Rho controls actin cytoskeletal assembly in renal epithelial cells during ATP depletion and recovery

Narayan Raman and Simon J. Atkinson

Department of Medicine, Indiana University School of Medicine, Indianapolis, Indiana 46202


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Actin cytoskeletal disruption is a hallmark of ischemic injury and ATP depletion in a number of cell types, including renal epithelial cells. We manipulated Rho GTPase signaling by transfection and microinjection in LLC-PK proximal tubule epithelial cells and observed actin cytoskeletal organization following ATP depletion or recovery by confocal microscopy and quantitative image analysis. ATP depletion resulted in disruption of stress fibers, cortical F-actin, and apical actin bundles. Constitutively active RhoV14 prevented disruption of stress fibers and cortical F-actin during ATP depletion and enhanced the rate of stress fiber reassembly during recovery. Conversely, the Rho inhibitor C3 or dominant negative RhoN19 prevented recovery of F-actin assemblies upon repletion. Actin bundles in the apical microvilli and cytosolic F-actin were not affected by Rho signaling. Assembly of vinculin and paxillin into focal adhesions was disrupted by ATP depletion, and constitutively active RhoV14, although protecting stress fibers from disassembly, did not prevent dispersion of vinculin and paxillin, resulting in uncoupling of stress fiber and focal adhesion assembly. We propose that ATP depletion causes Rho inactivation during ischemia and that recovery of normal cellular architecture and function requires Rho.

acute kidney tubular necrosis; cell adhesion; G proteins; ischemia; proximal kidney tubules


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

CELLULAR ATP DEPLETION THAT accompanies ischemic injury causes disruption of the normal architecture and function of the actin cytoskeleton in diverse cell types. The consequences of disruption to normal actin cytoskeletal function are particularly well studied in the kidney (7), in which ischemic injury and consequent acute tubular necrosis is characterized in its early phases by reversible cellular injury that affects particularly the proximal tubule cells of the nephron (16, 29, 37, 39, 55, 58). The polarized distribution of proteins and lipids is lost, and the Na+-K+-ATPase redistributes from the basolateral to the apical membrane (38). The tight junction barrier opens (6, 11, 33), and cell-cell and cell-substrate adhesion is lost. Formation of apical membrane blebs accompanies destruction of brush-border microvilli (19, 57). The sensitivity to injury of actin assemblies within the cell is unequal, with microvillar actin bundles being most sensitive and the cortical actin gel associated with the junctional complex being most stable (6). Many of the features of in vivo ischemic injury can be reproduced in a cell culture model by chemically induced ATP depletion, including disruption of brush-border microvillar actin bundles, cortical F-actin, and stress fibers (39). Remarkably, this wholesale rearrangement of cellular F-actin is accompanied by an increase, not a decrease, in the fraction of cellular actin that is polymeric; indicating that disassembly of normal structures does not result from a global depolymerization of actin (38) but rather may result from aberrant signaling events. During recovery, actin assemblies that were damaged by ischemic injury can be reconstructed.

The steady-state organization of actin in the cell is maintained as a result of the activities of a large number of actin binding proteins. Their functions include cross-linking actin filaments into higher-order assemblies of gels or bundles, linking actin filaments to the plasma membrane, and, most importantly, regulating the monomer-polymer equilibrium at filament ends (45). The cytoskeletal architecture of the cell results from coordination of the activity of actin binding proteins that are directly regulated by calcium ion, polyphosphoinositides, phosphorylation, or other mechanisms. ATP depletion in vitro, or ATP depletion resulting from ischemia in vivo, initiates events that disassemble actin-containing structures but also results in unregulated polymerization of monomeric actin (34). One explanation for the disruptive effect of ATP depletion on particular actin-containing structures is that it alters the signaling pathways that normally regulate the activity of actin binding proteins.

Regulation of most actin-dependent processes has been demonstrated to be mediated by Rho family GTPases in many different cell types, including epithelial cells (21). Rho family GTPases are members of the Ras gene superfamily of p21 GTPases that function as molecular switches by cycling between GTP- and GDP-bound states. Activity of Rho proteins is regulated by upstream pathways in response to growth factor and cytokine signaling, and their activity integrates control of downstream pathways that affect cell morphology and motility, transcription of specific genes, and cell cycle progression (56).

Three members of the Rho family GTPases, RhoA, Rac, and Cdc42, are widely expressed and have been extensively studied using microinjection and transfection of constitutively active and dominant negative mutants (17, 18) or, to selectively inhibit RhoA, by using C3 ADP-ribosyl transferase (1). In fibroblasts, Cdc42 induces formation of filopodia, and Rac induces membrane ruffling and formation of lamellipodia and "focal complexes" (structures antigenically related to but morphologically distinct from focal adhesions), whereas RhoA induces stress fiber and focal adhesion formation (40, 49, 50). These studies also established the potential for a hierarchy of Rho family activation in which activated Cdc42 promotes Rac activation, which in turn promotes Rho activation.

In epithelial cells, Rho family GTPases regulate structures associated with the actin cytoskeleton that are responsible for the maintenance of cell polarity and cohesiveness of the tissue (8, 9, 27, 28, 42, 54). Introducing C3 into cultured epithelial cells resulted in loss of E-cadherin, catenins, ZO-1, and actin from cell-cell junctions. Transfection with constitutively active RhoV14, however, induced accumulation of additional junctional complex components at junctions between transfected cells and also produced thickening of cortical F-actin in the perijunctional region. Scattering of Madin-Darby canine kidney cells upon stimulation with hepatocyte growth factor (HGF) was inhibited by constitutively active RhoV14 or dominant negative RacN17 (48), suggesting a need for differential regulation of the two pathways in certain cellular responses.

To test the potential involvement of Rho family GTPases in actin cytoskeletal disruption induced by ATP depletion and in reorganization during recovery, we manipulated Rho signaling in transfection and microinjection experiments and measured the effect on actin and associated structures in control, ATP-depleted, and recovering cells by quantitative fluorescence microscopy. Our results indicate that constitutive activation of Rho protects actin cytoskeletal structures during ATP depletion and that inactivation of Rho prevents recovery of normal actin cytoskeletal architecture. We suggest that ATP depletion attenuates and repletion stimulates the activity of Rho family GTPases and that this offers an explanation for much of the damage to proximal tubule cell structure and function in renal ischemic injury.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. LLC-PK porcine proximal tubule cells [American Type Culture Collection (ATCC), Manassas, VA] were maintained in 1:1 DMEM/F-12 (Sigma Chemical, St. Louis, MO) medium containing 10% FCS at 37°C in a humidified atmosphere of 5% CO2. LLC-PK cells were seeded at 2 × 105 cells/35-mm-diameter dish containing an 18-mm square coverslip. Transient transfections were performed on cells grown 1 day to 50% confluency. Microinjection experiments on LLC-PK cells were performed on cells grown for 3 days to >90% confluency.

Expression and purification of recombinant proteins. Plasmids pEXV-RhoA(V14)-Myc (constitutively active) and pEXV-RhoA(N19)-Myc (dominant negative), for transient expression of myc epitope-tagged proteins, were the kind gift of Dr. Marc Symons (Onyx Pharmaceuticals, Richmond, CA). Plasmids for expression of recombinant pGEX-2T-RhoV14 and pGEX-2T-C3 in bacteria were a kind gift from Dr. Alan Hall (University College London). Glutathione S-transferase-Rho fusion proteins were expressed in Escherichia coli strain DH5alpha and isolated from bacterial extracts by binding to glutathione-agarose beads (Sigma) as described (53). Rho and C3 proteins were released from the bead-bound glutathione S-transferase moiety by treatment with activated thrombin, and the protease was then removed by binding to p-aminobenzamidine-conjugated agarose (Sigma). A filter-binding assay using [3H]GTP was used to determine the concentration of active RhoV14 protein as described (53). C3 transferase concentrations were determined by the protein dye binding method using Bio-Rad (Hercules, CA) reagent and standard curves calibrated with BSA. Activity of C3 preparations was checked by ADP-ribosylation of purified recombinant RhoV14 using [32P]NAD as substrate (2). Purified proteins were dialyzed exhaustively against PBS [disodium hydrogen orthophosphate (84 mM), potassium dihydrogen orthophosphate (14 mM), sodium chloride (138 mM), and potassium chloride (26 mM), pH 7.4] and concentrated using Centricon 30 filter units (Amicon, Beverly, MA). Proteins were stored at -80°C.

Transfection and microinjection. For transient transfection of LLC-PK cells, Lipofectamine (GIBCO-BRL, Gaithersburg, MD) was used as a carrier. Plasmid DNA (2 µg), purified using Qiagen (Valencia, CA) Maxiprep kits, was mixed with 5 µl of Lipofectamine in 200 µl of serum-free DMEM and incubated at room temperature for 45 min. After addition of 800 µl of serum-free DMEM, the mix was added to cells that had been washed with serum-free DMEM. Cells were incubated with the Lipofectamine-DNA mixture for 4 h at 37°C in a humidified atmosphere of 5% CO2, after which the mixture was replaced with DMEM/F-12 medium containing 10% FCS. In preliminary experiments, we found that transfection efficiency peaked between 24 and 48 h after transfection, and so all experiments on transiently transfected cells were performed 48 h posttransfection.

For microinjection, the protein of interest was coinjected with fluorescein-dextran (80,000 mol wt, lysine-conjugated; Molecular Probes, Eugene, OR) to identify injected cells after processing for immunofluorescence. The proteins RhoV14 and C3 were at concentrations of 0.06 and 0.01 mg/ml, respectively.

ATP depletion and ATP assays. Cells 48 h posttransfection (transfection experiments) or cells 72 h postseeding (microinjection experiments) were exposed to either control medium (DMEM) or substrate-free (glucose and amino acids omitted) DMEM containing 0.1 µM antimycin A (11) for the time stated. To allow recovery of ATP concentrations following depletion, cells were washed and returned to normal growth medium with serum.

ATP levels were assayed by luminometry using FireLight reagents (Analytical Luminescence Laboratories, San Diego, CA) according to the manufacturer's instructions. Cells were extracted with 6% perchloric acid for 15 min on ice. The perchloric acid extract was centrifuged for 5 min in a microcentrifuge, and the supernatant was neutralized with potassium carbonate (5 M). The pellet was used to determine the protein content of the extract. The ATP concentration in the supernatant was measured using the luciferase assay in a Monolight luminometer (Analytical Luminescence Laboratories). ATP (Sigma) was used to prepare a standard curve. The pellet was dissolved in sodium hydroxide (1 N), and protein was assayed using the micro-bicinchoninic method (Pierce Chemical, Rockford, IL). BSA was used a standard. The amount of ATP in each sample was standardized relative to the protein content.

Fluorescence cytochemistry. Cells were fixed with 3.7% formaldehyde in PBS for 10 min at room temperature and permeabilized with 0.05% Triton X-100 in PBS for 5 min. For most experiments, coverslips were blocked in PBS containing 10% goat serum and 0.2% BSA for 30 min (block solution). F-actin was labeled by incubation of coverslips with 0.1 µg/ml rhodamine- or Texas red-labeled phalloidin (Molecular Probes) in block solution for 1 h at room temperature. Transfected cells that expressed RhoV14 or RhoN19 were identified by expression of the myc epitope tag. Coverslips were incubated with a 1:100 dilution of mouse monoclonal antibody 9E10 (ATCC) for 1 h, followed by a brief wash and further incubation with a 1:50 dilution of fluorescein isothiocyanate-conjugated goat anti-mouse IgG (Jackson Immunoresearch Laboratories, West Grove, PA).

To analyze focal adhesion assembly in transfected cells, we used simultaneous three-color labeling for F-actin, the myc epitope tag, and one of the focal adhesion proteins vinculin and paxillin (10). Cells were fixed and permeabilized as before but were blocked in 1:10 BlokHen (Aves Labs, Tigard, OR) solution for 60 min. Cells were labeled by incubating coverslips in chicken anti-myc epitope antibody (Aves Labs) diluted 1:1,000 together with mouse monoclonal antibodies at the indicated dilutions in 0.01% Triton X-100 in PBS for 1 h at room temperature. Anti-vinculin antibody VIN-11-5 (Sigma) and anti-paxillin antibody M165 (Transduction Laboratories, Lexington, KY) were diluted 1:50. Coverslips were washed for 20 min with 0.01% Triton-PBS and incubated with a 1:50 dilution of Cy-5-conjugated goat anti-mouse IgG (Jackson), 1:100 FITC-conjugated goat anti-chicken IgY (Aves Labs), and 0.1 µg/ml rhodamine phalloidin for 60 min. Coverslips were then washed as before. All coverslips were mounted in 50% glycerol-PBS containing diamino-bicyclo-[2.2.2] octane (100 mg/ml; Sigma) to minimize photobleaching.

Microscopy, image processing, and quantitative analysis. Images were collected with a Bio-Rad MRC 1024 confocal microscope mounted on a Nikon Diaphot stand using a ×100 (1.4 numerical aperture) objective lens and 3-mm confocal aperture. Z-series stacks of images were collected that included the entire cell volume with a separation of 0.5 µm between each confocal section. The background fluorescence signal was estimated by collecting planes from areas of the slide without cells and was subtracted from all sections before analysis. Images were analyzed using Metamorph software (Universal Imaging, West Chester, PA). Figures 1-6 were prepared using Photoshop (Adobe Systems, Mountain View, CA).

Quantitative analysis of F-actin accumulation was performed using Metamorph software. Measurement of stress fiber fluorescence was made on extended focus images made by summing five sections from the base of the cell that included all in-focus stress fiber fluorescence. Stress fiber structures were segmented from surrounding fluorescence in the image that interfered with estimation of stress fiber fluorescence (52). First, a sharpening filter was applied to the image. The resulting image was thresholded and converted to a binary mask. An erode function was applied to remove small particulate structures from the mask while retaining all filamentous structures. The mask was then multiplied against the original image, with the result that stress fiber fluorescence was retained with pixel values unaltered, whereas non-stress fiber fluorescence was removed (see example images in Fig. 4, E and F). F-actin accumulation in the perijunctional region and in the perinuclear cytoplasm was measured on extended focus images of the next five sections toward the apical side of the cell. For all measurements, the area containing the structures of interest was delineated, and the mean fluorescence intensity within the area was determined.

Data were analyzed for significance by Student's t-test (one tail, assuming unequal variances) and ANOVA using DataDesk software (Data Descriptions, Ithaca, NY).


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

ATP depletion and actin organization. ATP depletion of LLC-PK cells by treatment with substrate-depleted medium containing antimycin A (0.1 µM) resulted in disruption of the normal distribution of F-actin structures (Fig. 1A), as previously shown by others (29, 33, 38). Disruption of stress fibers and focal adhesions was nearly complete by 90 min of depletion, and cells frequently detached from the substrate, even when collagen was used to coat the coverslip. Alterations induced by ATP depletion were reversible. By 2 h of recovery, stress fibers and focal adhesions had re-formed, but substantially longer recovery periods were required for full recovery of the normal complement of basal actin structures. We observed an accumulation of amorphous phalloidin-staining aggregates in the perinuclear cytosol of ATP-depleted cells, as described previously (38). F-actin in the cortical region underlying the lateral borders of these cells was somewhat more ragged in appearance than in control cells, but the amount of F-actin labeling in this region did not appear to be significantly altered by ATP depletion. ATP depletion also resulted in disruption of apical actin. Microvillar actin bundles were no longer visible following ATP depletion (not shown).


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Fig. 1.   Effect of ATP depletion on stress fibers in LLC-PK cells. A: LLC-PK cells were incubated in control medium (Control), substrate-depleted medium with antimycin A for 90 min (-ATP), or for 90 min in depleted medium with antimycin followed by 60 min in control medium (-,+ ATP). Cells were fixed, permeabilized, and labeled with rhodamine-phalloidin. Note reversible disruption to organization of stress fibers with ATP depletion. Bar, 20 µm. B: time course of ATP depletion and recovery. ATP concentrations were measured by luminometry using luciferase in extracts of control cells (time 0), cells depleted for 10, 30, 60, and 90 min, or cells depleted for 90 min and allowed to recover for 30 or 60 min (90/30, 90/60). Values are normalized to controls (100%).

ATP content of LLC-PK cells treated with antimycin A and substrate-depleted media was measured using a luciferase assay by luminometry. ATP levels fell to ~20% of control values within 10 min (Fig. 1B). ATP levels recovered to >60% of controls after 60 min in normal medium. Transfection with RhoV14 plasmid vectors did not significantly alter the rate or extent of ATP depletion (not shown).

Microinjection of C3 exoenzyme and RhoV14. We microinjected RhoV14 or C3 transferase into LLC-PK cells to test the effect of constitutive activation or inactivation of Rho on actin cytoskeletal organization (Fig. 2, A and B). As expected from previous work (42, 49), microinjection of C3 transferase (0.01 mg/ml) resulted in disruption of stress fibers and altered cell morphology (Fig. 2B). Microinjected cells frequently had a more irregular shape than uninjected cells and became detached from adjacent cells in the monolayer. Amorphous aggregates that labeled with phalloidin were observed in the perinuclear cytosol, reminiscent of those that accumulate following ATP depletion. We (20) have previously reported that C3 transferase injection also caused tight junction disassembly, as shown by others in intestinal epithelial cells (42).


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Fig. 2.   Effect of microinjecting RhoV14 (constitutively active) or C3 exoenzyme on stress fiber organization in LLC-PK cells. LLC-PK cells were microinjected with RhoV14 (A, C, and E) or C3 exoenzyme (B, D, and F) and incubated for 45 min before exchange of medium to fresh control medium for 90 min (A, B), depleted medium with antimycin A for 90 min (C, D), or depleted medium with antimycin for 90 min followed by 60-min recovery in control medium (E, F). Cells were permeabilized, fixed, and labeled with rhodamine-phalloidin. Microinjected cells, identified by coinjection of FITC-conjugated dextran, are outlined in images. Bar, 20 µm.

Microinjection of RhoV14 caused an increase in the number and thickness of stress fibers and a corresponding accumulation of F-actin in focal adhesions (Fig. 2A). Microinjected cells appeared to be taller and to have a smaller cross-sectional area than other cells. Microinjection of RhoV14 did not alter the number or size of apical F-actin microvillar bundles. Microinjection of PBS-dextran alone did not alter cell morphology or the distribution of F-actin.

Transient transfection of constitutively active and dominant negative Rho. We transiently transfected LLC-PK cells with RhoV14 (constitutively active) and RhoN19 (dominant negative). Expression of mutant Rho proteins, which are myc-tagged, was monitored by immunofluorescence using a monoclonal antibody (9E10). Maximum expression was obtained between 24 and 48 h after transfection. Transfection efficiency appeared to be high, with 10-20% of cells showing high levels of expression and up to 75% showing detectable levels of myc antibody labeling.

Cells expressing high levels of RhoV14 showed changes comparable to those induced by microinjection of recombinant protein (Fig. 3, A and B). Stress fibers increased in number and thickness, and there was increased accumulation of F-actin in the cortical region near the tight junctions that was particularly noticeable at junctions between pairs of transfected cells (arrows in Fig. 3B). Microvillar actin bundles were not affected by RhoV14 expression (not shown).


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Fig. 3.   Transient transfection of LLC-PK cells with RhoV14 (constitutively active). LLC-PK cells were transiently transfected with plasmid pEXVRhoV14 using Lipofectamine. Experiments were performed 48 h after transfection. Medium was exchanged for fresh control medium for 90 min (A, B, C), depleted medium with antimycin for 90 min (D, E, F), or depleted medium for 90 min followed by 60-min recovery (G, H, I). Cells were labeled with rhodamine phalloidin (A, B, D, E, G, H) or anti-myc epitope antibody (C, F, I). Extended focus confocal images were collected at basal surface of cells (A, D, G) and at level of tight junctions (B, E, H). Arrows in B indicate an example of a junction between a pair of transfected cells. Bar, 20 µm.

Cells transfected with RhoN19 did not appear to express at as high a level as those transfected with RhoV14, as judged by the intensity of labeling with the anti-myc antibody (not shown). RhoN19-transfected cells tended to have a peculiar morphology, the base of the cell having an irregular outline, often with long processes that extended between neighboring cells. However, RhoN19 expression did not distinctly alter the pattern of F-actin structures in these cells. Stress fibers were as prominent in RhoN19-expressing cells as in control cells, and the amount of F-actin in the cortical region at cell-cell junctions was not diminished.

Effect of Rho mutants and C3 on ATP-depleted and recovering cells. We examined the effect of ATP depleting LLC-PK cells that had been microinjected with constitutively active RhoV14 or C3 transferase or transfected with RhoV14 or dominant negative RhoN19. Cells were maintained in normal growth medium for 60 min following microinjection or for 48 h following transfection. LLC-PK cells that had been microinjected with RhoV14 or C3 (Fig. 2) or transfected with RhoV14 (Fig. 3) or RhoN19 (not shown) were ATP depleted for 90 min before processing for immunofluorescence. Uninjected or untransfected cells lost all organized stress fibers and focal adhesions. However, cells that had been microinjected or transfected with RhoV14 (constitutively active) still retained organized stress fibers (Figs. 2C and 3D). RhoV14 also increased the degree of stress fiber organization relative to controls in cultures that had been ATP depleted for 90 min and then allowed to recover for 60 min (Fig. 3G).

C3 exoenzyme prevented recovery of the normal pattern of stress fibers in the cultures that were allowed to recover after ATP depletion, whereas uninjected cells in the same culture formed normal stress fibers after 60 min of recovery (Fig. 2F). RhoN19 transfection did not appear to affect stress fiber organization or reorganization following ATP depletion (not shown).

We also examined the effect of RhoV14 transfection on F-actin organization in the perinuclear cytosol and in the cortical region at cell-cell junctions. RhoV14 did not alter the pattern of F-actin accumulation in the cytosol of ATP-depleted cells (Fig. 3, E and H), nor did it protect microvillar actin bundles from disintegration, but it caused a considerable increase in the amount of F-actin at cell-cell junctions, particularly at those between two transfected cells. RhoN19 transfection did not alter the observed pattern of actin cytoskeletal organization at cell-cell junctions or the pattern of cytosolic F-actin in a way that was obvious by inspection of the images (but see the results of quantitative analysis below).

Quantitative analysis of stress fibers and cytosolic and cortical F-actin. We quantified the effects of microinjecting C3 and RhoV14 or transfecting RhoV14 or RhoN19 by measuring the mean fluorescence intensity within defined regions of experimental cells compared with controls in the same cultures. Stress fiber F-actin fluorescence in the basal region of the cell was measured in extended focus images summed from five planes that included all in-focus fluorescence from structures at the base of the cell. F-actin fluorescence in these images was largely attributable to stress fibers and focal adhesions, but there was a significant signal from diffuse fluorescence that filled the spaces between stress fibers. This signal was removed by using image processing to generate a mask that included the stress fibers but excluded diffuse or particulate fluorescence lying between the stress fibers. A typical field before and after processing is illustrated in Fig. 4, E and F. The mean fluorescence intensity within an area that excluded cell-cell junctions was measured (Fig. 4F). Cortical F-actin fluorescence and cytosolic F-actin were measured from extended focus images composed of five planes at a height that included cell-cell junctions. Mean intensities were measured along cell-cell junctions and in a region of the cytosol that excluded the cortical actin.


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Fig. 4.   Measurement of stress fiber formation by quantitative analysis of confocal micrographs. Stress fiber fluorescence was measured in LLC-PK cells under control conditions, after 90-min ATP depletion, or after 90-min ATP depletion followed by 60-min recovery. A-D: mean fluorescence intensities for stress fibers in RhoV14 (constitutively active) microinjection (A), RhoV14 (constitutively active) transfection (B), C3 exoenzyme microinjection (C), and RhoN19 (dominant negative) transfection (D). Controls were untransfected and uninjected cells in same fields as transfected and injected cells. Values are means ± SD derived from 4 separate experiments for each condition; 4-10 cells were analyzed for each experiment. * Comparisons that showed a statistically significant difference (P < 0.01, t-test). Images were processed before measurement to remove non-stress fiber fluorescence. E and F: images before and after processing, respectively. Out-of-focus fluorescence in basal plane was removed by processing, leaving stress fibers relatively unchanged. Typical mean intensity measurements are depicted for a control cell (F, bottom left; arrow in E) and a RhoV14 microinjected cell (F, top right; arrowhead in E). Boxes show mean intensity values associated with marked regions.

Microinjection or transfection of cells growing in normal medium with RhoV14 significantly increased F-actin fluorescence from stress fibers and focal adhesions compared with uninjected controls (Fig. 4, A and B, and Table 1). The difference between cells with and without RhoV14 was even greater in cultures that had been ATP depleted. The ratio between the amount of stress fiber fluorescence in RhoV14-transfected and untransfected cells was 22 in normal medium, but increased to 40 with ATP depletion. When RhoV14 was introduced by microinjection, the ratio was 9.2 in normal medium and increased to 22 with ATP depletion. The increased ratio under ATP-depleted conditions was due to both a decrease in stress fiber fluorescence of controls with ATP depletion (Fig. 4A) and a greater accumulation of stress fibers in RhoV14-expressing cells following ATP depletion. The synergistic effect of ATP depletion and Rho activation was not apparent from simple inspection of the micrographs.

                              
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Table 1.   Quantitative analysis of stress fiber formation

The stress fibers that were observed in RhoV14-containing, ATP-depleted cells could be the residue of excess stress fibers formed during the time before initiation of ATP depletion. However, when we only preincubated the microinjected cultures for 2 min before initiating ATP depletion, injected cells still maintained stress fibers, even though RhoV14 injection does not detectably increase stress fiber levels before 15 min after injection (data not shown). The stress fibers in the RhoV14-injected, ATP-depleted cells must therefore have been maintained or formed during ATP depletion.

C3 microinjection significantly reduced stress fiber fluorescence in microinjected cells compared with uninjected controls when cells were incubated in normal medium, but the difference was less significant when cells were ATP depleted, because the stress fiber fluorescence in control cells was low (Fig. 4C). C3 injection prevented the recovery of stress fiber fluorescence to a significant degree when cells were returned to normal medium following ATP depletion (Fig. 4C). However, RhoN19 transfection did not significantly affect measured stress fiber fluorescence under any of the conditions tested (Fig. 4D)

Transfection with RhoV14 did not significantly alter the amount of F-actin fluorescence in the perinuclear cytosol of control-treated cultures, in contrast to its effect on stress fiber fluorescence (Table 2). RhoV14-transfected, ATP-depleted cells had slightly higher cytosolic F-actin fluorescence than untransfected controls, but in cultures that were allowed to recover following depletion there was no significant difference between control and transfected cells. RhoN19 transfection and C3 microinjection both significantly decreased cytosolic F-actin fluorescence in control and recovered cells.

                              
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Table 2.   Quantitative analysis of cytosolic F-actin

We measured cortical F-actin accumulation at the level of the junctional complex, between two transfected (or microinjected) cells, between transfected and untransfected cells, and between two untransfected cells (Table 3). RhoV14 caused F-actin to accumulate significantly at junctions between two transfected cells and at junctions between transfected and untransfected cells compared with junctions between untransfected cells. This increase was more marked in ATP-depleted cells than in control or recovered cells. The fluorescence intensity ratios between doubly transfected junctions and untransfected junctions was approximately double that between singly transfected junctions and untransfected junctions, indicating that the effect on actin assembly across junctions was probably not cooperative.

                              
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Table 3.   Quantitative analysis of cortical F-actin

Junctions between pairs of cells microinjected with C3 transferase had diminished F-actin fluorescence compared with uninjected pairs (Table 3). This difference was most significant for doubly transfected junctions, and the effect appeared to be enhanced by ATP depletion [compare the ratio of doubly injected junctions to uninjected junctions for control conditions (0.56) and ATP depletion (0.49)]. RhoN19 transfection also caused a significant diminution in cortical actin fluorescence compared with controls.

Effect of RhoV14 microinjection on the time course of stress fiber recovery. To test the ability of RhoV14 to enhance recovery of actin cytoskeletal structures following ATP depletion, we microinjected LLC-PK cells during ATP repletion and compared the rate of recovery of stress fiber fluorescence in microinjected and uninjected cells. LLC-PK cells were ATP depleted for 90 min and then placed in normal medium to allow ATP levels to begin to recover. Cells were microinjected 10 min following initiation of repletion and then allowed to continue repletion for 5, 10, 20, 30, or 60 min, after which times they were processed for immunofluorescence (Fig. 5A). The rate of reassembly of stress fibers was dramatically accelerated in RhoV14 microinjected cells (Fig. 5B). Stress fibers were prominent in these cells by 20 min after microinjection (30 min of repletion), whereas surrounding uninjected cells had not reassembled any stress fibers after 30 or 40 min of repletion.


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Fig. 5.   A: effect of RhoV14 on time course of stress fiber reassembly during ATP repletion. LLC-PK cells were ATP depleted for 90 min and then returned to normal medium. Cells (outlined on micrographs) were microinjected 10 min into repletion and allowed to continue recovery for 5, 10, 20, 30, and 60 min before fixation and rhodamine phalloidin labeling. Micrographs are labeled with total time of ATP repletion. Bar, 20 µm. B: quantitative analysis of experiment shown in A. Data are means ± SD of 15-20 measurements from 3 independent experiments. Circles, uninjected cells; squares, RhoV14 microinjected.

Effect of RhoV14 and ATP depletion on focal adhesions. We used monoclonal antibodies against vinculin and paxillin to examine the effect of Rho signaling and ATP depletion on these focal adhesion proteins. As expected, transfection of RhoV14 into cells that were maintained in normal growth medium resulted in increased stress fiber formation and augmented assembly of vinculin and paxillin into focal adhesions (Fig. 6, A and B). ATP depletion disrupted focal adhesions as well as stress fibers, resulting in a diffuse labeling of cells with anti-vinculin or anti-paxillin. Cells that expressed significant levels of RhoV14 maintained stress fibers after ATP depletion, but neither paxillin nor vinculin was localized to structures at the ends of these stress fibers (Fig. 6, C and D). After recovery following ATP depletion, vinculin and paxillin were detected at stress fiber ends, and expression of RhoV14 resulted in increased labeling of focal adhesions with these antibodies (Fig. 6, E and F). Similar results were obtained in experiments in which RhoV14 was microinjected (not shown), except that in a few cases we observed some


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Fig. 6.   Effect of RhoV14 expression and ATP depletion on assembly of focal adhesion proteins vinculin and paxillin. LLC-PK cells were transfected with RhoV14. At 48 h following transfection, cells were control treated for 90 min (A, B), ATP-depleted (C, D), or ATP-depleted for 90 min and allowed to recover for 60 min (E, F). Cells were fixed, permeabilized, and labeled for F-actin with rhodamine-phalloidin (pseudocolored red), mouse monoclonal antibodies (pseudocolored green) directed against paxillin (A, C, E) or against vinculin (B, D, F), and chicken anti-myc epitope antibody (insets) to detect RhoV14-expressing cells. Under control conditions, paxillin and vinculin colocalize with F-actin at focal adhesions, producing a yellow color. Although RhoV14-injected cells maintained prominent stress fibers with ATP depletion, neither vinculin nor paxillin was assembled into focal adhesions. Bars, 20 µm.

accumulation of vinculin at stress fiber ends in ATP-depleted cultures, but paxillin was never detected at stress fiber ends. Microinjection of C3 exoenzyme resulted in disassembly of focal adhesions as well as stress fibers and prevented reassembly in cells recovering from ATP depletion (not shown).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Ischemic renal injury affects particularly the cells lining the proximal tubule. Disruption of the normal architecture of the actin cytoskeleton contributes to physiological alterations in renal functions that ensue from injury. To understand the role that signaling pathways play in actin alterations induced by ischemic injury, we used chemical ATP depletion in the porcine proximal tubule cell line LLC-PK (11) and investigated Rho GTPase signaling in view of its previously established role in regulating actin (21). We used microinjection and transient transfection to introduce constitutively active and dominant negative mutants of Rho and the Rho inhibitor C3 exoenzyme into LLC-PK cells. We show that constitutively active RhoV14 prevented the disruption of actin stress fibers that otherwise results from ATP depletion and also enhanced the rate of stress fiber reassembly in recovering cells. RhoV14 also enhanced cortical F-actin accumulation at cell-cell junctions even under ATP-depleted conditions. The effect was specific for stress fibers and cortical F-actin, because Rho signaling did not affect cytosolic F-actin or microvillar actin bundles. Inactivation of Rho by microinjection of C3 exoenzyme demonstrated that Rho is required for re-formation of stress fibers and cortical actin as cells recover from ATP depletion. Our results are consistent with the hypothesis that ischemia or ATP depletion inactivates Rho and that reactivation is required for recovery of the full complement of normal actin structures.

Transfection with RhoN19 affected cortical F-actin assembly and cytosolic F-actin but did not alter stress fiber formation (although the relatively low level of stress fiber fluorescence in all cells examined is noteworthy, perhaps indicating that RhoN19 transfection even at levels below the threshold of the confocal images had some effect), whereas C3 exoenzyme caused stress fiber disassembly and prevented their re-formation. RhoN19 transfection levels were generally lower than were those for RhoV14, and the dominant negative mutant is not as potent an inhibitor of endogenous Rho activity as is C3 (compare Ref. 27 with Ref. 42). We used a long period of ATP depletion (90 min) in all our experiments, and it is possible that RhoN19 might affect the time course of stress fiber reassembly or their reassembly following shorter periods of ATP depletion. It is important to note that in this experiment quantitative comparisons of uninjected or untransfected cells between control, ATP-depleted, and recovering cells cannot be made. At the low levels of stress fiber fluorescence in these cells, there is significant variation between coverslips and between experiments.

Quantitative analysis of stress fiber formation revealed an interesting synergistic effect of ATP depletion and Rho activation that was not obvious merely from inspection of the micrographs. Although under normal growth conditions microinjection or transfection of RhoV14 caused stress fiber formation, the increase was far greater when cultures were ATP depleted. Control cells in the ATP-depleted cultures had very low stress fiber fluorescence, which contributed to the high fluorescence ratio between RhoV14-containing and control cells, but the absolute value for stress fiber fluorescence of RhoV14-containing cells was consistently higher following ATP depletion than under control conditions. ATP depletion has been shown to result in net conversion of G-actin to F-actin (25, 38), and the presence of constitutively active RhoV14 apparently causes much of this excess F-actin to assemble into stress fibers. This suggests that ATP depletion separately affects actin polymerization and stress fiber formation and that Rho activation selectively overcomes only the effect on stress fiber formation and cortical F-actin accumulation. The mechanism by which ATP depletion leads to excess actin polymerization might then involve a direct biochemical effect of excess ADP-actin on the equilibrium between sequestered and unsequestered actin monomers. Actin itself binds ATP or ADP, and the affinity of actin binding proteins for actin differs, sometimes substantially, depending on the state of the bound nucleotide (12, 23, 36, 44). A shift in the cellular concentrations of ATP- and ADP-bound actin may directly cause unregulated polymerization by perturbing the equilibrium between the sequestered and unsequestered pools of actin monomers. An alternative mechanism might involve another signaling pathway, perhaps involving other Rho family GTPases. Rac activation, for example, can induce actin polymerization, whereas Rho activation typically does not (35).

Microinjection of RhoV14 significantly enhanced the rate of stress fiber reassembly when the RhoV14 was injected during ATP repletion. This effect was observed even at time points very shortly after injection, indicating that activated Rho can rapidly stimulate the signal transduction machinery necessary for stress fiber formation. Moreover, RhoV14-injected cells maintained significantly more stress fibers than control cells under ATP-depleted conditions, even when ATP depletion was initiated as little as 2 min following microinjection. Therefore, our results indicate that constitutively active Rho not only maintains stress fibers when ATP levels are low but also causes additional stress fiber assembly during ATP depletion. We attempted to microinject LLC-PK cells with Rho during the actual period of ATP depletion. However, even very early in the time course of depletion (2-5 min) penetration of the cells apparently caused irreversible damage to the plasma membrane, so that the cellular contents (and the fluorescent dextran marker) leaked out. We suspect that microinjection of Rho into cells that had been depleted of ATP for long periods would not result in stress fiber assembly.

One likely model for Rho-dependent stress fiber formation is that the Rho effector Rho kinase (4, 5, 30) activates myosin II (either by suppressing myosin-specific phosphatase activity or by directly phosphorylating the myosin regulatory light chain). The contractile activity of myosin results in stress fiber assembly by collecting actin filaments into bundles (14, 47). It will be interesting to determine whether the pathway through Rho kinase can function under ATP-depleted conditions, when the substrate (ATP) for Rho kinase is at low concentration and kinase activity in general is low (31) or whether Rho maintains stress fibers through an alternative pathway.

Assembly of focal adhesions is normally coupled to stress fiber formation (10). Rho activation under normal conditions increases the number and size of focal adhesions and results in additional accumulation of vinculin and paxillin (35). However, when we examined RhoV14-injected cells following ATP depletion, we were surprised to find that, although the cells contained numerous stress fibers, paxillin and vinculin were not localized to stress fiber ends. Thus when Rho is activated under ATP-depleted conditions, stress fiber formation is uncoupled from normal assembly of focal adhesion components. Perhaps the signaling pathway leading to focal adhesion assembly diverges from that leading to stress fiber assembly at a point downstream of Rho. Indeed, recent work demonstrates distinct requirements for Rho and another small GTPase, ARF1, in stress fiber and focal adhesion assembly (41). The branch leading to focal adhesion assembly might require a high level of kinase activity, whereas that leading to stress fiber assembly does not. Further work will be required to determine how stress fibers are anchored to the membrane in the absence of the normal complement of focal adhesion proteins.

The protective effect of RhoV14 was selective for stress fibers and cortical F-actin. F-actin structures at the apical plasma membrane were not protected against the effects of ATP depletion. This is consistent with results in fibroblasts and epithelial cells, in which Rho (and also Rac and Cdc42) has been shown to affect the organization of a particular subset of the F-actin in the cell (21). Rac or Cdc42 may have specific protective roles for other cytoskeletal assemblies, such as microvilli, that are disrupted by ATP depletion.

Indirect evidence of altered Rho family GTPase signaling in ischemic renal injury was provided previously by analysis of the c-jun NH2-terminal kinase [JNK; also known as stress-activated protein kinase (SAPK)] pathway, which showed a substantial increase in SAPK activity with reperfusion following ischemia (46). Similarly, ribosomal S6 kinase activity is increased during postischemic reflow (3). The SAPK pathway is reportedly downstream of Rac and Cdc42 (15), which also mediate activation of S6 kinase (13). Rho, Rac, and Cdc42 regulate transcriptional activation by serum response factor, which regulates genes whose promoters contain the serum response element (24). Among these are c-fos, which is one of the immediate-early genes activated following ischemic injury (43, 51), suggesting that Rho family GTPases may be involved in this response. Taken together with our studies, this indicates that Rho signaling pathways could integrate rebuilding of normal cellular architecture with activation of stress response mechanisms.

Recent studies have focused on the potential therapeutic effect of growth factors to ameliorate the injury resulting from renal ischemia (reviewed in Refs. 22, 26, 32). HGF has received considerable attention, as it is a potent renotrophic factor. Rho GTPases are important components of signaling pathways downstream of growth factors (56), including HGF, and our work suggests that Rho itself is affected by ischemic injury. Rho signaling is clearly a crucial mechanism in injury and recovery, and development of successful therapeutic approaches will require an understanding of these important proteins.


    ACKNOWLEDGEMENTS

We thank Bill Mokanyk, Matt Muterspaugh, and Paul Brown for technical assistance, Ken Dunn for expert help with quantitative image analysis, and Jim Marrs, Bob Bacallao, Mark Wagner, Lawrence Quilliam, Fred Pavalko, and Shobha Gopalakrishnan for critical reading of the manuscript and helpful discussion. We are particularly grateful to Bruce Molitoris for advice, encouragement, and support throughout this work.


    FOOTNOTES

This work was supported in part by the Ralph W. and Grace M. Showalter Research Trust. N. Raman is a postdoctoral fellow of the American Heart Association (Indiana Affiliate).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Address for reprint requests and other correspondence: S. J. Atkinson, Dept. of Medicine, Div. of Nephrology, Indiana University School of Medicine, Fesler Hall 115, 1120 South Dr., Indianapolis, IN 46202 (E-mail: satkinso{at}iupui.edu).

Received 29 January 1999; accepted in final form 11 March 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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