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Am J Physiol Cell Physiol 281: C1858-C1870, 2001;
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Vol. 281, Issue 6, C1858-C1870, December 2001

Characteristics of EYFP-actin and visualization of actin dynamics during ATP depletion and repletion

Stefan Herget-Rosenthal1,2, Melanie Hosford1, Andreas Kribben2, Simon J. Atkinson1, Ruben M. Sandoval1, and Bruce A. Molitoris1

1 Indiana Center for Biological Microscopy, Division of Nephrology, Department of Medicine, Indiana University School of Medicine and Roudebush Veterans Affairs Medical Center, Indianapolis, Indiana 46202; and 2 Division of Nephrology, Department of Medicine, University Hospital Essen, Essen, Germany


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Disruption of the actin cytoskeleton in proximal tubule cells is a key pathophysiological factor in acute renal failure. To investigate dynamic alterations of the actin cytoskeleton in live proximal tubule cells, LLC-PK10 cells were transfected with an enhanced yellow fluorescence protein (EYFP)-actin construct, and a clone with stable EYFP-actin expression was established. Confluent live cells were studied by confocal microscopy under physiological conditions or during ATP depletion of up to 60 min. Immunoblots of stable transfected LLC-PK10 cells confirmed the presence of EYFP-actin, accounting for 5% of total actin. EYFP-actin predominantly incorporated in stress fibers, i.e., cortical and microvillar actin as shown by excellent colocalization with Texas red phalloidin. Homogenous cytosolic distribution of EYFP-actin indicated colocalization with G-actin as well. Beyond previous findings, we observed differential subcellular disassembly of F-actin structures: stress fibers tagged with EYFP-actin underwent rapid and complete disruption, whereas cortical and microvillar actin disassembled at slower rates. In parallel, ATP depletion induced the formation of perinuclear EYFP-actin aggregates that colocalized with F-actin. During ATP depletion the G-actin fraction of EYFP-actin substantially decreased while endogenous and EYFP-F-actin increased. During intracellular ATP repletion, after 30 min of ATP depletion, there was a high degree of agreement between F-actin formation from EYFP-actin and endogenous actin. Our data indicate that EYFP-actin did not alter the characteristics of the endogenous actin cytoskeleton or the morphology of LLC-PK10 cells. Furthermore, EYFP-actin is a suitable probe to study the spatial and temporal dynamics of actin cytoskeleton alterations in live proximal tubule cells during ATP depletion and ATP repletion.

actin cytoskeleton; green fluorescent protein; live imaging; renal proximal tubule cell; enhanced yellow fluorescent protein


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

ISCHEMIA IN VIVO and cellular ATP depletion in vitro severely disrupt the actin cytoskeleton of renal proximal tubule cells (1, 12, 14, 15, 20). Filamentous actin structures within the microvilli, in the meshwork beneath the junctional complexes and in actin stress fibers, are fragmented (1). Disruption of the actin cytoskeleton initiates further structural cellular changes, such as loss of surface membrane polarity and microvilli, membrane destruction, opening of tight junctions, dissociation of junctional complexes, and detachment of cells from the basement membrane (13, 17, 18). These alterations are detrimental to the reabsorbtive and secretory function of proximal tubule cells since their physiological function depends on an intact polarized structure (18, 26). In addition, these cellular alterations have been related to abnormalities in tubuloglomerular feedback, backleak of glomerular filtrate, and tubular obstruction, as mechanisms for decreased glomerular filtration rate and acute renal failure (19, 23). Therefore, disruption of the actin cytoskeleton in proximal tubule cells is a key factor in the pathophysiology of acute renal failure.

Despite the important insights provided by previous studies, the precise dynamics of actin cytoskeletal alterations in proximal tubule cells during ATP depletion and repletion are poorly understood. Previous studies of actin cytoskeletal alterations in proximal tubule cells have predominantly been of static nature, i.e., performed on fixed samples (1, 11, 12, 17, 20). To analyze the spatial and temporal details of highly dynamic, actin cytoskeletal alterations, studies in live cells are of great benefit (2, 9, 25). Dynamic studies with microinjection of fluorescently labeled actin have been limited by the short experimental time due to proteolysis of labeled actin, by mechanical injury to cells, and by the small number of cells microinjected. Fusion of proteins with green fluorescent protein (GFP) has become a useful method to observe proteins in living cells (10, 16). GFP fusion proteins enable direct visualization of protein localization and dynamics in a large number of unimpaired cells in real time (2, 5, 8, 9, 25, 27). However, GFP fusion proteins need to be characterized before use, as they may display nonphysiological properties and may impair the properties of endogenous proteins (27, 29).

Therefore, the purpose of the present study was to evaluate the application of enhanced yellow fluorescent protein (EYFP)-actin fusion protein as a probe for actin in live proximal tubule cells. EYFP is a mutant form of GFP with enhanced fluorescent intensity. We hypothesized that EYFP-actin would colocalize with endogenous F-actin and G-actin, show the characteristics of these actin fractions, and provide visualization of spatial and temporal alterations of different actin cytoskeletal structures during ATP depletion and repletion. Furthermore, EYFP-actin would not interfere with the behavior of the endogenous actin cytoskeleton.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture, reagents, and ATP depletion and repletion. LLC-PK10 cells, a clone of porcine proximal tubule LLC-PK1 cells, were utilized for all studies. LLC-PK10 cells were maintained in 1:1 DMEM/Ham's F-12 medium (GIBCO BRL, Gaithersburg, MD) supplemented with 10% FCS, 100 IU/ml penicillin, 100 µg/ml streptomycin, 14 mM NaHCO3, and 12.5 mM HEPES in a 5% CO2 incubator at 37°C. For live imaging and transfection, cells were plated on poly-D-lysine-coated glass coverslips mounted on culture dishes. For imaging of fixed cells, LLC-PK10 cells were grown on glass coverslips. Experiments were performed on confluent cells. Reagents were from Sigma (St. Louis, MO), unless otherwise indicated. For ATP depletion, LLC-PK10 cells were incubated with substrate-free medium (no glucose, amino acids, pyruvate, FCS, or geneticin) containing 0.1 µM antimycin A (17). To allow ATP repletion, substrate-free medium was replaced with complete medium containing FCS.

Transfection procedures and selection of EYFP-actin-expressing cells. LLC-PK10 cells were plated at 5 × 104 cells/plate and transfected 48 h later with 1 µg of purified plasmid DNA encoding for EYFP or with 1 µg of plasmid DNA encoding for EYFP-tagged beta -actin (both from Clontech, Palo Alto, CA), mixed with 2 µg Novafector reagent (Venn Nova, Pompano Beach, FL) per dish in 200 µl of FCS-free DMEM. EYFP is linked via its COOH-terminal end to the NH2-terminal end of actin with a seven-amino-acid linker. Cells were incubated with the DNA-Novafector mixture for 6 h at 37°C, and then complete medium was added. A cell population entirely expressing EYFP-actin was selected with medium containing geneticin (200 µg/ml; GIBCO BRL). Two consecutive selection steps were performed, and EYFP-actin expression was confirmed by fluorescence microscopy. This cell population was maintained in geneticin-containing medium. Transfection procedures for transient transfections were performed similarly. Experiments were performed 24-96 h after transfection.

Fluorescent staining. Cells were fixed in 4% paraformaldehyde in PBS overnight at 4°C, permeabilized in 0.1% Triton X-100 in PBS for 10 min, and blocked in PBS containing 2% BSA for 1 h at room temperature. F-actin was labeled with 0.1 µg/ml Texas red or Alexa-647-conjugated phalloidin (Molecular Probes, Eugene, OR) for 1 h at room temperature. G-actin was labeled with the G-actin-specific monoclonal mouse antibody JLA-20 (1:100; obtained from Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA) (4) for 1 h at room temperature. This was followed by a 1-h incubation with Cy-5 conjugated goat anti-mouse IgM (Jackson Immuno Research, West Grove, PA). The samples were mounted in 50% glycerol/PBS with 100 mg/ml 1,4-diamino-bicyclo[2,2,2]octane (Sigma).

Immunofluorescence microscopy. Images were collected with an MRC-1024 laser scanning confocal microscope (Bio-Rad, Hercules, CA) on a Nikon Diaphot 200 inverted stand using ×40 numerical aperture (NA) 1.3 or ×100 NA 1.4 oil-immersion objectives (Nikon, Melville, NY). During live studies temperature was kept constant at 37°C with a warm stage and pH 7.4 was maintained by gassing with 5% CO2. To avoid possible spectral overlap, all signals were excited and acquired sequentially. Through focus, optical series were collected from entire cell volumes with separations of 0.2-0.4 µm between focal planes. Images were processed with Metamorph 4.01 imaging software (Universal Imaging, West Chester, PA). We quantified stress fibers in live cells under physiological conditions and during ATP depletion by measuring the mean EYFP-actin fluorescence intensity from representative stress fibers, summed from four basal planes, according to an established protocol (21). Background fluorescence from cytoplasmic actin was subtracted from EYFP-actin fluorescence. A threshold function was applied to the resulting image and converted to a binary mask. An erode function was applied to remove small particulate structures while retaining 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 nonstress fiber fluorescence was removed. For all consecutive measurements during one time sequence, the fluorescence intensity was determined within the same area of the respective stress fiber.

Microscopic colocalization of EYFP-actin and Alexa-647 phalloidin. Corresponding images from EYFP-actin and Alexa-647 phalloidin acquired simultaneously were processed using Metamorph software v 4.1 (Universal Imaging). Because the question of colocalization of the two signals was the main issue to be addressed, the images were first enhanced in contrast and brightness using the "autoenhance" function. Next, a 3 × 3 low-pass filter was applied. The average intensity of the EYFP signal was calculated. The Alexa-647 phalloidin image was then subtracted from the EYFP-actin, and the average intensity from the subtracted image was calculated. Background readings were taken from several subconfluent areas, and the values were averaged and then subtracted from both average intensity readings to yield a background corrected value. For each corresponding image, intensity was normalized to the EYFP-actin fluorescence. A value for percent colocalization was derived by subtracting the residual intensity from the EYFP-actin intensity.

SDS-PAGE and immunoblotting. To recover cell homogenate for total actin, cells were extracted in hot SDS buffer (1% SDS, 10 mM Tris, pH 7.5, 2 mM EDTA), and lysates were boiled for 3 min and sonicated. To recover supernatant samples for G-actin, cells were extracted with a PBS extraction buffer containing 0.1% Triton X-100, 10 mM EDTA, 2 mM MgSO4, 0.5 mM phenylmethylsulfonyl fluoride, 0.1 mM dithiothreitol, and 10 µg/ml each of chymostatin, leupeptin, aprotinin, and pepstatin A for 30 s on ice. Cells were scraped and lysates centrifuged for 15 min at 12,000 g at 4°C. Samples were resuspended in equal volumes of 2× sample buffer (4% SDS, 20% glycerol, 10% beta -mercaptoethanol, 0.5 M Tris, pH 6.8, containing bromophenol blue) and immediately frozen at -20°C. We measured total protein concentrations by bicinchoninic acid assay (Pierce, Rockford, IL). Equal quantities of total protein were loaded on each lane, and proteins were electrophoretically separated on 14% polyacrylamide gels. Proteins were transferred to polyvinylidene difluoride membranes (Millipore, Bedford, MA) in a buffer containing 10% methanol, 0.1% SDS, 40 mM glycine, and 120 mM Tris, pH 8.2. Membranes were blocked in wash buffer (0.1 M NaCl, 0.01 M Tris, 0.05% Tween 20, pH 7.4) containing 10% newborn calf serum at 4°C overnight and incubated with the monoclonal panactin antibody C4 (1:2,000; Roche, Indianapolis, IN) or a monoclonal GFP antibody (1:3,000; BABCO, Richmond, CA) for 2 h at room temperature. Incubation with horseradish perioxidase-conjugated goat anti-mouse IgG (1:40,000; Southern Biotechnology, Birmingham, AL) for 1 h at room temperature followed. The antigen-antibody complexes were detected with enhanced chemiluminescence (Pierce) and exposed to film (Eastman Kodak, Rochester, NY). For quantification, films were scanned using a Silverscanner III (LaVie, Beaverton, OR) and analyzed using Bio Image Intelligent Quantifier software (BI Systems, Ann Arbor, MI). The concentration of endogenous actin concentration in supernatant and homogenate was determined by densitometrically quantifying the respective bands and comparing them to actin standards. The concentration of EYFP-actin in supernatant and homogenate was approximated by densitometrically quantifying the respective bands and comparing them to rGFP standards (Clontech).

F-actin determination. F-actin content was determined in confluent cells grown on 96-well plates and fixed and stained with 0.1 mg/ml tetramethylrhodamine isothiocyanate (TRITC)-phalloidin and 50 µg/ml 4',6-diamidino-2-phenylindole (DAPI; procedures as above) (3). TRITC fluorescence intensity was measured on the Cytofluor II fluorescence plate reader (PerSeptive Biosystems, Framingham, MA). TRITC fluorescence intensity was corrected for cell number by division by DAPI fluorescence intensity of the same sample.

Statistics. Data are presented as means ± SD. Results are expressed either as absolute values or as percent of the control levels. A minimum of four values were collected for each condition in each experiment. Differences between groups were evaluated using ANOVA, and significance was defined as P < 0.05.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

EYFP-actin expression and labeling of the actin cytoskeleton in LLC-PK10 cells. EYFP-actin expression was initially detectable 8 h after transfection of LLC-PK10 cells with pEYFP-actin. We obtained maximum expression of EYFP-actin, monitored by fluorescence microscopy, 24-96 h after transfection. Under physiological conditions, distinct fluorescent structures were present in the cytoplasm but not in the nucleus of LLC-PK10 cells (Fig. 1, A and B), and the fluorescence intensity differed between EYFP-actin-expressing cells. We noted less difference of EYFP-actin expression levels in cells with stable EYFP-actin expression (Fig. 1B). In contrast to the fluorescence pattern of EYFP-actin, LLC-PK10 cells transfected with pEYFP showed a more homogenous distribution of EYFP (Fig. 1C). EYFP was present in all cell compartments, including the nucleus but excluding some vesicular cytoplasmic structures, while filamentous actin was not labeled by EYFP.


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  		Fig. 1.   Localization of enhanced yellow fluorescent protein (EYFP)-actin or EYFP in live LLC-PK10 cells under physiological conditions by fluorescence microscopy. Emission from EYFP-actin expressed in transient transfected LLC-PK10 cells (A) and in cells of the stably transfected LLC-PK10 subclone (B) is shown. Approximately only 10% of the transient transfected cells expressed EYFP-actin, whereas EYFP-actin was expressed by nearly all cells of the LLC-PK10 subclone. EYFP-actin was distributed in the entire cell excluding the nucleus. Emission from EYFP expressed in transient transfected LLC-PK10 cells is shown in C. EYFP was distributed over the entire cell including the nucleus. Bars in A and B = 30 µm; bar in C = 20 µm.

Imaging at higher magnification revealed that EYFP-actin was primarily located to stress fibers, to cortical actin, and to the actin core within microvilli (Fig. 2, A-C). As shown in middle (Fig. 2B) and apical planes (Fig. 2C) of LLC-PK10 cells, a less intense fluorescence signal from EYFP-actin was present in the cytoplasm. This cytoplasmic distribution pattern of EYFP-actin was diffuse and almost homogenous.


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  		Fig. 2.   Localization of EYFP-actin in stable transfected live LLC-PK10 cells under physiological conditions. The localization of EYFP-actin is shown in 3 representative focal planes of the same image field: basal (A), middle (B), and apical (C). Note predominant labeling of EYFP-actin of stress fibers (A, arrowheads), of cortical actin (B), and microvilli (C). In addition, a homogenous cytoplasmic fluorescence signal was present that resembled the distribution of G-actin. Again the nucleus showed no labeling with EYFP-actin. Bars = 10 µm.

To confirm these observations, we compared the localization of EYFP-actin to the localization of G- and F-actin. On basal planes EYFP-actin colocalized predominantly with F-actin (Fig. 3, A and C). In apical planes, the close colocalization of EYFP-actin and F-actin was also present, as seen with cortical actin and microvillar actin (Fig. 3, B and D). The diffuse fluorescence distribution of EYFP-actin resembled the pattern of endogenous G-actin. The cytoplasmic localization of EYFP-actin and G-actin was similar in middle and apical planes (Fig. 3, D and F). Somewhat different from G-actin, EYFP-actin demonstrated a more homogenous cytoplasmic distribution. The exclusion of EYFP-actin fluorescence from the nucleus was in agreement with both the G- and the F-actin staining. The expression of EYFP-actin did not appear to affect the morphology of LLC-PK10 cells, and the incorporation of EYFP-actin into the cytoskeleton did not alter the organization or distribution of the actin cytoskeleton compared with endogenous F-actin or G-actin in nontransfected cells (data not shown; actin not labeled with EYFP is referred to as endogenous actin).


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  		Fig. 3.   Localization of EYFP-actin (A, B), F-actin (C, D), and G-actin (E, F) in confluent LLC-PK10 cells under physiological conditions on representative basal (A, C, E) and apical focal plane (B, D, F). Colocalization of EYFP-actin with F-actin in stress fibers (A and C, arrows). Apically, EYFP-actin colocalized with F-actin in microvilli (B and D, arrowheads) and cortical F-actin (arrow, D), and there was cytoplasmic colocalization of EYFP-actin with G-actin (B and F). Bars = 20 µm.

The content of total and globular endogenous actin and of total and globular EYFP-actin was estimated by immunobloting techniques in nontransfected cells, transiently transfected cells, and cells with stably transfected EYFP-actin under physiological conditions (Fig. 4 and Table 1). Total endogenous actin and total EYFP-actin were determined from cell homogenates and from endogenous and EYFP-labeled G-actin from cell supernatants. When quantified, nontransfected cells, transiently transfected cells, and cells with stable expression of EYFP-actin did not significantly differ in total endogenous actin. The difference of endogenous G-actin between nontransfected and stable transfected cells was significant (P < 0.05), but no difference was noted between nontransfected and transiently transfected cells (Table 1). The monoclonal antibody against G-actin did not detect EYFP-actin; therefore, an antibody to GFP was used to quantify GFP-actin. As expected, stable transfected cells expressed the greatest amount of total EYFP-actin and the G-actin fraction from EYFP-actin. Markedly smaller amounts were expressed in transiently transfected cells, as determined by immunoblot analysis with a monoclonal antibody against GFP. Total EYFP-actin accounted for ~4% of total cellular actin in cells with stable expression of EYFP-actin. A ratio of ~2:1 was observed for endogenous F-actin to G-actin in nontransfected cells and in transiently and stable transfected cells. A similar ratio was found for F-actin to G-actin fractions of EYFP-actin in transient and stable transfected cells expressing EYFP-actin.


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  		Fig. 4.   Immunoblot of endogenous actin and EYFP-actin in LLC-PK10 cells. Ten micrograms of total protein from the supernatant of Triton X-100-solubilized cells or total cell homogenate from nontransfected cells, transient transfected cells, and cells of the stable subclone expressing EYFP-actin were separated by SDS-PAGE, transferred to polyvinylidene difluoride (PVDF) membranes, and immunoblotted to detect actin or EYFP-actin as described in METHODS. Expression of EYFP-actin reduced total cellular endogenous actin and G-actin in the supernatant in transient transfected cells. This suppression was more marked in cells of the stable subclone expressing EYFP-actin compared with nontransfected cells. Cells of the stable subclone showed higher EYFP-actin than transient transfected cells.


                              
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  		Table 1.   Endogenous actin and EYFP-actin concentrations under physiological conditions

Effect of ATP depletion and repletion on EYFP-actin and endogenous actin. The effect of various time periods of ATP depletion and repletion on EYFP-actin and endogenous actin was evaluated in LLC-PK10 cells with stable expression of EYFP-actin. These cells maintained constant amounts of both total EYFP-actin and total endogenous actin during ATP depletion as well as ATP repletion, compared with controls (Fig. 5, A and B). ATP depletion resulted in a marked decrease of the G-actin fraction of EYFP-actin (Fig. 6A). The largest decrease occurred during the first 5 min of ATP depletion to levels significantly lower than control levels (P < 0.01). The G-actin fraction of EYFP-actin remained at low levels throughout 60 min of ATP depletion. After 4 h of ATP repletion the amount of unpolymerized EYFP-actin had slightly increased again. Twenty-four hours of ATP repletion were required for the G-actin fraction of EYFP-actin to return to baseline levels. As demonstrated in Fig. 6B, ATP depletion resulted in a substantial increase of the combined F-actin fractions of EYFP-actin and endogenous actin in LLC-PK10 cells. F-actin had increased significantly after 15 min (P < 0.05) and 60 min of ATP depletion (P < 0.01). However, after 4 h of ATP repletion, F-actin levels were still significantly higher compared with control levels (P < 0.05). By 24 h of ATP repletion, the combined F-actin fractions of EYFP-actin and endogenous actin had returned to the F-actin level found during physiological conditions.


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  		Fig. 5.   Effect of ATP depletion and repletion on total endogenous actin and total EYFP-actin. A: total cell homogenates from stable transfected LLC-PK10 cells were separated by SDS-PAGE, transferred to PVDF membranes, and immunoblotted to detect actin or EYFP-actin as described in METHODS. These 2 immunoblots are representative of 6 separate experiments. B: total endogenous actin and total EYFP-actin levels from total cell homogenate of stably transfected LLC-PK10 cells were quantitated densitometrically from immunoblots. Values are expressed as a percentage of the control values and are means ± SD of 6 separate experiments. Samples were taken under physiological conditions (control), after 5, 15, 30, and 60 min of ATP depletion, and 4 and 24 h after ATP repletion succeeding 60 min of ATP depletion. As demonstrated in A and B, total endogenous actin and total EYFP-actin levels were unaltered by ATP depletion and ATP repletion compared with physiological conditions.



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  		Fig. 6.   Effect of ATP depletion and repletion on G-actin fraction of EYFP-actin and total endogenous and EYFP-labeled F-actin. A: determination of G-actin fraction of EYFP-actin from supernatant of Triton X-100 solubilized, stably transfected LLC-PK10 cells by immunoblot (see METHODS) during ATP depletion and repletion. B: quantification of total endogenous and EYFP-labeled F-actin in stably transfected LLC-PK10 cells. Samples in A and B were measured under physiological conditions (control), after 5, 15, 30, and 60 min of ATP depletion, and 4 and 24 h after ATP repletion succeeding 60 min of ATP depletion. Values in A and B are expressed as a percentage of the control values. Values are means ± SD of 8 (A) or 16 (B) separate experiments. *P < 0.05 and ** P < 0.01 compared with control groups.

Dynamics of EYFP-actin during ATP depletion. To visualize the dynamics of actin cytoskeletal alterations, we observed live LLC-PK10 cells with stable expression of EYFP-actin over a period of 60 min with or without prior ATP depletion. Because alterations of the actin cytoskeleton were most prominent in stress fibers, basal images were obtained, and the fluorescence intensity of stress fibers were quantitatively analyzed. The time sequence in Fig. 7, A-D, provides representative images of the alterations of stress fibers. After 15 min of ATP depletion (Fig. 7B), the fluorescence intensity of stress fibers with EYFP-actin was markedly reduced and the stress fibers were severely disrupted. Disruption of stress fibers became more pronounced after 30 min of ATP depletion (Fig. 7C). By 60 min of ATP depletion (Fig. 7D), stress fibers had almost completely disintegrated and could hardly be detected by immunofluorescence microscopy. Control cells showed no alterations of stress fibers under physiological conditions during 60 min of repeated observations (data not shown).


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  		Fig. 7.   Dynamics of stress fiber alterations during ATP depletion. Representative basal images from the same live, stably transfected LLC-PK10 cells under physiological conditions (A), and after 15 min (B), 30 min (C), and 60 min (D) of ATP depletion. Images were taken from one basal focal plane, maintaining the same microscopy and laser settings. Note the marked disruption of stress fibers labeled with EYFP-actin, which was present at 15 min and proceeded over the entire time sequence. Bars = 10 µm. Alterations of stress fibers were quantified from their fluorescence on confocal micrographs (E). Fluorescence of the same individual stress fibers was measured in live, stably transfected LLC-PK10 cells at 0, 15, 30, and 60 min under physiological conditions (control), and during ATP depletion. Values are means ± SD of 120 (ATP depletion) or 100 (control) separate stress fibers imaged over the same time sequence. *P < 0.05 and **P < 0.01 compared with the control group.

To quantify these changes, the mean EYFP-actin fluorescence intensity of stress fibers was consecutively measured within the same defined cell regions over a 60-min time course (Fig. 7E). The fluorescence intensity of stress fibers decreased substantially during ATP depletion. The most pronounced decrease of stress fiber fluorescence occurred during the first 15 min of ATP depletion. In the next consecutive 45-min interval, fluorescence intensity diminished further but at a slower rate than before. In parallel to qualitative fluorescence image analysis, control cells without ATP depletion exhibited no marked change in stress fiber fluorescence intensity during the 60 min of observation (Fig. 7E).

Besides disruption of stress fibers and cortical and microvillar actin, multiple EYFP-actin aggregates accumulated in live LLC-PK10 cells during ATP depletion (Fig. 8, A and B). These aggregates formed a globular, perinuclear pattern and colocalized with F-actin stained with Texas red phalloidin. No characteristic F-actin structures, such as stress fibers or microvillar bundles, were visible in the aggregates. We initially observed perinuclear F-actin after as little as 10 min of ATP depletion. The size and the number of aggregates increased continuously to a maximum at 60 min of ATP depletion.


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  		Fig. 8.   Dynamics of actin cytoskeleton alterations during ATP depletion. A and B: basal images from the same live, stably transfected LLC-PK10 cells under physiological conditions (A) and after 30 min of ATP depletion (B). Besides the disruption of stress fibers, EYFP-actin was incorporated into multiple, cytosolic aggregates (arrowheads) formed during ATP depletion. These aggregates were widely distributed in the cells and demonstrated high fluorescence intensity. C and D: apical images from live, stably transfected LLC-PK10 cells under physiological conditions (C) and after 60 min of ATP depletion (D). The microvillar actin core (arrows) was lost during ATP depletion. Images were taken from the same basal (A and B) or apical (C and D) focal plane, respectively, maintaining the same microscopy and laser settings. Bars = 10 µm.

Little alteration of microvillar and cortical actin was present during this early time period (15 min) of ATP depletion. Apical planes of EYFP-actin labeling in the microvilli core were present with only moderate decreases in fluorescence intensity for 45 min of ATP depletion (Fig. 8C). However, we hardly detected any microvillar EYFP-actin after 60 min of ATP depletion, and no actin was detected. Only a weak homogeneous cytoplasmic distribution of EYFP-actin remained on apical planes in the former location of the actin microvillar cores (Fig. 8D). In addition, the apical cell membrane of these cells extended outward but was still intact after 60 min ATP depletion. In parallel to microvillar F-actin, the fluorescence intensity of EYFP-actin incorporated in cortical actin diminished later compared with that of stress fibers. We observed a decrease of cortical actin only after 60 min of ATP depletion. Unlike stress fiber and microvillar actin, the fluorescence intensity of cortical EYFP-actin never decreased substantially.

Microscopic colocalization of EYFP-actin and Alexa-647 phalloidin. To address the issue of utilization of EYFP-actin as a marker of endogenous actin, colocalization studies were conducted in LLC-PK10 cells with stable expression of EYFP-actin using Alexa-647 phalloidin, a far-red-emitting fluorophore. The use of the Alexa-647 fluorophore enabled the simultaneous acquisition of EYFP-actin and phalloidin signals without the possibility of spectral emission overlap. Cells stained with Alexa-647 phalloidin after 30 min of ATP depletion (Fig. 9A) showed excellent correlation with the corresponding EYFP-actin signal (Fig. 9B). There was ~83 ± 13.0% colocalization of EYFP-actin and filamentous actin structures, occurring as either stress fibers or actin aggregates (arrows in Fig. 9, A and B). The number of aggregates previously seen in abundance during 30 min of depletion, are greatly reduced after 4 h of ATP repletion in both the EYFP-actin (Fig. 9C) and Alexa-647-phalloidin (Fig. 9D) channels. More filamentous actin was seen in both channels as either stress fibers (arrows) or cortical actin (arrowheads in Fig. 9, C and D). The average colocalization between the two channels falls to ~74 ± 16.5% after the 4 h of ATP repletion. The reduction in colocalization, compared with controls, is in part due to our inability to factor out EYFP-G-actin from filamentous actin.


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  		Fig. 9.   Colocalization of EYFP-actin with Alexa-647 phalloidin during ATP depletion and repletion. Basal images from stably transfected LLC-PK10 cells after 30 min of ATP depletion showing EYFP-actin (A) and Alexa-647 phalloidin (B) fluorescence. Quantitative colocalization yielded an overlap value of 83 ± 13.0% between the 2 signals at 30 min of ATP depletion. EYFP-actin aggregates (A, open arrow) show good colocalization with the signal from filamentous actin stained with phalloidin (B, open arrow). A general breakdown of the stress fibers and cortical actin was seen in both photomicrographs. In cells allowed to recover for 4 h after ATP depletion, both EYFP-actin (C) and filamentous actin fluorescence (D) exhibited reformation of both stress fibers (thin arrows) and the cortical actin network (arrowheads). Colocalization values for the EYFP-actin and phalloidin channels for this group was 74 ± 16.5%. Bars = 10 µm.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Our study demonstrates that the fusion protein EYFP-actin expressed in LLC-PK10 cells exhibits major structural and functional characteristics similar to endogenous F-actin and G-actin. EYFP-actin incorporated well into the endogenous actin cytoskeleton and did not disturb its characteristic structural properties. Our quantitative and qualitative data indicate that EYFP-actin is a suitable probe for the actin cytoskeleton in live proximal tubule cells. Compared with previous studies with cells fixed and stained for actin, or microinjected with fluorescently labeled actin, we present more detailed data of spatial and temporal dynamics of actin cytoskeletal alterations in proximal tubule cells during ATP depletion and repletion over an extended observation period. The actin cytoskeleton in epithelial cells is a highly dynamic structure that undergoes rapid and frequent alterations (7, 19, 30). Ischemia and reperfusion are potent causes of actin cytoskeletal alterations in proximal tubule cells (1, 13, 14, 20-22). LLC-PK10 cells are a valid model to study the actin cytoskeleton in proximal tubule cells (1, 6, 11, 20, 24) with ischemia mimicked by antimycin A-induced ATP depletion (17).

EYFP-actin predominantly colocalized with F-actin in LLC-PK10 cells and incorporated especially into microvilli, stress fibers, and the cortical actin ring. This result was in agreement with recent studies showing the distribution of GFP-actin in other mammalian cells (2, 5, 9, 25). Besides labeling F-actin structures, EYFP-actin also localized diffusely in the cytoplasm. This diffuse cytoplasmic pattern of EYFP-actin resembled the distribution of endogenous G-actin and presumably represents the distribution of EYFP-actin as monomers (4). In contrast, EYFP alone was not associated with cytoskeletal structures but distributed diffusely in the cytoplasm and nucleus of LLC-PK10 cells. This indicates that EYFP itself does not specifically label actin cytoskeletal structures.

In the subpopulation of LLC-PK10 cells with stable EYFP-actin expression, EYFP-actin accounted for ~4% of total intracellular actin. This result was in agreement with the amount of 5-6% GFP-actin of total actin, which has been reported recently for other cells (5, 27). Besides qualitative characteristics of F- and G-actin fractions, EYFP-actin also demonstrated quantitative characteristics of both endogenous actin fractions. Under physiological conditions, the distribution between endogenous F-actin and G-actin in epithelial cells was tightly regulated, with a ratio of ~2:1 (F-actin to G-actin) (19). This ratio was reflected correctly by the F- to G-actin ratio of EYFP-actin in our study. In LLC-PK10 cells expressing EYFP-actin, the ratio of endogenous F-actin to G-actin of 2:1 was also preserved under physiological conditions. Additionally, total endogenous actin concentrations did not markedly differ between nontransfected LLC-PK10 cells and those expressing EYFP-actin.

Although tagging a protein with EYFP may interfere with the protein's structure and/or function (16, 27, 29), EYFP-actin did not seem to impair the endogenous actin cytoskeleton under physiological conditions. No major differences were present between entirely endogenous F-actin structures and F-actin with EYFP-actin incorporated in LLC-PK10 cells. We observed no differences in form, organization, or distribution of the actin cytoskeleton between transfected and nontransfected LLC-PK10 cells, comparing our data with previous actin cytoskeleton studies (13, 18, 20, 21). The overexpression of EYFP-actin did not result in overt cytotoxic effects, since the morphology of transfected and nontransfected LLC-PK10 cells did not differ.

The behavior of EYFP-actin was consistent with known actin cytoskeleton alterations during ATP depletion in live proximal tubule cells (1, 12, 14, 28). However, our results extend previous findings, because we obtained more detailed immunofluorescent analysis of the spatial and temporal dynamics of actin disassembly with EYFP-actin. A short period of ATP depletion caused dramatic disruption of stress fibers in live LLC-PK10 cells, while no changes in microvilli and cortical actin were apparent at this time point. In parallel to further shortening of stress fibers, we observed destruction of microvilli and cortical actin as the time of ATP depletion was extended. After an extended period of ATP depletion, stress fibers and microvilli were disassembled while cortical actin was only moderately affected by disassembly. The perinuclear aggregation of EYFP-actin, observed during ATP depletion, was consistent with previously described F-actin structures (15, 20, 28). Differential subcellular disassembly of F-actin structures may be due to differences in activity of different actin-binding proteins. During ATP depletion, actin-severing proteins are possibly recruited to different subcellular regions. Therefore, differential disassembly of stress fibers and cortical and microvillar actin tagged with EYFP-actin may illustrate spatially and temporally separate regulatory mechanisms of different actin-binding proteins present in proximal tubule cells. EYFP-actin incorporation into cellular F-actin structures during ATP repletion was also consistent with the behavior of endogenous actin, as shown by the high degree of colocalization of EYFP- and phalloidin-labeled F-actin. This was not so apparent for the intracellular aggregates of EYFP-actin during ATP depletion. However, this may relate to excessive actin depolymerizing factor (ADF) binding and inhibition of phalloidin binding (22).

The incorporation of the EYFP-actin into the actin cytoskeleton did not affect characteristics of endogenous actin during ATP depletion and repletion. During ATP depletion, transfected LLC-PK10 cells maintained constant amounts of total endogenous actin and EYFP-actin during ATP depletion and repletion. Meanwhile, the concentration of F-actin and G-actin, as endogenous and EYFP-actin, varied simultaneously. Endogenous F-actin and EYFP-actin in the F-actin state increased, while endogenous G-actin and EYFP-actin in the G-actin state decreased, and these changes reversed during ATP repletion.

The strong emission of EYFP-actin permitted monitoring of the actin cytoskeleton alterations in LLC-PK10 cells for an extended period. Maximal fluorescence intensity of EYFP-actin was observed in LLC-PK10 cells over 48 h, and experiments can be performed at least for that time period. The stable emission of EYFP-actin permits serial excitation without marked quenching of the EYFP-actin fluorescence intensity. Therefore, diminished fluorescence signal intensity indicates true changes of the actin cytoskeleton and seems not to be due to photobleaching. This further underscores the usefulness of EYFP-actin as a marker for actin dynamics.

In summary, our data indicate that EYFP-actin is a suitable probe for actin in live proximal tubule cells. EYFP-actin incorporates into all components of the actin cytoskeleton and demonstrates the characteristics of endogenous F-actin and G-actin without altering the endogenous actin cytoskeleton. EYFP-actin provides detailed information regarding spatial and temporal dynamics of actin cytoskeletal alterations in live proximal tubule cells during ATP depletion and repletion. Furthermore, EYFP-actin enables one to quantify these alterations, with stress fibers undergoing the most rapid and complete disassembly during ATP depletion and rapid reassembly during ATP repletion.


    ACKNOWLEDGEMENTS

The monoclonal antibody JLA-20, developed by Dr. J. J. Lin, was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the National Institute of Child Health and Human Development and maintained by the University of Iowa, Department of Biological Sciences, Iowa City, IA 52242.


    FOOTNOTES

Address for reprint requests and other correspondence: B. A. Molitoris, Division of Nephrology, Indiana Univ. School of Medicine, 1120 South Dr., FH 115, Indianapolis, IN 46202-5116.

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. Section 1734 solely to indicate this fact.

Received 30 April 2001; accepted in final form 24 August 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Bacallao, R, Garfinkel A, Monke S, Zampighi G, and Mandel LJ. ATP depletion: a novel method to study junctional properties in epithelial tissues. J Cell Sci 107: 3301-3313, 1994[Abstract].

2.   Ballestrem, C, Wehrle-aller B, and Imhof BA. Actin dynamics in living mammalian cells. J Cell Sci 111: 1649-1658, 1998[Abstract].

3.   Cano, ML, Lauffenburger DA, and Zigmond SH. Kinetic analysis of F-actin depolymerization in PMF leucocyte lysates indicates that chemoattractant stimulation increases actin filament number without altering filament length distribution. J Cell Biol 115: 677-687, 1991[Abstract/Free Full Text].

4.   Cao, L, Fishkind DJ, and Wang Y. Localization and dynamics of nonfilamentous actin in cultured cells. J Cell Biol 123: 173-181, 1993[Abstract/Free Full Text].

5.   Choidas, A, Jungbluth A, Sechi A, Murphy J, Ullrich A, and Marriott G. The suitability and application of a GFP-actin fusion protein for long-term imaging of the organization and dynamics of the cytoskeleton in mammalian cells. Eur J Cell Biol 77: 81-90, 1998[Web of Science][Medline].

6.   Chuman, L, Fine LG, Cohen AH, and Saier MH. Continuous growth of proximal tubular kidney epithelial cells in hormone-supplemented serum-free medium. J Cell Biol 94: 506-510, 1982[Abstract/Free Full Text].

7.   Cooper, JA, and Schafer DA. Control of actin assembly and disassembly at filament ends. Curr Opin Cell Biol 12: 97-103, 2000[Web of Science][Medline].

8.   Doyle, T, and Botstein D. Movement of yeast cortical actin cytoskeleton visualized in vivo. Proc Natl Acad Sci USA 93: 3886-3891, 1996[Abstract/Free Full Text].

9.   Fischer, M, Kaech S, Knutti D, and Matus A. Rapid actin-based plasticity in dendritic spines. Neuron 20: 847-854, 1998[Web of Science][Medline].

10.   Gerdes, HH, and Kaether C. Green fluorescent protein: applications in cell biology. FEBS Lett 389: 44-47, 1996[Web of Science][Medline].

11.   Golenhofen, N, Doctor RB, Bacallao R, and Mandel LJ. Actin and villin compartmentation during ATP depletion and recovery in renal cultured cells. Kidney Int 48: 1837-1845, 1995[Web of Science][Medline].

12.   Kellerman, PS, and Bogusky RT. Microfilament disruption occurs very early in ischemic proximal tubule cell injury. Kidney Int 42: 896-902, 1992[Web of Science][Medline].

13.   Kellerman, PS, Clark RAF, Hoilien CA, Linas SL, and Molitoris BA. Role of microfilaments in maintenance of proximal tubule structural and functional integrity. Am J Physiol Renal Fluid Electrolyte Physiol 259: F279-F285, 1990[Abstract/Free Full Text].

14.   Kellerman, PS, Norenberg SL, and Jones GM. Early recovery of the actin cytoskeleton during renal ischemic injury in vivo. Am J Kidney Dis 27: 709-714, 1996[Web of Science][Medline].

15.   Kroshian, VM, Sheridan AM, and Lieberthal W. Functional and cytoskeletal changes induced by sublethal injury in proximal tubular epithelial cells. Am J Physiol Renal Fluid Electrolyte Physiol 266: F21-F30, 1994[Abstract/Free Full Text].

16.   Ludin, B, and Matus A. GFP illuminates the cytoskeleton. Trends Cell Biol 8: 72-77, 1998[Web of Science][Medline].

17.   Molitoris, BA, Dahl R, and Hosford M. Cellular ATP depletion induces disruption of the spectrin cytoskeletal network. Am J Physiol Renal Fluid Electrolyte Physiol 271: F790-F798, 1996[Abstract/Free Full Text].

18.   Molitoris, BA. Ischemia-induced loss of epithelial polarity: potential role of the actin cytoskeleton. Am J Physiol Renal Fluid Electrolyte Physiol 260: F769-F778, 1991[Abstract/Free Full Text].

19.   Molitoris, BA. Putting the actin cytoskeleton into perspective: pathophysiology of ischemic alterations. Am J Physiol Renal Physiol 272: F430-F433, 1997[Abstract/Free Full Text].

20.   Molitoris, BA, Geeredes A, and McIntosh JR. Dissociation and redistribution of Na+/K+ ATPase from its surface membrane actin cytoskeletal complex during cellular ATP depletion. J Clin Invest 88: 462-469, 1991.

21.   Raman, N, and Atkinson SJ. Rho controls actin cytoskeletal assembly in renal epithelial cells during ATP depletion and recovery. Am J Physiol Cell Physiol 276: C1312-C1324, 1999[Abstract/Free Full Text].

22.   Schwartz, N, Hosford M, Sandoval RM, Wagner MC, Atkinson SJ, Bamburg J, and Molitoris BA. Ischemia activates actin depolymerizing factor: role in proximal tubule microvillar actin alterations. Am J Physiol Renal Physiol 276: F544-F551, 1999[Abstract/Free Full Text].

23.   Sheridan, A, and Bonyentre JV. Cell biology and molecular mechanisms of injury in ischemic acute renal failure. Curr Opin Nephrol Hypertens 9: 427-434, 2000[Web of Science][Medline].

24.   Venkatachalam, MA, Bernard DB, Donohoe JF, and Levinsky NG. Ischemic damage and repair in the rat proximal tubule: differences among the S1, S2 and S3 segments. Kidney Int 14: 31-49, 1978[Web of Science][Medline].

25.   Verkhusha, VV, Tsukita S, and Oda H. Actin dynamics in lamellipodia of migrating border cells in the Drosophila ovary revealed by a GFP-actin fusion protein. FEBS Lett 445: 395-401, 1999[Web of Science][Medline].

26.   Wagner, MC, and Molitoris BA. ATP depletion alters myosin Ibeta cellular location in LLC-PK1 cells. Am J Physiol Cell Physiol 272: C1680-C1690, 1997[Abstract/Free Full Text].

27.   Westphal, M, Jungbluth A, Heidecker M, Mühlbauer B, Heizer C, Schwartz JM, Marriott G, and Gerisch G. Microfilament dynamics during cell movement and chemotaxis monitored using a GFP-actin fusion protein. Curr Biol 7: 176-183, 1997[Web of Science][Medline].

28.   White, P, Doctor RB, Dahl RH, and Chen J. Coincident microvillar actin bundle disruption and perinuclear actin sequestration in anoxic proximal tubule. Am J Physiol Renal Physiol 278: F886-F893, 2000[Abstract/Free Full Text].

29.   Wittenmayer, N, Rothkegel M, Jockusch BM, and Schluter K. Functional characterization of green fluorescent protein-profilin fusion proteins. Eur J Biochem 267: 5247-5256, 2000[Web of Science][Medline].

30.   Yeaman, C, Grindstaff KK, and Nelson WJ. New perspectives on mechanisms involved in generating epithelial cell polarity. Physiol Rev 79: 73-98, 1999[Abstract/Free Full Text].

Am J Physiol Cell Physiol 281(6):C1858-C1870
0363-6143/01 $5.00 Copyright © 2001 the American Physiological Society


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