Understanding the driving force for cell migration plasticity

Abstract

Cell migration is a complex phenomenon. Not only do different cells migrate in different default modes, but the same cell can also change its migration mode to adapt to different terrains. This complexity has riddled cell biologists and biophysicists for decades in that, despite the development of many powerful tools over the past 30 years, how cells move is still being actively investigated. This is because we have yet to fully understand the mystery of cell migration plasticity, particularly the reciprocal relation between force generation and migration mode transition. Herein we explore the future directions, in terms of measurement platforms and imaging-based techniques, to facilitate the undertaking of elucidating the relation between force generation machinery and migration mode transition. By briefly reviewing the evolution of the platforms and techniques developed in the past, we propose the desirable features to be added to achieve high measurement accuracy and improved temporal and spatial resolution, permitting us to unveil the mystery of cell migration plasticity.

Introduction

Echoing Descartes’ notion made in the 17^th century that animals are complicated living machines (1), the rapid advances in locomotive robotics recently are largely inspired by animal movements (2). For example, robots can traverse complex terrains mimicking the moving strategies of snakes, sensing contact forces to discern terrain characteristics (3,4). However, the bioinspiration for locomotive robotics is currently limited to the organismal levels, despite the wondrous range of physical environment through which cells are capable of navigating. Take macrophages for instance: they can be found in circulating blood, rolling on endothelium of the blood vessel, which is a warped 2D surface. Macrophages also move in 3D microenvironments, and they can be found in very hard tissues such as bones (15 GPa) (5,6) or in very soft ones such as the lung (1 kPa). This versatility is not unique to macrophages, as cancer cells hijack such capability, circulating through the blood stream and then invading bones and lungs through metastatic migration. Understanding the mechanism by which cells adapt to vastly different landscapes can lead to novel therapeutic opportunities. For example, by manipulating the mechanosignaling pathways involved in adaptive migration, immune cells can be more rapidly recruited to eliminate pathogens (7). Similarly, cancer metastasis can be suppressed by decreasing their adaptability to move efficiently in different tissue landscapes (8). One impactful application might be a micron-sized robot (micro-robot) delivering anticancer drugs to disseminated tumor cells at multiple organs in precision. For example, by incorporating the adaptability of migrating cancer cells to traverse vastly different tissue landscapes, the micro-robot can reach the metastatic foci in the distant organ, targeting cancer cells locally. Precisely releasing drugs in the proximity of diseased cells is desirable because systemic adverse effects of the drugs can be minimized. To accomplish these, comprehensive understanding must first be established regarding how cells can sense the physical attributes of their environment important to cell motility and how cells adapt their force generation pattern accordingly to power the migration. Indeed, pioneering works of drug delivery micro-robots are limited to performing simple tasks of drug release in simple geometry in vitro (9), not capable of traversing complex tissue landscape in vivo. The insights gained from studying the plastic and dynamic migrating behaviors can lend momentum to propel the development of drug delivery micro-robots. In this article, we will discuss the opportunities and challenges in studying the mechanics of cell migration.

Environmental adaptation by migration mode transition

Recently, many in vitro platforms have been devised to study cell migration in more complex microenvironments, mimicking the various geometries found in vivo. These include cells seeded on compliant surfaces (10), cells encapsulated in collagen gels (11) or cell-derived matrices (12), cells confined vertically by agarose gels (13), or confined in a narrow microchannel (14), among others. To study cell migration in various platforms, optical microscopy is commonly used. The displacement of the cells can be tracked in brightfield. The dynamics of proteins relevant to cell migration can be recorded and analyzed using fluorescence microscopy, provided the proteins are fluorescently labeled. Force generated by cells can be estimated by imaging the deformation of the substrate on which cells migrate (for details, please see “force generation and transmission in different migration modes” section). Plasma membrane tension, which is related to the mode of cell migration and its speed, can also be evaluated by measuring the fluorescence lifetime of recently developed membrane tension sensors (15).

Studies from this wide variety of geometries showed that cells move in different modes when subjected to different geometries (16,17). For example, it was reported that cells migrate on unconfined 2D surfaces through dynamic focal adhesion (FA) engagement to the extracellular matrix (ECM)-coated surfaces (18), known as mesenchymal mode, or through small transient blebs that facilitate gliding (19), known as ameboid mode (Fig. 1). When imposed with dorsal confinement, cells were shown to crawl through the 3D space with FAs (20), or to use friction to sustain leading-edge protrusion (21), or to form a stable bleb to propel the displacement of the cell body (22), or to use intracellular hydrostatic pressure to drive such displacement, where a lobular structure forms, known as lobopodium, at the leading edge of the cell (Fig. 1) (23). Moreover, cells tend to migrate in mesenchymal mode when in contact with relatively stiff substrate but in ameboid mode when in contact with relatively soft substrate (24). It is thought that myosin activity is important in determining migration modes (25). It is also widely acknowledged that the extent of the resistance of the nucleus passing through a confining 3D environment governs the amounts of adhesion and contractility required for cell motility and subsequently the cell’s migration mode (17). However, it is not known which mechanosignaling pathways trigger the cell to transition from one mode to another as the cell navigates through different physical environments. The cytoskeleton dynamics during such transition remains a mystery too.

Figure 1.

Different cell migration modes are associated with various biophysical properties of the cytoskeleton. Migrating in 2D, cells can move via mesenchymal migration using FAs or via ameboid gliding migration with small transient blebs. The latter results in lower traction forces transmitted to the substrate. In 3D, cells use FAs or lobopodium, where the intracellular pressure difference is employed to drive cell displacement, or friction from actin retrograde flow, which sustains leading-edge protrusion, or a stable leader bleb, which displaces the cell body by gliding. When cells are confined in narrow microchannels, cells also exert normal forces of relatively large magnitude (14), in addition to traction forces, to propel the cell body forward. Furthermore, when cells are confined in 2D under micropillar array, cells change their migration mode from mesenchymal to ameboid (13). How the transition from one to another mode is initiated at the cytoskeletal level is yet to be elucidated. To see this figure in color, go online.

Cytoskeleton dynamics of different migration modes and during mode transition

It was first shown over three decades ago that the actin cytoskeleton is the driving force for cell migration (26). Actin proteins (G-actin) polymerize to form filamentous (F-actin) structures at the cell’s leading edge and push the plasma membrane forward (27). As the leading edge protrudes further, the increased tension in the plasma membrane stops the polymerizing actin from moving forward. In the case of 2D mesenchymal migration, the F-actin flows in the retrograde direction as a result, generating traction forces that facilitate force-dependent FA turnover (28,29). The dynamic F-actin distribution at the cell leading edge at high membrane tension in other migration modes is poorly understood. Moreover, F-actin underneath the plasma membrane plays an important role in determining the cell shape, which varies depending on the migration mode. The differences in cell morphology and locomotion in different modes illustrate that the cytoskeleton machinery configures differently in different tissue landscapes. Indeed, fast, reversible transitions between mesenchymal and ameboid modes have been reported by many (19).

Though it was shown the degree of cell-substrate engagement is associated with mode transition (17), little is known about how such transition is regulated and relayed to alter actin cytoskeleton dynamics. For example, it is unclear what the most upstream trigger is when cells transition from 2D mesenchymal to 3D lobopodium-based migration (Fig. 1) (23). Is it the integrin engagement on the dorsal side of the cell that initiates actin reorganization? Or is it the confinement that deforms the actin cortex leading to global reorganization of the cell cortex? One of the hurdles preventing further understanding is the imageability of the platforms mimicking nonplanar geometries of in vivo tissues. Indeed, although detailed organization of actin cytoskeletons can be successfully visualized in experiments where cells adhere to glass or other thin and flat substrates (10,13), there have been few successes, in any, in visualizing cells migrating in 3D. For example, this difficulty is especially prominent when studying confined migration in microchannels (Fig. 1). The four walls of the compliant microchannels are of 5–10 mm thickness (30). Consequently, the refraction caused by the microchannel walls severely distorts the images, and photon scattering caused by the thick walls dims the fluorescence signals. Resolving fine features such as actin networks in cells in a microchannel is therefore impossible. The images of migrating cells in microchannels often are of low spatial resolution. One cannot discern actin filaments in these images, not to mention measuring actin polymerization and retrograde flow rates. The low-resolution images only allow one to compare cell shapes (the cell length, for example) or total fluorescence intensity of certain fluorescently labeled proteins (F-actin, for example) between groups without resolving fine features at the subcellular level (14).

It should be noted that in addition to actin, microtubules (31) and intermediate filaments (32) have also been implicated in cell migration. However, studies regarding the roles of microtubules and intermediate filaments in cell motility have mostly been focused on mesenchymal migration. Without the capability of resolving fine structures of the cytoskeletons, it is impossible to learn the importance of microtubules and intermediate filaments in other migration modes or lack thereof. Additionally, the dynamic reorganization of the cytoskeleton generates forces that reshape and propel cells during migration. The forces are often transmitted to the microenvironment, resulting in the mechanical remodeling of the ECM. Therefore, measuring the forces generated by migrating cells can inform us of the physical interaction between the cell and its environment (see “force generation and transmission in different migration modes” section for discussion). Resolving the dynamic cytoskeleton organization in all migration modes, as well as identifying how it is regulated, will lead to in-depth knowledge regarding how cells undergo shape change as they navigate different physical environments, as well as how cells generate and transmit forces in a spatially optimized manner. This knowledge in turn can inspire better design of locomotive micro-robots by mimicking how cells move. It also can inspire more effective treatment strategies to suppress metastasis by inhibiting specific cytoskeleton-associated enzymes promoting migration plasticity.

Force generation and transmission in different migration modes

Cells are known to reorganize the ECM via mechanical forces. For example, migrating cells can transmit forces onto the ECM, creating space for their passage. Such forces, depending on the magnitude, can temporarily or permanently change the ECM organization (20,33). Currently, we do not have the full picture of how force generation is related to different modes of cell migration or how such a link is manifested in the alteration of tissue landscape. Moreover, we do not know how force-generating machinery regulates or is regulated by the switch of migration modes. To obtain such an understanding will require experimental methods allowing force measurement intra- and extra-cellularly, as the migration mode transitions from one to another, at high spatiotemporal resolution and with minimal disruption to the process. Highly sensitive force spectroscopy techniques have been widely adopted to measure forces at the single-molecule or cellular levels, including atomic force microscopy (34), optical tweezers (35), magnetic tweezers (36), and microneedle (37), among others. The aforementioned methods are powerful in determining forces generated by a single cytoskeleton component, e.g., the kinesin motor (35) or actin filament (34,37), but they cannot be applied in a highly multiplexing manner. Further, it should be noted that these methods actively impose forces onto the sample, thus likely measure force generation as a response of the cytoskeletal component to such externally applied force. The actively imposed external force is different from the resistive force, such as viscous drag encountered by a freely moving motor protein or a polymerizing actin filament pushing against the cell membrane. Given many force-generating processes in the cell are load dependent (38), one should take caution not to interpret the measurement readout as the force generated under negligible loading, which is the case in many physiological conditions.

On the other hand, traction force microscopy (TFM) maps the stress field across the cell through force-dependent deformation of compliant substrates on which the cell migrates, without disturbing the process (Fig. 2). Traction forces in 2D can also be measured using microfabricated elastic posts (44) as a substrate for cell movement. TFM has advanced the knowledge of 2D mesenchymal (39) and ameboid (45) migration significantly. Notably, TFM results elucidated how mesenchymal cells sense substrate stiffness gradients with force fluctuation within FAs to guide durotactic migration (10). TFM has also been performed in 3D hydrogels (40) but is only capable of measuring nonmoving cells. Note that when cells are embedded in substrates closely mimicking in vivo conditions, the force field calculation is severely complicated by the heterogeneous viscoelastic properties typical of physiological ECM and by the constantly changing viscoelastic properties of the substrate as cells remodel it. An ideal system capable of force measurement on cells migrating in 3D with minimal perturbation and high spatiotemporal resolution is yet to be developed to enhance our knowledge regarding the interplay between force generation and 3D migration.

Figure 2.

Dynamic force generation can be evaluated using various force measurement methods. Micropillar arrays provide discrete information as individual pillars deform under cell traction forces. The degree of bending of each pillar relates proportionally to cellular traction forces, allowing the mapping of the magnitudes of forces across the cell in 2D. Traction force microscopy (TFM) works by observing the displacement of beads that are incorporated on or embedded in the continuum substrate, which may be 2D (39) or 3D (40). The displacements of all the beads are recorded to construct a finite element analysis network, for mapping the stress field across the cell body (39). To directly measure traction force at single molecule level, tension sensor (TS) modules are incorporated into the molecule of interest (41). The tensile force from the actin cytoskeleton is inversely proportional to the FRET efficiency from the TS modules, which allows one to measure force transmission. In addition, DNA-based probes have also been developed (42,43). Shown here are the DNA-based quenched tension gauge tethers (qTGTs), where a quencher-fluorophore pair is conjugated to the two strands of the DNA respectively. One strand of the DNA is immobilized onto the substrate. The DNA double strand dehybridizes when traction force exceeds the magnitude required to break all the hydrogen bonds between the double strands. As the two DNA strands separate, the fluorophore emits photons to be detected as the readout. To see this figure in color, go online.

In the past 15 years, the development of DNA- or elastic protein-based force probes (Fig. 2) have allowed one to measure force transmission at high-throughput and single-molecule level (41,42,43). Because force-induced dehybridization in DNA or conformational changes in elastic proteins subsequently alters the intensity of the FRET pair or quencher-fluorophore pair conjugated to the DNA or protein, simultaneous imaging of hundreds to thousands of probes using fluorescence microscopy can be readily accomplished. Consequently, force generation per molecule can be mapped dynamically across a migrating cell. The force map can be examined in correlation to other measurements to identify the regulators and/or effectors of the force generation. For example, by correlating the local intensity changes over time in fluorescently labeled proteins of interest, a potential force-sensitive protein can be identified (46). The force measurement employing the single-molecule probes revealed that for a mesenchymal cell on 2D surfaces, most integrins exist in a minimally tensioned state, and a small fraction of ligand-engaged integrins experience loads larger than 11 pN (47). This finding implies that cells transmit forces in a highly regulated, spatially selective manner during 2D mesenchymal migration. It was shown that DNA-based probes can independently quantitate traction forces at various physiologically relevant substrate stiffness values, without postmeasurement correction (43), in contrast to TFM, where the stress field evaluation depends on determining the substrate stiffness value a priori. Therefore, it is possible to measure the force generation of migrating cells in 3D, by conjugating these single-molecule force probes onto the substrate (e.g., ECM proteins, synthetic polymers), despite the heterogeneous nature of the 3D environment.

Concluding remarks

Cells are capable of traversing and adapting to a wide range of physical environments from 2D to 3D, from very soft to bone rigid, possibly by changing the migration mode. Although cytoskeleton dynamics of migrating cells in some of the modes has been extensively studied, that of others (e.g., confined migration in microchannels) is yet to be elucidated. Moreover, the mechanism that governs the mode transition as cells adapt to a new physical microenvironment is still unknown. Many questions need to be answered before a complete picture of migration mode switching emerges. For example, how is the intracellular hydrostatic pressure regulated as the cell switches to and from lobopodium-based migration? An intracellular pressure sensor that can map hydrostatic pressure across the cell body with submicron resolution, which does not exist currently, will enable us to answer such questions. More generally, advances in experiment platforms allowing high-resolution imaging, force mapping, as well as a host of other biophysical measurements (e.g., pressure, tension, temperature, etc.) will enable us to build a more comprehensive understanding regarding migration mode transition.

Author contributions

Y.C. conceived the concept of the article. J.C., D.Y., and Y.C. wrote the article.

Editor: Meyer Jackson.