Micropatterned mammalian cells exhibit phenotype-specific left-right asymmetry

Abstract

Left-right (LR) asymmetry (handedness, chirality) is a well-conserved biological property of critical importance to normal development. Changes in orientation of the LR axis due to genetic or environmental factors can lead to malformations and disease. While the LR asymmetry of organs and whole organisms has been extensively studied, little is known about the LR asymmetry at cellular and multicellular levels. Here we show that the cultivation of cell populations on micropatterns with defined boundaries reveals intrinsic cell chirality that can be readily determined by image analysis of cell alignment and directional motion. By patterning 11 different types of cells on ring-shaped micropatterns of various sizes, we found that each cell type exhibited definite LR asymmetry (p value down to 10^-185) that was different between normal and cancer cells of the same type, and not dependent on surface chemistry, protein coating, or the orientation of the gravitational field. Interestingly, drugs interfering with actin but not microtubule function reversed the LR asymmetry in some cell types. Our results show that micropatterned cell populations exhibit phenotype-specific LR asymmetry that is dependent on the functionality of the actin cytoskeleton. We propose that micropatterning could potentially be used as an effective in vitro tool to study the initiation of LR asymmetry in cell populations, to diagnose disease, and to study factors involved with birth defects in laterality.

Left-right (LR) asymmetry is seen in the development of numerous living organisms, including climbing plants (1), helices of snail shells (2), and the human body (3, 4). Genetic diseases and prenatal exposure to teratogens can cause birth defects in laterality (5, 6). Using embryos of Drosophila, zebrafish, frog Xenopus, mouse, and chicken, several models have been proposed for the establishment of LR asymmetry, such as directional nodal flow driven by primary cilia (7–9), voltage gradients resulting from asymmetric expression of ion channels (10, 11), and asymmetric vesicular transport via myosin 1D along actin cable networks (12–14). In vertebrates, while some of the mechanisms appear to be conserved, important differences are observed between species such as differential regulation of asymmetric ion flux in frog and chicken embryos (10), and a distinct role of FGF-8 in mouse and rabbit embryos as a sidedness determinant (15). Two main models describe the establishment of LR assymetry. The primary cilium model holds that the LR asymmetry is a phenomenon initiated by the formation of a ciliated embryonic structure known as a node, which generates directional fluid flow being transduced into asymmetric gene expression and giving rise to LR asymmetry (7–9, 16). The second model holds that LR asymmetry is a fundamental property of the cell, based on the observations that LR asymmetry can be established without primary cilia, node, or fluid flow (10, 17). Recapitulating chiral morphogenesis using in vitro models with human cells would thus help further interrogate the existing competing models, toward better understanding of tissue morphogenesis in embryo.

The initiation of LR asymmetry in development is often first observed in populations of cells of the same type, such as snail embryonic cells at four-cell and eight-cell stages and mouse cells at embryonic nodes. The establishment of LR asymmetry or chirality within such cell clusters may rely on some intracellular structure, such as the hypothetical F-molecule or actin/microtubule cytoskeleton (18, 19) that can distinguish left from right by orienting the third axis with respect to predetermined dorsal–ventral and anterior–posterior axes. In addition, during development, the specification and self-organization of migrating cells are mediated by physical boundaries imposed by the extracellular matrix and the surrounding cells and tissues. We, therefore, hypothesized that the populations of cells of the same type, if cultured within patterns with well-defined boundaries, would express directional alignment and motion associated with the establishment of LR asymmetry.

We find that the cells form an invariant chiral alignment, depending on the cell phenotype and actin function. We infer that all cells are intrinsically chiral and that the function of actin may play a significant role in switching the intrinsic chirality of cells and their LR asymmetry in developmental biology. Due to its simplicity, our 2D microscale system provides an effective method to study the LR symmetry of tissue development in vitro and may provide a useful tool for identifying biological and environmental factors involved in bilateral birth defects.

Results and Discussion

Cells Exhibit Chiral Morphogenesis on Micropatterned Geometries.

We cultured the C2C12 murine myoblasts on ring-shaped micropatterns (Fig. S1 and Table S1 in SI Appendix) (20, 21). Phase contrast images, in which cell contour appears brighter than the inner cell region, were used to measure cell alignment (22) (Fig. 1A), as indicated by green lines in Fig. 1B. Each green line was assigned a biased angle between -90^° and 90°, based on its deviation from the circumferential direction (blue dash line); a positive value represented a counter clockwise (CCW) alignment, while a negative value represented a clockwise (CW) alignment (Fig. 1C). An angular histogram (Fig. 1D) and the radial distribution (Fig. 1E) of the measured angles revealed preference for positive angles, corresponding to the CCW bias. In > 30 independent series of experiments using > 1,000 individual rings (Fig. 1F and Table S2 in SI Appendix), the C2C12 cells showed CCW alignment with a biased angle of 8.47° ± 0.20° (mean ± SEM), with very strong statistical significance (p = 9.3 × 10^-186).

Fig. 1.

Mouse myoblasts (C2C12) show distinct chirality on micropatterned surfaces. Scale bars: 100 μm. (A) Asymmetric cell alignment on ring patterns (phase contrast image). (B) Cell alignment directions (green lines). (C) The biased angle of cell alignment (green lines) was defined as either CW or CCW, based on the deviation from the circumferential direction (blue dash line). (D) The circular histogram of biased angles shows CCW chirality. (E) Circumferential averages of the subregional biased angles at different radial positions on the ring (mean ± SEM). (F) The histogram of the mean biased angles of C2C12 cells (33 experiments, > 1,000 ring patterns).

Other geometries such as circles, squares, and linear strips were also tested. Biased cell alignment was observed on linear strips and rings but not on circles and squares (Fig. S2 A–F in SI Appendix), suggesting the importance of appositional boundaries for the expression of chirality. Because equivalent bias in cell alignment was observed on linear strips and rings of different sizes (Fig. S2G in SI Appendix), rings with an inner diameter of 250 μm and a distance of 200 μm between the inner and outer boundary were used in subsequent studies.

Cell Chirality on Patterned Surfaces Depends on Cell Phenotype.

We then tested a panel of mouse, rat, and human cells derived from skeletal muscle, bone, adipose tissue, skin, heart, and blood vessels (Fig. 2 and Table S1 in SI Appendix). Cell phenotypes included myoblasts, osteoblasts, endothelial cells, fibroblasts, and mesenchymal stromal/stem cells. All cells exhibited distinct chirality after they reached a confluency of approximately 75% and were cultured overnight. Interestingly, mouse (C2C12) and human skeletal muscle cells (hSkMCs) showed a CCW alignment, while all other cell types exhibited a CW alignment (Fig. 2 and Fig. S3 and Tables S2 and S3 in SI Appendix). Cancer skin cells also exhibited a CCW alignment, different from the CW alignment of skin fibroblasts from the same individual. These data suggest that adherent mammalian cells exhibit an invariant chirality determined by the cell phenotype and disease condition.

Fig. 2.

Cell chirality on patterned substrates depends on cell phenotype and disease condition. (A) Phase contrast images of various cell types. (B) Cell source and their chirality on micropatterned surfaces. Here, “−” indicates that cells were isolated in the lab; CW: clockwise alignment; CCW: counterclockwise alignment; N/S: not significantly biased to CW or CCW. The C2C12 cells and human skeletal muscle cells exhibit a counterclockwise alignment while other cells that were studied show a clockwise alignment. The human skin cancer fibroblast cell line show a counterclockwise alignment, opposite from healthy human skin fibroblasts. Scale bars: 100 μm.

LR Asymmetry on Micropatterns Is Established by Mechanisms Involving Cellular Directional Migration on Boundaries.

For cells on a patterned geometry to display their chirality and to distinguish between left and right (y axis), the polarity of the z axis (up-down) and x axis (front-back) must be established (Fig. 3A). Notably, the z axis was established independent of gravity direction, as the same biased alignment (relative to the cellular apical-basal axis) was observed experimentally in regular and vertically inverted cell cultures (Table S4 in SI Appendix). The geometric boundaries determined organelle positioning (i.e., x axis), with centrosomes (bright green, Fig. 3B and Fig. S4 in SI Appendix) and Golgi apparatus (red, Fig. 3C) being positioned closer to the boundaries than the cell nucleus (blue) (23), independent of the gravitational direction. In addition, cell chirality was maintained on patterns with different surface chemistry (24), after the disruption of cadherin function by reducing calcium levels (25), and the inhibition of cell proliferation (Fig. S1 and Tables S5–S7 in SI Appendix).

Fig. 3.

Cellular LR asymmetry on micropatterned rings is established by mechanisms involving boundary effects. (A) Cells on a ring “sense” the z axis through attachment to the substrate and the x axis through the ring boundaries. The cell alignment bias of the y axis (dash red lines) creates the observed cellular chiral behavior or LR asymmetry. (B) Centrosomes (bright green) are positioned closer to each boundary than nuclei (blue) in C2C12 cells (actin: red, tubulin: green). Scale bars: 50 μm. (C) Golgi apparatus (red) is positioned closer to ring boundaries than nuclei (blue) in hUVEC cells. Scale bars: 50 μm. (D) Phase contrast images of the cells (top) at 5, 10, 20, and 30 hr after cell seeding and the corresponding histograms (bottom) of biased angles from the subregions for each image. Scale bars: 100 μm. (E) The time history of the mean biased angle of C2C12 cells on a ring, with the insert for cell number increasing exponentially with time. (F) Average velocity and direction of C2C12 cells are indicated by arrow length and direction, respectively. (G) Average velocity of C2C12 cell migration in the circumferential (V_θ) and the radial direction (V_r) as a function of radial position. (H) Average velocity and (I) the circumferential migration velocity of hUVEC cells at the inner and outer ring boundary as a function of time.

For further insight into the biased cell alignment, we analyzed C2C12 cell motion. When the cells were seeded sparsely, no clear bias in motion or alignment was observed over more than 20 hrs of culture (Movie S1). For a higher cell density seeding, cell alignment did not show a clear bias before confluency (after 15 hrs; Fig. 3 D and E and Movies S2 and S3). These data suggest that significant inhibition of the random walk of the cells through physical cell–cell contact was necessary for the cells to express subtle directional migration and biased alignment. The cells were labeled and tracked along the inner and outer ring pattern (Movie S4), and the cell migration velocity was estimated by digital image correlation (26). The speed of migration was higher at the inner and outer ring boundaries than within the interior region (25 μm/hr vs. 10 μm/hr; p < 0.05). The average velocity (Fig. 3F) and the velocity changes in radial and circumferential directions (Fig. 3G) demonstrated that the cells migrated in the CW direction (at 8 μm/hr) at the inner ring boundary (p < 0.05), and in the CCW direction (at 4 μm/hr) at the outer ring boundary (p < 0.05).

At the level of an individual cell, this seemingly opposite circular motion of cells on the inner and outer boundary is in fact consistent with biased migration. Based on cell polarization at boundaries, the x axis can be defined as the direction from the nucleus to centrosome (27), as shown in Fig. 3A. In other words, the cells “face” outward on the outer ring, and inward to the center on the inner ring. Thus, for the C2C12 cells, the biased migration can thus be considered as “leftward bias” along both the inner and outer ring boundary. Also, observed biased alignment of the cells on micropatterns is related to the directional migration at the boundaries, as seen for a C2C12 cell migrating toward a boundary and adopting the leftward biased migration. Because cell polarization and biased migration occur at the boundaries of micropatterns, cell proximity to a boundary is necessary for the expression of chirality. This finding was further supported by the biased cell alignment being most clearly seen in the regions close to the boundaries, especially for less elongated cells such as cardiac fibroblasts (Fig. 2A).

The mean biased angle of human umbilical cord endothelial cells (hUVEC) on rings was similar in magnitude to that of C2C12 cells but was negative (-8.47° ± 0.33°, n = 388), indicating a CW alignment (Table S3 in SI Appendix). By the time the cell chirality was established (15–20 h after seeding; Movies S5–S7), cells along the boundaries had a significantly higher migration speed than those in the interior regions (35 μm/hr vs. 20 μm/hr; p < 0.05). In contrast to C2C12 cells, hUVEC migration was in the CCW direction (15 μm/hr) at the inner ring boundary (p < 0.05) and in the CW direction (20 μm/hr) at the outer ring boundary (p < 0.05) (Fig. 3 H and I). Based on the x axis directed from the nucleus to the centrosome/Golgi apparatus, the migration of hUVECs exhibited a “rightward” bias.

Chirality of Muscle Cells Requires Functional Actin but not Tubulin.

To investigate the roles of actin and tubulin, cyoskeletal proteins putatively linked to cell chirality (1, 2, 12, 14, 27), we used drugs to alter the dynamics of their polymerization and depolymerization (Fig. 4 A and B). For C2C12 and hUVEC cells, cell alignment on micropatterned rings followed a dose-dependent response (Table S8 in SI Appendix). Low concentrations of the actin treadmilling inhibitors (Latrunculin A, cytochalsin D, Jasplakinolide), which did not completely inhibit actin polymerization/depolymerization, reversed the chirality of C2C12 cells from CCW into CW. In contrast, inhibitors of tubulin dynamics (Nocodazole, Taxol) at concentrations below those resulting in cell apoptosis or inhibition of cell migration did not change cell chirality. Similar results were obtained for human skeletal muscle cells (Table S8 in SI Appendix). The CCW bias of the cells depended on the function of actin but not nonmuscle myosin II (Fig. S5 in SI Appendix). In contrast to C2C12 cells, the drugs tested could not reverse the chirality of hUVEC cells or rat cardiac fibroblasts (Table S8 in SI Appendix). Collectively, these data suggest that functional actin is required for the muscle cells exhibiting CCW bias but not the cells exhibiting CW bias.

Fig. 4.

Chirality of muscle cells requires functional actin but not tubulin. (A) Phase contrast images and (B) chirality of C2C12 cells on micro-patterned rings in the presence of 50 nM Latrunculin A, 0.2 μg/mL Cytochalasin D, 30 nM Jasplakinolide, 200 nM Nocodazole, and 10 nM Taxol. Scale bars: 100 μm. (C) Latrunculin A does not change the polarity of C2C12 cells, as the cells positioned their centrosome (bright green), rather than the nucleus (blue), closer to ring boundaries. Scale bars: 50 μm. (D) Migration of C2C12 cells in the presence of Latrunculin A, with the direction and magnitude of velocity indicated by arrow direction and length, respectively. (E) Average velocity of the cells along the circumferential direction (V_θ) and the radial direction (V_r) as a function of radial position.

Interestingly, C2C12 cells treated with Latrunculin A polarized in the same fashion as the untreated cells (Fig. 4C), as evidenced by organelle positioning relative to boundaries, suggesting that the drug did not alter cell polarization. Analysis of cell migration (Movies S8 and S9) showed reversal at the boundaries (Fig. 4 D and E), with the cells migrating CW along the outer ring and CCW along the inner ring at 15 μm/hr (p < 0.05). Similar analyses showed that C2C12 cells treated with 200 nM Nocodazole polarized and migrated in the same fashion as the untreated controls (Fig. S6 in SI Appendix). Thus, inhibition of actin function reversed the CCW chirality and the biased migration of C2C12 cells.

The establishment of cell asymmetric alignment on micropatterns required definite polarization and directional migration of the cells on appositional boundaries of geometries such as rings and long strips. Compared to the cells cultured on large surfaces, and microscale circles or squares, rings and strips provided narrow space for the cells to align and to sense boundaries. In addition, appositional boundaries offered a primary direction for the cells to elongate and migrate, such that any bias in cellular LR decision would be amplified, leading to directional motion. Such a mechanism required at least two-cell width between opposing boundaries (> 20 μm). Otherwise, a single cell would receive confusing signals by simultaneously touching both boundaries, and not express definite polarization or directional migration.

During native development, boundaries are established by cell populations compartmentalized into distinct functional units (e.g., in Drosophila wing), in response to gradients of morphogens (28), and differential adhesion and cortical tension (29). Interestingly, even at the four-cell and eight-cell stages, the snail embryos exhibit a biased alignment, which has a similar response to the drugs (i.e., Latrunculin A and Nocodazole) as mammalian muscle cells (2).

We propose that cell chiral alignment is associated with cell migration based on the following observations: (i) The cells are randomly oriented right after seeding, and the establishment of cell alignment required cell motion; (ii) biased alignment was initiated at the same time as the directional motion of cells on boundaries; and (iii) the chirality of cell alignment was consistently paired with the leftward or rightward bias of cell motion at the boundaries. Following Latrunculin A treatment, the chirality of C2C12 cells was reversed, as was the biased motion of cells on boundaries.

From the chirality data of muscle cells treated with Latrunculin A, we infer that it is possible that two competing mechanisms coexist within cells to determine their LR decisions or chirality. One mechanism would require actin function and lead to the intrinsic leftward bias at the boundaries and subsequent CCW alignment in chiral morphogenesis. The second mechanism would induce intrinsic rightward bias at the boundaries and a CW alignment in chiral morphogenesis. Our drug treatment studies do not exclude the role of microtubule/centrosome in determining cell chirality, especially for the rightward bias, which is worthy of further investigation. The differences in chirality may be due to the higher expression of actin in muscle cells than other cells (30, 31), necessitating identification of cell-type-related determinants of chirality in tissue development. Interestingly, actin-dependent mechanisms were reported to account for chiral properties of the Xenopus egg cortex (32), early development of asymmetry in Xenopus embryos (33), and cardiac looping in vertebrate embryos (34, 35). In addition, alternations in actin polymerization were reported to regulate phonotypical events in malignant cells (36). Further studies are necessary to evaluate the role of actin expression levels in the establishment and reversal of cell chirality, possibly mediated through the noncanonical Wnt signaling pathway, which plays a critical role in pattern determination during embryonic development (37).

In summary, cells cultured on micropatterns with well-defined appositional boundaries exhibit chiral morphogenesis that can be readily determined by analysis of cell alignment and directional motion. For cell populations to express chirality on micropatterns, it was necessary to provide: (i) close appositional boundaries for the cells to sense and polarize; (ii) certain confluence (75% or more) to inhibit cell random walk; and (iii) sufficient time (≥15 hours) for the cells to migrate. In studies of 11 different cell types cultured on thousands of ring-shaped patterns, we observed that the cell chirality was defined by the cell phenotype, and that the loss of actin but not microtubule function could reverse the CCW cell chirality. The simple and highly accurate in vitro platform developed in these studies could potentially be used to study the initiation of chiral morphogenesis and identify genetic, biochemical, and environmental factors leading to malformations.

Methods

Microcontact Printing.

Cell patterning was done by using polydimethylsiloxane (PDMS) elastomeric stamps and self-assembly monolayers (SAMs) (20, 21, 24). A master mold was first fabricated with SU-8 2050 photoresist (MicroChem Corp.) and chromium masks with desired geometric features. The mixture (10∶1) of PDMS prepolymer and curing agent (Dow Corning) was poured into the mold and cured at 70 °C for 4 h.

In most studies (and unless indicated otherwise), an adhesive SAM octadecanethiol (Sigma) was transferred onto the gold-coated (150 Å in thickness) glass slide with the PDMS stamp (Fig. S1 in SI Appendix) (20). The slide was then immersed in a nonadhesive ethylene glycol-terminated SAM (HS-(CH₂)₁₁-EG₃, Prochimia) for 3 h. Finally, patterned surfaces were washed with ethanol and coated with 10 μg/mL fibronectin (Sigma) for 30 min.

Alternatively (Fig. S1 in SI Appendix), the PDMS stamp was coated with 50 μg/mL fibronectin for 30 min, aspirated, and dried in the air for 1 min (24). The stamp was then gently placed onto tissue culture-treated dishes for 2 min and incubated in a nonadhesion SAM, 100 μg/mL poly-L-lysine-polyethylene glycol (PLL-g-PEG; Susos AG), for 1 h. Finally the surface was washed with phosphate buffered saline (PBS).

Cell Culture.

Cells were maintained in tissue flasks with culture media specified in Table S1 in SI Appendix. After reaching 70% confluency, cells were trypsinized and seeded onto protein-coated patterned surfaces. Once the cells attached, extra cells were washed off with fresh medium. At this time drugs were added into the culture medium if necessary. Phase contrast images were taken after overnight incubation at 37 °C and 5% CO₂ when cells reached confluency on the ring patterns.

Drug Treatment.

To examine the role of actin in LR asymmetry, cells were tested with 20–500 nM Latrunculin A, 0.05–1.0 μg/mL Cytochalasin D, and 1–100 nM Jasplakinolide. Latrunculin A inhibits actin polymerization by forming a 1∶1 molar complex with G-actin, thereby inhibiting its ability to polymerize into F-actin (38). Cytochalasin D inhibits actin polymerization by binding to the growing ends of F-actin chains and thus preventing the attachment and addition of G-actin monomers (39). Jasplakinolide is known to bind to and stabilize actin filaments in vitro (40).

To examine the role of microtubules in LR asymmetry, 0.2–2 μM Nocodazole, and 0.3–30 nM Taxol were used. Nocodazole suppresses microtubule dynamics by destabilizing and disassembling microtubules (39). Taxol inhibits the microtubule depolymerization and stabilizes microtubules (41).

To examine the role of the actomyosin motor, cells were tested with 0.2–10 mM Y-27632, 1–20 mM ML-7, and 0.5–10 mM Blebbistatin. Y-27632 works as a selective inhibitor to prevent the phosphorylation of the myosin regulatory light chain (42). ML-7 acts as a selective inhibitor of the myosin light chain kinase (43). Blebbistatin forms a low actin affinity complex through binding to myosin heads, causing the inhibition of nonmuscle myosin II ATPase activity (44).

Immunofluorescence Staining.

After imaging, cells were fixed with 4% formaldehyde in cytoskeletal buffer (10 mM MES, 138 mM KCl, 3 mM MgCl₂, 2 mM EGTA, and 0.32 M sucrose) for 25 min. For actin/tubulin double staining, the cells were incubated with phalloidin-TRITC (1∶400; Invitrogen) and anti-Tubulin-FITC (1∶50; Sigma) for 1 h. For the Golgi apparatus positioning inside patterned rings, the cells were incubated in 1 μg/ml antihuman golgin-97 (Invitrogen) for 1 h. After secondary antibodies, cell nuclei were stained with 200 ng/mL 40, 6-diamidino-2-phenylindole (DAPI; Sigma) for 10 min. Finally, the cells were mounted with Fluoromount-G medium (SouthernBiotech).

Analysis of Cell Alignment.

High-resolution phase contrast images of live patterned cells were taken at a resolution of approximately 0.645 μm/pixel, and analyzed using a custom-written code in MatLab (MathWorks), based on the automated detection of intensity gradient and circular statistics (22). In this algorithm, the intensity gradient was determined pixel by pixel with a Gaussian differential filter. In each subregion of the image, the dominant local direction was determined using an accumulator scheme, in which the orientation of each pixel follows a von Mises distribution, a circular analogue of the linear normal distribution. Subsequently, the orientation in each subregion was converted into an angle bias based on its deviation from the circumferential direction (see Fig. 1C). Mean angle and standard deviation of LR asymmetry were calculated for all subregions, using circular statistics (45). We verified that the variation of subregion size from 10 by 10 pixels to 60 by 60 pixels would not significantly alter the judgment of cell chirality on rings. The subregion size was therefore set to 20 by 20 pixels (i.e., 13 by 13 μm).

Analysis of Cell Migration.

For time-lapse videos, cells were patterned on a gold-coated glass-bottom Petri dish. After cells attached, the dish was transferred into an environmental chamber (37 °C and 5% CO₂) and image time series were recorded every 5 min at a resolution of 1.56 μm/pixel for a total time of 20–40 h. As the image capture rate is much higher than the characteristic time for cell migration, digital image correlation, together with subpixel displacement estimation, were used to determine the displacements of cell migration. A fast Fourier transform (FFT)-based method was first utilized to match regions of two sequential phase contrast images. The calculated displacement values were then used as inputs for a more accurate estimation of displacement fields at a subpixel level, using a second-order image correlation algorithm described previously (26). To evaluate the bias of cell migration, the obtained velocity field was further projected in the circumferential (V_θ) and radial (V_r) direction.

Statistical Analysis.

Cell chirality on ring patterns (i.e., clockwise or counterclockwise alignment) was determined from calculated biased angles in local regions with one sample test for the mean angle, analogous to the one sample t test in linear statistics (45). The overall biased behavior of the cells was tested based on the number of rings exhibiting either clockwise or counter clockwise alignment in the rank test. The directionality of cell migration on boundaries was determined with the one-tailed Student t test. The confidence level was set to 0.05 for all statistical tests.