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. Author manuscript; available in PMC: 2025 Aug 1.
Published in final edited form as: Biomech Model Mechanobiol. 2024 Mar 19;23(4):1289–1298. doi: 10.1007/s10237-024-01839-1

A homeostatic role of nucleus-actin filament coupling in the regulation of cellular traction forces in fibroblasts

Naoya Sakamoto 1,2, Keisuke Ito 1, Satoshi Ii 1,2, Daniel E Conway 3, Yuki Ueda 1, Jiro Nagatomi 2,4
PMCID: PMC11932025  NIHMSID: NIHMS2062844  PMID: 38502433

Abstract

Cellular traction forces are contractile forces that depend on the material/substrate stiffness and play essential roles in sensing mechanical environments and regulating cell morphology and function. Traction forces are primarily generated by the actin cytoskeleton and transmitted to the substrate through focal adhesions. The cell nucleus is also believed to be involved in the regulation of this type of force; however, the role of the nucleus in cellular traction forces remains unclear. In this study, we explored the effects of nucleus-actin filament coupling on cellular traction forces in human dermal fibroblasts cultured on substrates with varying stiffness (5, 15, and 48 kPa). To investigate these effects, we transfected the cells with a dominant-negative Klarsicht/ANC-1/Syne homology (DN-KASH) protein that was designed to displace endogenous linker proteins and disrupt nucleus-actin cytoskeleton connections. The force that exists between the cytoskeleton and the nucleus (nuclear tension) was also evaluated with a fluorescence resonance energy transfer (FRET)-based tension sensor. We observed a biphasic change in cellular traction forces with a peak at 15 kPa, regardless of DN-KASH expression, that was inversely correlated with the nuclear tension. In addition, the relative magnitude and distribution of traction forces in nontreated wild-type cells were similar across different stiffness conditions, while DN-KASH-transfected cells exhibited a different distribution pattern that was impacted by the substrate stiffness. These results suggest that the nucleus-actin filament coupling play a homeostatic role by maintaining the relative magnitude of cellular traction forces in fibroblasts under different stiffness conditions.

1. Introduction

Cells are able to sense and respond to the mechanical properties of the surroundings. For example, the substrate stiffness is a critical environmental stimulus that plays roles in cell behaviors including cell morphology (Solon et al. 2007), migration, proliferation, and differentiation (Denis et al. 2021; Engler et al. 2006), all of which are involved in physiological and disease processes (Ladoux and Mege 2017; Tschumperlin et al. 2018). Understanding the mechanisms by which cells respond to their mechanical environment can help develop cell-based regenerative medicine therapies.

Contractile forces that cells exert on a substrate, the so-called cellular traction forces, depend both on the substrate stiffness (Lo et al. 2000) and the mechanical cellular stress (Eckert et al. 2021; Ting et al. 2012), and play an essential role in sensing mechanical stimuli (Engler et al. 2006). Traction forces are primarily generated by the actin cytoskeleton and transmitted to the substrate via focal adhesions (Goffin et al. 2006). The cell nucleus is thought to also be involved in the regulation of traction forces, thus directing cellular morphology and related functions. The nucleus is directly connected to the actin cytoskeleton by the linker of nucleoskeleton and cytoskeleton (LINC) complex, an evolutionarily conserved structure located on the nuclear envelope, which consists of nesprins and SUN proteins (Crisp et al. 2006; Stewart-Hutchinson et al. 2008). Previous studies have reported that knocking down nesprin-1, a protein directly linked to actin filaments, suppresses cellular responses to external mechanical stimuli (Anno et al. 2012; Sakamoto et al. 2017), and induces changes in cellular traction forces (Chancellor et al. 2010; Graham et al. 2018). In addition, changes in the cell adhesion area and cell morphology affect not only cellular traction forces, but also forces between the nucleus and the cytoskeleton (Arsenovic et al. 2016). Therefore, it has been proposed that nuclear forces play important roles in the regulation of cellular traction forces, although details of its role in these processes are still not well understood.

Herein, we investigated changes in the cellular traction force in cells with impaired LINC complex activity on different subtracted stiffnesses, and also evaluated cellular forces acting on the nucleus to understand the role of nucleus-actin filament coupling in the regulation of traction force.

2. Methods

2.1. Cell culture and DN-KASH transfection

Human skin fibroblasts (Riken BRC, Japan) were cultured in MEMα medium (Fujifilm Wako, Japan) containing 10% heat-inactivated fetal bovine serum (Sigma-Aldrich, USA) and 100 unit/mL penicillin/streptomycin (Fujifilm Wako). Cells were transfected with a vector specific for the dominant-negative nesprin KASH (DN-KASH) protein (Arsenovic et al. 2016; Denis et al. 2021), using a Lipofectamine LTX transfection reagent (Thermo Fisher Scientific, USA). One day after the transfection, the expression and localization of the DN-KASH in the nuclear envelope of transfected cells were confirmed by fluorescence microscopy (Fig. 1A).

Fig. 1.

Fig. 1

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Typical fluorescence images of wild-type (WT) and dominant-negative nesprin (DN-KASH, mCherry) fibroblasts. (A) Nuclear images of WT and DN-KASH-transfected cells. Bars indicate 20 μm. (B) Confocal fluorescence images of the nucleus and actin filaments in a WT fibroblast cultured on a 5 kPa polyacrylamide gel. Bars indicate 50 μm

2.2. Polyacrylamide gel preparation and traction force microscopy

To prepare hydrogel substrates with varying stiffness levels, different concentrations of acrylamide and bis-acrylamide were polymerized between silanized and siliconized glass coverslips using spacers of 0.03 mm thickness. The elastic modulus of each substrate was determined via atomic force microscopy indentation (described below), and substrates with moduli 5.1 ± 0.2 kPa (mean ± standard deviation (SD)), 15.2 ± 0.9 kPa, and 47.6 ± 1.7 kPa, hereinafter referred to as 5, 15, and 48 kPa gels, respectively, were used. The gel surface was activated with sulfo-SANPAH (Thermo Fisher Scientific) and then coated with 0.1 mg/mL collagen type I (CellMatrix Type I-C, Nitta Gelatin, Japan) prior to cell seeding.

Polyacrylamide gels containing polystyrene beads (Thermo Fisher Scientific) of 0.2 μm in diameter were prepared and traction force microscopy was performed in a stage-top incubator (Tokai Hit, Japan) mounted on a microscope stage. Images of the beads with cells on the gel substrate and after cell removal by trypsin-EDTA treatment were captured with an inverted fluorescent microscope (Olympus, Japan). The displacement and strain fields of the beads were obtained, and the magnitude of the cellular traction stress was inversely estimated using an adjoint method (Ambrosi et al. 2009; Ii et al. 2019). Average and maximum traction stresses were calculated for the whole cell area.

2.3. Image acquisition and analysis of the FRET nesprin-2G tension sensor

The binding forces between the cytoskeleton and the cell nucleus (nuclear tension) were quantified using a fluorescence resonance energy transfer (FRET)-based tension sensor for nesprin-2G (nesprin-TS) (Arsenovic et al. 2016). Cells were transfected with nesprin-TS in the same manner as DN-KASH transfection. Images of a live cell expressing mTFP1 (donor) and venus (acceptor) were acquired using an inverted fluorescent microscope equipped with a 3-CCD camera (Hamamatsu, Japan) and the stage-top incubator. Excitation light through a mercury lamp (100 W) with an excitation filter (427/10 nm) was used in this study. The ratio image was then calculated by dividing the acceptor intensity image over the donor intensity image and multiplying it with a binary image mask that was made from the outline of the nuclear image with ImageJ. Fluorescent intensities of the inside of the nucleus were excluded in this study because they possibly included the signals from the out-focus regions. The donor and acceptor images were background subtracted before calculating the ratio. The FRET ratio is inversely proportional to the nuclear tension, i.e., the higher the nuclear tension, the lower the FRET ratio. The average FRET ratio in the region of interest was obtained and reported.

2.4. Atomic force microscopy indentation

To determine the elastic moduli of polyacrylamide gel substrates and cell nuclei, indentation analysis was performed using an atomic force microscope (AFM, JPK, Germany). Force-indentation curves were obtained and fitted using the following Sneddon model equation (Sneddon 1965):

F=2Etanαπ1-ν2δ2, (1)

where E is elastic modulus, v Poisson’s ratio of the substrate (= 0.5), F loading force, δ indentation depth, α opening angle of the cantilever tip. V-shaped silicon nitride cantilevers (pyramidal tip, a spring constant = 0.08 N/m, Olympus) were used. The curves obtained in the maximum indentation depth of approximately 1.0 μm were used for analyses.

To assess nuclear properties, cells were treated with 160 nM Latrunculin B (Fujifilm Wako) for 30 min before indentation tests to depolymerize the actin cytoskeleton on the apical surface of the nuclei (Khatau et al. 2009). Although treatments for actin depolymerization may have an effect on nuclear stiffness, actin cytoskeletons mechanically and dominantly contribute to the cell membrane and the nucleus (Wang et al. 2018) and these treatments have been adopted for measuring the nuclear stiffness in previous studies (Kasas et al. 2005; Nagayama 2021). Actin cytoskeleton depolymerization was confirmed by fluorescence microscopy (data not shown). Force-indentation curves for near central nuclear regions were obtained to calculate the elastic moduli values reported.

2.5. Fluorescence staining and image analysis

Cells on polyacrylamide gel substrates were fixed with 4% paraformaldehyde and costained with Hoechst 33342 (Invitrogen) and either phalloidin-iFluor488 (Abcam, UK) or Alexa Fluor 546 phalloidin (Thermo Fisher Scientific). Confocal microscopic images were acquired at 0.3 μm z-intervals along the apical-basal cell axis (Fig. 1B). To examine cellular and nuclear morphology, z-stack fluorescence images were binarized using the Otsu method, outlines of each cell and its nucleus traced with the ImageJ program (National Institutes of Health, USA) and fitted to an equivalent ellipsoid. To assess nuclear and cellular morphology, the inverse aspect ratio was defined as the ratio between the minor and major axes of the ellipsoid cell outline. The inverse aspect ratio ranges from zero (0) to one (1) and is equal to 1 for a perfect circle approaching 0 for elongated shapes. Cellular and nuclear-projected areas and nuclear heights were also obtained from fluorescence images.

2.6. Statistics

Data were presented as mean ± SD or as mean ± the standard error of the mean (SEM), and derived from at least three independent experiments. Comparisons among means of experimental groups were made using one- or two-way analysis of variance (ANOVA). p-values less than 0.05 were considered statistically significant. The Tukey-Kramer post hoc test was used for comparison among groups when significant differences were detected.

3. Results

Results of cellular and nuclear morphological parameters are shown in Fig. 2. Nontreated wild-type cells (WT cells) exhibited significantly higher cell areas on the stiffer 48 kPa gel than on the softer 5 kPa gel (Fig. 2A). The transfection of DN-KASH significantly decreased cell-projected areas (p < 0.001, ANOVA). The inverse aspect ratio was similar in WT cells on substrates with different stiffness, while DN-KASH-transfected cells exhibited rounded morphology on the stiffer substrate (Fig. 2B). The substrate stiffness was not a determining factor, but DN-KASH transfection led to a decrease in nuclear-projected areas (Fig. 2C). Neither substrate stiffness nor DN-KASH transfection affected the nuclear inverse aspect ratio (Fig. 2D). Nuclear height decreased in WT cells when cultured on the stiffer substrate, while no change was observed in DN-KASH-transfected cells (Fig. 2E).

Fig. 2.

Fig. 2

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Cellular and nuclear morphological analyses. (A) Cell area, (B) Cell inverse aspect ratio, (C) Nuclear area, (D) Nuclear inverse aspect ratio, and (E) Nuclear height for wild-type (WT) and dominant-negative nesprin (DN-KASH) transfected fibroblasts cultured on polyacrylamide gel substrates for 5 kPa, 15 kPa, and 48 kPa elastic modulus formulations. Results are expressed as mean + SEM. Number of cells (n) for WT cells: n = 31 (5 kPa, 15 kPa), n = 29 (48 kPa) and for DN-KASH cells: n = 34 (5 kPa, 15 kPa), n = 29 (48 kPa). *: p < 0.05, †: p < 0.01, ‡: p < 0.001

Figure 3 shows representative confocal fluorescence images at different focal planes of actin filaments in fibroblasts cultured on 15 kPa gel. Actin filaments were observed on the nuclear apical surface of all cells although no detectable differences in the actin filament structure were found between WT and DN-KASH-transfected cells. No evident differences in the actin filament structure were noticed between substrates with different stiffness (Supplementary Fig. S1).

Fig. 3.

Fig. 3

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Typical confocal fluorescence images of actin filaments in wild-type and dominant-negative nesprin (DN-KASH) transfected fibroblasts cultured on a 15 kPa polyacrylamide gel. Cell images were taken at the apical, middle, and basal focal planes. Bars indicate 50 μm

Figure 4A shows the typical overlay of cellular traction stress vectors and differential interference contrast images for fibroblasts cultured on 15 kPa gel. Greater traction stresses were concentrated at the cell periphery in both WT and DN-KASH-transfected cells, which was consistent with the results of previous studies (Ghosh et al. 2007; Li et al. 2008). A similar distribution of cellular traction stress was also observed on 5 and 48 kPa gel substrates (data not shown). The maximum traction stress in both WT and DN-KASH-transfected cells differed depending on the substrate stiffness (Fig. 4B) (p < 0.05, ANOVA) and tended to be higher on the 15 kPa gel than on the 5 and 48 kPa substrates. Similarly, the average traction stress was significantly higher on the 15 kPa gel than on other gels. There were no significant differences in either the maximum or average traction stresses when compared between WT and DN-KASH-transfected cells.

Fig. 4.

Fig. 4

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Results of cell traction force microscopy. (A) Typical traction force vectors overlayed on differential interference contrast images for wild-type (WT) and dominant-negative nesprin (DN-KASH) transfected fibroblasts cultured on a 15 kPa polyacrylamide gel substrate. Cell outlines are shown as white lines. Bars indicate 50 μm. (B) Maximum and average cellular traction stress for WT and DN-KASH expressing fibroblasts cultured on polyacrylamide substrates with different elastic moduli. Data are presented as box and whisker plots. Number of cells (n) for WT cells: n = 7 (5 kPa) n = 12 (15 kPa), n = 6 (48 kPa) and for DN-KASH cells: n = 7 (5 kPa), n = 8 (15 kPa), n = 9 (48 kPa). *: p < 0.05, †: p < 0.01

To examine the role of the cell nucleus in the relative magnitude distribution of the cellular traction stress, we plotted as a histogram normalized traction stress divided by the maximum stress observed in each cell (Fig. 5). WT cells and DN-KASH-transfected cells on the 15 kPa gel substrate showed a similar magnitude distribution of normalized cellular traction stresses regardless of the substrate stiffness. In contrast, the lower ( < = 0.1) normalized cellular traction stresses in DN-KASH-transfected cells on the 5 kPa gel were significantly greater than those on other substrates. Moreover, the relative magnitude distribution of stresses in DN-KASH-transfected cells on the stiffer 48 kPa gel was more uniform than on other substrates.

Fig. 5.

Fig. 5

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Histograms of normalized cellular traction stresses for (A) wild-type (WT) and (B) dominant-negative nesprin (DN-KASH) transfected fibroblasts cultured on glass or different polyacrylamide gel substrates with varying elastic moduli (5 kPa, 15 kPa, and 48 kPa). Number of cells (n) for WT cells: n = 7 (5 kPa) n = 12 (15 kPa), n = 6 (48 kPa) and for DN-KASH cells: n = 7 (5 kPa), n = 8 (15 kPa), n = 9 (48 kPa). Data are expressed as mean + SEM. *: p < 0.05, †: p < 0.01, ‡: p < 0.001

The nuclear tension was determined by the FRET ratio in nesprin-TS expressing cells (Fig. 6). We found that the nuclear tension in fibroblasts was influenced by the stiffness on the PA gel substrates in a biphasic manner (p < 0.01, ANOVA). The FRET ratio on the 15 kPa gel was higher than that on 5 and 48 kPa gels, indicating that the nuclear tension in fibroblasts on the 15 kPa gel was lower than the observed value under different stiffness conditions. By the AFM indentation test, we obtained force-indentation curves for the nuclear regions of cells (Fig. 7A) and determined the elastic moduli of nuclei. Nuclear elastic moduli also showed a biphasic change (p < 0.05, ANOVA) (Fig. 7B). Fibroblast nuclei on the 15 kPa gel showed a lower elastic modulus than cells on 5 and 48 kPa gels.

Fig. 6.

Fig. 6

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FRET ratio for nesprin tension sensor-expressing fibroblasts cultured on polyacrylamide substrates with different elastic modulus (5 kPa, 15 kPa, and 48 kPa). Data are presented as box and whisker plots. Number of cells (n): n = 8 (5 kPa), n = 6 (15 kPa), n = 10 (48 kPa). †: p < 0.05

Fig. 7.

Fig. 7

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Results of AFM indentation tests. (A) A typical force-indentation curve for the nuclear region of a WT cell. (B) Nuclear elastic moduli of fibroblasts cultured on polyacrylamide substrates with varying stiffness levels (5 kPa, 15 kPa, and 48 kPa). Data are presented as box and whisker plots. Number of cells (n): n = 20 (5 kPa), n = 25 (15 kPa), n = 22 (48 kPa). *: p < 0.05

4. Discussion

Based on the results reported in previous studies, we expected to observe greater traction forces in cells cultured on stiffer substrates than on softer substrates (Goffin et al. 2006; Lo et al. 2000). However, we found that the maximum and the average cellular traction stress tended to increase between 5 and 15 kPa gels, but then decreased between the 15 and 48 kPa gel (Fig. 8). This type of biphasic cell response to substrate stiffness was not completely unexpected as it has been reported in several previous studies. For example, it has been reported that embryonic fibroblast and smooth muscle cell migration speed initially increases as substrate stiffness increases until reaching a threshold level, and then decreases on stiffer substrates (Graham et al. 2018; Peyton and Putnam 2005). Moreover, Kong et al. (Kong et al. 2005) showed that the cellular traction force in MC-3T3 osteoblasts peaks at intermediate levels of substrate stiffness and is lower on softer and stiffer substrates. We need further investigation to understand the reasons behind the biphasic change in the cellular traction force in response to substrate stiffness. However, since a similar pattern was found in DN-KASH-transfected cells, it is speculated that nucleus-actin filament coupling does not play a major role in the cellular traction forces as a function of substrate stiffness.

Fig. 8.

Fig. 8

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Schematic summary of the results of this study

The results of the present study suggest a role for nucleus-actin filament coupling in the distribution of normalized cellular traction stresses. The histogram of the normalized cellular traction stress in WT cells showed a similar gamma-like distribution under different stiffness conditions (Fig. 5). In contrast, the distribution in DN-KASH-transfected cells was of an exponential distribution, which gradually flattened as substrate stiffness increased. This result indicates that cytoskeletal actin-mediated binding plays an important role in the regulation of relative cellular traction forces. In addition, we found that the cytoskeletal actin-mediated forces transmitted to the nucleus by binding proteins show an inverse tendency to the change in the traction force induced by substrate stiffness (Figs. 6 and 8). In a preliminary experiment, we confirmed that the treatment with myosin light chain kinase inhibitor led to a decrease in traction forces shown by an increase in the FRET index, indicating that both nuclear tension and cellular traction forces depend on actomyosin contractility (Supplementary Fig. S2). In this study, we only evaluated FRET signals of the nesprin-TS at around the middle focal plane of the nucleus due to the limitation of the axial resolution of epifluorescence microscopy. This did not allow us to examine the role of the actin filament structure above the nuclear apical surface, the so-called actin cap, which is believed to play a crucial role in the transmission of mechanical forces to the nucleus (Khatau et al. 2009). Nonetheless, our results imply the existence of a control system that independently regulates the contractility of actin filaments linked to focal adhesions in the basal cell surface and to the cell nucleus in the cytoplasm. Furthermore, it is possible that nucleus-actin filament coupling plays a role in maintaining the relative distribution of the cellular traction force at a constant level (i.e., the homeostasis of relative traction forces) under different stiffness conditions.

Previously, it was shown that knocking down nesprin-1, a core component of the LINC complex, induces increases in the traction force of vascular endothelial cells (Chancellor et al. 2010). Although we detected an increasing tendency in higher normalized traction stresses for DN-KASH cells on the 48 kPa gel compared to those on the 15 kPa gel, there are no statistical significances in maximum and average traction forces between WT and DN-KASH cells. Since nesprin-2 also mediates the nucleus and actin filament coupling and potentially compensate for the loss of nesprin-1 (Zhang et al. 2007), the effect of DN-KASH transfection may not be the same as the knockdown of nesprin-1. Therefore, in a separate preliminary study, we also performed siRNA-mediated nesprin-1 knockdown in fibroblasts (Supplementary Fig. S3). Similar to the case of the DN-KASH, the knockdown of nesprin-1 resulted in a tendency to increase the higher normalized traction stresses, but siRNA-transfected cells also exhibited a biphasic change in the traction force in response to substrate stiffness, and similar magnitudes at the maximum and average cellular traction stress as observed in WT cells. Our previous results suggested differential effects of nesprin-1 knockdown on the morphology of fibroblasts and vascular endothelial cells under cyclic strain conditions (Anno et al. 2012; Sakamoto et al. 2017). Together these results suggest that the role of nucleus-actin filament coupling in the generation and regulation of traction forces may also be dependent on the cell type.

The results provided additional evidence of a biphasic change in the nuclear elastic modulus under different stiffness conditions, showing the same tendency as the nuclear tension with nesprin-TS. Previous studies have reported that externally applied mechanical stimuli to both whole cells and nuclei induce changes in nuclear mechanical properties. In vascular endothelial cells, Deguchi et al. (Deguchi et al. 2005) observed that nuclear elastic modulus increased by the application of fluid shear stress. The change in nuclear mechanical properties is thought to be caused by forces transmitted from the cell surface to the nucleus via the cytoskeleton. Moreover, Guilluy et al. (Guilluy et al. 2014) reported that cyclic tension forces exerted through the attachment of nesprin-1 to the nuclear membrane increased nuclear mechanical properties in isolated nuclei from HeLa cells. In addition, Swift et al. (Swift et al. 2013) showed that the expression of Lamin A, a structural protein of the nuclear lamina which is well-known to contribute to nuclear stiffness, increases the intracellular tension as substrate stiffness increases. We found that the nuclear elastic modulus and the force acting on the nucleus tended to be inversely correlated with the magnitude of the cell traction force. Our results support the idea that changes in nuclear mechanical properties may be in response to the nuclear tension mediated by nesprins, and such a tension force depends on the effect exerted by the substrate stiffness.

Changes in LINC complex protein expression have been reported for cells from Hutchinson–Gilford progeria syndrome (HGPS) patients (Dahl et al. 2006; Hale et al. 2008). HGPS cells exhibit reduced traction forces (Booth-Gauthier et al. 2013) as well as decreased nuclear tensions (Arsenovic et al. 2016) even though their nuclei are stiffer than those of normal cells (Verstraeten et al. 2008). Although it is difficult to directly compare the mechanobiological behaviors of normal cells with those of patient cells with genetic mutations, this relationship is contrary to the results of the present study. HGPS is typically caused by autosomal dominant mutations in the LMNA gene which encodes the nuclear intermediate filament proteins lamin A and C, and the HGPS cells show not only increased nuclear stiffness but also abnormal nuclear shapes (Arsenovic et al. 2016; Verstraeten et al. 2008), which could also affect the regulation of traction forces. It would be interesting to investigate the homeostasis of relative traction forces in HGPS cells and the contribution of the structural proteins and shapes of nuclei in future works.

Previous studies have shown that, in fibroblasts, the increase in the cellular area and the decrease in the nuclear height may be in response to an increase in substrate stiffness (Engler et al. 2006; Lovett et al. 2013) and the correlation between the cell area and the nuclear height (Vishavkarma et al. 2014). In the present study, a similar relationship between the cell area and nuclear height in WT cells was observed. Specifically, we found that cellular and nuclear-projected areas were greater in WT cells than in cells where nucleus-actin filament coupling was disrupted by DN-KASH transfection. Furthermore, DN-KASH transfection suppressed changes in the cellular/nuclear-projected area and nuclear height depending on the substrate stiffness. These are in line with the findings reported by Lovett et al. (Lovett et al. 2013) demonstrating that the effect of substrate mechanical properties on the cellular/nuclear morphology can be abolished by inhibition of nucleus-actin filament coupling. However, in the present study, no correlation was observed between cellular/nuclear morphology, cellular traction, and nuclear forces, contrary to previous findings that changes in the cytoskeletal actin tension are associated with changes in the cellular/nuclear morphology. Other factors including cytoskeletal networks (i.e., microtubules and intermediate filaments), and focal adhesions may also be involved in the regulation of the cellular/nuclear morphology in response to substrate stiffness.

It has been suggested that the force transmitted through the LINC complex is further transferred to the inside of the nucleus via structural molecules, such as emerin and lamin A/C, and then involved in local transcriptional activities (Graham and Burridge 2016). It is also reported that force transmission to the nucleus through nesprins is essential for nuclear translocation of a transcription factor YAP (Driscoll et al. 2015). Thus, the changes in the nuclear tension found in the present study may be part of transcriptional responses to the stiffness of the extracellular matrix. Although we need to be careful not to disrupt the balance between these nuclear molecules, the knockdown of interrelated proteins of the LINC complex using siRNA can help reveal the role of nucleus-actin filament coupling in force regulation. Moreover, understanding the role of the changes in the nuclear tension may contribute to the research of skin regeneration. It was reported that the actomyosin tension in response to the tissue stiffness is involved in extracellular matrix reorganization and fibroblast proliferation (Fu et al. 2024). Tensional homeostasis at the cellular level, as well as force transmission to the cell nucleus, has also been suggested to be crucial for skin functions and structures (Kimura et al. 2020; Samuel et al. 2011). Indeed, the biphasic change of the cellular traction force observed in fibroblasts may be associated with differential cellular responses to substrate stiffness. The elastic modulus of dermis tissues measured by AFM indentation is approximately 20 kPa (Achterberg et al. 2014; Petrie et al. 2012), and normal cultured fibroblasts are able to maintain a WT phenotype on substrates with an elastic modulus of up to 15 kPa but lose on stiffer substrates in response to stimulation of transforming growth factor-β1 (Achterberg et al. 2014). Further investigation on the homeostatic regulation of the relative traction forces and the nuclear tension will contribute to the elucidation of the detailed mechanisms and regeneration of physiological functions of the skin.

5. Conclusion

In the present study, cellular traction and nuclear tension forces in fibroblasts cultured on substrates at varying stiffness levels were evaluated. The cellular traction force showed a biphasic response to stiffness regardless of nucleus-actin filament coupling, and WT traction forces were inversely correlated with nuclear tension. In addition, the magnitude of normalized traction forces in nontreated cells was similar across different stiffness conditions, while the transfected cells with disrupted actin-mediated binding function exhibited a different distribution pattern that was impacted by the substrate stiffness. The results of the present study suggest that nucleus-actin filament coupling has a homeostatic role in regulating the relative magnitude and distribution of cellular traction forces in cells under different stiffness conditions.

Supplementary Material

Supplementary Material

Acknowledgements

This study was supported in part by National Institutes of Health R35GM119617, Grants-in-Aid for Scientific Research from the MEXT of Japan (Nos. 18H03521, 18K19934, and 22K19898), a Tokyo Metropolitan Government Advanced Research Grant (R2–2), and the TMU Research Project for Emergent Future Society.

Footnotes

Competing interests The authors declare no competing interests.

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/s10237-024-01839-1.

Data availability

No datasets were generated or analysed during the current study.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Material
NIHMS2062844-supplement-Supplementary_Material.docx (14.9MB, docx)

Data Availability Statement

No datasets were generated or analysed during the current study.

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