Abstract
Adhesion between animal cells and the underlying extracellular matrix is challenged during wounding, cell division, and a variety of pathological processes. How cells recover adhesion in the immediate aftermath of detachment from the extracellular matrix remains incompletely understood, due in part to technical limitations. Here, we used acute chemical and mechanical perturbations to examine how epithelial cells respond to partial delamination events. In both cases, we found that cells extended lamellipodia to establish readhesion within seconds of detachment. These lamellipodia were guided by sparse membrane tethers whose tips remained attached to their original points of adhesion, yielding lamellipodia that appear to be qualitatively distinct from those observed during cell migration. In vivo measurements in the context of a zebrafish wound assay showed a similar behavior, in which membrane tethers guided rapidly extending lamellipodia. In the case of mechanical wounding events, cells selectively extended tether-guided lamellipodia in the direction opposite of the pulling force, resulting in the rapid reestablishment of contact with the substrate. We suggest that membrane tether-guided lamellipodial respreading may represent a general mechanism to reestablish tissue integrity in the face of acute disruption.
Significance
As the outermost layer of organs, epithelia must withstand and respond to various physical and chemical insults. However, relatively few studies to date have examined how cells act to repair damage on the seconds timescale. Here, using in vitro and zebrafish models, we subjected epithelia to acute detachment from the underlying extracellular matrix. We found that even during large-scale detachments, cells remained attached to the extracellular matrix via long membrane tethers. Cells rapidly extended lamellipodia along these tethers, allowing them to reestablish contact with the underlying substrate on the seconds timescale. We suggest that tether-guided lamellipodia such as we observe may constitute an understudied mechanism used by epithelial cells to rapidly repair the damage caused by localized wounding events.
Introduction
Robust physical adhesion between neighboring cells and to the underlying extracellular matrix (ECM) is essential to the stability of living tissues and hence for the existence of multicellular life. The maintenance of tissue cohesion is particularly important in the case of epithelia, which are subject to mechanical forces that threaten cellular and tissue integrity. Previous work shows that cells and tissues in general act as strain-stiffening materials and show a remarkable degree of mechanical toughness (1,2). However, when the adhesive forces that link cells to each other and to the ECM are overcome, the tissue ruptures. Two major mechanisms for repairing the resulting wound have been described: a purse string mechanism, in which actin- and myosin-driven ring contraction closes the wound, and collective or individual lamellipodia-based migration of cells toward the wound site (3,4). Both of these mechanisms occur on the minutes to days timescale. How, and even whether, cells may act to repair wounds on a more rapid seconds timescale remains considerably less explored.
Epithelial tissues are also disrupted by a variety of normal physiological processes, the most prominent of these being cell division. In many cases rounded, mitotic cells remain tethered to the basal ECM by sparse, tubular membrane extensions termed retraction fibers. In some cases, these retraction fibers guide daughter cells to respread in the mother cell's premitotic footprint, which potentially ensures optimal repositioning of the daughter cells within the tissue (5, 6, 7, 8) (Fig. S1 b). Similar tubular structures are extruded from cells as the cell body pulls away from persistent adhesive contacts, for example during cell migration (9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20). Less is known about the presence and role of retraction fibers or related, membranous tethers in vivo. However, structures of this type have been documented during progenitor cell movement in zebrafish gastrulation, where leftover membrane tethers were observed at separating cell-ECM and cell-cell contacts (21,22). Whether retraction fibers or related membrane tethers have a lamellipodial guidance role outside of mitotic respreading is, to our knowledge, unexplored.
Materials and methods
Plasmid generation
To generate a plasmid that would fluorescently label F-actin filaments and plasma membrane, GFP-tagged Lifeact (Lifeact-GFP, Addgene #51010; gift from Rusty Lansford) (23) followed by a P2A ribosomal skipping sequence (24) and a myristoylation motif of Src (Myr-RFP, Addgene #32604; gift from Anna-Katerina Hadjantonakis) were cloned into a Piggybac vector (pJ509-02 construct from DNA 2.0) by Epoch (Missouri City, Texas).
Cell transfection
Wild-type Madin-Darby canine kidney (MDCK)-II cells (MDCK WT) were acquired from Millipore Sigma (00062107-1VL) at passage 27 and further expanded to make stocks within ∼5–10 passages. To generate the stable MDCK Lifeact-GFP/Myr-RFP cell line, Lifeact-GFP-P2A-myr-RFP plasmid was transfected using Lonza Nucleofector 4D X-Unit mostly following manufacturer's optimized protocol for MDCK cells. Our changes to the protocol were the usage of 106 cells in 100 uL Cell Line Nucleofector Solution SE and program CM-113. Cell stocks were selected using puromycin at ∼5 μg/mL and checked for expression using by live-cell epifluorescence. Cells stocks were frozen using 10% DMSO (ATCC, 4-X) and passaging media (see below).
Cell passaging between imaging experiments
MDCK Lifeact-GFP/Myr-RFP cells were thawed from stocks, passaged no more than seven times after expansion in passaging media. Passaging media comprised sterile-filtered DMEM media containing phenol red (15 mg/L), low glucose (1000 mg/L), sodium pyruvate (110 mg/L), L-glutamine (584 mg/L) (Life Technologies, 11885092), with added 10% FBS (Corning, 35-011-CV), and 1% penicillin-streptomycin (ThermoFisher, 15140122). Cells were passaged via rinsing by PBS (ThermoFisher, 10010049) and splitting by dissociation using 0.25% Trypsin-EDTA (Life Technologies, 25200056) and subculturing.
Extracellular matrix coating
Experiments were conducted using two types of extracellular matrices as indicated in the relevant sections.
Fibronectin-coated dishes
Fibronectin (Corning, 356008) stock aliquots (5 mg/mL) were diluted 1:100 in PBS (ThermoFisher, 10010049), and this working solution was incubated for 70–90 min at 37°C in 35-mm glass-bottom dishes with 10-mm microwells (#1.5 coverslip) from Cellvis (D35-10-1.5-N). The dishes were then washed with roughly 3x volume PBS solution before seeding cells.
Collagen-coated dishes
35-mm collagen-coated (type I, rat tail) glass-bottom dishes with 14-mm microwells (#1.5 coverslip) were purchased from Mattek (P35GCOL-1.5-14-C).
TIRF microscopy
The total internal reflection fluorescence (TIRF) microscopy was done with an inverted microscope (Nikon TiE) using an Apo TIRF x100 oil, NA 1.49, objective lens (Nikon) and controlled through the software Micromanager (25). Experiments were conducted with an objective heater (Bioptechs) equilibrated at 37°C. The scope's Perfect Focus System was utilized. A green laser (532 nm, Crystalaser), a blue laser (473 nm Obis, Coherent), and a red laser (635 nm, Blue Sky Research) were used for RFP, GFP, and CellMask Deep-Red (Thermo Fisher, C10046) excitation respectively. The emitted light went through a quad-edge laser-flat dichroic beam splitter with center/bandwidths 405 nm/60 nm, 488 nm/100 nm, 532 nm/100 nm, and S5 635 nm/100 nm from Semrock (Di01-R405/488/532/635-25x36) and a corresponding quad-pass filter with center/bandwidths 446 nm/37 nm, 510 nm/20 nm, 581 nm/70 nm, 703 nm/88 nm bandpass filter (FF01- 446/510/581/703-25). An additional filter cube (550 nm LP for RFP imaging; 470 nm/40 nm, 495 nm LP, 525 nm/50 nm for GFP imaging; 679 nm/41 nm, 700 nm/75 nm for CellMask Deep-Red imaging) was included before the camera in the light path. All processed images were taken using the full chip of a Hamamatsu Orca Flash 4.0 camera (chip size 2048 x 2048 pixel2). Fluorescence videos were captured with 200-ms continuous acquisition except for the time lapse shown in Fig. S1, where 4-s intervals between 200-ms exposures were used for the mitotic cell (Fig. S1 b and c) imaging.
Imaging experiments
In descriptions below, we will refer to “imaging media,” which is sterile-filtered Leibovitz's L-15 Medium (ThermoFisher, 21,083,027) with additional 1% FBS (Corning, 35-011-CV), 1% ITS-A supplement (ThermoFisher, 51,300,044), and 1% penicillin-streptomycin (ThermoFisher, 15,140,122).
For the mitotic cell imaging in Fig. S1 a–c, we used cells at a heterogeneous confluency (20%–100%) seeded on a collagen-coated dish overnight. Cells were changed into 1.5 mL imaging media 1 h before imaging start.
Chemical perturbation via EDTA
For experiments in Fig. 1, dishes at 80%–100% confluency (seeded 1 to 2 days overnight) were changed from passaging media to 1.5 mL imaging media 1–3 h before experiment start. The 0.5 M ethylenediaminetetraacetic acid (EDTA) stock (pH 8.0 in water, 03,690-100ML, Sigma) was diluted 18:100 in imaging media equilibrated at 37°C, and 100 uL of this working solution was added to the dishes to give ∼4.8 mM final EDTA concentration. Imaging either started within ∼2–3 min, or cells were kept in the incubator for a delayed time before imaging, which was noted for each experiment. Videos of ∼5–20 different regions within a dish were taken, with each video typically consisting of ∼500–900 frames. Brightfield snapshots were often taken before and after each video capture. Generally after 40–60 min, there were many regions with lifted off cells, so we searched for regions where the basal surface of cells was still mostly in the TIRF field.
Figure 1.
Disruption of cell adhesion induces retropodial respreading. (a) F-actin (Lifeact-GFP) signal of a peeling region 15 min after addition of 5 mM EDTA (background subtracted). (b and c) F-actin (Lifeact-GFP) signal of a peeling region 35 min after addition of EDTA. Time course from zoomed-in section highlights retropodial extension (yellow arrows). (d) i. Time course from zoomed-in section from (a), showing a retropodium that extends between a membrane tether pair. ii. Kymograph of the spreading front in i (orange dashed line). (e) F-actin (Lifeact-GFP) signal of retropodia 60 min after EDTA addition. Orange dashes were drawn as a fixed boundary at time zero to guide the eye. (f) Fraction of cells within imaged regions that contained retropodia (220 cells, six time lapses, three samples for collagen-coated dishes; 170 cells, four time lapses, two samples for fibronectin-coated dishes). Scale bars, 5 μm. Throughout, the total time cells were exposed to EDTA is indicated to the nearest 5 min. Time is shown as min:sec. To see this figure in color, go online.
Mechanical perturbation via micropipette
Setup
Micropipettes were pulled from glass capillaries of outer diameter 1.0 mm (World Precision Instruments) using a micropipette puller (P-97, Sutter). To manually control the micropipette during experiments, an XYZ translation stage with Standard Micrometers or with Differential Adjusters (with coarse range used) (Thorlabs MT3 and MT3A) was mounted on the microscope using posts. The micropipette was inserted into a microinjection pipette holder combined with a 3D printed plastic part, and this was connected to post assemblies (Thorlabs) mounted on the translation stage itself.
Mechanical displacement of cell sheets
For multi-cell experiments in Fig. 2, cells were changed into 1 to 1.5 mL imaging media 1–4 h before experiments. The micropipette was lowered until seen in brightfield, and its tip was brought to the field of view and above the cells using the micromanipulators. We started the application of a pull during the capturing of the Lifeact-GFP or Myr-RFP signal. To apply a pull, we lowered the z axis manipulator until we saw the micropipette poking a cell in the field of view and continued lowering in the z axis until the micropipette tip was dragged in the plane of the glass by ∼50 μm and then left the micropipette at this final position. The application of a pull took ∼4 s, after which the imaging of the region continued.
Figure 2.
Retropodia help recover contacts of mechanically perturbed epithelia. (a) A glass micropipette displaces the cells that are simultaneously imaged in brightfield or TIRF mode. (b) Membrane (Myr-RFP) signal of a multicellular region before and after peeling (bleach corrected). The second panel corresponds to ∼11 s after micropipette motion ceases. Time course from zoomed-in section highlights retropodia. (c) i. F-actin (Lifeact-GFP) signal of a multicellular region before and after peeling (bleach corrected). A montage shows several examples of retropodial extension. ii. Kymograph of a single retropodium (i; orange dashed line). (d) Fraction of observable cells that formed retropodia (146 cells, seven time-lapses, one sample for collagen-coated dishes (see Materials and methods); 188 cells, 10 time-lapses, two samples for fibronectin-coated dishes). (e) F-actin (Lifeact-GFP) signal of a single cell with no neighbors, before and after peeling. Scale bars, 5 μm. Time is shown as min:sec. To see this figure in color, go online.
When cells were successfully dragged by this protocol, the retropodia phenotype was robust. However, there were cases when we failed to drag cells and ended up just scraping the cells in front of the micropipette, or inducing a quick large rupture, some of which we chose not to acquire data from to save time. Anecdotally we found more success by applying the pull closer to borders of epithelial colonies (within ∼1–200 cells of edge) and also potentially when cells were seeded for fewer days. Thus we conducted our experiments with confluences 40%–90% with cells seeded 1 to 2 days overnight. We did not notice a difference in the phenotype in our experiments with longer days of seeding as long as dragging was successful. We also note that there was a variability in the ability to drag based on the micropipette tip shape. Before each experiment, we made sure we were not dragging a region that was already compromised by the micropipette.
Mechanical displacement of single cells
For single cell experiments, where we displaced cells lacking cell-cell contacts, cells were seeded on collagen-coated glass for 2–4 h in 0.4–1.5 mL imaging media before imaging start (>70 time-lapses, four samples). Since this was a delicate experiment due to the ease with which single cells were damaged (judged by heavy blebbing) or completely detached, most of the time we chose to displace the cells with the micropipette while watching in brightfield mode, and we started imaging in TIRF as soon as we were done moving the micropipette. This was the case for the experiment shown in Fig. S4, and as indicated, we estimate there was a delay of ∼5 s between the displacement and imaging. We should note that some of the single cells were already lamellipodial and showed tubular and lamellipodial protrusions. Thus we took control images of the Lifeact-GFP signal of the cells before any mechanical perturbation to be able to make out the membrane tethers that were generated due to the displacement and were followed by distinct lamellipodial activity. We estimate below 10% rate for retropodia production by our protocol, since we chose not to continue imaging cases where cells were fully detached or looked too damaged. Out of >70 time-lapses, at least seven cells generated clear retropodia, with each of the four different samples containing at least one cell that generated retropodia.
Maintenance and imaging of JEG-3 cells
Wild-type and ERM-free JEG-3 cells (a generous gift from Bretscher lab) were obtained as stocks at passage 8 and passage 10 respectively. Cells were thawed from stocks and cultured in sterile-filtered MEM media containing phenol red and Glutamax supplement (ThermoFisher, 41090036), with added 10% FBS (Corning, 35-011-CV), and 1% penicillin-streptomycin (ThermoFisher, 15140122). During passaging, cells were rinsed with PBS (ThermoFisher, 10010049) and split by dissociation using 0.25% Trypsin-EDTA (Life Technologies, 25200056) and subculturing. Cells were passaged no more than four times.
For mechanical pulling experiments, cells were seeded overnight directly on 35-mm glass-bottom dishes with 10-mm microwells (#1.5 coverslip) from Cellvis (D35-10-1.5-N). Before imaging, cells were labeled with 3:10,000 dilution of CellMask Deep-Red in culture media for 20 min, then washed four times with MEM media containing HEPES, phenol red, and Glutamax supplement (ThermoFisher, 42,360,032), with added 1% FBS (Corning, 35-011-CV), and 1% penicillin-streptomycin (ThermoFisher, 15,140,122), in which they were imaged after 20 min of recovery in the incubator.
Zebrafish experiments
Animals and husbandry
Transgenic TgBAC(ΔNp63:Gal4)la213; Tg(UAS:LifeAct-EGFP)mu271 zebrafish (a generous gift from A. Sagasti) were raised according to standard procedures, outcrossed to TAB5 wild-type fish (26). Crossing and rearing of embryos was as previously described (27). All experiments were performed on larvae 72–108 h postfertilization to 3–4 days postfertilization (dpf). No significant differences in the cellular response were observed between 3 and 4 dpf. Experiments were approved by the University of Washington Institutional Animal Care and Use Committee (protocol 4427-01).
Preparation of larvae for imaging
At 3 dpf, larvae were screened for LifeAct expression. At 3 or 4 dpf, larvae were anesthetized in E3 medium + 160 mg/L tricaine (Sigma E10521) + 1.6 mM Tris (pH 7), referred to as E3 + tricaine. Larvae were mounted in 35-mm #1.5 glass-bottom dishes (CellVis D35C4-20-1.5N) with 1.2% low-melt agarose (Invitrogen) in E3 + tricaine. Excess agarose was removed from around the tail of each larva with a #11 scalpel and mounted on the microscope for imaging at room temperature.
EDTA experiments
Before imaging, the media around mounted larvae were exchanged for E3 + tricaine +135 mM NaCl + 2 mM EDTA. Without prior injury, the addition of EDTA did not generate an acute effect on basal cells. However, even a relatively small injury to the skin would eventually lead to basal epidermal cell rounding and retraction. To control the onset of this process, one or two small punctures in the epidermis were introduced with a glass needle formed from pulling a 1-mm solid glass rod with a Brown-Flaming type micropipette puller (Sutter P-87). The addition of 135 mM NaCl in the medium prevented an extensive migratory wound response that would obscure the retraction induced by EDTA (27).
Zebrafish confocal imaging
Images were acquired using a Plan Apo 60× NA 1.27 objective on a Nikon Ti2 inverted microscope, equipped with a piezo-z stage (Applied Scientific Instruments, PZ-2300-XY-FT), a Yokogawa CSU-W1 spinning disk confocal with Borealis attachment (Andor), a laser launch (Vortran VersaLase) with 50-mW 488-nm diode laser (Vortran Stradus), 405/488/561/640/755 penta-band dichroic (Andor), and a 535/50m emission filter (Chroma). A 2× optovar within the spinning disk unit was used to increase the magnification. Full-chip 16-bit 1024 x 1024 pixel images were acquired with a back-thinned EMCCD camera (Andor DU888 iXon Ultra) with Frame Transfer mode and EM Gain applied. MicroManager v1.4.23 was used to control all equipment (25). Z-stacks with 0.2–0.25 nm spacing were acquired every 3 s, and maximum intensity projections along the Z axis were used for further analysis.
Image processing and analysis
Images were processed in FIJI (28). Image montage sequences at different time intervals and the videos were generated either directly or after downsampling image stacks with the “Reduce” command. Montages were generated using the “Make Montage” command or by separately saving images within a stack. Images within a given montage sequence were kept at the same Brightness & Contrast scaling, whereas montages of zoomed-in regions were in general scaled differently than the montages of the larger regions they belonged to. Although this contrast adjustment may lead to some oversaturation, we found that it was helpful in highlighting the features in question. The zoomed-in region in Fig. S1 c was contrast-enhanced. Background Subtractor at a sliding window of 100 pixels was applied to images in Fig. 1 a except for the kymograph, which was made using the raw image sequence that extended in time before and after the montage sequence of the figure. Exponential bleach correction was applied in the montages of Fig. 2 b and c and in videos when indicated in the legend. The right panel of Video S8 was made by applying to the raw video “Laplace Filter” from MosaicSuite (29) and then the “Smooth” and “Invert” commands respectively.
Kymographs were made with the Kymograph Builder plugin in FIJI, using raw F-actin (Lifeact-GFP) images captured continuously at 200-ms exposure. Linewidths of 3, 3, and 5 pixels were used in Figs. 1 d, S2, and 2 c respectively.
Quantification
For the chemical disruption (EDTA) experiments with MDCK cells, multiple F-actin (Lifeact-GFP) time-lapses from three separate samples of collagen-coated dishes and two separate samples of fibronectin-coated dishes were collected, with the time of initial EDTA addition noted. Among the videos within each sample, we analyzed the first two experimental regions (full field of view of camera) where the cells started separating enough to a point we could relatively easily make out different cells. We counted 39 cells in each region on average. For each video, the first 510 frames acquired continuously at 200 ms were reduced by 50 frames in FIJI to ease viewing. There were some ambiguous or small cell footprints, for which we used the same criteria across regions when annotating. For the analyzed videos, we labeled all the cells whose footprint was mostly within the field of view, and we counted the cells with retropodia by going through the cells in order. Cells were counted as having retropodia 1) when a lamellipodium spread through a pair of membrane tethers in a different manner from the outside of the tethers or 2) a lamellipodium preferentially traced a single membrane tether. Some of the cells within a region had blebs, which we also annotated: 0%–40% of the cells in each region were found to contain blebs.
For the multi-cell micropipette experiments with MDCK cells, among the captured videos, we analyzed the first five Lifeact-GFP imaging and pulling time lapses for each of the two fibronectin-coated dish samples, and the first seven Lifeact-GFP imaging and pulling time lapses of the one collagen-coated dish sample. The first three time lapses of one sample were discarded due to having low contrast, which we suspect was due to a combination of poor TIRF angle and focusing. Among the videos, there was a varying extent of ripping, but all videos showed membrane tether formation and thus had mechanically displaced (peeled or separated) cells. We labeled all the cells that had any part of their footprint affected by the pull (deformed and/or displaced), including those partially outside the field of view. We included cells that had noticeable gaps opening with the deformed or displaced cells, even if they were otherwise not affected by the pull. On average, 20 cells per analyzed experimental region (full field of view of camera) were affected by the pull. In two of the 17 analyzed videos, multiple border cells of a colony were also significantly displaced by the mechanical pull, in addition to the cells in the bulk, and some of those produced retropodia. We chose to not distinguish between border cells and bulk cells in our quantifications of these two videos.
Results
We examined how epithelial cells reestablish cell adhesion following a variety of chemical and mechanical perturbations. Cell adhesion to the ECM is largely dependent on integrins, a class of transmembrane adhesion proteins whose full activity requires Mg2+ (30,31). We treated MDCK cells with 5 mM EDTA, a Mg2+ chelator, to acutely weaken integrin-based adhesion to the ECM (31,32) and imaged the cellular basal surface using TIRF microscopy, which allowed us to visualize basal protrusions with high contrast and temporal resolution (Fig. S1 a). We found that, within 40 min of EDTA addition, many cells formed long membrane extensions that, for generality, we hereafter term membrane tethers (Fig. 1; Video S1). Lamellipodia bridging adjacent tethers extended outward from the cell body toward the original points of adhesion (Fig. 1 a–e; Video S1). We hereafter refer to tether-guided lamellipodia as “retropodia” to denote their role in respreading. EDTA-induced detachment of cells from the substrate was not synchronous across a single field of view. In regions where cells showed partial delamination (see Materials and methods), we recorded retropodia in 20%–60% of the cells, with each region containing ∼40 cells visible in TIRF (Fig. 1 f). Under these conditions retropodia could extend up to >4 μm and move at speeds of up to >150 nm/s (Fig. 1 d; Fig. S2), which is similar to lamellipodial edge spreading speeds, for example in migrating fish keratocytes (33).
F-actin (Lifeact-GFP) signal of a peeling region starting at 34 minutes after EDTA addition. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
We next examined how cells responded to mechanically induced detachment from the ECM. We built a micropipette pulling setup with which we could simultaneously and stably image in TIRF at high spatiotemporal resolution (Fig. 2 a). By laterally pulling on cell sheets with the micropipette, we were able to rapidly (∼3 s) displace small regions of the MDCK sheet. Under these conditions, cells extruded multiple membrane tethers, some of which reached ∼20 μm in length without snapping (Fig. S3). As in the chemical disruption experiments, cells produced retropodia that extended >4 μm and moved at speeds of >150 nm/s (Fig. 2 b–d; Figs. S2 and S6; Videos S2, S3, S4, S5, S6, and S7).
Membrane (Myr-RFP) signal of a multicellular region before and after peeling (bleach corrected). Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal from a multicellular region (within Video S6) before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Scale bar, 1 μm. Time is shown as min:sec.
We next tested whether cell-ECM junctions were sufficient for the formation of mechanically induced retropodia, or if instead cell-cell contacts were required. With care, we managed to push single cells that lacked cell-cell junctions sufficiently gently to displace them from the coverslip without killing or fully detaching them. Although the behavior was heterogeneous (see Materials and methods), we saw unambiguous examples of retropodia formation (Fig. 2 e; Fig. S4; Video S8). Thus, cell-cell junctions are not necessary for retropodia formation.
Left panel: F-actin (Lifeact-GFP) signal of a single cell with no neighbors, before and after peeling. Arrows indicate select regions with retropodia. Right panel: Laplace filtered, smoothed, and inverted F-actin (Lifeact-GFP) signal to highlight the thin membrane tethers in the displaced region (see Materials and methods). Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-EGFP) signal of a region of basal epidermal cells 7 minutes after puncture of the larval epidermis incubated with 2 mM EDTA. Arrow indicates an emerging retropodium. Scale bar, 5 μm. Time is shown as min:sec.
Retropodia predominantly appeared on the side of cells facing away from the direction of pull in our mechanical pulling experiments (Fig. S6). We used small-molecule inhibitor experiments to examine whether specific signaling pathways were likely to be required specifically for retropodia formation (Table S1). Signaling through phosphoinositide 3-kinase (PI3K) plays a central role in polarized cell migration in numerous systems. However, treatment with the PI3K inhibitor wortmannin did not block retropodia formation (Video S12), nor did treatment with the phosphatidylinositol (4,5)-trisphosphate sequesterer neomycin (Table S1). Similarly, inhibition of phospholipase C, which often acts in concert with and downstream of PI3K in setting up directional migration, did not affect retropodia formation (Table S1). As positive controls, disruption of actin filaments with cytochalasin D blocked retropodia formation (Videos S13 and S16) and disruption of Arp2/3-mediated actin polymerization with CK666 resulted in a clear reduction in retropodia formation (Table S1; Video S17). Retropodia were able to form in absence of IRSp53, a curvature sensing regulator of protrusions in MDCK cells (34,35) (Video S14). Indeed, no single perturbation targeting some typical upstream players in protrusive processes (Table S1) blocked retropodia formation.
Ezrin (Ezrin-GFP) signal of a multicellular region before and after mechanical peeling. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
Membrane (CellMask Deep-Red) signal of a multicellular region before and after mechanical peeling in ER-KO JEG-3 cells. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region before and after mechanical peeling, after 1 hr 20 min treatment with 10 μM wortmannin. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region before and after mechanical peeling, after 30 min treatment with 10 μM cytochalasin D. Scale bar, 5 μm. Time is shown as min:sec.
Membrane (CellMask Deep-Red) signal of a multicellular region before and after mechanical peeling in IRSp53 knockout MDCK cells. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region before and after mechanical pull occurs outside the field of view (lower right corner) at ∼16 sec. Scale bar, 5 μm. Time is shown as min:sec.
Membrane (Myr-RFP) signal of a multicellular region before and after mechanical peeling, after >30 min treatment with 10 μM cytochalasin D. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
It is possible that the formation of retropodia is mediated by redundant pathways, or by an atypical or altogether novel upstream pathway. However, an alternative explanation is that a physical cue, for example membrane tension or curvature, present at the base of membrane tethers acts to localize retropodia formation. Consistent with this possibility, in many instances the retropodial edge curved inward (Fig. S5 a), and flanking tethers bent inward as the retropodium advanced (Fig. S5 b); both observations suggest that membrane stresses at the base of tethers may help localize extension (36). We noted as well that cells in the immediate vicinity of a pulled region often exhibited nondirectional membrane ruffling (Videos S6 and S15). This latter observation argues that although membrane tethers direct retropodial extension, they are not strictly required for lamellipodia formation.
Several biophysical mechanisms apart from membrane tension could in principle act to direct the formation of retropodia. One such possibility is alterations in membrane-cortex attachment, which is mediated in part by ezrin-radixin-moesin (ERM) proteins (37,38). We confirmed that ezrin, which links the cell membrane to the actin cortex, was indeed recruited to the base of membrane tethers as well as retropodia nucleation points, though in a heterogeneous fashion (Video S10). To explore this observation further, we used a recently developed ezrin-radixin double knockout in the JEG-3 human epithelial cell line (39) (hereafter, ER-KO JEG-3) to test the role of ERM proteins in retropodia formation. Since JEG-3 cells do not express detectable levels of moesin, ER-KO JEG-3 cells are essentially free of ERM proteins. Both ER-KO and wild-type JEG-3 cells generated retropodia (Video S11). Thus, retropodia formation does not require ERM proteins, though we cannot rule out the involvement of other cortical linkers. Instead, and consistent with previous studies (40), it seems more probable that ezrin, and by extension other cortical linkers, are recruited to reinforce membrane-cortex attachment once retropodia form.
Finally, we examined whether retropodia helped to mediate wound healing in vivo, using the larval zebrafish as a model system. The zebrafish epidermis is formed of two layers—an outer layer containing tight junctions, and a basal layer that adheres to the basement membrane (Fig. 3 a). Micropipette-based pulling experiments, analogous to those in cell culture, proved to be technically challenging (data not shown). However, EDTA-based chemical wounding was compatible with high-speed imaging. The chelation of divalent cations by EDTA mimics an acute drop in “water hardness,” a critical parameter for aquatic organisms reflecting the concentration of dissolved divalent cations in the environment (41, 42, 43). Focusing on the basal cells, we found that introducing a small puncture into the skin allowed EDTA to penetrate past the tight-junction-containing outer layer and induce cell rounding and retraction, comparable to MDCK monolayers (Fig. 3 b). Focusing on actin dynamics using a transgenic fish in which LifeAct-EGFP was expressed in the basal layer, we found that upon EDTA treatment cells initially retracted, leaving behind actin-rich tethers. As in cell culture, instances of retropodia could be observed emerging between and around membrane tethers in multiple zebrafish (Fig. 3 c; Video S9), before broader, fan-shaped protrusions dominated. These observations confirm that retropodia occur in zebrafish epithelia, suggesting that they may help guide cell reattachment in vivo.
Figure 3.
Retropodial readhesion is a recovery mechanism in vivo. (a) Basal epidermis of the zebrafish larva was imaged in live animals 3 to 4 days postfertilization. TJ: tight junction, localized in the superficial epidermal layer. (b) F-actin (Lifeact-EGFP) signal of a region of basal epidermal cells at three timepoints, 6 s after puncture of the larval epidermis in the presence of 2 mM EDTA in the surrounding media. Cell rounding and retraction comparable to MDCK cells were both observed. (c) F-actin (Lifeact-EGFP) signal of a region 9 min after puncture of the epidermis in the presence of 2 mM EDTA. Time course from a zoomed-in section highlights retropodial extension (yellow arrows). Scale bars, 5 μm. Time is shown as min:sec. To see this figure in color, go online.
Discussion
As the outermost layer of organs, epithelia experience large, localized forces due to pathogen infiltration, immune cell entry, wounding, and cell rearrangements. Epithelial cells have evolved multiple strategies to respond to these and other circumstances to maintain tissue integrity. We propose that retropodial respreading constitutes an additional, understudied means by which cells can reestablish tissue integrity on the seconds timescale.
The formation of retropodia could in principle be regulated by intracellular signal transduction pathways. In some circumstances, for example during neuronal growth cone extension (44), cell migration is guided by filopodia (45, 46, 47). It has been proposed that filopodia and retraction fibers are interconvertible (18), suggesting that tubular membrane protrusions may play a general role in guiding lamellipodial extension. However, to our knowledge, the underlying mechanisms by which this guidance occurs remain obscure. In our study, an examination of the pathways known to regulate polarized lamellipodial extension, for example lipid signaling via PI3K, did not reveal obvious candidates for upstream molecular players, nor did the perturbation of kinases with roles in mechanical signaling (focal adhesion kinase, integrin-linked kinase, epidermal growth factor receptor; Table S1). IRSp53, which can play a prominent role in lamellipodium initiation (34), and ERM proteins, which help anchor actin filaments to the membrane, were likewise not required for the formation of retropodia. Although these experiments argue against a requirement for the above pathways for retropodia formation, we do not exclude the possibility that more subtle features of retropodia, such as their morphology or frequency per cell, may be dependent on these pathways. The robustness of retropodia formation to perturbation of any one signaling pathway is nonetheless striking.
A plausible explanation for the mechanism of retropodia formation is that retropodia nucleation and extension is primarily guided by the physical presence of membrane tethers. The appearance of many concave retropodial fronts suggests the possibility of significant interfacial tension. A possible explanation is that decreased membrane-cortex attachment (37,38) and/or membrane tension gradients (48) along retropodial fronts may act to recruit or stabilize F-actin polymerization factors, decrease the resistance to the growth of the branched F-actin network, or both. Another recent study suggests that the geometric interplay of membrane curvature and Arp2/3-medated branched actin polymerization can accelerate membrane protrusion in regions of negative membrane curvature, as occurs between membrane tethers, even absent upstream signaling (49). These possibilities are nonexclusive, and it is plausible that their relative importance may be context dependent.
It will be useful to understand the commonalities and differences between retropodia and other examples in which concave membrane fronts sustain lamellipodia, such as in between filopodia in neuronal growth cones, and between suspended noncellular fibers on which cells can migrate (50). Lamellipodial ruffling can in some cases emerge directly on filopodia to form “dactylopodia” (51), which may also be mechanistically related to retropodia formation. It is likewise possible that retropodia may possess similarities to the lamellipodia-free concave edges of cells adhering to stripes of ECM (52), or the actin structures that form during the repair of holes in lamellipodia (53). These possibilities await further investigation.
In this study we examined retropodia formation in the specific context of epithelial detachment from the extracellular matrix. Although intercellular cadherin trans interactions were not required for retropodia formation in our study (Fig. 2 e; Fig. S4; Video S8), it is possible that they may play an analogous role as anchor points or force transducers in other contexts (54). More broadly, it is possible that force-generated membrane tethers may have a general though understudied role as “safety lines” that preserve a physical linkage to the cell's original attachment points. The rate at which these tethers extended under external force, at least in our study, was striking: we estimate extension rates of >5 μm/s, or greater than fivefold faster than the extreme speeds of Dictyostelium filopodial growth (55). We suggest that membrane tethers function analogously to the dragline of spiders and a variety of insects (56,57), which can also extend rapidly when the animal loses its footing. Overall, our results add to the diverse functions provided by membrane tethers, which include aiding in neutrophil rolling (58) and in forming tunneling nanotubes that communicate intercellular information (59).
Author contributions
E.K. and A.R.D. conceived the idea. E.K. performed the experiments with MDCK and JEG-3 cells. A.S.K. performed the experiments with zebrafish. C.G.C. and C.G.V. contributed early training, and C.G.V. contributed reagents. E.K. and A.R.D wrote and all authors edited the manuscript.
Acknowledgments
E.K. is supported by the Stanford Bio-X Graduate Fellowship and the Biophysics Ph.D. Program at Stanford University. A.S.K. is supported by NIGMS Training Grant T32GM008294. C.G.C is supported by a long-term postdoctoral fellowship from the Human Frontier Science Program. C.G.V. is supported by the NIH NIGMS (1F32GM125113-01). The IRSp53 knockout and control MDCK cell lines were generous gifts from Giorgio Scita. The ezrin-radixin double knockout JEG-3 cell line with undetectable moesin expression and the control cell line were generous gifts from Anthony Bretscher. A.R.D. acknowledges the HHMI (Faculty Scholar Award), and the NIH (R35GM130332). We are grateful to Dr. Christina Hueschen, Dr. Steven Tan, Vipul Vachharajani, other members of the Dunn lab, Dr. Angela Barth, Dr. Lucy O'Brien, and Dr. Mark Peifer for discussion and comments on the manuscript.
Editor: Philip LeDuc.
Footnotes
Supporting material (Video S18. Retropodia formation or ablation in presence of different perturbations, Video S19. Retropodia formation or ablation in presence of different perturbations, Video S20. Retropodia formation or ablation in presence of different perturbations, Video S21. Retropodia formation or ablation in presence of different perturbations, Video S22. Retropodia formation or ablation in presence of different perturbations, Video S23. Retropodia formation or ablation in presence of different perturbations, Video S24. Retropodia formation or ablation in presence of different perturbations, Video S25. Retropodia formation or ablation in presence of different perturbations, Video S26. Retropodia formation or ablation in presence of different perturbations, Video S27. Retropodia formation or ablation in presence of different perturbations, Video S28. Postprocessing highlights membrane tethers formed upon mechanical perturbation) can be found online at https://doi.org/10.1016/j.bpj.2022.02.006.
Supporting material
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
To highlight membrane tethers, Laplace filtering, smoothing, and inversion were applied to the bleach-corrected F-actin (Lifeact-GFP) signal of the multicellular region corresponding to Video S6. Scale bar, 5 μm. Time is shown as min:sec.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
F-actin (Lifeact-GFP) signal of a peeling region starting at 34 minutes after EDTA addition. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
Membrane (Myr-RFP) signal of a multicellular region before and after peeling (bleach corrected). Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of multicellular regions before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Pulling induces lamellipodial ruffling in cells without retraction fibers as well, for example as in upper right corner of Video S6. Membrane tethers in Video S6 are highlighted in Video S28 through postprocessing. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal from a multicellular region (within Video S6) before and after mechanical peeling (bleach corrected). Arrows indicate select regions with retropodia. Scale bar, 1 μm. Time is shown as min:sec.
Left panel: F-actin (Lifeact-GFP) signal of a single cell with no neighbors, before and after peeling. Arrows indicate select regions with retropodia. Right panel: Laplace filtered, smoothed, and inverted F-actin (Lifeact-GFP) signal to highlight the thin membrane tethers in the displaced region (see Materials and methods). Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-EGFP) signal of a region of basal epidermal cells 7 minutes after puncture of the larval epidermis incubated with 2 mM EDTA. Arrow indicates an emerging retropodium. Scale bar, 5 μm. Time is shown as min:sec.
Ezrin (Ezrin-GFP) signal of a multicellular region before and after mechanical peeling. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
Membrane (CellMask Deep-Red) signal of a multicellular region before and after mechanical peeling in ER-KO JEG-3 cells. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region before and after mechanical peeling, after 1 hr 20 min treatment with 10 μM wortmannin. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region before and after mechanical peeling, after 30 min treatment with 10 μM cytochalasin D. Scale bar, 5 μm. Time is shown as min:sec.
Membrane (CellMask Deep-Red) signal of a multicellular region before and after mechanical peeling in IRSp53 knockout MDCK cells. Arrows indicate select regions with retropodia. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region before and after mechanical pull occurs outside the field of view (lower right corner) at ∼16 sec. Scale bar, 5 μm. Time is shown as min:sec.
Membrane (Myr-RFP) signal of a multicellular region before and after mechanical peeling, after >30 min treatment with 10 μM cytochalasin D. Scale bar, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
F-actin (Lifeact-GFP) signal of a multicellular region either before and after mechanical pull, or after EDTA treatment. See Table S1 for the specific perturbations corresponding to each video and the results. Scale bars, 5 μm. Time is shown as min:sec.
To highlight membrane tethers, Laplace filtering, smoothing, and inversion were applied to the bleach-corrected F-actin (Lifeact-GFP) signal of the multicellular region corresponding to Video S6. Scale bar, 5 μm. Time is shown as min:sec.