
==== Front
Biophys J
Biophys J
Biophysical Journal
0006-3495
1542-0086
The Biophysical Society

S0006-3495(23)00274-6
10.1016/j.bpj.2023.04.023
Articles
Changes in cell surface excess are coordinated with protrusion dynamics during 3D motility
Kapustina Maryna mkapust@med.unc.edu
1∗
Li Donna 1
Zhu James 1
Wall Brittany 1
Weinreb Violetta 1
Cheney Richard E. 12
1 Department of Cell Biology and Physiology, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina
2 Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina
∗ Corresponding author mkapust@med.unc.edu
19 9 2023
18 5 2023
122 18 36563677
8 9 2022
20 4 2023
© 2023 Biophysical Society.
2023
Biophysical Society
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
To facilitate rapid changes in morphology without endangering cell integrity, each cell possesses a substantial amount of cell surface excess (CSE) that can be promptly deployed to cover cell extensions. CSE can be stored in different types of small surface projections such as filopodia, microvilli, and ridges, with rounded bleb-like projections being the most common and rapidly achieved form of storage. We demonstrate that, similar to rounded cells in 2D culture, rounded cells in 3D collagen contain large amounts of CSE and use it to cover developing protrusions. Upon retraction of a protrusion, the CSE this produces is stored over the cell body similar to the CSE produced by cell rounding. We present high-resolution imaging of F-actin and microtubules (MTs) for different cell lines in a 3D environment and demonstrate the correlated changes between CSE and protrusion dynamics. To coordinate CSE storage and release with protrusion formation and motility, we expect cells to have specific mechanisms for regulating CSE, and we hypothesize that MTs play a substantial role in this mechanism by reducing cell surface dynamics and stabilizing CSE. We also suggest that different effects of MT depolymerization on cell motility, such as inhibiting mesenchymal motility and enhancing amoeboid, can be explained by this role of MTs in CSE regulation.

Editor: Yu-Li Wang.
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pmcSignificance

Many cells exhibit rich phenotypic plasticity during locomotion and adapt their mechanisms of migration to fit the three-dimensional microenvironment, which includes the ability of a cell to achieve rapid large-scale changes in shape. We believe that a key part of this mechanism is a highly convoluted cell surface populated with blebs, microvilli, filopodia, and folds that can accommodate a large amount of surface surplus. The process of release or accumulation of this surface surplus provides a simple and effective way of transforming cell morphology. We anticipate that our work will find applicability to various types of cell shape transformations including cell division and migration. It also has implications for understanding cell biomechanics.

Introduction

Crucial cell functions including migration, cytokinesis, and differentiation involve major changes in cell shape. Morphological transformations often result in dramatically different cell shapes, requiring cells to develop efficient mechanisms to manipulate surface area while keeping volume nearly constant. The periphery of the cell, which we term the cell surface, must be very flexible to accommodate fast shape changes during migration while maintaining the integrity of the cell. It is comprised of the plasma membrane (PM) with a thin (∼<200–500 nm) layer of cytoskeleton structure known as the cell cortex, which is coupled to the PM by adaptor proteins and consists of an F-actin network cross-linked by actin-binding proteins (1,2,3,4,5,6). Importantly, despite the flexibility of the PM, it can only undergo ∼3% areal extension without rupturing (7,8,9,10). Thus, to adapt to diverse environmental requirements, a sufficient reservoir of the PM that is immediately deployable is critically important for cell survival.

Pioneering work with scanning electron microscopy (SEM) (11,12,13,14) revealed that the surface of rounded cells is covered with numerous small surface projections such as blebs, microvilli, filopodia, and folds. These surface projections largely disappear upon cell spreading, leading to the hypothesis that they serve as a reservoir for storing the excess of the PM that can be flattened and reused during surface expansion. Studies from the past decades provided evidence that cell surface excess (CSE) can be utilized to cover increasing surface areas or to buffer mechanical stresses occurring in diverse physiological processes such as phagocytosis, cellularization in Drosophila embryos, cell spreading, and many others. (13,14,15,16,17,18).

In our previous work (19,20,21) we investigated the striking morphological oscillations of rounded cells that manifest themselves as highly periodic and repeatable shape changes that can last for many hours. These oscillations in cell shape can appear spontaneously after large morphological changes such as transformation from spread to rounded cell shape and can be enhanced by microtubule (MT) depolymerization. Presuming that the cell volume stays constant, conversion from a spread to a rounded state requires a cell to accommodate the large amount of cell surface above the minimum required to cover the rounded body. Using light and electron microscopy we demonstrated that cells store this excess by the formation of small surface projections and folds. The oscillations are due to the periodic accumulation and then flattening of these small projections and folds, which often create an appearance of a traveling wave of cortex density that moves around the cell with a period of 40–200 s (Video S1). Note, that the intersection between the direction of the traveling wave and microscope focal plane creates a variety of microscope images of cell oscillations. Importantly, we showed that during this process the PM and underlying actin mesh remain intact, making CSE a highly dynamic structure that cells can rapidly deploy during changes in shape. We also suggested that this mechanism for rapidly altering cell surface area is essential for many morphological transformations and can provide a model for cell shape transformation during migration.

Video S1. Traveling wave of cortex density

The video presents two oscillatory CHO cells with stable transfection of Lifeact-GFP for F-actin (green) and myosin light chain-mRFP for Myosin II (red). The traveling wave is only obvious on cell 2, where the dense portion of the cortex moves around the cell periphery. Importantly, despite the visual illusion of density movement around the cell, no mass is actually being transported; rather, a compression wave is propagated with new folds and small surface protrusions generated at the leading edge of the wave and flattening at its trailing edge (A). The morphological oscillations are a 3D phenomenon, and the traveling wave can propagate in any direction on the cell periphery and not always be apparent in confocal sections. For example, the morphological changes and traveling wave around cell 1 look completely different because the traveling wave propagates in a direction normal to the substrate. If we would section cell 2 in the perpendicular direction to imaging on (A) we would see an image similar to cell 1 (B)

Many cells exhibit rich phenotypic plasticity during locomotion and adapt their mechanisms of migration to fit their 3D tissue microenvironment (22,23,24). The two major modes of motility, mesenchymal and amoeboid, have different cytoskeletal, biomechanical, and signaling properties (25,26,27). Cells utilizing the mesenchymal mode of motility rely on acto-myosin fibers to produce contractile forces, on strong focal adhesions to the extracellular matrix (ECM) to apply traction forces, and on ECM-degrading enzymes to perform ECM remodeling and generate a path for translocation (28). Amoeboid migration commonly refers to the movement of more rounded cells that lack mature focal adhesions and stress fibers (27,29,30). This migration style was first observed in amebae and later found in many other cells, including leukocytes and certain types of tumor cells. A characteristic feature of amoeboid migration is a high level of acto-myosin contractility at the cell cortex, which is widely accepted to be primarily mediated by Rho signaling (31). The amoeboid mode of migration allows cells to move at high velocities by adapting their bodies to the pre-existing spaces and squeezing through the gaps in the ECM fibers without developing mature adhesions or inducing ECM degradation (32,33).

In this paper, we define CSE as the difference between actual cell surface area, which includes the surface area of all small projections, and the area required to cover the cell volume with a smooth surface layer. We demonstrate that cells in a 3D collagen environment maintain a large amount of CSE stored in small surface projections. We also demonstrate that the process of releasing stored surface by flattening small surface projections is coordinated with the extension of large cell protrusions. Conversely, the process of forming small surface projections, primarily small spherical projections that have the appearance of blebs, is coordinated with the retraction of the cell protrusions. We also demonstrate that an intact MT system supports proper CSE regulation and stability. Using several inhibitors, we showed that the mechanism by which MT affects CSE dynamics can be more diverse than the known regulation of Rho-GTPase activity (34). We suggest that, despite many differences between mesenchymal and amoeboid motility, both modes of motility rely on the proper regulation of CSE.

Methods

CHO-K1, MDA-MB-231, DU-145, and U937 cells were obtained from the Tissue Culture Facility of Lineberger Comprehensive Cancer Center, UNC at Chapel Hill. Walker carcinoma (WC) cells were acquired from the ATCC (Manassas, VA). CHO cells stably expressing Lifeact-GFP (the small 17-amino acid peptide, Lifeact, fused to GFP) and CHO-Lifeact-GFP cells expressing myosin regulatory light chain fused to RFP (MLC-RFP) were a gift from the James Bear laboratory (UNC at Chapel Hill). All types of culture media were obtained from GIBCO/Thermo Fisher Scientific, Waltham, MA, USA. MDA-MB and DU-145 cells were cultured in DMEM medium with 10% FBS (Gibco/Thermo Fisher Scientific). CHO cells were grown in DMEM/F12 medium containing 10% FBS and 4 mM L-glutamine. WC and U937 cells were grown in RPMI medium containing 10% FBS. All media contained 100 U/mL penicillin/streptomycin. Colchicine (Sigma-Aldrich,St. Louis, MO, USA), LPA (Sigma-Aldrich), Rho inhibitor C3 (Cytoskeleton Inc., Denver, CO, USA), Calyculin (Cell Signaling, Danvers, MA, USA), RhoA inhibitor Rhosin, and Rho kinase inhibitor Y-27632 (Tocris Bioscience, Minneapolis, MN, USA) were used for cell treatment during the experiments.

Electron microscopy

The samples for SEM and transmission electron microscopy (TEM) were prepared according to the protocol published previously (21,35).

Collagen preparation

Bovine collagen I solution from Advanced BioMatrix (Carlsbad, CA, USA): PureCol (3 mg/mL, no. 5005) or PureCol EZ Gel (5 mg/mL, no. 5074) were used for experiments. NaOH (0.5 M) was used to adjust the pH in the case of PureCol solution. For the experiments, we used glass-bottomed dishes, 14 mm glass, N = 0 (Mattek, Ashland, MA,USA or CellVis, Mountain View, CA, USA). The thin glass (N = 0) allowed us to visualize a thicker slice of the collagen matrix. All gel preparations were carried out on a table cooler located inside a tissue culture hood. To ensure that experimental cells were located inside the collagen matrix and were far from the hard substrate, we embedded cells in a collagen “sandwich.”

For the first layer of the sandwich we used 40 μL of cold collagen solution adjusted appropriately for the cell line medium to the concentration used in the experiment. This solution was distributed evenly on the bottom glass and allowed to polymerize for 10 min in the incubator. Then 100 μL of collagen solution with the cells (∼106/mL) were placed on top of the first layer and polymerized for 1 h in the incubator. After 1 h, 500 μL of medium was added to the top of the collagen sandwich and the dish was either kept in the incubator or used for recording.

For visualization of collagen structure, 10% of the collagen volume was replaced with collagen-FITC (Sigma-Aldrich) and used for creating the collagen matrix.

Collagen fixation and cell labeling

The collagen matrix with cells was fixed using 2 mL of warm (37°C) paraformaldehyde solution (4%, w/v, in PBS [pH 7.4]) for 20 min, washed twice with PBS, and incubated with blocking/permeabilization solution containing 5% goat serum, 1% BSA, and 0.1% saponin in PBS for 1 h at room temperature. Samples were incubated overnight at 4°C with primary antibodies followed by three washes and incubation for 1 h at room temperature with appropriate secondary antibodies.

All primary antibodies and their matching species-specific Alexa Fluor-conjugated secondary antibodies were diluted in PBS supplemented with 1% BSA and 0.1% saponin. Excess antibodies were removed by three washes with PBS containing 0.05% saponin, and the nuclei were counterstained with Hoechst 33342 (1:2000) (Thermo Fisher Scientific, Waltham, MA, USA) for 8 min at room temperature. After washes with PBS containing 0.075% saponin, collagen with cells were washed with PBS, covered in 1 mL PBS, and directly visualized under a confocal microscope as described below. Collagen fixation, labeling, and washing were all performed extremely gently to avoid perturbing the collagen structure or detaching the matrix from the glass.

Primary antibodies used in the experiments: Anti-vinculin (1:200) (Sigma-Aldrich, SAB4200729), Anti-tubulin (Invitrogen/Thermo Fisher Scientific, PA5-85133, TU-01, 1:200; and Novus 6-11B-1, 1:100) To visualize F-actin in fixed cells we used Alexa Fluor 488 or 568 Phalloidin (Invitrogen/Thermo Fisher Scientific) (3:100); species-specific Alexa Fluor-conjugated secondary antibodies were purchased from Thermo Fisher Scientific (nos. A-11001, A-11008, A-11011, A-11004, A-21235, and A-21244) and used at 1:1000. The PM in fixed cells was stained by CellMask Plasma Membrane Stain (Thermo Fisher Scientific no. C10046) at 1:1000 for 5 min before fixation.

For PM visualization in live cells, we transiently expressed the PMT-mRFP, which is a lipid-linked protein that resides almost exclusively on the inner leaflet of the PM (36). For MT visualization in live cells, we transfected CHO cells with EMTB-3xGFP (Addgene,Watertown, MA, USA, no. 26741). The transfections were performed using Mirus (Marietta, GA, USA) transfection reagent following the manufacturer’s instructions (TransIT-X2).

Microscope imaging

The long-term time lapses in DIC mode were recorded on an Olympus VivaView incubator microscope (Tokyo, Japan), which provides the ability to record up to eight samples simultaneously with full control over position and acquisition parameters. The images with fluorescence signals of cells in the collagen were recorded with a 40× silicon immersion objective using an Olympus FluoView1200 (Tokyo, Japan) laser scanning confocal microscope with an environmental chamber. The super-resolution images were acquired on Zeiss 880 microscope (Carl Zeiss Microscopy, LLC., White Plains, NY, USA) with Airyscan detector using a 40× oil objective.

Image analysis

Image analysis was performed using ImageJ and Imaris 9.7 software.

When cells are imaged using DIC mode within a large volume of collagen matrix and for extended periods of time, the analysis of cell movement becomes complicated. This is especially true for cell shape segmentation due to the low signal/noise ratio of the gray DIC signal. Therefore we term this method a speed estimation rather than a speed calculation. We estimated cell speed using the Spot detection tool with time tracking in the Imaris 9.7 software. For our analysis we used representative videos of cells moving inside the collagen matrix recorded for three different Z planes during two separate experiments for each inhibitor. The frame rate for all analyzed recordings was adjusted to be around 60 s. Under visual control, spots with a 10 μm diameter were created based on the intensity of DIC signal and their tracks were built using the autoregressive algorithm. We set the maximum gap size equal to 2 and the maximum distance to 20 μm. The track was considered successful and used for analysis if its length was larger than 50 μm and the displacement from the origin longer than 25 μm. The output provided an average instantaneous speed of all successful tracks for each time point. After tracking, the statistical data were collected and presented to MS Excel 365 and GraphPad Prism 9.5. We analyzed the effect of reagents 10 min after application of treatment to allow for the drug distribution inside the collagen. To verify our semi-automatic speed estimation we re-analyzed a set of cells using the Manual Tracking plugin from ImageJ and obtained similar results.

Statistical analysis

Statistical tests were performed using GraphPad Prism 9.5 software. The statistical tests applied to each quantification are indicated in each Figure legend. “+/−“ indicates standard deviations (SD).

Estimation of surface area

For rounded cells

To estimate the apparent surface of rounded cells we measured the cell perimeter on the cell equator after manual segmentation. The diameter was calculated as d = perimeter/π and the surface of a rounded cell was calculated as Surface=π∗d2.

Cigar-shaped cells

To estimate the surface area of a polarized cell with protrusions we assumed that the shape of the cell body can be approximately presented as a prolate ellipsoid (cigar shape) and be calculated after measurement of its axis: long (c) and short (a).Surfaceofprolateellipsoid=2∗π∗a2∗(1+ca∗e∗sin−1e)

wheree2=1−a2c2

The surface of the protrusion was estimated as a surface of a tube with a diameter measured in the middle of the protrusion length. For the measurement, we selected cells with symmetrical, cigar-shaped bodies that were fully visible on the focal plane during at least 10 consecutive frames.

Estimation of cell surface for irregularly shaped DU-145 cells in collagen

Using ImageJ software the cell shape was manually segmented and morphological parameters (area and perimeter) were measured. Cell surface was approximated as twice the visible area (2 ∗ area, apical, and ventral surface) plus the length of the perimeter multiplied by 1 μm (1 ∗ perimeter) to approximate the surface area due to cell thickness: S = 2 ∗ area + 1 ∗ perimeter.

2D area of amoeboid cells

To estimate the area for the amoeboid cells we manually outlined the cell periphery and measured the enclosed area for each time point in which the cell was fully visible in the focal plane.

Results

Rounded cells possess large amounts of CSE

When a spread cell (Fig. 1 A) is proteolytically detached, it rapidly rounds on a timescale of ∼30–60 s with a significant reduction in apparent cell surface area (Fig. 1 B). We estimated previously (35) that, despite limited time for membrane endocytosis, this reduction for CHO cells was 3.5 ± 2.25-fold and up to 12-fold for Swiss 3T3 cells. Importantly, the fluorescence signal from the cortical F-actin, which is practically unresolved in spread cells (37), becomes visible as a thick, bright band at the margins of rounded cells (Fig. 1 B and D). The same cells visualized using high-resolution SEM reveal a highly convoluted surface morphology that is capable of storing a large amount of surface area (Fig. 1 C, E, and F). Note that only fully spread cells have a smooth surface lacking small membrane projections. (Fig. 1 C and G). The morphology of rounded cells is characterized by folds and several types of small projections (Fig. 1 H): short spherical protrusions, which we call blebs, short and slender microvilli or filopodia-like structures, and ruffles. The typical length of small filopodia-like projections is between 200 and 2000 nm, with an average length of 626 ± 360 nm (n = 114). In our previous study we estimated that the average radius of a small bleb-like structure on a cell surface is below 250 nm with a wide distribution (35). TEM images of anti-GFP immunogold staining of rounded CHO cells stably expressing a biomarker of filamentous actin, Lifeact-GFP, revealed that all these small projections contain F-actin (Fig. 1 I and J). These small structures are tightly packed on the cell periphery (see the plot in Fig. 1 J) and are often unrecognizable as separate structures at optical microscope resolution (Fig. 1 D), where they appear as a thick homogeneous actin layer and are frequently assumed to be a continuous cortical actin network. A simple estimate of the perimeter length on a random position of a rounded cell from a TEM image (Fig. 1 I) demonstrates that the area with small projections has approximately threefold longer perimeter than a smooth surface without the projections, highlighting the immense amount of CSE contained in rounded cells.Figure 1 Rounded cells on 2D substrate possess ample amounts of cell surface excess. (A and B) The same CHO cells stably expressing Lifeact-GFP shown spread on glass (A) and immediately after rounding (B). (C) Scanning electron microscopy (SEM) image demonstrates the difference in the surface morphology between spread and rounded CHO cells. (D) Image of a single CHO cell stably expressing Lifeact-GFP in the rounded state (left, DIC; right, fluorescence) demonstrates the apparent thickness of actin cortex visible at optical microscope resolution. (E and F) Examples of SEM images of CHO (E) and DU-145 (F) cells fixed 20 min after rounding. (G) Transmission electron microscopy (TEM) image of fully spread CHO cells sliced perpendicularly to the substrate as shown on the cartoon above the image. This image demonstrates the absence of membrane folds or any other small structures on the surface. (H) Magnified view of three different types of small surface structures (blebs, ridges, and filopodia-like) on rounded CHO cells. (I and J) TEM image of GFP immunogold staining of rounded CHO cells with stable expression of Lifeact-GFP. Black dots, which correspond to gold particles, show the position of actin labeling. (I) Green and red lines illustrate the difference in the length cell periphery with small protrusions (red) and without (green) (ratio between two length is ∼3). (J) Plot of image intensity taken along the red line on (J). The plot demonstrates that the distance between small surface projections is in the range of 50–200 nm, which is below the optical resolution of a regular confocal microscope. Scale bars, 50 μm (A and B), 10 μm (C), 5 μm (D), 1 μm (E, F, H, and J), 2 μm (G), 500 nm (I). To see this figure in color, go online.

Electron micrographs demonstrate surface smoothing during protrusion

We hypothesize that a fast-growing cell protrusion can acquire most of its surface from the surface excess stored around the cell body (see the diagram in Fig. 2 A). This mechanism would be analogous to the initial stages of cell spreading, where cell surface stored in small surface projections on rounded cells is employed to cover the spreading areas. To find examples of such a mechanism, we used light microscopy and SEM to visualize the morphology of DU-145 cells (a human metastatic prostate cancer line) during the initial stages of cell attachment to a glass substrate. Light microscopy shows that the rounded DU-145 cells often develop short protrusions (Fig. 2 B, blue arrows; Video S2) that connect the cells with the substrate before cell spreading. Although these short protrusions can look like elongated blebs, they often persist for over an hour, much longer than the ∼1–3-min lifetime reported for blebs (38,39). We assume that these short protrusions can serve as an analogy to the protrusion in 3D collagen. SEM images in Fig. 2 C–F demonstrate that the surface of similar short protrusions is relatively smooth (green arrows) and their edges are characterized by elongated narrow filopodia-like or blunted projections that contact the coverslip (red arrows). The blebs that remained at the leading margin of the protrusion (red arrows) are in a position to provide a surface for further growth. Similar examples of bleb-like protrusions making contact with the substrate are shown in Fig. 2 G and H.Figure 2 CSE release during protrusion development and spreading. (A) Diagram illustrating the hypothetical process of protrusion development and retraction in correlation with CSE release and accommodation. (B) DIC image of DU-145 cells 30 min after plating on the glass. Note the short protrusions (blue arrows) that serve as the initial points of attachment. The cell bodies can hang on these “hands” for several hours before spreading occurs (see Video S2). (C–H) SEM images of DU-145 cells 30 min after plating on glass. The enlarged parts of image (D) in (E) (yellow box) and (F) (red box) provide several examples of the flat membrane surface on the main protrusion (green arrows) with blebs that seem to undergo transformation into small protrusive structures (red arrows). (C, G, and H) Images demonstrate a plasma membrane flattening and possible flow during spreading. Part of the cell is still covered with small surface structures, whereas on the opposite side these structures disappear and could provide cell surface for spreading. We presume that blebs close to substrate become elongated (red arrows) to induce cell spreading. (I and J) SEM images of CHO cell 30 min after plating on glass. Images demonstrate the same process of small surface structure flattening and possible plasma membrane flow during spreading as for the DU-145 cells. Scale bars, 20 μm (B), 5 μm (C, D, G, and I), 2 μm (H and J), and 1 μm (E and F). To see this figure in color, go online.

Video S2. Initial steps of spreading of DU-145 cells after plating on glass-bottomed dishes for 30 min

Note that rounded cell bodies move back and forth, whereas the protrusions extending from them stay adhere to the substrate

SEM images in Fig. 2 C and G also demonstrate that, whereas the cell surface distal to the extending region is covered with small projections holding a large amount of surface excess, these structures are flattened and the cell surface becomes much less convoluted in regions adjacent to the extension. The SEM images of rounded CHO cells revealed similar changes in cell surface morphologies during the initial steps of spreading (Fig. 2 I and J). The presented SEM images illustrate how small surface structures could be flattened, with the released surface reused for protrusion development and extension of the cell periphery.

CHO cells embedded in a 3D matrix preserve surface excess for successful morphological transformations

In our previous work (20), using detailed computer analysis of the traveling wave of cortex density on rounded cells placed on a 2D substrate we proved that the density of PM folds and cortex actin density are correlated in space and time. To confirm that the fluorescence signal from the cortical actin (Lifeact-GFP) can be used as a marker for the cell surface in 3D collagen, we analyzed the fluorescence signal from CHO cells with stably expressing Lifeact-GFP and transiently expressing the PM marker (PMT-RFP). Fluorescence images and plots of fluorescence intensity in Fig. S1 show that these signals are highly correlated during protrusion dynamics. The high-intensity signal from Lifeact-GFP that marks the position of F-actin colocalizes with the high-intensity signal from the PM marker, and both signals synchronously decrease within the subsequent 20 min. The correlation coefficient between the two signals was R = 0.89 (Pearson correlation, two experiments, six cells, five time points each). Therefore we concluded that, for rounded cells in 3D collagen, Lifeact-GFP provides a reliable marker for membrane-cortex density and morphology.

While cell rounding after detachment from a coverslip clearly demonstrates the phenomenon of CSE, it is not a naturally occurring condition, so we next explored whether different types of cells maintain CSE under physiologically relevant conditions such as during migration in a 3D collagen matrix.

First, we investigated the morphology and behavior of CHO cells. These cells do not possess the specific integrin (α2) for binding to collagen I fibers (40,41). In a 3D collagen matrix, they maintain either a rounded or a slightly polarized shape with infrequent protrusions (Fig. 3 A). Our analysis of CHO cells embedded in collagen matrices with different densities revealed that the cells develop lobopodial-type protrusions substantially more often in the dense collagen matrix (Fig. 3 C). This can be explained by the difference in the size of collagen pores, implying that a slim lobopodial protrusion can penetrate a narrower space between collagen fibers than a large rounded bleb. The morphology of CHO cells transiently transfected with α2-integrins shows that the presence of integrins increases the probability of developing stable protrusions even in the low-density collagen matrix. It also significantly (p < 0.001) increases the length of protrusions (Fig. S2).Figure 3 Cells embedded in a 3D matrix preserve surface excess and use it for protrusion development. (A) DIC image shows different morphology of CHO cells embedded in the 3D collagen for 24 h. (B) Cells embedded in a 3D matrix preserve CSE. The morphology of rounded CHO cell transfected with membrane marker mRFP-PMT and Lifeact-GFP shows that cells embedded in a 3D matrix are covered with CSE. (C) The chart presents the percent of CHO cells with lobopodial protrusions in different densities of collagen (p < 0.001, one-way ANOVA, error bars represent SD, N = 215, 267, 258, n = 2 experiments). (D) Merged DIC and F-actin fluorescence images of the CHO cell during protrusion retraction which results in cell blebbing and oscillations (Video S7). Note the smooth cell surface while protrusion is fully extended. (E and F) DIC image (E) and kymograph (F) of CHO cell in collagen during several cycles of protrusion-retraction (Video S8). (G and H) DIC and Lifeact-GFP fluorescence signal during protrusion retraction and rounding in the collagen matrix (time in seconds). (I) A kymograph at the position shown by the red arrow on (G). Note the presence of a smooth surface on the cell with large rounded protrusion, blebbing during retraction, and the beginning of oscillations after protrusion retraction (Video S9). The yellow arrow points at the position of CSE accumulation after protrusion withdrawal. The enlarged part of image (G) on (J) demonstrates how the morphological oscillations help to redistribute local CSE around the cell body. (K) Maximum intensity projection image of CHO-Lifeact-GFP cells in collagen before and during development of the protrusion. The image demonstrates the disappearance of blebbing during protrusion. (L–N) The charts visualize the results of quantitative analysis of cell protrusive activity in correlation with surface dynamics. The bar graph on (L) demonstrate high positive correlation between blebbing and protrusion retraction and high negative correlation between blebbing and protrusion development (p < 0.001, Spearman nonparametric test, n = 258 occurrences, 37 cells, N = 5). Scale bars, 10 μm (A, C, D, G, and H) and 5 μm (J and K). To see this figure in color, go online.

Surprisingly, rounded CHO cells that had been embedded for longer than 24 h in 3D collagen retained a convoluted surface morphology that can store a large amount of CSE, similar to that of newly rounded cells on a 2D substrate. Small surface projections labeled with both Lifeact-GFP and the PM tag are often clearly visible on the surface of the cells (Fig. 3 B). The cortical actin associated with these structures is underlain by a band of myosin II, very similar to CHO cells after rounding on a coverslip (Fig. S3 B).

The radius of rounded CHO cells embedded in collagen for more than 24 h was between 6 and 9.5 μm, with an average of 7.91 ± 1.41 μm (N = 128), which is not significantly different from CHO cells after rounding on a 2D substrate (8.07 ± 1.62 μm, N = 226, p > 0.1, t-test). These results clearly show that the CSE phenomenon is not an artifact of the experimental conditions, but is a part of normal cell physiology and therefore should be regulated by specific mechanisms.

In the absence of strong collagen adhesions and matrix-degrading proteases, CHO cells can move either by developing blunt cylindrical protrusions (Video S3) similar to lobopodial motility (42) or by using protrusive bulges (blebby type of amoeboid migration, Video S4). The latter has similarities to the compression and dilation seen in oscillatory cells where the combined image of DIC and F-actin (Lifeact-GFP) fluorescence signals shows stretching of the cell membrane-cortex layer during bulge extension (Fig. S3 C–E; Videos S5 and S6) and appearance of the traveling wave of cortex density around the cell periphery (Fig. S3 D) (21).

Video S3. An example of CHO cell migration in 3D collagen using a lobopodial type of protrusion

Note the periods of cell surface quietness (no visible blebbing) during protrusion extension

Video S4. An example of CHO cell migration in 3D collagen using blebby type ameboid motility

One cell is moving by developing a bulge and pushing through the collagen while another cell is oscillating without translocation. The green fluorescence signal on the stationary cell shows the localization of the F-actin marker (Lifeact-GFP)

Video S5. Oscillation of a CHO cell after embedding in 3D collagen for more than 24 h

The F-actin (Lifeact-GFP) fluorescence signal demonstrates stretching of the cell cortex during bulge extension

Video S6. CHO cell oscillations in collagen

In the collagen matrix, the periodic cell protrusions during the oscillation can occur in different directions. The top cell oscillates in the XY direction, whereas the bottom cell oscillates in the Z direction (in and out of the focal plane). The maximum intensity projection image illustrates the unfolding and stretching of the cortex (arrow) during shape transformations in 3D. The green fluorescence signal shows the localization of the F-actin marker (Lifeact-GFP)

To better understand the amount of CSE maintained by CHO cells during migration, we estimated how much surface excess is produced by retraction of the lobopodial protrusion, similar to the one shown in Fig. 3 D (Video S7). Assuming that the cell volume is conserved during protrusion retraction, our estimate shows that approximately 64% (240 μm2) of the protrusion surface in this particular cell is no longer needed for covering cell volume after protrusion retraction and therefore must be stored as CSE or endocytosed (Fig. S4). Our analysis of the large population of CHO cells with lobopodial protrusions (103 protrusions on 53 cells, average length of 23.4 ± 13.5 μm and width of 4.94 ± 1.4 μm) shows that between 60 and 80% of protrusion surface rapidly became CSE after protrusion withdrawal.

Video S7. This video of a CHO cell in 3D collagen demonstrates a smooth surface with fully extended protrusion and blebbing during and after protrusion withdrawal. Oscillations are initiated at the end of protrusion retraction

The green fluorescence signal shows the localization of the F-actin marker (Lifeact-GFP)

CHO cells employ CSE to cover the extension of protrusion while CSE is accumulated during retraction

According to our hypothesis, the flattening of pre-existing surface projections to release CSE should be observed in CHO cells during 3D migration. To test this, we imaged CHO cells embedded in collagen matrices for prolonged periods and analyzed their surface and protrusive dynamics. Fig. 3 E and Video S8 demonstrate a protrusive phenotype of CHO cells in collagen matrices where these cells undergo morphological oscillations and blebbing before the extension of a lobopodial-like protrusion. Simultaneously with the initiation of a protrusion, the morphological oscillations ceased and the blebbing decreased. After the extension of a large, stable protrusion, the cell body exhibited low surface dynamics and only a few blebs. A kymograph (Fig. 3 F) displaying changes in cell surface position as a function of time demonstrates a striking alteration between periods of high cell surface activity with oscillation of the cell body and surface relaxation during the extension of a protrusion.

Video S8. CHO cell in the collagen during phases of oscillation and protrusion. The video demonstrates a protrusive phenotype of CHO cells in collagen matrices in which these cells undergo morphological oscillations and blebbing before the extension of a lobopodial protrusion

Simultaneously with the initiation of a new protrusion, the morphological oscillations ceased and the blebbing decreased. After the extension of a large, stable protrusion, the cell exhibited low surface dynamics and only a few blebs

The transition from active blebbing to a smooth surface after protrusion growth is also clearly visible on confocal fluorescence imaging of F-actin (Fig. 3 K). In contrast, in the initial stages of protrusion withdrawal, the cell surface became covered with dynamic blebs, which were highly pronounced after retraction and during morphological oscillations initiated at the later stages of protrusion retraction (Fig. 3 G and H; Video S9). The retraction of a protrusion usually leaves a region on the cell periphery densely populated with blebs, which remain local until the excess surface stored in those blebs is redistributed around the cell surface. Fig. 3 J and Video S9 demonstrate how the CSE can be rearranged by the traveling wave of cortex density during the cell oscillations. Blebs and morphological oscillations often characterize the rounded non-polarized cell until a new protrusion emerges.

Video S9. Correlation between protrusion and blebbing. A CHO cell with a fluorescence signal from the F-actin marker (Lifeact-GFP) appears with a smooth surface while having a large round protrusion. In the initial stages of protrusion withdrawal, the cell surface became covered with dynamic blebs which were highly pronounced after retraction

During the later stages of protrusion retraction, morphological oscillations began. The retraction of protrusion leaves a region on the cell periphery densely populated with blebs that locally store a high amount of CSE. The traveling wave of cortex density during morphological oscillations redistributes this local CSE around the cell

Using the time-lapse recording of CHO cells (37 cells in 5 experiments) we collected information about changes in cell protrusions, motility, surface blebbing, and morphological oscillations (258 events total). The data analysis (Fig. 3 L–N) confirmed statistically significant positive correlations between cycles of high surface activity (apparent blebbing) and protrusion retraction (r = 0.45, p << 0.001, Spearman correlation). It also showed a highly significant negative correlation between protrusion development and blebbing (r = −0.85, p << 0.001, Spearman correlation), confirming our hypothesis that the cell body’s surface relaxes and flattens during lobopodial protrusion development.

Cells with mesenchymal motility also employ CSE for the protrusion in 3D collagen

Next, we asked if the cells that are known to employ a mesenchymal mode of motility also maintain CSE and use it for protrusion development during 3D motility. To test this, we used several different cell lines. First, we used WC cells, a rat mammary carcinoma line that is highly motile. The average apparent radius of rounded WC cells on a 2D surface and after embedding into 3D collagen were very similar, R2D = 6.02 ± 0.65 μm, N = 53 vs. R3D = 6.04 ± 0.51 μm, N = 34. Shortly after embedding in 3D collagen, WC cells developed multiple very long (>100 μm) and thin (0.4–1 μm) protrusions (Fig. 4; Video S10) and were capable of rapidly migrating at speeds of up to 2 μm/min. The fast migration of these cells involved substantial collagen contraction (Fig. 4 B; Video S11). Rounded WC cells had a convoluted surface morphology (Fig. 4 D), covered with blebs and small projections holding a large amount of CSE, whereas the surfaces of polarized cells possessing long protrusions were smooth with no apparent blebbing (Fig. 4 C). Visual and kymographic analysis demonstrate that blebbing on rounded cells decreased simultaneously with protrusion initiation (Fig. 4 E; Video S12). Blebbing and oscillations return during full or partial protrusion retraction; the blebbing may be global, involving the entire cell surface, or local, only in the area near the retracting protrusion (Fig. 4 F; Video S13). If a new protrusion started to emerge simultaneously with the withdrawal of an old protrusion, blebbing was often diminished, suggesting that the surface surplus was immediately re-used to allow the extension of the new protrusion (Fig. 4 G).Figure 4 Correlation between protrusion dynamics and apparent blebbing in Walker Carcinoma cells. (A–G) WC cells in 3D collagen. (A and B) DIC recording shows WC morphology and strong collagen contraction during cell motility (Videos S10 and S11). (C) Maximum intensity projection image of a single WC cell with Lifeact-GFP signal. The MIP image was built from Z-stack images taking each 1 μm through 47 μm of collagen thickness. (D) Merged, DIC, and fluorescence images of WC cells with Lifeact-GFP stable expression demonstrate a blebby cortex-membrane morphology in rounded state. (E) A polarized WC cell has a smooth surface with little blebbing (left and right images), whereas a rounded cell actively blebs (center) until a new protrusion appears (Video S12). The correlated kymograph (10 min long) presented below each image taken at the position shown on (E). (F) Fluorescence images of a WC cell with Lifeact-GFP expression demonstrate the appearance of a blebby surface morphology only on the side of a protrusion that partially retracted (Video S13). (G) An example demonstrating decreased blebbing during protrusion retraction (red arrows) when a new protrusion is growing simultaneously (green arrows). Scale bars, 30 μm (A), 20 μm (B, C, and E), 5 μm (D), and 10 μm (F). To see this figure in color, go online.

Video S10. Walker carcinoma cells demonstrate fast motility in 3D collagen

Video S11. The protrusions and migration of WC cells involve substantial collagen contraction

The video shows how the WC cell, which is located inside the collagen but out of a focal plane, is developing two long protrusions (60 and 50 μm) that contract the collagen fibers

Video S12. The video demonstrates that WC cells in 3D collagen have substantial blebbing if they obtain a rounded shape during motility

The blebbing on rounded cells ceases simultaneously with protrusion initiation

Video S13. A recording of WC cell with the stable expression of F-marker (Lifeact-GFP) demonstrates the development of blebs only on the retracting part of the protrusion

Next, we investigated the presence of CSE on the surface of the highly aggressive and motile MDA-MB-231 human breast cancer cells (Fig. S5 A). High-resolution fluorescence imaging shows that, similar to the CHO and WC cells investigated before, rounded MDA-MB cells in 3D collagen possess a highly convoluted surface morphology with many blebs (Fig. 5 A). After polarization, these cells develop strong attachments to collagen fibers and are capable of substantial matrix deformation (Fig. 5 C). Polarized cells in 3D collagen have elongated shapes with well-defined long actin cables beneath the PM and, in contrast to rounded cells, exhibit little or no apparent surface blebbing (Fig. 5 B and D). The ends of the extending protrusions in these mesenchymal cells often have blebs or bulges that may serve as a membrane reservoir for protrusion growth. An example of how a large bleb on an MDA-MB-231 cell decreases in size simultaneously with the protrusion extension is presented in Fig. S5 B and Video S14.Figure 5 Correlation between protrusion dynamics and apparent blebbing in MDA-MB-231 cells. Presence of MTs in the long protrusions. (A) Airyscan high-resolution fluorescence images of rounded MDA-MB cell fixed and stained with Phalloidin (green) and membrane dye (red). Merged (top) and single channels (bottom) images depict the small structures on cell surface at two different positions along the cell body. The left image (Z = 1.6 μm) shows numerous small projections extending out of the cell body into the collagen matrix. Note the clear colocalization of F-actin and membrane signals, which demonstrates that the F-actin signal originated from surface projections. (B) 3D reconstruction of a polarized MDA-MB cell in collagen. Inset on (K) (magnified part depicted by the yellow rectangle; scale bar, 5 μm) presents a magnified single slice near the ventral part of the cell, depicted by the yellow box. The image shows an absence of small projections on the cell surface and the presence of stress fibers around the cell body near the interface with collagen fibers. (C) Merged confocal image of fluorescently labeled collagen (cyan) and an MIP image of F-actin (green), vinculin (red), and nucleus (magenta). (D) DIC image of a single MDA-MB cell in collagen presents a cell that transforms from a rounded cell with apparent blebbing to polarized cell with a smooth surface. (E) The chart presents the comparison between apparent surface area of rounded and polarized cells for different cell lines embedded in collagen. The average values were derived for CHO, MDA-MB, WC, and DU-145 cells using recordings of 122, 178, 82, and 88 rounded cells and of 98, 25, 28, and 43 polarized cells, respectively (p << 0.001, multiple unpaired t-test, error bars represent SD). (F) The chart presents the percentage of cells that experienced apparent surface blebbing in two groups: rounded cells and polarized cells in collagen matrix. (G–L) 3D volume reconstruction from high-resolution Airyscan images of cells embedded in 3D collagen and magnified regions depicted by the corresponding arrows demonstrate the MT system in the polarized cells. MDA-MB (G–I) and WC (J–L) cells fixed after 24 h embedding in collagen and stained with Phalloidin (green) and anti-tubulin antibodies (red). Scale bars, 10 μm (A, B, and K), 5 μm (C and H), 1 μm (J), 0.5 μm (L), and 20 μm (D and G). To see this figure in color, go online.

Video S14. The video demonstrates how the MDA-MB cell uses a large bleb to extend the protrusion

We next investigated DU-145 cells (Fig. S6), which in 3D collagen remained mostly stationary with only rare, random translocations observed over the entire duration of the recordings (>24 h). After embedding into the collagen matrix, a rounded DU-145 cell with a large amount of stored CSE (Fig. S6 D and E), typically initiates multiple short protrusions from blebs around the periphery (Fig. S6 B) similar to those we observed on SEM micrographs (Fig. 2 G). After polarization, the blebbing activity disappears around the cell body and mostly remains apparent only on the edges of the protrusions (Fig. S6 C). Polarized DU-145 cells in 3D collagen have mostly irregular shapes with many short protrusions and smooth surfaces as evident from high-resolution fluorescence microscopy (Fig. S6 G). Some elongated cells revealed well-defined long actin cables around the entire cell body beneath the PM similar to MDA-MB cells (Fig. S6 H and I).

Using DIC images of cells in collagen, we estimated and compared the apparent surface area for the WC, MDA-MB, and DU-145 cells when they were in the rounded state with the area of polarized cells. These estimates show that the apparent area of polarized WC was up to 3 times larger than the apparent surface area of rounded WC cells (the average ratio was 1.8, N = 28). The apparent surface of the polarized MDA-MB cell was 1.8 times larger than the apparent surface of a rounded cell, and it was 2 times larger for DU-145 cells (Fig. 5 E). The analysis of our video recordings again confirmed a positive correlation between apparent surface blebbing and rounded cell shape for mesenchymal cells in our experiments (Fig. 5 F).

Many mesenchymal cells embedded in collagen, similar to CHO cells, exhibit oscillatory behavior. Oscillations often arose after the retraction of a long protrusion, which is analogous to the initiation of oscillations after cell rounding from the spread state. This provides additional evidence that cells in 3D collagen maintain a large amount of CSE that can be used for rapid shape transformations.

Morphology of mesenchymal cells depends on intact MTs in 3D collagen

Prior studies have shown that the protrusion development and 3D migration of mesenchymal cells, in contrast to 2D migration, is highly dependent on MTs, although the exact role of MTs is not completely understood (43,44,45). In our previous investigations of cell oscillations (19,20), we demonstrated that the amplitude and speed of cell morphological changes are substantially amplified by MT depolymerization, which implies a role of MTs in the regulation of CSE. Therefore, we next evaluated the effect of MT depolymerization on changes in shape and surface dynamics in cells embedded in 3D collagen.

All three cell lines using mesenchymal motility after polarization in collagen demonstrated the presence of MTs throughout the entire length of the protrusions, even in extremely long protrusions such as those in WC cells (Fig. 5 G–J and S7). The high-resolution Airyscan imaging also showed that MTs were present in some short projections in the direction perpendicular to the main polarization axis (Fig. S7 E). High-resolution magnified images also revealed that MTs can be very closely intertwined with cortical actin beneath the cell surface suggesting their supporting and stabilizing function (Fig. 5 L).

When an MT depolymerizing drug was added at the time of embedding cells, all investigated cell types that normally exhibit mesenchymal motility (WC, MDA-MB, and DU145) remained stationary with rounded morphology and were unable to develop long and stable protrusions (Fig. S8 B; Video S15). However, short protrusions (∼5–10 μm) were still able to randomly emerge from the cell periphery. These short protrusions were highly dynamic and were unable to form a stable shape with a smooth surface. Often these protrusions became fragmented, leaving large numbers of small actin-containing vesicles in the matrix. Many of these fragments moved substantial distances after detachment from the cell. If MT depolymerizing treatment was added after cells were embedded in the collagen, cells stopped protruding and withdrew the majority of their protrusions while fragmenting some others (Fig. S8 C–E).

Video S15. WC cells embedded in collagen with 1 μM of colchicine for 18 h

In CHO cells, MTs are withdrawn before lobopodial protrusion retraction

To investigate the structure and dynamics of the MT system in CHO cells during migration in 3D collagen matrices, we transfected cells with the MT marker, EMTB-3xGFP. We use for the visualization only cells with low to moderate expression of EMTB-3xGFP judging by the fluorescence intensity as suggested previously (46). For CHO cells, the MT system inside the wide lobopodial protrusions is organized as a complex network (Fig. 6 A and B). Surprisingly, during protrusion withdrawal, the MT filaments inside the retracting protrusion did not appear to depolymerize, but rather were retrieved intact and folded inside the cell (Fig. 6 D; Video S16). After MT withdrawal, a substantial part of the remaining protrusion without an MT signal was still visible on the cell periphery in DIC images (Fig. 6 C) and collapsed with some delay after full MT withdrawal. The time delay between complete MT retrieval and disappearance of the protrusions varied between 5 and 15 min (N = 7 cells in 3 experiments). After 24 h in the collagen matrix, the MTs in isolated rounded cells often showed a distinctive MT architecture in which MTs formed a basket-like structure with some MTs in close proximity to the cell surface (Fig. 6 E and G).Figure 6 Microtubule architecture and dynamics in CHO cells in 3D collagen. (A) DIC image, fluorescence signals from anti-tubulin staining (red) and phalloidin staining (green) illustrating the organization of the MT network and actin filaments in CHO cells in 3D collagen. (B–D) Example of MT architecture in CHO cells transfected with EMTB-3xGFP (MT-marker, green) and Lifeact-RFP (red) and embedded in 3D collagen, DIC, and merged fluorescence image (maximum intensity projections) of MTs and F-actin (B). (C) The image demonstrates the presence of a thin protrusion, which retracted with delay after MT withdrawal. (D) MTs folding during protrusion retraction in the area shown by the yellow rectangle on (B) (see Video S16). (E and F) Examples of MT structures in the rounded CHO cell (EMTB-3xGFP (green) and Lifeact-RFP (red). Single confocal slide of merged (DIC, F-actin, MTs) (left), merged F-actin and MTs (center), and MIP of MT signal (right). The arrow on (E) points to a protrusion that does not have visible MTs inside. (G) Lifetime of protrusions with or without visible MT signal and under colchicine treatment (p < 0.001, ANOVA test, error bars represent SD, N = 11, 32, and 45). (H) Length of protrusions with and without apparent MT signal and under colchicine treatment (p < 0.001, ANOVA, N = 126, 45, 60). To see this figure in color, go online.

Video S16. CHO cells with stable expression of Lifeact-RFP (red) and transient transfection with EMTB-3xGFP embedded in the collagen demonstrate a cage-like organization of MTs in the cell body and the presence of a complicated MT network within lobopodial protrusions

Note that withdrawal of MTs precedes protrusion retraction. Interestingly, MTs undergoing retraction and folding inside the cell body sometimes appear to coil into ring-like structures that were also occasionally observed in CHO cells stained with anti-tubulin without transfection EMTB-3xGFP

With an intact MT system, the migration of CHO cells often depends on multiple pseudopods, perhaps employed in an exploratory mode to find suitable openings in the matrix. Our observations revealed that not all of these pseudopods contain MTs (Fig. 6 E, arrow). We found that the protrusions containing detectable MT filaments have a much longer lifetime than the protrusions without MTs (Fig. 6 F). The length of the protrusions with visible MT fluorescence signals was also significantly longer in both live CHO cells with MT-marker expression (Fig. 6 H) and in fixed CHO-wild-type cells prepared for MT immunofluorescence. Similar to the mesenchymal cells, MT depolymerization in CHO cells prevented the formation of stable lobopodial protrusions (Fig. S9). However, without MTs, CHO cells were still able to spontaneously translocate in an amoeboid fashion.

Cell surface excess during amoeboid motility of CHO cells

The amoeboid type of motility, despite the general name, has at least two very distinct modalities (47). The so-called “blebby-type” amoeboid motility occurs when a cell develops protrusive bulges that push through interstices in the extracellular matrix. The other type of amoeboid motility, which is prominent in leukocytes, is characterized by protrusive short sheets of lamellipodium for navigation in the extracellular environment (27,47,48,49).

To investigate the magnitude of the changes in surface area during blebby-type amoeboid motility, we studied the morphology and behavior of the sub-population of CHO cells that employs this type of migration (Fig. 7 A; Videos S4 and S17). We noted above that blebby motility shares many similarities with the compression-dilation mode seen in oscillating cells (Videos S5 and S6), where a part of a convoluted cell surface stretched and extended during morphological oscillations in collagen (Figs. 7 B, S3 C, D, and E). We postulate that the same mechanism allows a cell to extend, squeeze, and eventually translocate through the collagen fibers (Fig. 6 C and D).Figure 7 Morphological changes and CSE in CHO cells during blebby-type amoeboid motility. (A) Video recording of two CHO cells in 3D collagen. The top cell is translocating using the amoeboid type of motility (Video S2). While pushing through the collagen fibers, the surface of the protruding bulge appears fully stretched and smooth (arrow) for a short period of time. The second cell with an F-actin fluorescence signal demonstrates blebbing and oscillations. (B) DIC and F-actin fluorescence images show stretching/unfolding of the cell cortex that leads to the development of rounded protrusions (arrows). (C) The proposed mechanism of cell protrusion. The signal received by the extracellular receptor triggers small projection and folds flattening and CSE release, which locally enlarges the cell surface creating additional volume. This volume is immediately filled with cytosol promoting a new protrusion. To preserve the total cell volume the other parts of the cell periphery shrink, adding more CSE on their surface. (D) 3D reconstruction of F-actin fluorescence signal from the Z-stack of confocal images (see Video S16) of CHO cell during amoeboid motility in collagen demonstrates protrusion of surface bulge with high actin density (arrow). (E) Comparison of average 2D area values between rounded cells (N = 120) and polarized cells with amoeboid movement (p < 0.001, t-test, error bars represent SD, N = 10 cells, 20 time frames for each, 3 experiments). (F) Morphological changes with the example of area segmentation of a CHO cell during amoeboid motility (Video S15). (G) Changes in 2D area of CHO cells before and after MTs were depolymerized. Values were normalized on the average value for each group (N = 10 cells). (H) Time plot of 2D area changes for two individual cells with and without MTs. (I) Examples of shape dynamics of the CHO cell with depolymerized MTs (Video S17). Scale bars, 10 μm (A, F, and I) and 5 μm (B). To see this figure in color, go online.

Video S17. Ameboid motility of a CHO cell visualized by merging DIC images with Lifeact-GFP fluorescence

This video demonstrates that the surface is constantly blebbing with local extensions that drive relocations

During amoeboid movement, the CHO cells become more elongated, with an average circularity of 0.81. The complicated cell shape during their motility made it difficult to calculate the apparent surface area from the DIC signal. Therefore, we used as a proxy for these changes the apparent 2D area (XY) measured by segmentation of the periphery near the equatorial section of cells fully visible in the focal plane (Fig. 7 F). We found that the difference in the average 2D area of an equatorial section between rounded and polarized cells is around 20% (S2D_pol = 245 ± 63 μm2, N = 24 vs. S2D_rounded = 203 ± 86 μm2, N = 126 for rounded cells) while changes of an individual cell during amoeboid motility are below 14% (with the average 5.8 ± 4.1%, N = 6). The visual evaluation showed that, during amoeboid motility, cells always have high surface dynamics with apparent blebbing. Similar to an oscillating cell, a large rounded protrusion that eventually can lead to cell translocation appears on the cell periphery mostly in or near the position with a large amount of CSE visually evident by the increased actin density (Fig. 7 D; Video S18). This large spherical protrusion during a short period of extension (Fig. 7 A–D) appeared smooth and devoid of blebbing.

Video S18. 3D reconstruction of F-actin fluorescence signal from a Z-stack of confocal images of a CHO cell during ameboid motility in collagen

The movie demonstrates that a large rounded protrusion that leads to cell translocation appears mostly on or near the position with a high amount of CSE marked by increased actin density (Lifeact-GFP)

While the structure and function of MTs in mesenchymal cells have been studied intensively, especially on 2D substrates (50,51,52,53), much less is known about the role of MTs during amoeboid migration (44,45). In our experiments we found that, although MT depolymerization in CHO cells did not completely abrogate the random amoeboid translocation of these cells, the morphological changes became erratic, demonstrating faster dynamics with greater amplitude (Fig. 6 G, H, and I; Video S19) This result is consistent with the changes developed after MT depolymerization on the oscillating cells on a solid substrate (19).

Video S19. CHO cells embedded in collagen with depolymerized MTs demonstrate the large-scale, fast, and erratic cell shape dynamics induced by colchicine treatment

The movie was recorded 24 h after free CHO cells were embedded in the collagen and 2 h after 1 μM colchicine was added to the medium that covered the collagen matrix. On the left is the maximum intensity projection of F-actin fluorescence (Lifeact-GFP) imaging and on the right is a DIC imaging of the same cell

CSE and MT structure during amoeboid motility of leukocytic cells

To study the dynamics of cell shape and associated changes of surface area in cells with a type of amoeboid migration observed in leukocytes (47,49), we used U937 cells, a human monocytic cell line. Similar to CHO cells with amoeboid motility, the surface of U937 cells embedded in collagen was highly dynamic. The presence of blebs and small fin-like surface projections was also confirmed by fluorescence imaging of U937 cells in the collagen matrix (Fig. 8). The rounded U937 cells have an average apparent radius of R = 7 ± 0.7 μm. After several hours in collagen, a sub-set of cells becomes spontaneously polarized and motile (Fig. 8 A; Video S20). The motile cells have a slightly elongated or irregular shape (average circularity index of 0.78 ± 0.1) with an average speed of 0.98 ± 0.70 μm/min, N = 20. The measurement of the 2D area during motility shows that the difference between rounded and polarized cells during migration on average is below 25% (Fig. 8 B).Figure 8 Shape dynamics and MT system in U937 cells. (A) Time steps of U937 cell shape transformation during motility in 3D collagen. (B) Average 2D area of rounded and polarized U937 cells (N = 30, 114; P << 001, nonpaired t-test, error bars represent SD). (C and D) DIC and confocal fluorescence images of a U937 cell stained with Phalloidin (green) and anti-tubulin antibody (red) (single plane in C and 3D MIP view in D). (E–J) Airyscan super-resolution imaging of U937 cells in collagen: F-actin (green), MTs (red), nucleus (blue). (E and F) Single slice of merge fluorescence signals (top) and 3D reconstruction (bottom) (built from 160 slices, 25.5 μm total thickness) of the MT network of a rounded U937 cell. (F) The section (thickness 0.9 μm) depicted by yellow rectangle on (E) show the intertwining of cortical actin and MTs. Note that a nonpolarized rounded cell has an MT system that is equally distributed and wrapped around the nucleus, similar to MTs in rounded CHO cells. (G) 3D volume reconstruction image (built from 101 slices, 16.5 μm total thickness) and magnified section (H) (2 μm thickness) showing MT presence inside the small protrusion. (I) 3D volume reconstruction image (built from 134 slices, 21 μm total thickness) shows two polarized cells with a highly asymmetrical signal from F-actin and MTs. Merged fluorescence signals (left) MIP for MT signal (center), and a single Airyscan slide (J) from the middle of Z-stack. Note that the positions with high F-actin signal have no detectable signal from MTs in the vicinity (arrows). To see this figure in color, go online.

Video S20. Example of U937 cells exhibiting the ameboid type of motility in 3D collagen

Fluorescence imaging demonstrated that U937 cells have a complex MT network and a cell body covered with small surface projections, which are capable of storing large amounts of additional surface needed for shape transformation during migration (Figs. 8 and S11). Because of cell speed and fast shape dynamics, we were not able to resolve MT structure in live U937 cells during migration, so our conclusions are based on fixed cells with immunofluorescence staining, which, unfortunately, does not allow us to establish with confidence the direction of particular cell movement. The MT fluorescence signal revealed that, in rounded nonpolarized U937 cells, the MT system is distributed evenly around the nucleus (N = 15), creating a structure similar to rounded CHO cells (Figs. 8 E and S11 A). The MT structure in protruding U937 cells was substantially polarized with some visible gaps in MT presence near the cortex where the dense patch of cortical actin is visible (Fig 8 J, arrows, N = 14 cells from 25 polarized cells imaged). Surprisingly, a bright MT signal was found inside some small bleb-like protrusions (r = 1 μm) (Figs. 8 H and S11 C), which might be evidence of an MT mechanism to provide directionality of cell movement. By penetrating and supporting some protrusions versus another the MT is stabilizing cell shape in the chosen direction providing guidance for motility. The Airyscan magnified images of the cell section near the PM (Fig. 8 F) demonstrate how MTs and the actin-cortex are possibly interlinked near the PM, similar to what we saw in WC cells (Fig. 5 L).

MT depolymerization in U937 cells embedded in collagen (Fig. 9 A and B) resulted in substantial changes in cell morphology, surface dynamics, and speed within 30 min after application of the depolymerizing agent (Video S21). Around 90% of cells transformed from rounded or slightly elongated shapes into a morphology that combined bulges of 3–5 μm in diameter with long tubules up to ∼30 μm in length (Fig. 9 E–G). Fluorescence imaging of F-actin showed that a layer of actin cortex underlaid the cell surface, whereas the small surface projections, including blebs, almost completely vanished after MT depolymerization (Fig. 7 H and G). The abrupt disappearance or flattening of numerous small surface structures inevitably resulted in an abundance of surface area that was stored in them. Our measurements of the 2D cell surface area of some individual cells show increases of up to 300% 30 min after the addition of colchicine (Fig. 9 C and D). This sudden release of CSE appears to force cells to adopt the morphology of long tubules, allowing for the accommodation of this CSE without using the surface projections storage (cartoon in Fig. 9 K).Figure 9 U937 cell motility in 3D collagen and effect of MT depolymerization. (A and B) DIC image of U937 cells embedded in 3D collagen for 24 h with intact MT system (A) and the same cells 30 min after addition of 1 μM of colchicine (B) (Videos S20 and S21). (C) Average normalized 2D area of U937 cells during amoeboid type of migration with intact MTs and after 1 h with depolymerized MTs. (D) The trajectory of the normalized 2D area changes after MT depolymerization for three different U937 cells in comparison with the area changes for the cell with intact MTs. (E) Time steps of U937 cell shape transformation after a colchicine treatment at t = 0. (F) DIC record shows a U937 cell with depolymerized MTs, which has an elongated shape with a long tail (arrow). The tail often gets stuck between collagen fibers, hindering cell movement until it breaks away from the cell (Video S22). (H and G). Two examples of U937 morphology in 3D collagen after MT depolymerization demonstrate smooth surface without blebs. Merged DIC and maximum intensity projection of fluorescence confocal image of U937 cells stained with Phalloidin (green) and DAPI (blue). (I) Percentage of mobile U937 cells in 3D collagen under different treatments (p << 0.001 for colchicine [N = 387 cells, n = 5] and calyculin [N = 158, n = 2]; p > 0.05 for FBS [N = 319, 317, n = 2] and LPA [N = 227, 236, n = 2]. (J) Average speed of cell random motility in 3D collagen under different treatments (P << 0.001 for colchicine [N = 130, 54 cells, n = 3]; p > 0.05 for FBS [N = 25, 20, n = 2] and LPA [N = 25, 15, n = 2]. (K) Cartoon illustrating our hypothesis of how the rapid release of stored CSE after MT depolymerization transforms cell morphology. (L) Percentage of U937 cells randomly moving in 3D collagen after incubation with the inhibitors: 1 μM/mL of C3 (N = 22, 468, n = 2), 30 μM of Rhosin (N = 494, 335, n = 2), and 10 μM of Y27632 (N = 142, 180, n = 2), and after additional application of colchicine (C3, N = 326, n = 2; Rhosin, N = 335, n = 2; Y27632, N = 560, n = 3). (M) Chart presenting changes in average cell speed under the same combination of treatments (C3, N = 25, 31, 150, n = 2; Rhosin, N = 53, 50, 230 n = 2; Y27632, N = 56, 12, 590, n = 2). ANOVA test was used for (I, J, L, and M). Error bars represent SD. Scale bars, 10 μm (A and H), 50 μm (C and D), 20 μm (G), and 5 μm (I and J). To see this figure in color, go online.

Video S21. Microtubule depolymerization greatly increases the 3D motility of U937 cells

U937 cells were embedded in 3D collagen with intact MTs for 8 h before the recording. During the recording, 1 μM colchicine was added to the medium that covered the collagen matrix

Concomitant with the changes in morphology after MT depolymerization, the percentage of U937 cells randomly moving inside the collagen matrix increased markedly up to 92% and the cells more than doubled in speed (average speed 2.2 ± 0.37 μm/min) (Fig. 9 I and J). The changes in cell morphology and dynamics persisted for the time of experimental recording (up to 10 h after colchicine treatment).

The fast moving cells often adopted a long tubular shape (Fig. 9 F; Video S22), with which these cells appeared to “flow” through the matrix. These cells also frequently acquired long tails that were often enmeshed in the collagen fibers constraining cell movement until the tails broke off. It is possible that such tails appeared in the absence of intact MTs as storage for CSE that cannot be distributed around the cell body.

Video S22. Example of U937 motility in 3D collagen with depolymerized MT system

It was established previously that the amoeboid migration is dependent on the Rho/ROCK signaling pathway (33,54), which regulates the contractility of the acto-myosin cortex. The Rho guanine nucleotide exchange factor, GEF-H1, is the known mediator that connects RhoA activation, actin polymerization, and myosin contractility to MT dynamics. GEF-H1 is inactive when it is sequestered by MTs and upon release from MTs becomes active (34,55). Therefore, the MT depolymerization sharply increases the level of active GEF-H1 followed by the increase of active RhoA and Myosin II contractility. To test whether the strong effect of MT depolymerization on the shape and motility of U937 can be solely attributed to increased myosin II contractility via the GEF-H1-RhoA-ROCK pathway, we stimulated the activity of RhoA and myosin II by 30% FBS or 2 μM LPA treatments after cell starvation. However, neither treatment led to shape changes, membrane tubules, an increase in the number of migrating cells, or migration speed (Figs. 9 I, J, and S11 A–C). The treatment with 40 nM of calyculin, which increases Myosin II contractility, led to cells rounding, extensive large blebs, and complete abolishment of motility (Fig. S11 D).

Application of two different Rho inhibitors: C3 (1 μg/mL for 6 h) and Rhosin (30 μM for 24 h) did not diminish the effect of MT depolymerization on U937 cell morphology and motility (Figs. 9 L, M, S11 E, and F). Surprisingly, the incubation with ROCK inhibitor Y27632 (10 μM) for 1 h before the application of colchicine, although almost abolishing cell motility, did not change the overall effect produced by MT depolymerization on cell shape and on the substantial increase of speed (Fig. 9 L and M). These inhibitor experiments indicate that the effects of MT depolymerization reported are unlikely to be due to the GEF-H1 > Rho > ROCK pathway and suggest the existence of additional mechanisms of MT regulation of cell shape and protrusion in 3D environment.

Discussion

Cells require a reserve of surface area to successfully execute fast shape transformations without threatening PM integrity. Despite its importance, the regulation of cell surface area has not attracted as much attention as the regulation of cell volume or membrane tension since it has been assumed that the surface area is a passive partner in changes in cell shape, simply following the morphology dictated by membrane tension and the cytoskeleton. However, cells generally have much more surface than is required to simply cover the cell volume, implying that specific mechanisms of surface area regulation should exist in addition to endo-exocytosis. We suggest that the simplest and most efficient mechanism for surface area regulation is a controlled development of small surface projections: blebs, ruffles, filopodia, microvilli, and others. These projections can store massive amounts of cell surface and are highly dynamic, which means they can be rapidly dismounted with the release of local CSE and immediately rebuilt in other locations on the cell periphery.

Our investigation of cell dynamics during 3D migration in collagen reveals that cycles of high cell surface activity (blebbing and oscillations) were followed by quiescence and visible surface smoothing. Similar cycling behavior between protruding and migrating cells (“running”) and more static cells that remain in the same position but with a loss of cell polarity (“tumbling”) was found during the migration of primordial germ cells which protrusions depend on the unfolding of membrane invaginations (56,57,58,59), mammary epithelial cells (60), invasive metastatic cells (61,62), and during bacterial chemotaxis (63). It was suggested that this running and tumbling behavior represents an intrinsic property of migrating cells (64); however, the mechanism underlying this behavior was not established.

Our analysis shows that intervals of surface smoothing on the cell body correspond with the extension of large protrusions, while active blebbing corresponds to cell rounding after the withdrawal of large protrusions. Fluorescence analysis of mesenchymal cells in collagen matrices revealed that, whereas rounded cells have a large number of blebs and other surface projections, polarized cells with long protrusions have substantially smoother surfaces, which is consistent with our hypothesis that cells release CSE stored around the cell body to cover the newly developing protrusion. During the protrusion retraction, the resulting CSE requires rapid packing, which is achievable by forming new blebs as they present the fastest attainable form of storage.

For the past two decades, blebs have often been considered to be an artifact arising from a compromised connection between the cortex and PM, which is pushed out by high pressure in the cytosol (65,66). It should also be noted that the majority of information about blebs and cortical actin structure has come from studies of cells spread on a solid 2D substrate or during artificially induced blebbing (67,68,69,70,71,72,73,74). Moreover, for purposes, such as the estimation of cortex thickness or modeling the biomechanical properties (74,75,76,77,78), it was often assumed that the membrane and actin cortex are two perfectly smooth concentric layers with uniform density. However, bearing in mind that the PM is essentially not stretchable, under this assumption the process of blebbing would not be possible because, to develop one typical bleb with a radius of r = 1 μm, an additional PM area of ∼12 μm2 is required. The fact that the average time of bleb expansion is less than 30 s leads us to the conclusion that the additional surface must come from some readily available source in the close vicinity of the growing bleb. Cell rounding or protrusion withdrawal each generates large amounts of CSE, which creates a favorable condition for producing numerous blebs. For cells with a stable morphology, the most obvious source for a new bleb is a shrinking bleb or filopodium nearby, which frees surface area for reuse. Several studies already confirmed that the probability of a new bleb appearing is higher around the position where another bleb is shrinking or after the disappearance of filopodia (39,79,80).

In this paper we emphasize that surface blebs provide an efficient mechanism for the prompt storing of surface excess. Because a typical lifetime of a single bleb is 1–3 min, this mechanism also allows the rapid redistribution of surface area and therefore should be considered as an important part of normal cell physiology with specific regulatory pathways that need to be established.

In this paper, we also suggest an important role of MTs in maintaining cell shape stability. Although there is a large knowledge database about MT organization and dynamics in cells migrating on a 2D substrate, much less is known about the organization and function of MTs for cells in 3D matrices. Our imaging results show that, in fixed mesenchymal cells, the bright fluorescence signal from MTs spans through very long and thin protrusions (Figs. 5 G–K and S7) and in some cells we detected that MTs located in very close proximity to cortical F-actin beneath the PM. Our imaging also revealed a complicated network of MTs in the lobopodial protrusions, the basket-like structure built from MTs inside the rounded body of CHO and U937, and the presence of MTs inside some small bleb-like projections in U937 (r = 1 μm) (Figs. 8 H and S11 C).

Importantly, our live cell imaging with CHO cells showed that not all lobopodial protrusions have a detectable MT signal. The protrusions lacking MTs were smaller and had substantially shorter lifetimes than protrusions with MTs, similar to protrusion after MT depolymerization. Furthermore, monitoring MT signals from live cells we detected that MTs can retract, bend, and fold inside the cell before protrusion withdrawal in CHO cells (Fig. 6; Video S16).

It was shown previously (43,44,45,81,82) that MT depolymerization in 3D culture had different effects on cell migration depending on the cell type: suppressing mesenchymal motility while unaffecting or enhancing leukocytic motility. In our experiments, MT depolymerization indeed prevented motility in WC, MDA-MB-231 breast carcinoma, and DU-145 prostate cancer cells, all of which normally exhibit mesenchymal motility in 3D collagen. Importantly, MT depolymerization also led to the loss of stable protrusions and to substantial changes in morphology in these cells. In CHO cells with mixed types of motility (blebby amoeboid and lobopodial), MT depolymerization also led to the loss of protrusions and associated motility and increased erratic surface dynamics, but did not substantially affect the blebby amoeboid mode of translocation.

Together these results indicate that, in the 3D matrix the withdrawal or depolymerization of MTs led to cell morphological changes with protrusion abrogation and that the presence of intact MTs stabilizes protrusions and cell morphology.

A striking finding of this work is that MT depolymerization greatly stimulated the migration of U937 cells in a 3D collagen matrix. These monocytic cells normally use a leukocytic mode of amoeboid migration, but MT depolymerization triggered them to change morphology and begin moving at approximately twice their normal speed.

To investigate the mechanism by which MT depolymerization stimulates 3D motility in U937 cells, we used inhibitors to test if the effect was solely due to stimulating the well-known GEF-H1 > RhoA > ROCK > Myosin II pathway. Surprisingly, none of the three different inhibitors that target the Rho pathway—C3 to inhibit Rho, Rhosin to inhibit RhoA, or Y27362 to inhibit ROCK—blocked the effects of MT depolymerization regarding shape changes and surface dynamics, although reducing speed under Y27362 application. Furthermore, two different treatments to activate Rho—the addition of a high concentration of serum or LPA—did not lead to changes in cell shape or motility. Adding calyculin to activate Myosin II by blocking dephosphorylation of its regulatory light chain had the opposite effect: it completely stopped cell motility and surface dynamics, although it induced intensive transient blebbing within the first 30 min after application. These results indicate that many effects of MT depolymerization on U937 cells are unlikely to be due to the GEF-H1 > RhoA > ROCK > Myosin II pathway.

To explain such substantial changes in cell morphology and surface dynamic in all different types of cells that we investigated, we hypothesize that MT depolymerization leads to uncontrolled release and destabilization of CSE, which, in turn, increases surface dynamics and accelerates shape changes. Consistent with this hypothesis, it was reported that tethers extracted from cell membranes after MT depolymerization became significantly longer than in control cells and even longer than in cells with depolymerized actin filaments (83,84,85). Similarly, micropipette aspiration studies of living cells revealed substantial cell softening after MT depolymerization (86,87).

There are several possible mechanisms by which MT depolymerization might act besides the regulation of contractility through the GEF-H1 > RhoA > ROCK > Myosin II pathway. These mechanisms can be valid for different cell types and motilities.

Because small surface projections that store CSE are actin-based structures, one possible mechanism is crosstalk between cortical actin and MTs through shared regulatory factors such as profilin or similar proteins capable of regulating actin polymerization. (88,89) Profilin, for example, can bind actin monomers or tubulin dimers and therefore regulate the pool of monomers available for polymerization. This can possibly explain the disappearance of actin-based projections on the cell surface and transformation to a more blebby morphology.

Another possible mechanism is a direct stabilization and reinforcement of cell surface by cross-linking and bundling directly between F-actin and MTs or by proteins that have specific binding sites for both filaments, for example, tau, fascin, motor proteins, and others (90,91). Our images showing examples of closely intertwined MTs and actin staining just below the cell surface are also consistent with either molecular links or physical entanglement of MTs and cortical actin as a mechanism to stabilize the cell surface.

Another mechanism by which MTs might facilitate CSE storage in small actin-based surface structures would be a direct induction of actin polymerization from MT ends (92). As we mentioned previously, spherical projections (blebs) provide a rapid way to accommodate surface excess; however, they are also the least stable. Early work by Kinn and Allen (12) showed that, within an hour after rounding, the blebbed surface reorganized to a more stable microvillous topography. MTs could potentially accelerate this type of transformation through processes such as actin nucleation.

However, how we can explain why the uncontrolled CSE with increased surface dynamics has different effects on different types of motility?

We demonstrate here that CSE is involved in both amoeboid and mesenchymal motility in 3D. Because the morphological changes are slow for mesenchymal migration and rapid for amoeboid, these two modes require the deployment of CSE within different timescales. When mesenchymally migrating cells are surrounded by elastic fibers in the collagen matrix, slow cell surface dynamics and regulated CSE are important factors for building stable protrusions and developing new adhesions. We hypothesize that dysregulation of CSE and destabilization of the cell surface after MT depolymerization prevent the execution of all these functions. In the absence of a coordinated supply of CSE, cells are only able to extend short protrusions, and without the stabilizing function of MTs, these protrusions are likely to collapse shortly after development.

MTs may be not as important for 2D mesenchymal migration as for 3D because the solid substrate stabilizes cell shape and surface, allowing the development of strong adhesions, adhesion-based signaling, and exertion of force for movement.

In contrast to mesenchymal motility, during amoeboid migration, the rapid shape transformation necessary for squeezing between ECM filaments involves fast CSE dynamics. For fast-moving cells undergoing amoeboid motility with short protrusions, local reserves of CSE appear sufficient for providing the needed cell surface. In this case, the MT network stabilization function is important for restricting CSE release everywhere on the cell periphery except for localized release in the direction of movement. This locally released CSE is inflated by cytosol and followed by actin polymerization inside the protrusion and stabilization by newly grown MTs. By penetrating and supporting some growing protrusions versus others, the MTs might encourage cell protrusions in the chosen direction, providing guidance for motility. When MTs are depolymerized, CSE is released without spatial regulation, forcing cells to adopt a new morphology, creating conditions for rapid erratic movement, leading in some cases to cell fragmentation.

As the result of our investigation, we put forward a hypothesis that intact MTs can stabilize cell shape and regulate cell surface stability by controlling and restricting CSE dynamics and by providing support to a cortical actin network. We also present evidence suggesting that CSE and its control by MTs may regulate the cell protrusion, stabilization, and retraction required for cell migration in 3D.

Author contributions

M.K. conceived and designed the study, performed experiments, analyzed data, and wrote the manuscript draft with help from Ken Jacobson. D.L. and J.Z. performed experiments, analyzed data, and edited the manuscript. B.W. and V.W. performed experiments and analyzed data. R.E.C. provided resources, participated in the discussions, and edited the manuscript.

Supporting material

Document S1. Figures S1–S11

Document S2. Article plus supporting material

Acknowledgments

We dedicate this manuscript to our friend, colleague, and coauthor, Dr. Ken Jacobson, who passed away during the preparation of the final drafts of this manuscript. This work was supported by 10.13039/100000057 National Institute of General Medical Sciences grant NIH R01GM134531 (REC). The microscopy was performed at the UNC Hooker Imaging Core Facility. The facility is supported in part by NIH/National Cancer Institute P30 CA016086 Cancer Center Core Support Grant to the UNC Lineberger Comprehensive Cancer.

Declaration of interests

The authors declare no competing interests.

Donna Li’s present address is Cancer Biology Graduate Program, McArdle Laboratories for Cancer Research, University of Wisconsin-Madison, Madison, Wisconsin

James Zhu’s present address is Quantitative Biomedical Research Center, Department of Population and Data Sciences, University of Texas Southwestern Medical Center, Dallas, Texas

Brittany Wall’s present address is Department of Periodontics, School of Dentistry, University of California at Los Angeles, Los Angeles, California

Supporting material can be found online at https://doi.org/10.1016/j.bpj.2023.04.023.
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