Site-Specific Alteration of Actin Assembly Visualized in Living Renal Epithelial Cells during ATP Depletion
Eric A. Shelden*,
Joel M. Weinberg,
Dorothy R. Sorenson*,
Chris A. Edwards* and
Fiona M. Pollock*
*Department of Cell and Developmental Biology, and Department of Internal Medicine, Division of Nephrology, University of Michigan Medical School, Ann Arbor, Michigan.
Correspondence to Dr. Eric A. Shelden, Assistant Professor, Cell and Developmental Biology, University of Michigan Medical School, Ann Arbor, MI 48109-0616. Phone: 734-764-0271; Fax: 734-763-1166;
ABSTRACT. Disruption of normal actin organization in renal tubularepithelial cells is an important element of renal injury inducedby ischemia. Studies of fixed cells indicate that the cytoskeletonis disrupted by both ischemia and ATP depletion in a site-specificmanner. However, few studies have examined these effects inliving cells, and the relationship between the time course ofATP reduction and alteration of the cytoskeleton remains unclear.Here, time-lapse video images of cultured renal epithelial cellsexpressing an enhanced green fluorescent protein (EGFP)-actinfusion protein were obtained, and the kinetics of fluorescenceactin distribution before and during ATP depletion is quantifiedand compared with measured ATP levels. This study found thatassembly of lamellar actin is inhibited rapidly as cellularATP levels are reduced, whereas disruption of actin in stressfibers is more gradual and persistent. Actin associated withfocal adhesions is largely resistant to ATP depletion in theseexperiments, and, consistent with previous studies, particulateaggregates of actin were formed within the cytoplasm of ATP-depletedcells. Most surprisingly, time-lapse imaging of EGFP-actin distribution,quantitative fluorescence imaging of phalloidin-stained cells,and ultrastructural studies indicate that assembly of actinfilaments occurs at sites of epithelial cell-cell attachmentin ATP-depleted cells. This assembly is initiated early duringATP depletion and continues after ATP levels are maximally reduced.Assembly of actin at sites of cell-cell attachment may be anelement of the pathology of injury induced by ischemia, or alternatively,could reflect the function of a protective mechanism. Thesestudies directly demonstrate site-specific alteration of actinassembly in living epithelial cells during ATP depletion. Theresults also reveal that actin reorganization continues afterATP levels are maximally decreased and that epithelial cell-cellattachments are sites of actin assembly in ATP-depleted cells.Email: shelden@umich.edu
Normal epithelial function is dependent on the integrity ofactin cytoskeletal arrays and complexes mediating cell-celland cell substrate attachment. In the kidney, disruption ofthese arrays in renal tubular epithelial cells (RTE) is thoughtto be an important mediator of ischemic acute renal failure(for review, see references 1–3). For example, disruptionof microvillar actin arrays in RTE can be detected within 5min of renal artery occlusion in vivo (4). Dissolution of basalactin filament bundles, or stress fibers, has also been observedin a variety of ATP-depleted cells (5–9), and this mayweaken cell-substrate attachment. In the kidney, it is thoughtthat the loss of cell-substrate attachment during ischemia resultsin the shedding of cells into tubule lumens and contributesto impaired renal function (10–12). Similarly, disruptionof actin filament at sites of cell-cell attachment during ATPdepletion has been described (9,13,14) and may play a role incompromising epithelial barrier function during ischemia. Evidencefrom recent in vitro studies indicates that disruption of actinfilament arrays during ischemia may be mediated by upregulationin ADF/cofilin actin severing activity (15) as well as lossof Rho kinase-mediated assembly of actin at cell-substrate attachments(13) and other elements of cell-cell junctions (16).
Because many actin filament arrays found in cells under controlconditions are disrupted by ATP depletion, it is surprisingthat the total cellular content of filamentous actin (F-actin)in epithelial cells increases during ATP depletion (8,14,17,18).Several groups have shown that actin aggregates appear withinthe cytoplasm of ATP-depleted cells, but the mechanism of theirformation and their significance remain under investigation.Recently, we have also documented that hsp27, a putative stress-activatedactin-associated protein, is recruited to sites of cell-celladhesions during ATP depletion of renal epithelial cells (19).Overexpression of hsp27 has previously been shown to increasestability of actin arrays in cells subject to a variety of injuries(for review, see references 20–22), and previous investigatorshave noted that actin arrays associated with cell junctionsare more resistant to disruption during ATP depletion than thoseat other sites (9,23). These studies generally indicate thatsite-specific alteration of actin stability or assembly occursin ATP-depleted cells and support the hypothesis that preferentialstabilization of actin filaments at epithelial cell junctionsmay be an important aspect of the cellular response to ATP depletion.However, previous studies examining fixed, fluorescently stainedactin arrays have been limited in their ability to address thisissue because they could not correlate the distribution of actinboth before and during ATP depletion in individual cells.
In the present study, we conducted time-lapse fluorescence imagingof a renal epithelial cell line (LLC-PK1) stably expressingan enhanced green fluorescent protein (EGFP)-actin fusion proteinbefore and during ATP depletion and during ATP repletion. Wefind that actin in lamellar protrusions is rapidly disruptedby ATP depletion, whereas disruption of stress fibers occursmore gradually. Actin associated with terminal focal adhesionswas resistant to disruption by ATP depletion. As expected, aggregatesof actin were formed within the cytoplasm of ATP-depleted cells.Most surprisingly, our videos of EGFP-actin in living cellsreveal that fluorescence actin accumulates at sites of epithelialcell-cell attachment in ATP-depleted cells. The time courseof this accumulation correlates well with a quantitative reductionin background cytosolic fluorescence, and presumably actin monomer,quantified here and noted in previous studies (8), suggestingthat the two events may be causally linked. Accumulation offluorescence actin at sites of cell-cell attachment continuedafter ATP levels were maximally reduced. Ultrastructural analysisof cell junctions and association of phalloidin with sites ofcell-cell attachment in fixed cells before and during ATP depletionfurther indicate that the observed accumulation of fluorescenceactin in ATP-depleted cells represents actin filament assembly.
Together, these results reveal that individual types of actinfilament arrays are distinctly altered by ATP depletion. Ourresults demonstrate that the inhibition of actin assembly inlamellae is an early consequence of ATP depletion and that assemblyof actin filament at sites of cell-cell attachment can playa role in the formation of actin polymer in ATP-depleted cells.We propose that assembly of actin at epithelial cell junctionscould be one element of a protective response of epithelialcells to ischemic injury or, alternatively, may be an aspectof the pathology of renal injury induced by ischemia.
Cell Culture
LLC-PK1 epithelial cells were purchased from American Type CultureCollection (Manassas, VA) and cultured in Dulbecco’s ModifiedEagle Medium (Invitrogen Life Technologies, Carlsbad, CA) containing25 mM glucose, 10% fetal bovine serum, and antibiotics at 37°Cin an environment containing 5% CO2. Cells were transfectedwith 2 µg of an expression vector coding for a fusionprotein of actin and EGFP (Clontech, Palo Alto, CA) using lipofectamine(Invitrogen) according to the manufacturer’s instructions.Transfected cells were selected and subcloned as described inour previous study (19), generating a stable cell line expressingthe EGFP-actin fusion protein.
Measurement of ATP Levels
ATP levels were measured by plating LLC-PK1 into 24 well platesand allowing them to grow to 90% confluence over a 2-d period.Triplicate wells were rinsed with HEPES-buffered saline (HBS;20 mM HEPES, 135 mM NaCl, 4 mM KCl, 1 mM Na2HPO4, 2 mM CaCl2,1 mM MgCl2, pH 7.2) and treated with ATP depletion medium (HBScontaining 1 µM antimycin A and 10 mM 2-deoxyglucose,pH 7.2) for indicated times. All reagents were purchased fromSigma Chemical Co., St. Louis, MO, except as indicated. Triplicatecontrol wells were left untreated, and a second triplicate setof wells was rinsed twice with HBS and incubated for 1 h withHBS containing 25 mM glucose. ATP was extracted in a 6% solutionof TCA, acidity was neutralized by vortexing with tri-N-octylamine/Freon,and ATP content of samples was analyzed using HPLC as describedpreviously (24).
Imaging of Living Cells
Observation chambers for live cell imaging were made by drillinga 15-mm-diameter hole in the bottom of 35-mm petri dishes andgluing number 1 microscope coverslips over the hole using Sylgardelastomer (Dow Corning Corp.). Chambers were sterilized by treatmentwith 70% ethanol before use. Before the conduction of experiments,cells were trypsinized and plated in observation chambers ata density sufficient to reach approximately 50% confluence after1 to 2 d of culture. Cells were imaged with a Zeiss Axiovert135 microscope (Carl Zeiss Inc., Thornwood, NY) equipped witha 40 x 1.4 NA oil immersion objective lens. Temperature wasmaintained at 37°C by using an Airtherm air stream incubator(World Precision Instruments, Sarasota, FL). Images of cellswere obtained at 2-min intervals by using a cooled integratingCCD camera (DAGE RT3000, DAGE-MTI Inc. Michigan City, IN), usinga 0.5-s integration time. Illumination was provided with anAttoarc 100W mercury arc lamp (Carl Zeiss Inc.) attenuated usingneutral density filters and shuttered using a Uniblitz shutterand controller (Vincent Associates, Rochester, NY). Camera integrationtimes, shutters, and image capture were coordinated by macrocommand sets using NIH-Image running on an Apple Macintosh G4computer equipped with an image capture board (LG3; Scion Corp,Frederick, MD). For initial imaging of cells, culture mediumwas replaced with HBS, pH 7.2, containing 25 mM glucose. Afterimaging cells in this medium, ATP levels were depleted by rinsingchambers twice with HBS, pH 7.2, without glucose followed byaddition of HBS containing 1 µM antimycin A and 10 mM2-deoxyglucose at pH 5.5 or pH 7.2. ATP repletion was conductedby removing solution with inhibitors, rinsing cells twice withHBS, pH 7.2, and adding HBS, pH 7.2, containing 25 mM glucose.Imaging of cells was continued during medium changes. Representativemovies in Quicktime format may be found on the Internet at http://www.umich.edu/~shelden/JASN2002b.html.
Confocal Imaging of EGFP-Actin and Total Actin
Cells were cultured for 24 to 48 h on glass coverslips untilconfluent and subjected to ATP depletion with or without recoveryas described above, then fixed in 4% paraformaldehyde at roomtemperature and stained with rhodamine phalloidin, as describedin our previous study (19), and 1 µg/ml Hoechst to revealnuclear morphology. Coverslips were mounted for observationon microscope slides using Prolong mounting medium (MolecularProbes, Eugene, OR). Imaging of EGFP-actin and rhodamine phalloidinwas conducted using a Zeiss LSM510 confocal microscope (CarlZeiss Inc.) equipped with a 63 x 1.2 NA water immersion objective.Laser output and detector gain and black level settings wereoptimized using a preparation of ATP-depleted cells and thenheld constant for all imaging. Images of all three fluorescenceprobes were obtained simultaneously using a multichannel scanningprocedure in which each line of the final image was scannedthree times, using excitation and imaging filters specific foreach individual fluorophore.
Quantitative Analysis and Statistics
Actin assembly kinetics in lamellar protrusions were quantifiedusing an "image difference analysis" developed by our laboratoryfor analyzing lamellar ruffling dynamics from phase contrastimages (25). Briefly, sequential video images were digitallysubtracted from each other and regions varying by more than5% selected. Camera noise was reduced using a median filter,and the final area of difference was measured for each imagepair. We believe that this study represents the first use ofthis method to quantify actin array turnover in living cells.
Fluorescence intensity of stress fibers and background fluorescenceof the cytoplasm was calculated essentially as described previouslyby others (8,13). For each image, a duplicate image was created,and background fluorescence was removed using a 2D rolling ballbackground subtraction algorithm. The resultant image of stressfibers (and other sharp detail) was used as a digital mask andmultiplied by the original image, creating an image in whichonly stress fiber fluorescence was retained. The region of eachcell containing stress fibers was selected with a cursor, andthe average brightness of the region was measured. The brightnessof the background cytoplasm in the same region was obtainedusing the inverse of the stress fiber mask to select regionslacking stress fibers. To measure the brightness of cell-cellattachments in live cells, a duplicate image of each video framewas created and sites of cell-cell attachment were traced witha digital brush set to a unique value. The trace was used asa mask to select grayscale values in the original image, andthe area and brightness of the final selected region in eachimage was measured. This approach was also used to measure thefluorescence intensity of cell junctions in fixed, triple-labeledcells imaged using confocal microscopy. The fluorescence intensityof actin in non-junctional regions was measured from confocalimages by selecting each cell with a cursor, excluding junctionalareas, and computing the average fluorescence intensity of selectedregions. For analysis of confocal images, only the rhodamine-phalloidinstaining intensity (red channel in Figure 5) was analyzed, thus,this analysis specifically examines only polymerized actin.Measurements of junctions and of cytoplasmic actin in non-junctionalregions were analyzed separately for cells expressing and lackingdetectable expression of EGFP-actin. In all of our studies,maximum brightness would correspond to a measured value of 256,while a black background would have a grayscale value of zero.Confocal images of control and ATP-depleted cells were alsoscored for the presence of actin aggregates within the cytoplasmby a blinded observer, but no attempt was made to analyze changesin the size or number of aggregates in cells in the presentstudy.
Figure 5. Distribution of EGFP-actin and total filamentous actin in confluent monolayers of LLC-PK1 cells before, during, and after ATP depletion. EGFP-actin (left column, green) is incorporated into rhodamine phalloidin-stained actin filament arrays (middle column, red) in control cells (A), after 1 h of ATP depletion at pH 5.5 (B) and after 90 min of recovery (C). Incorporation of EGFP-actin into stress fiber bundles is particularly evident in cells marked with asterisks. However, not all cells express detectable EGFP-actin (arrowheads). Comparison of the brightness of junctional staining in control cells (A) expressing (arrows) and lacking (arrowheads) EGFP-actin with the intensity of junctions in corresponding ATP-depleted cells (B) suggests that junctional regions increase in thickness and intensity of actin fluorescence during ATP depletion. Actin aggregates are found in the cytoplasm of cells under all conditions (arrowheads, right panels) but were most evident in ATP-depleted cells (B). Enhanced actin fluorescence at site of cell-cell attachment persists after 90 min of ATP repletion in some cells (arrowhead, C). Scale bar = 20 µm.
Finally, the fluorescence intensity of focal adhesions was measuredby first outlining each focal adhesion using a cursor. For thepurposes of this study, focal adhesions were defined as oblong,often somewhat triangular, areas of actin fluorescence at thebottom of cells, which terminated and were slightly larger andbrighter than an attached stress fiber. Isolated, stationaryfluorescence actin structures similar in size, orientation,and brightness to neighboring focal adhesions with attachedstress fibers were also considered to be focal adhesions (Figure 8).The area and average intensity of each selected region wasquantified using software commands available within the NIH-Imageprogram. For each focal adhesion, the average brightness wascalculated from four images obtained before the onset of ATPdepletion, and four images obtained after 1 h of ATP depletion.
Figure 8. Fluorescence intensity of cytoplasm and stress fibers but not focal adhesions is reduced by 1 h of ATP depletion. Actin distribution in a cell before (A and C) and after 1 h (B and D) of ATP depletion, showing loss of stress fibers (arrow, A) but not focal adhesions (C and D). Scale bar = 20 µm in panels A and B. Quantitative analysis shows the time course of fluorescent intensity decreases within stress fibers (E; four cells from two videos) and the background fluorescence of the cytoplasm (F), but a plot of focal adhesion fluorescence before (horizontal axis) versus after 1 h (vertical axis) of ATP depletion (G) reveals no average change in focal adhesion fluorescence intensity.
Statistical analyses of data were conducted using MicrosoftExcel. Comparison of population means was conducted using at test assuming equal variance.
Electron Microscopy
Cells were cultured in 35-mm dishes containing 200-mesh nickelgrids with a Formvar/carbon coating (Electron Microscope Sciences,Fort Washington, PA) and either ATP depleted at pH 5.5 or processedwithout ATP depletion (controls) as described above. Cells weredetergent-lysed for 5 min to remove non-cytoskeletal componentsessentially as described by Svitkina and Borisy (26) in an actin-stabilizinglysis buffer (50 mM imidazole, 50 mM KCl, 0.5 mM MgCl2; 0.1mM EDTA; 1 mM EGTA, 4% polyethylene glycol [8000 MW], and 200µg/ml rhodamine phalloidin, pH 6.8), then fixed in 0.1M phosphate buffer containing 2.5% glutaraldehyde, pH 7.2. Gridswere rinsed three times with distilled water and stained for3 min with 2% aqueous phosphotungstic acid. Excess stain wasremoved and grids dried slowly for about 10 min in a humid chamber.Cells were imaged using a Phillips CM100 transmission electronmicroscope operated at 60 kV equipped with a Kodak Megapluscamera, model 1.6.
Time Course of ATP Depletion
To characterize the effects of our procedures on ATP levelsin LLC-PK1 cells expressing EGFP-actin and to permit the directcomparison of changes in actin cytoskeletal organization withATP levels, ATP levels were measured in cells treated for upto 1 h with 1 µM antimycin A and 10 mM 2-deoxyglucose.As expected, ATP levels were rapidly reduced after the applicationof these inhibitors. Relative to the control time zero, values(± SD) were 71.4 ± 6.2% at 2.5 min, 34.9 ±1.6% at 5 min, 12.7 ± 1.2% at 10 min, 3.9 ± 0.5%at 20 min, 1.2 ± 0.2% at 40 min, and 1.0 ± 0.1%at 60 min. (Figure 1). In contrast, replacement of culture mediumwith HBS containing 25 mM glucose, pH 7.2, produced no changein ATP levels after 1 h of treatment.
Figure 1. ATP levels in LLC-PK1 cells expressing enhanced green fluorescent protein (EGFP)-actin during ATP depletion and control experiments. ATP levels fall rapidly in cells treated with HEPES-buffered saline (HBS) containing 1 µM antimycin A and 10 mM 2-deoxyglucose, but not when incubated with HBS containing 25 mM glucose. Values shown are means and SD calculated from three replicate experiments.
Lamellar Protrusion Is Inhibited Rapidly during ATP Depletion
EGFP-actin was imaged at 2-min intervals in LLC-PK1 cells undercontrol conditions, during ATP depletion at an extracellularpH of either 7.2 or 5.5, and during ATP repletion at pH 7.2in the presence of 25 mM glucose. In total, actin fluorescenceand the kinetics of actin array turnover were examined in 19videos showing 143 ATP-depleted cells and 3 videos showing 18control cells. To quantify effects of ATP depletion on actinassembly dynamics, an analysis of image difference was appliedto videos (see Materials and Methods). Figure 2 shows two sequentialimages of fluorescent actin in a cell before ATP depletion (Figure 2A)with the resultant difference image (Figure 2C). Regionsundergoing more than 5% change in grayscale value are observedat sites of lamellar protrusion at the leading edge of the cells(arrows, Figure 2B) and are black in the final difference image(Figure 2C). After 15 min of ATP depletion, no changes in actindistribution are detected in this same cell over a 2-min interval(Figure 2, D and E), and no area of change is detected in thedifference image (Figure 2F). In Figure 2G, difference valuesobtained over the time course of ATP depletion for four cellsand for one control cell are shown. Measured dynamic turnoverof the actin cytoskeleton is greatly reduced within 10 min ofthe addition of metabolic inhibitors (transient increases atthe start of ATP depletion [arrow, Figure 3G] and other pointsare due to focus and stage position changes [data not shown]),whereas replacing the initial medium with additional glucosecontaining medium (cntrl, Figure 2G) has no effect on the dynamicsof actin reorganization.
Figure 2. Analysis of actin array turnover in lamellar protrusion using image difference calculations. Images of a cell under control conditions taken 2 min apart (A and B) are subtracted and areas changing by more than 5% in grayscale value selected to produce a resultant image (C) in which sites of array turnover (arrows, B) are black. The same method applied to images of this cell after 15 min of ATP depletion (D and E) produce a difference image (F) showing no regions of change. Scale bar = 20 µm. (G) The time courses of normalized difference values (area of black regions in panels C and F) calculated before and during ATP depletion for four cells and a control cell (open circles) in which medium was exchanged without ATP depletion are shown. The peak at the time of medium change reflects shifts in culture chamber position (arrow). Axes are min of ATP depletion (horizontal) and proportional change in image difference value (log scale, vertical).
Figure 3. Rapid inhibition of actin assembly in lamellar protrusions during early ATP depletion. (A) Video images taken 2 min apart of lamellae formed between two attached cells were combined to show areas of actin array extension or assembly in green and areas of retraction or disassembly in red. A diagram of the first image (panel I) shows junctional actin arrays that are stable over the 2-min interval in yellow. Dynamic reorganization of actin arrays in lamellar protrusions is seen before the onset of ATP depletion but is inhibited within 4 min of the start of ATP depletion. (B) ATP depletion also induces loss of actin fluorescence within lamellar protrusions (arrow) without retraction. Numbers are min before (negative) or after (positive) application of metabolic inhibitors. Scale bar = 10 µm.
Figure 3A shows images of lamellae formed at a site of cell-cellattachment between two LLC-PK1 cells. Images obtained 2 minapart were combined such that the later image is green and theearlier image is red (Figure 3A). Sites of actin assembly aregreen in the resultant image, whereas structures that disappearduring this interval are red. The site of cell-cell attachmentand other stable actin-containing structures are unchanged inboth images and are therefore yellow (quiescent cell boundariesare shown in gray in the accompanying diagram of the first image).Both green (arrows, Figure 3A) and red areas are seen in theseimages before the addition of inhibitors, indicating dynamicturnover. In contrast, images obtained within 4 min of inhibitoraddition show complete overlap of red and green, indicatingthat time-dependent change in the distribution of fluorescentactin is no longer occurring. Figure 3B also shows grayscaleimages of fluorescent actin in a lamellar protrusion (arrows)in which loss of actin fluorescence is observed without lamellarretraction. Together, data shown in Figures 1, 2, and 3 revealthat actin assembly ceases in lamellar protrusions within minutesof the start of ATP depletion and at time points before maximalreduction of ATP levels.
Accumulation of EGFP-Actin at Sites of Epithelial Cell-Cell Attachment in ATP-Depleted Cells Figure 4A shows images of fluorescent actin at a site of epithelialcell-cell attachment in a cell treated with inhibitors of ATPproduction. Unlike lamellar protrusions, EGFP-actin fluorescenceassociated with this structure increased gradually and persistentlyin fluorescence intensity and apparent thickness over about2 h of ATP depletion. Figure 4B shows higher magnification imagesof an attachment site between two cells (shown at low powerin the inset) after the application of inhibitors of ATP productionat time zero. The image series illustrates that the increasein apparent thickness of fluorescent cell-cell junctions seenin Figure 4A is due, at least in part, to the formation or elongationof fluorescent structures resembling microspikes or filopodia.In Figure 4C, normalized fluorescent actin intensities quantifiedfor five sites of cell-cell attachment in ATP-depleted cellsand a control cell are shown. Increased actin fluorescence isdetected at cell-cell attachments during ATP depletion, andcomparison of these graphs with data shown in Figure 1 revealsthat much of this increase occurs after ATP levels have fallento less than 2% of control. No change in actin fluorescenceis detected at site cell-cell attachment in cells before thestart of ATP depletion (Figure 4B) and in a 1 h control experiment(cntrl 1, Figure 4B).
Figure 4. EGFP-actin fluorescence intensity at epithelial cell junctions during ATP depletion. (A) Images of a site of cell-cell attachment (from a series at 2-min intervals) after the start of ATP depletion. Increased brightness of the junctional region is observed. (B) Higher magnification of the region between two cells expressing EGFP-actin (inset, panel B) showing assembly of structures resembling filopodia or microspikes. Numbers are min after the start of ATP depletion. Scale bars: 10 µm in panel A; 4 µm in panel B. (C) Graph of normalized EGFP-actin fluorescence intensity at sites of cell-cell attachment (n = 5 cells from two videos) during ATP depletion and from a control experiment in which medium was changed at time 0 without ATP depletion (open circles). Axes are time after medium exchange (horizontal) and relative change in average brightness (vertical).
ATP Depletion Induces Accumulation of Actin Filaments at Sites of Epithelial Cell-Cell Attachment
Videos of cells expressing EGFP-actin described above were madeof well-spread cells cultured at moderate (50%) confluence toclearly visualize cell junctions, lamellar protrusions, andstress fibers in the same image. Additionally, structures containingfluorescence EGFP-actin might contain either assembled actinfilaments or monomeric actin. Therefore, to determine if accumulatedEGFP-actin represented the presence of actin filaments, andto address whether junctional actin accumulated during ATP depletionin cells cultured at higher densities, cells were plated atconfluent cell densities, experimentally treated, and then fixedand stained with rhodamine phalloidin, a specific marker forassembled actin filaments. Multichannel confocal fluorescenceimaging was used to examine and compare the distribution andintensity of the EGFP-actin and rhodamine phalloidin probesat sites of cell-cell attachment (Figure 5). Inspection of theseimages shows that EGFP-actin (left column and green, Figure 5)is incorporated into all structures stained with rhodaminephalloidin (middle column and red, Figure 5). Incorporationof EGFP-actin into basal stress fibers is particularly evidentin cells marked with asterisks. Junctional regions in controlcells expressing EGFP-actin (arrows, center, Figure 5A) andneighboring cells, which lack detectable EGFP-actin (arrowhead,center, Figure 5A), are thin and stain relatively dimly withrhodamine phalloidin when compared with cells fixed after 1h of ATP depletion (Figure 5B). Junctional regions in ATP-depletedcells expressing EGFP-actin (arrow, center, Figure 5B) and lackingdetectable EGFP-actin (arrowhead, center, Figure 5B) are comparativelybright, thick, and fibrous. Images of cells obtained after 2h of recovery from ATP depletion (Figure 5C) show some junctionalareas with normal fluorescence intensity (arrow, center, Figure 5C),while other junctional areas remain fibrous and brightlyfluorescent (arrowhead, center, Figure 5C). Aggregates of phalloidinstained actin were observed within the cytoplasm of cells underall conditions (representative examples are indicated by arrowheadsin the color panels, right, Figure 5), but they were more commonand numerous in cells after 1 h of ATP depletion than in controlcells. For example, 25.8% of control cells (73 of 283) expressingEGFP-actin and 22.6% of control cells (51 of 226) lacking detectableEGFP-actin displayed some actin aggregates, whereas 57.6% cells(177 of 307) expressing EGFP-actin and 51.4% of cells (126 of245) lacking detectable EGFP-actin displayed aggregates after1 h of ATP depletion. Because our imaging parameters were optimizedfor detection of our probes in ATP-depleted cells (Figure 5B),phalloidin staining in controls cells (Figure 5A) is comparativelylow (Figure 5A, center column; red, right column, Figure 5A).Additionally, background fluorescence of monomeric EGFP-actinwould not be expected to stain with rhodamine phalloidin, andthe presence of monomeric EGFP-actin probably accounts for theoverall green color in the color images of fixed control cellsand cells fixed during recovery from ATP depletion. All imagesshown in Figure 5 were obtained using identical imaging parameters,and brightness and contrast levels were adjusted for all EGFP-actinimages and rhodamine phalloidin images together.
Quantitative analysis of the brightness of phalloidin stainingat cell junctions was conducted from confocal images of cellsfixed without ATP depletion and cells fixed after 1 h of ATPdepletion (Figure 6), and results obtained from cells expressingEGFP-actin (green in Figure 5) compared with those obtainedfrom the analysis of cells lacking detectable EGFP-actin (redin Figure 5). Triplicate experiments were conducted, and atleast ten random fields of cells were imaged for each trial.Fluorescence phalloidin intensities of all visible junctionalregions associated with well-spread, non-mitotic cells weremeasured. The average brightness (± SD) of junctionalregions for control cells expressing EGFP-actin was 73.3 ±20.7 (n = 820) and 99.5 ± 28.4 (n = 641) for ATP-depletedcells expressing EGFP-actin (Figure 6A). For cells lacking detectableEGFP-actin, the average fluorescence intensity of junctionalregions for control and ATP-depleted cells was 75.3 ±20.8 (n = 275) and 111.3 ± 31.3 (n = 331), respectively(Figure 6B). The increases in fluorescence intensity observedin ATP-depleted cells expressing EGFP-actin and cells lackingEGFP-actin were both significantly higher than values measuredfor corresponding control cells (P 0.01). We conclude thatthe increased phalloidin intensity, and thus actin filamentcontent, of junctional regions in ATP-depleted cells is independentof EGFP-actin expression. Indeed, cells lacking detectable EGFP-actinshowed a small but significantly greater increase in fluorescenceintensity of junctional regions after ATP depletion (P 0.01).For comparison, the average fluorescence intensity of rhodaminephalloidin staining was analyzed for the total cell area, excludingcell junctions (Figure 6, C and D). In contrast to actin stainingat sites of cell-cell attachment, a significant decrease (P .01) in the average rhodamine phalloidin brightness of non-junctionalareas was measured for cells after ATP depletion. The averagebrightness of the non-junctional regions (± SD) of controlcells was 32.1 ± 8.2 (n = 194) for EGFP-expressing cellsand 37.5 ± 8.4 (n = 180) for cells lacking EGFP-actin.The average brightness of the non-junctional regions (±SD) of ATP-depleted cells was 27.0 ± 6.4 (n = 191) forEGFP-expressing cells and 31.7 ± 8.4 (n = 165) for cellslacking EGFP-actin. Because actin aggregates are formed in non-junctionalregions of ATP-depleted cells, the decrease in average phalloidinstaining measured in ATP-depleted as compared with control cellswas unexpected, but it may reflect a comparatively large decreasein the polymer content of stress fibers in ATP-depleted cells(Figure 8E).
Figure 6. Quantitative analysis of phalloidin staining intensity in control and ATP-depleted cells expressing and lacking detectable EGFP-actin. Normalized histogram distributions of phalloidin staining intensity are shown. (A) Intensity of rhodamine phalloidin staining was measured at sites of cell-cell attachment in EGFP-actin expressing cells fixed under control conditions (black bars, n = 820) or after 1 h of ATP depletion (gray bars, n = 641). A population shift toward higher (brighter) values is observed after ATP depletion. (B) Analysis of junctional phalloidin staining in cells lacking detectable EGFP-actin also shows an increase in brightness of junctions in ATP-depleted cells (gray, n = 331) as compared with control cells (black, n = 275). In contrast, normalized histogram distributions of the average intensity of actin staining in non-junctional regions in control cells (black bars) and ATP-depleted cells (gray bars) show no overall increase in brightness of ATP-depleted cells versus control cells in cells expressing (C) or lacking (D) EGFP-actin.
Actin Assembly at Sites of Cell-Cell Attachment during ATP Depletion Is Independent of the Degree of Cellular Injury
Previous studies have shown that cell survival during recoveryfrom ATP depletion is strongly inhibited when ATP depletionis conducted at neutral or greater extracellular pH, and enhancedwhen ATP depletion is conducted at acidic extracellular pH (27,28).Therefore, to determine whether actin assembly at cell junctionscould be correlated with the amount of cellular injury inducedduring ATP depletion, we analyzed and compared the increasein actin brightness at cell-cell attachments from videos ofLLC-PK1 cells undergoing ATP depletion at extracellular pH valuesof 7.2 and 5.5. Most cells undergoing ATP depletion at pH 5.5were subsequently observed to recover normal lamellar protrusionbehavior after addition of HBS containing glucose, whereas thoseundergoing ATP depletion at pH 7.2 failed to recover lamellarruffling behavior. These cells instead underwent dramatic lossof actin cytoskeletal integrity after the addition of HBS andglucose and may have undergone necrotic cell death during theexperiment (see videos available on our web site, as describedin Materials and Methods). The analysis of the brightness ofactin at cell-cell attachments in these experiments was alsocomplicated by the substantial reduction of fluorescence thatGFP exhibits at acidic pH (29). We therefore determined theaverage fluorescence intensity of cell-cell attachments in thefirst four images obtained during ATP depletion with the averageintensity of these same junctions in four images taken after1 h of ATP depletion (Table 1). The increase in brightness ofcell-cell attachments in cells observed to recover lamellarprotrusion (pH 5.5), and those that did not recover during asimilar observation period (pH 7.2) did not differ significantly(P > 0.1).
Table 1. Average brightness increases during ATP depletion of actin arrays at sites of cell- cell attachment but notfocal adhesionsa
Electron Microscopy of Actin Filaments at Sites of Cell-Cell Attachment in Control and ATP-Depleted Epithelial Cells
The increases of EGFP-actin fluorescence intensity in livingcells during ATP depletion and phalloidin staining in fixedcells after ATP depletion (above) support the hypothesis thatactin polymer mass increases at sites of epithelial cell-cellattachment during ATP depletion. To further assess the morphologyand distribution of actin polymer at these sites, we conductedultrastructural studies of cells after detergent extractionusing an actin stabilizing lysis buffer (Figure 7). Initialstudies conducted by thin sectioning epon-embedded cells followedby uranyl acetate and lead citrate staining were less informativethan we hoped, perhaps because of the difficulty of visualizingthree dimensional actin filament arrays in 70-nm-thick sections(not shown). However, examination of whole cells cultured onEM grids after detergent lysis, fixation, and negative stainingusing phosphotungstic acid reveals the extensive presence ofnegatively stained filaments at sites of cell-cell attachmentin control cells (Figure 7, A through C). Similar filamentsare observed along the margin of sites of cell-cell attachment(white arrows, Figure 7F) and in fibrous protrusions associatedwith sites of cell-cell attachment (black arrows, Figure 7F)in cells fixed after 1 h of ATP depletion at pH 5.5. In bothcases, filaments observed here are morphologically similar tothose observed by previous investigators at the leading edgeof fibroblasts, stress fibers, and microspikes (30) and at thesesites in the present study (not shown). Images shown are representativeof 11 junctional regions of ATP-depleted cells and 9 junctionalregions of control cells, and no attempt was made to distinguishbetween cells that expressed or lacked detectable EGFP-actinin these studies. The width of filaments shown in Figures 7C and 7Fmeasured between 7 and 8 nm, the predicted thicknessof actin filaments in negatively stained preparations (datanot shown), and the orientation of fibers is consistent withthat expected for actin filaments associated with epithelialadherens junctions (31). Comparison of the electron densityof staining in cell junctions of control cells (Figure 7C) andATP-depleted cells (asterisk, Figure 7F) also suggests thatan increase in electron-dense material characterizes ATP-depletedcell junctions.
Figure 7. Electron microscopy of negatively stained actin filaments at regions of cell-cell attachment in control and ATP-depleted cells. Low magnification view (x2350, panel A) and medium magnification (x13,500, panel B) of the junctional region between two control cells, circled in panel A. (C) High magnification view (x130,000) of the region indicated with an arrow in panel B. The junctional region contains negatively stained filaments oriented along the length of the junctional region. Particularly clear examples are indicated with arrows. The width and orientation of these filaments is consistent with individual actin filaments associated with epithelial adherens junctions. Scale bar = 100 nm. Low magnification view (x2600, panel D) and medium magnification view (x25,000, panel E) of the junctional region between two ATP-depleted cells (circled in panel D). The junctional region is more electron-dense than that of control cells and displays numerous lateral protrusions that appear to be attached to the junctional region. (F) High magnification view of the region indicated by the arrow in panel E. Aligned, negatively stained filaments are detected in protrusions associated with junctional regions (black arrows, F) and at the margin of electron-dense junctional cell borders (white arrows, F). The inset shows the central region of this process after contrast enhancement. The electron density of the junctional region (asterisk) can be directly compared with that of control cells shown in panel C. Scale bar = 100 nm.
Focal Adhesions Are More Resistant to Early ATP Depletion than Stress Fibers
To determine if actin arrays at sites other than cell-cell attachmentsdisplay altered actin assembly during ATP depletion, we analyzedthe intensity of EGFP-actin fluorescence in stress fibers andfocal adhesions. Images of a cell before ATP depletion (Figure 8A)and after 1 h of ATP depletion (Figure 8B) show that stressfibers are partially disrupted by ATP depletion in our experiments.Reduced fluorescence intensity of stress fibers can be observedas well as the disappearance of some thin stress fibers (arrow,Figure 8A and surrounding region). These results are in agreementwith a previous analysis of stress fiber disruption in livingrenal epithelial cells during ATP depletion (8). In contrast,the fluorescence intensity of some individual focal adhesionsincreased during the same time interval (compare insets in Figures 8A and 8Bshown at higher magnification in Figures 8C and 8D).As expected, the measured, average fluorescence intensity ofstress fibers declined in ATP-depleted cells (Figure 8E). However,when the fluorescence intensities of focal adhesions beforeand after 1 h of ATP depletion are compared, both decreasesand increases in fluorescence intensity of focal adhesions areseen (Figure 8G), but no average change in the fluorescenceof focal adhesions is detected (Figure 6G and Table 1).
Finally, previous studies have correlated a decrease in fluorescenceactin intensity of the cytoplasm during ATP depletion with areduction in total actin monomer content (8). To confirm thatthe behavior of cells in our studies replicated that reportedpreviously and to compare the time courses of the increase influorescence of cell-cell attachments in our studies with thereduction in fluorescence of nonpolymerized actin, the fluorescenceintensity of the cytoplasm was analyzed here. We find that time-dependentdecreases in the fluorescence intensity of the cytoplasm occurin ATP-depleted but not control cells (Figure 8F), and thatthe time course of this reduction is similar to that observedfor the increase in intensity at cell-cell attachments (comparewith Figure 4B).
Results of the present study provide new data on actin distributionin living renal epithelial cells (RTE) during ATP depletionand permit the comparison of changes in actin organization withmeasured reductions in ATP levels. The most immediate effectof ATP depletion observed in our studies is the inhibition oflamellar turnover and the accompanying loss of actin from thesestructures (Figures 2 and 3). Presently, relatively little isknown about the contribution of processes mediating lamellarprotrusion to maintenance of epithelial tissues in vivo. Directvisualization of lamellar protrusion by RTE in vivo has notbeen accomplished, and the possibility that the generation ofthese structures by cells in our studies is an artifact of ourin vitro cell culture conditions should not be ruled out. However,lamellar protrusion by RTE is likely to play a significant rolein wound healing and re-epithelialization in vivo during recoveryfrom ischemic and other injuries. Additionally, recent evidencesuggests that mechanisms involved in mediating lamellar protrusionalso play roles in the maintenance of normal epithelial function.For example, lamellar ruffling is induced as a consequence ofactivation of the Rho family small GTPase Rac1 (32–34),and results of recent studies indicate that Rac1 plays a rolein the maintenance of epithelial cell junctions (35,36). Resultsof the present study indicate that such functions may be highlysensitive to disruption as a consequence of ATP depletion andthat even very brief periods or modest degrees of ischemia couldinhibit epithelial functions that are dependent on lamellarprotrusion.
Perhaps the most novel aspect of this study is the demonstrationof an increase in brightness of fluorescent actin probes atsites of cell-cell attachment during ATP depletion. The increasein EGFP-actin intensity observed in living ATP-depleted cells(Figure 4A), the increased intensity of phalloidin stainingafter ATP depletion, both in cells expressing and lacking detectableEGFP-actin (Figure 6), the formation of microspike-like structuresat sites of cell-cell attachment in ATP-depleted cells (Figure 4B),and the presence of 7- to 8-nm-diameter filaments at thesesites (Figure 7) all lead us to conclude that this increasein fluorescence intensity is due, at least in part, to actinfilament assembly during ATP depletion. Our findings agree wellwith results from previous analysis of fixed cells showing greaterresistance of junctional actin to ATP depletion than actin instress fibers (9,23). Indeed, although to our knowledge ourstudies provide the first direct demonstration of actin assemblyat sites of epithelial cell-cell attachment during ATP depletion,such assembly can be inferred from careful comparison of actindistribution patterns in fixed cells published in previous studies(23,28). Additionally, like results from previous studies offixed cells (28), our results show that actin cytoskeletal alterationis independent of extracellular pH during ATP depletion. Thesesimilarities indicate that the actin assembly observed in ourexperiments is likely representative of changes in actin assemblyoccurring in other experimental models of ischemia and in vivo.
The assembly of actin in ATP-depleted cells has been previouslydemonstrated using biochemical assays of actin polymer and monomer(8,14,17,18) and has been inferred from an increase in cytoplasmicactin aggregates within the cytoplasm and perinuclear area inseveral previous studies of ATP-depleted renal epithelial cellsand tissues (9,14,17,28). Unfortunately, although actin aggregateswere observed in fixed, phalloidin-stained cells in our studies(Figure 5), these structures were not observed in living cellsin the present study, probably because our observations of livingcells focused only on actin arrays close to the cell substrate.Because our analysis of actin polymer in fixed cells was limitedto individual confocal images focused at the level of cell junctions(Figures 5 and 6) these results also do not address the relativecontribution of assembly in aggregates and at sites of cell-cellattachment to the accumulation of actin polymer in ATP-depletedcells. Additionally, although our ultrastructural studies demonstratethe presence of morphologically normal actin filaments in structuresassembled at sites of cell-cell attachment in ATP-depleted cells,it also remains unclear whether actin in ATP-depleted cellsat either cell attachments or cytoplasmic aggregates polymerizesthrough a normal assembly mechanism; for example, through incorporationof ATP-associated actin monomer at filament barbed ends or throughsome other mechanism. However, because we determined that ATPlevels fell to less than 2% of control levels within 20 to 40min of the application of inhibitors in our study (Figure 1),we conclude that much of this assembly is either ATP-independentor requires extremely low ATP concentrations. Thus, actin assemblyobserved in ATP-depleted cells may occur as a consequence ofmechanisms that do not normally play a role in actin assemblyin control cells.
Assembly of actin filaments in junctional regions may be animportant aspect of the epithelial cell response to prolongedATP depletion. However, examination of the graphs shown in Figure 4Calso suggests that there is no significant lag period betweenthe start of ATP depletion and the time-point at which actinfluorescence begins to increase at sites of cell-cell attachment.Therefore, actin assembly at cell junctions also appears tobe an early consequence of ATP depletion. Interestingly, actinassembly occurs at cell-cell attachments in cells, but stressfibers and focal adhesions in the same cells did not exhibitassembly during ATP depletion in our study (Figure 8). Thus,not all actin filament arrays in cells are capable of promotingactin assembly during ATP depletion. These differences may reflectthe diversity of actin-associated proteins found at cell adhesioncomplexes or other cortical actin arrays. Our discovery thatactin filament content increases can occur at epithelial cell-cellattachments during ATP depletion raises the question of thefunctional significance of this behavior. Because cell adhesioncomplexes are a highly ordered assembly of cytoskeletal, regulatory,and transmembrane proteins, it is possible that abnormal actinassembly at these sites could play a role in the disruptionof epithelial barrier function accompanying ischemic injury.Alternatively, assembly of actin and recruitment of actin-associatedproteins to cell junctions may reflect the function of mechanismsinvolved in preserving epithelial integrity during injury.
Finally, it is of importance to consider whether the EGFP-actinprobe is an appropriate marker for actin cytoskeletal reorganization.Previous investigators have shown normal actin-dependent cellbehavior in cells expressing the EGFP-actin construct used inour studies (37). Additionally, Herget-Rosenthal et al. (8)have recently published an extensive analysis of the behaviorof EYFP-actin in LLC-PK1 renal epithelial cells and have concludedthat EYFP-actin expression is a faithful marker for the totalcellular pool of actin and that its expression does not alteractin-dependent cellular responses. The actin coding regionof the EGFP-actin construct used in our studies is identicalto that of the EYFP-actin construct used in this previous report;in total, the two expressed proteins differ by only 5 of 621amino acids. It seems likely that the assembly characteristicsof the EGFP-actin probe used in the current study are very similarto that of the EYFP-actin probe used in the previous study.Additionally, our examination of the phalloidin-staining intensityof cells expressing detectable EGFP-actin and those lackingEGFP-actin reveal similar increases in ATP-depleted cells ascompared with control cells (Figure 6). We conclude that EGFP-actinexpression is not a causal factor in generating the observedchanges in actin assembly in our studies.
In summary, we have quantified the kinetics of actin distributionin cultured renal epithelial cells before and during ATP depletionand correlated these data with measured ATP levels. Loss ofactin assembly in lamellar protrusion is an immediate consequenceof reducing ATP levels, and actin turnover in lamellae is completelyinhibited when ATP levels are reduced to less than 2% of controlvalues. Actin associated with stress fibers was also disruptedduring ATP depletion, albeit more slowly. In contrast, actinassembly is detected in cytoplasmic aggregates and observedat sites of epithelial cell-cell attachment. Assembly at sitesof cell-cell attachment is initiated early during ATP depletionbut persists after ATP levels are maximally reduced. These resultsillustrate that actin assembly is altered in a site-specificmanner during ATP depletion and suggest that actin assemblyat sites of epithelial cell-cell attachment is an importantaspect of the cellular consequence of ATP depletion.
Acknowledgments
We thank N. Roeser and R. Senter for assistance with ATP measurements,and Drs. S.A. Ernst and B. Margolis at the University of Michiganfor critical reading of this manuscript. Grant support fromthe National Institute of Environmental Health Science (ES11196–01)and the National Institute on Aging (AG19847–01) to EricA. Shelden and from the National Institute of Diabetes and Digestiveand Kidney Diseases (DK-34275 and DK-39255) to Joel M. Weinbergis gratefully acknowledged.
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Received for publication April 8, 2002.
Accepted for publication July 19, 2002.
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Abstract
Introduction
0.01). We conclude that the increased phalloidin intensity, and thus actin filament content, of junctional regions in ATP-depleted cells is independent of EGFP-actin expression. Indeed, cells lacking detectable EGFP-actin showed a small but significantly greater increase in fluorescence intensity of junctional regions after ATP depletion (P 
