Voltohmmeter was used to measure the transepithelial electrical resistance (before and after 24 h of initiating the treatment) in each sample.
Atomic force microscopy (AFM)
MDCK cells were seeded on 34 mm culture dishes (TPP) and grown overnight to confluency. Cells were treated with DMSO (0.1%), LatA (0.2 μM), CytoD (0.25 μM), Bis-T-23 (30 µM), or Dynole (10 µM) for 30 min before data acquisition. When indicated, cells were treated with identical concentrations of either DMSO or Bis-T-23 for 10 min before adding LatA for 20 min. When indicated, experiments were performed in cells expressing either Dyn1WT or Dyn2KE.
NanoWizard IV system
AFM imaging was performed using a NanoWizard IV (JPK Instruments) mounted on an Axiovert 200 inverted light microscope (Carl Zeiss). Microscopes were equipped with a heated stage (37 °C) and cells were imaged in HEPES-buffered medium. As previously reported for MDCK cells, indentation measurements were performed with silicon nitride cantilevers (MLCT, Bruker) with a nominal force constant of 0.01 N/m21. Automated calibration was performed in contact-free mode as provided by the calibration manager in the JPK SMP software before every series of experiments. Areas of 2500 µm2 were scanned in QI imaging mode to acquire 3600 independent force measurements per area. Cells were indented up to a force of 0.9 nN at a vertical speed of 18 µm/s. Determination of Young’s Modulus was performed using JPK data analysis software. Fit parameters were set to the Hertzian model according to Sneddon. Cantilever was specified as quadratic pyramid tip shape with a half-angle to edge of 15°. Image analysis and quantification of apical stiffness and stiffness at cell–cell contacts were performed using Fiji software. Topography maps of confluent MDCK cells show elevation at cell–cell junctions. Cell contact sites were marked as ROI manually in the grayscale topography map and ROIs were transferred to the associated force map to analyze stiffness at the respective sites. For determination of apical cell stiffness, grayscale topography maps were analyzed using Fiji threshold presets MaxEntropy/Dark background to mark cells and identify ROIs. ROIs were transferred to the associated force map to determine cell stiffness. Data from at least 10 cells in at least three culture dishes were included in the analysis. Statistical analysis was performed as univariate analysis, defining a p-value of <0.05 as statistically significant.
Bioscope II AFM system
AFM imaging was performed using a Bioscope II AFM (Veeco) mounted on an Olympus IX 73 inverted light microscope (Olympus). The microscope was equipped with a heated stage (37 °C) and cells were imaged in HEPES-buffered medium. Measurements were performed in contact mode using silicon nitride cantilevers with a nominal force constant of 0.01 N/m. Cantilevers were calibrated before every series of experiments by thermal tune using the manufacturer’s software. Force indentation curves were analyzed according to the model of Discher et al.23 computed with MATLab software. Data from at least 10 cells in at least three culture dishes were included in the analysis. Statistical analysis was performed as univariate analysis, defining a p-value of <0.05 as statistically significant.
Electron microscopy (EM)
All EM images were acquired at the Massachusetts General Hospital (MGH), Harvard Medical School, or Brandeis University EM core facility.
Scanning electron microscopy (SEM)
MDCK cells were treated as described in the “Immunocytochemistry for ZO-1 and F-actin” section, fixed for 2 h in 2% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4 (Electron Microscopy Sciences, EMS), rinsed with 0.1 M sodium cacodylate buffer, treated with 1% osmium tetroxide, rinsed in 0.1 M sodium cacodylate buffer and dehydrated through a graded series of ethanol reaching 100%. The samples were dehydrated in the critical point dryer (CPD) Autosamdri®-815 (Tousimis), coated with a 5 nm layer of platinum using a sputter coater (SC) EM ACE600 (Leica), and mounted on stubs using double-sided carbon conductive tape (EMS). Samples were examined at 3–5 kV using S-4700 FE-SEM (Hitachi). Cell height and microvilli length were measured using SEM software and ImageJ, respectively. Microvilli density in a constant area (0.7 in2) on the membrane (≥10 areas per condition) was counted manually.
Transmission electron microscopy (TEM) experiments
(1) Negative staining of proteins in the reconstituted system: G-actin, F-actin and gelsolin-actin filaments (gelsolin:actin = 1:50) were prepared as described previously10. Gelsolin-actin was diluted to 10 μM and incubated with 2 µM recombinant Dyn1, Dyn2, or Dyn1∆PRD for 60 min at 25 °C in the presence or absence of Bis-T-23 (0.4 μM). All samples were diluted to 0.2 µM final actin concentration, and absorbed to glow discharged formvar-carbon coated copper grids for 1 min, blotted to remove excess solution, negatively stained with 1.5% (w/v) uranyl acetate for 1 min, blotted again, and allowed to air-dry. Images were captured at an acceleration voltage of 80 kV using an FEI Morgani 268 or JEOL JEM 1011 transmission electron microscope (TEM) equipped with CCD camera. The brush tool with 50% opacity in Photoshop (Adobe), and “contour plotter” in ImageJ were used for data presentation.
(2) Platinum replica EM and immunogold EM in cells: MDCK cells were treated as described in the “Immunocytochemistry for ZO-1 and F-actin” section, detergent extracted and processed for PR-EM as described previously71. For dynamin localization, samples were fixed with 0.2% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, quenched with NaBH4 for 10 min, treated with Dyn2 antibody (1:100), and subsequently with 18 nm colloidal gold-secondary antibody (1:5). Cells were fixed and processed the same for PREM71. For PR-EM and immunogold PREM, samples were dehydrated in a CPD, coated with a 2 nm layer of platinum, and stabilized with 5 nm of carbon using SC. Samples were examined as described above and images were presented as inverted contrast.
(3) Immunogold of ultrathin sections in cells: The experiments were performed as described8. Hudy 1 (1:100) and goat anti-mouse IgG gold-conjugated (15 nm) antibodies (1:15) were used for identifying dynamin; and anti-actin (1:200) and rabbit anti-chicken IgG gold-conjugated (10 nm) antibodies (1:15) were used for identifying actin. Sections were examined at 80 kV in a JEOL JEM 1011 TEM with AMT digital camera.
Measurement of cellular bioenergetics
A Seahorse Extracellular Flux (XFe24) Analyzer (Agilent) was used to measure oxygen consumption rates (OCR) in real time47. HK-2 cells were treated with only media (control) or media containing human recombinant uPAR (10 ng/ml), human anti-uPAR antibody (50 ng/ml), or Bis-T-23 (10 µM) either alone or in combination for 24 h. Respiratory parameters were quantified, using the Cell Mito Stress Test Kit (Agilent), by subtracting respiration rates at times before and after the addition of mitochondrial activators and inhibitors47. Experiments were replicated in five wells, averaged for each treatment group and the data were presented as means ± standard errors of the means (SEM). One-way ANOVA using Prism was used to compare controls (CTL) with treatment groups (*P < 0.05, **P < 0.01, ***P < 0.001).
Rat kidney tissue slice experiment
Male Spraque-Dawley rats (strain code 400) were from Charles River. Rat kidney were prepared with minor modifications29. The slices were treated with DMSO (0.1%) or Bis-T-23 (30 μM, 0.1% DMSO) for 1 h before adding cisplatin (200 μM) for 8 h and fixing with periodate-lysine paraformaldehyde overnight at 4 °C. For the albumin uptake assay, kidney slices were incubated in the presence or absence of Bis-T-23 (30 μM) for 1.5 h before adding rhodamine-albumin (100 μg/ml) for 30 min. Slices were then rinsed with PBS, fixed, and processed. Animal experiments were approved by the institutional committee on Research Animal Care at MGH, in accordance with the NIH Guide for the care and use of the laboratory animals (protocol #2012N000004 (PI, Brown)).
1. Single-dose cisplatin induced kidney injury39
Male and female C57BL/J6 mice (9 weeks, 25–30 g) (Strain #000664, common name BL6, The Jackson Laboratory) were randomly divided into 3 experimental groups as follows: (1) cisplatin+DMSO; (2) cisplatin+saline; and (3) cisplatin+Bis-T-23. Mice in each group received a single intraperitoneal (i.p.) injection of 15 mg/kg cisplatin in 0.9% saline. Bis-T-23 dissolved in 1% DMSO was injected i.p. once 24 h before cisplatin injection, and daily at 20 mg/kg per injection. Body weight was monitored, and urine and blood parameters were assessed at baseline and after cisplatin injection. Mice were euthanized 4 days after cisplatin injection by cervical dislocation. Experiments were performed at Rush University and were approved by Rush University Institutional Animal Care and Use Committee (IACUC) (protocol #18-060 (PI, Reiser)).
2. Iohexol-induced acute kidney injury47
BL6 mice (males and females) were purchased and mice expressing suPAR from the fat tissue (suPAR-Tg)48 were bred at Rush University. Mice aged 9–12 weeks (25–30 g) were randomly divided in 3 experimental groups as follows: (1) contrast+DMSO; (2) contrast+saline; and (3) contrast+Bis-T-23. Blood and urine were collected for baseline measurements. Mice were denied free oral water access for 48 h. Then, group 1 was administered DMSO (1%, 200 µl), group 2 received 0.9% saline (200 µl) and group 3 was given Bis-T-23 (20 mg/kg in 200 µl of 0.9% saline). After 5 h, all the three groups received L-NAME (10 mg/kg in 100 µl of water), a nitric oxide synthase inhibitor, and indomethacin (10 mg/kg in 100 µl of water), an inhibitor of prostaglandin synthesis. After 1 h, all three groups received iohexol (5 g/kg of water), a non-ionic low osmotic contrast medium. Subsequently, oral rehydration was resumed. A second dose of Bis-T-23 (20 mg/kg in 200 µl of 0.9% saline) was administered to group 3 after 24 h of the first dose. Group 1 and group 2 received DMSO or saline as described above. All injections were given i.p. Urine and blood samples were collected 24 and 48 h after iohexol injection. Experiments were performed at Rush University and were approved by RU IACUC (protocol #19-014 (PI, Reiser)).
3. Alport syndrome mice
Alport strain (#002908 Jackson Labs) heterozygous mice were bred at the Rush University, and the homozygous mice (Col4a3−/−)55 were treated with either DMSO (1%) or Bis-T-23 (20 mg/kg) daily starting at 42 days of age (D0 of the treatment). Experiments were performed at Rush University and were approved by Rush University IACUC (protocol #18-060 (PI, Reiser)).
Log-Rank (Mantel-cox) test was performed in Prism to analyze survival curves of the two treatment groups and the control animals. The log-rank test reports chi-square value, to compute P-value testing of the null hypothesis that there is no linear trend between column order and median survival.
Assessment of renal function
BUN levels were measured using Quantichrom Urea assay kit (BioAssay Systems). Urinary albumin-to-creatinine ratio (ACR) was assessed by mouse albumin ELISA kit (Bethyl Laboratories) and creatinine colorimetric assay kit (Cayman Chemical). Assays were performed according to manufacturer’s protocols and read using EnSpire multimode plate reader (Perkin Elmer). Unpaired two-tailed t-tests were performed using Prism (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001; whereas P > 0.05 was considered not significant, ns).
The authors declare that all data supporting the findings of this study are available within the paper, its supplementary information files, and in the Source data file. There are no restrictions on any data availability. We did not use any accession codes, unique identifiers, or web links for publicly available datasets. No clinical datasets were used.
Basile, D. P., Anderson, M. D. & Sutton, T. A. Pathophysiology of acute kidney injury. Compr. Physiol. 2, 1303–1353 (2012).
Bonventre, J. V. & Yang, L. Cellular pathophysiology of ischemic acute kidney injury. J. Clin. Invest. 121, 4210–4221 (2011).
Campanale, J. P., Sun, T. Y. & Montell, D. J. Development and dynamics of cell polarity at a glance. J. Cell Sci. 130, 1201–1207 (2017).
Salbreux, G., Charras, G. & Paluch, E. Actin cortex mechanics and cellular morphogenesis. Trends Cell Biol. 22, 536–545 (2012).
Vicente-Manzanares, M., Ma, X., Adelstein, R. S. & Horwitz, A. R. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol. Cell Biol. 10, 778–790 (2009).
Chugh, P. et al. Actin cortex architecture regulates cell surface tension. Nat. Cell Biol. 19, 689–697 (2017).
Nguyen, L. T. & Hirst, L. S. Polymorphism of highly cross-linked F-actin networks: probing multiple length scales. Phys. Rev. E Stat. Nonlin Soft Matter Phys. 83, 031910 (2011).
Sever, S. et al. Physical and functional connection between auxilin and dynamin during endocytosis. EMBO J. 25, 4163–4174 (2006).
Antonny, B. et al. Membrane fission by dynamin: what we know and what we need to know. EMBO J. 35, 2270–2284 (2016).
Gu, C. et al. Direct dynamin-actin interactions regulate the actin cytoskeleton. EMBO J. 29, 3593–3606 (2010).
Gu, C. et al. Dynamin autonomously regulates podocyte focal adhesion maturation. J. Am. Soc. Nephrol. 28, 446–451 (2017).
Schiffer, M. et al. Pharmacological targeting of actin-dependent dynamin oligomerization ameliorates chronic kidney disease in diverse animal models. Nat. Med. 21, 601–609 (2015).
Gu, C. et al. Regulation of dynamin oligomerization in cells: the role of dynamin-actin interactions and its GTPase activity. Traffic 15, 819–838 (2014).
Hill, T. A. et al. Inhibition of dynamin mediated endocytosis by the dynoles—synthesis and functional activity of a family of indoles. J. Med. Chem. 52, 3762–3773 (2009).
Madsen, K. M., Verlander, J. W. & Tisher, C. C. Relationship between structure and function in distal tubule and collecting duct. J. Electron Microsc Tech. 9, 187–208 (1988).
Wayt, J. & Bretscher, A. Cordon Bleu serves as a platform at the basal region of microvilli, where it regulates microvillar length through its WH2 domains. Mol. Biol. Cell 25, 2817–2827 (2014).
Holzinger, A. Jasplakinolide: an actin-specific reagent that promotes actin polymerization. Methods Mol. Biol. 586, 71–87 (2009).
Mettlen, M., Pucadyil, T., Ramachandran, R. & Schmid, S. L. Dissecting dynamin’s role in clathrin-mediated endocytosis. Biochemical Soc. Trans. 37, 1022–1026 (2009).
Aguet, F., Antonescu, C. N., Mettlen, M., Schmid, S. L. & Danuser, G. Advances in analysis of low signal-to-noise images link dynamin and AP2 to the functions of an endocytic checkpoint. Dev. Cell 26, 279–291 (2013).
Chugh, P. & Paluch, E. K. The actin cortex at a glance. J. Cell Sci. 131, jcs186254 (2018).
Nehls, S., Noding, H., Karsch, S., Ries, F. & Janshoff, A. Stiffness of MDCK II cells depends on confluency and cell size. Biophys. J. 116, 2204–2211 (2019).
Shawky, J. H., Balakrishnan, U. L., Stuckenholz, C. & Davidson, L. A. Multiscale analysis of architecture, cell size and the cell cortex reveals cortical F-actin density and composition are major contributors to mechanical properties during convergent extension. Development 145, dev161281 (2018).
Sen, S., Subramanian, S. & Discher, D. E. Indentation and adhesive probing of a cell membrane with AFM: theoretical model and experiments. Biophys. J. 89, 3203–3213 (2005).
Collins, A., Warrington, A., Taylor, K. A. & Svitkina, T. Structural organization of the actin cytoskeleton at sites of clathrin-mediated endocytosis. Curr. Biol. 21, 1167–1175 (2011).
Chuang, M. C. et al. Tks5 and Dynamin-2 enhance actin bundle rigidity in invadosomes to promote myoblast fusion. J. Cell Biol. 218, 1670–1685 (2019).
Menon, M., Askinazi, O. L. & Schafer, D. A. Dynamin 2 organizes lamellipodial actin networks to orcestrate lamellar actomyosin. PLoS ONE 9, e94330 (2014).
Lin, S. S. et al. Dynamin-2 regulates postsynaptic cytoskeleton organization and neuromuscular junction development. Cell Rep. 33, 108310 (2020).
Zhang, R. et al. Dynamin regulates the dynamics and mechanical strength of the actin cytoskeleton as a multifilament actin-bundling protein. Nat. Cell Biol. 22, 674–688 (2020).
Bouley, R. et al. Nitric oxide and atrial natriuretic factor stimulate cGMP-dependent membrane insertion of aquaporin 2 in renal epithelial cells. J. Clin. Invest. 106, 1115–1126 (2000).
Holditch, S. J., Brown, C. N., Lombardi, A. M., Nguyen, K. N. & Edelstein, C. L. Recent advances in models, mechanisms, biomarkers, and interventions in cisplatin-induced acute kidney Injury. Int. J. Mol. Sci. 20, 3011 (2019).
Meng, X. M. et al. NADPH oxidase 4 promotes cisplatin-induced acute kidney injury via ROS-mediated programmed cell death and inflammation. Lab Invest. 98, 63–78 (2018).
Balta, E., Kramer, J. & Samstag, Y. Redox regulation of the actin cytoskeleton in cell migration and adhesion: on the way to a spatiotemporal view. Front Cell Dev. Biol. 8, 618261 (2020).
Gourlay, C. W., Carpp, L. N., Timpson, P., Winder, S. J. & Ayscough, K. R. A role for the actin cytoskeleton in cell death and aging in yeast. J. Cell Biol. 164, 803–809 (2004).
Moore, A. S. & Holzbaur, E. L. F. Mitochondrial-cytoskeletal interactions: dynamic associations that facilitate network function and remodeling. Curr. Opin. Physiol. 3, 94–100 (2018).
Kruidering, M. et al. Cisplatin effects on F-actin and matrix proteins precede renal tubular cell detachment and apoptosis in vitro. Cell Death Differ. 5, 601–614 (1998).
Vinaiphat, A., Charngkaew, K. & Thongboonkerd, V. More complete polarization of renal tubular epithelial cells by artificial urine. Cell Death Disco. 4, 47 (2018).
Mukherjee, K. et al. A novel fluorogenic assay for the detection of nephrotoxin-induced oxidative stress in live cells and renal tissue. ACS Sens 6, 2523–2528 (2021).
Corridon, P. R., Karam, S. H., Khraibi, A. A., Khan, A. A. & Alhashmi, M. A. Intravital imaging of real-time endogenous actin dysregulation in proximal and distal tubules at the onset of severe ischemia-reperfusion injury. Sci. Rep. 11, 8280 (2021).
Katagiri, D. et al. Protection of glucagon-like peptide-1 in cisplatin-induced renal injury elucidates gut-kidney connection. J. Am. Soc. Nephrol. 24, 2034–2043 (2013).
Resh, M. D. & Erikson, R. L. Development and characterization of antisera specific for amino- and carboxy-terminal regions of pp60src. J. Virol. 55, 242–245 (1985).
Baliga, R., Zhang, Z., Baliga, M., Ueda, N. & Shah, S. V. In vitro and in vivo evidence suggesting a role for iron in cisplatin-induced nephrotoxicity. Kidney Int. 53, 394–401 (1998).
Ozkok, A. & Edelstein, C. L. Pathophysiology of cisplatin-induced acute kidney injury. Biomed. Res. Int. 2014, 967826 (2014).
Lei, R. et al. Mitophagy plays a protective role in iodinated contrast-induced acute renal tubular epithelial cells injury. Cell Physiol. Biochem. 46, 975–985 (2018).
Seeliger, E., Sendeski, M., Rihal, C. S. & Persson, P. B. Contrast-induced kidney injury: mechanisms, risk factors, and prevention. Eur. Heart J. 33, 2007–2015 (2012).
Jeong, B. Y. et al. Oxidative stress caused by activation of NADPH oxidase 4 promotes contrast-induced acute kidney injury. PLoS ONE 13, e0191034 (2018).
Meinel, F. G., De Cecco, C. N., Schoepf, U. J. & Katzberg, R. Contrast-induced acute kidney injury: definition, epidemiology, and outcome. Biomed. Res. Int. 2014, 859328 (2014).
Hayek, S. S. et al. Soluble urokinase receptor and acute kidney injury. N. Engl. J. Med. 382, 416–426 (2020).
Wei, C. et al. uPAR isoform 2 forms a dimer and induces severe kidney disease in mice. J. Clin. Invest. 129, 1946–1959 (2019).
Gubler, M. et al. Alport’s syndrome. A report of 58 cases and a review of the literature. Am. J. Med. 70, 493–505 (1981).
Pescucci, C. et al. Autosomal-dominant Alport syndrome: natural history of a disease due to COL4A3 or COL4A4 gene. Kidney Int. 65, 1598–1603 (2004).
Kim, K. H. et al. Structural-functional relationships in Alport syndrome. J. Am. Soc. Nephrol. 5, 1659–1668 (1995).
Jedlicka, J. et al. Interstitial inflammation in Alport syndrome. Hum. Pathol. 41, 582–593 (2010).
Hood, J. C. et al. Correlation of histopathological features and renal impairment in autosomal dominant Alport syndrome in Bull terriers. Nephrol. Dial. Transpl. 17, 1897–1908 (2002).
Tanaka, M. et al. Loss of the BMP antagonist USAG-1 ameliorates disease in a mouse model of the progressive hereditary kidney disease Alport syndrome. J. Clin. Invest. 120, 768–777 (2010).
Cosgrove, D. et al. Collagen COL4A3 knockout: a mouse model for autosomal Alport syndrome. Genes Dev. 10, 2981–2992 (1996).
Yamada, H., Takeda, T., Michiue, H., Abe, T. & Takei, K. Actin bundling by dynamin 2 and cortactin is implicated in cell migration by stabilizing filopodia in human non-small cell lung carcinoma cells. Int. J. Oncol. 49, 877–886 (2016).
Warnock, D. E., Hinshaw, J. E. & Schmid, S. L. Dynamin self-assembly stimulates its GTPase activity. J. Biol. Chem. 271, 22310–22314 (1996).
Ross, J. A. et al. Dimeric endophilin A2 stimulates assembly and GTPase activity of dynamin 2. Biophysical J. 100, 729–737 (2011).
Ralto, K. M. & Parikh, S. M. Mitochondria in acute kidney injury. Semin Nephrol. 36, 8–16 (2016).
Murphy, M. P. How mitochondria produce reactive oxygen species. Biochem. J. 417, 1–13 (2009).
Duchen, M. R. & Szabadkai, G. Roles of mitochondria in human disease. Essays Biochem. 47, 115–137 (2010).
Marullo, R. et al. Cisplatin induces a mitochondrial-ROS response that contributes to cytotoxicity depending on mitochondrial redox status and bioenergetic functions. PLoS ONE 8, e81162 (2013).
Cheng, W. et al. Comparison of iohexol and iodixanol induced nephrotoxicity, mitochondrial damage and mitophagy in a new contrast-induced acute kidney injury rat model. Arch. Toxicol. 92, 2245–2257 (2018).
Kleih, M. et al. Direct impact of cisplatin on mitochondria induces ROS production that dictates cell fate of ovarian cancer cells. Cell Death Dis. 10, 851 (2019).
Xu, Q., Huff, L. P., Fujii, M. & Griendling, K. K. Redox regulation of the actin cytoskeleton and its role in the vascular system. Free Radic. Biol. Med. 109, 84–107 (2017).
Kurzhagen, J. T., Dellepiane, S., Cantaluppi, V. & Rabb, H. AKI: an increasingly recognized risk factor for CKD development and progression. J. Nephrol. 33, 1171–1187 (2020).
Faubel, S. et al. Ongoing clinical trials in AKI. Clin. J. Am. Soc. Nephrol. 7, 861–873 (2012).
Azam, T. U. et al. Soluble urokinase receptor (SuPAR) in COVID-19-related AKI. J. Am. Soc. Nephrol. 31, 2725–2735 (2020).
Hayek, S. S. et al. Soluble urokinase receptor and chronic kidney disease. N. Engl. J. Med. 373, 1916–1925 (2015).
Battistone, M. A. et al. Proinflammatory P2Y14 receptor inhibition protects against ischemic acute kidney injury in mice. J. Clin. Invest. 130, 3734–3749 (2020).
Svitkina, T. Imaging Cytoskeleton components by electron microscopy. Methods Mol. Biol. 1365, 99–118 (2016).
We would like to thank Astrid Schauss and Lisa Wirtz (CECAD Imaging Facility, University of Cologne, Germany) for their advice on AFM measurements and analysis. We would like to thank Benjamin Dellaripa for his technical assistance with cell culture and image quantification. We acknowledge support from the National Institutes of Health (R01 DK093773 and DK087985 to S.S.; R01 DK084195 to V.G. and S.S.; GM73165 to M.M.; CA227747 to S.L.B.). C.G. was supported by Rush University Medical Center. Electron microscopy was performed in the Microscopy Core of the Center for Systems Biology/Program in Membrane Biology, which is partially supported by an Inflammatory Bowel Disease Grant DK043351. P.T.B. and H.H. received funding from the DFG (KFO329, BR-BR 2955/8-1), the Koeln Fortune Program of the University of Cologne.
S.S. and J.R. are co-founders and shareholders of Walden Biosciences, a biotechnology company that develops novel kidney-protective therapies. S.L.B. and K.M. are inventors on a pending patent application pertaining to the detection of oxidative stress-induced carbonylation. The remaining authors declare no competing interests.
Peer review information
Nature Communications thanks the anonymous reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Mukherjee, K., Gu, C., Collins, A. et al. Simultaneous stabilization of actin cytoskeleton in multiple nephron-specific cells protects the kidney from diverse injury. Nat Commun 13, 2422 (2022). https://doi.org/10.1038/s41467-022-30101-4
Nature Reviews Nephrology (2022)