Abstract
As cells enter mitosis, cell cortex contraction generates surface tension to establish a geometry feasible for division in a physically confined environment. Cell surface tension rises in prophase and continues to stay constant during metaphase to support mitosis. How the cell surface tension is maintained throughout mitosis is not well explored. We show that the cell surface tension is actively maintained by a mechanosensitive RhoA pathway at the cell cortex during mitosis. Mechanical activation of RhoA leads to non-muscle myosin IIB (NMIIB) stabilization and mechanosensitive accumulation at the cell cortex via Rho kinase (ROCK) regulation of the NMIIB head domain. Interestingly, when the NMIIB tail domain regulation is perturbed, the NMIIB has reduced ability to generate tension but could still support mitotic cells to withstand compressive stress by undergoing mechanosensitive accumulation at the cell cortex. Thus, mechanical RhoA activation drives NMIIB mechanoresponse via its head domain regulation to maintain cell surface tension during mitosis.
Introduction
Most animal cells undergo a dramatic change of cell geometry when they enter mitosis (Ramkumar and Baum, 2016). Upon entry into mitosis, cells increase their intracellular hydrostatic pressure by water influx, and concomitantly, contracts the cell cortex to generate higher cell surface tension to counteract the hydrostatic force (Chugh et al., 2017; Ramanathan et al., 2015; Roubinet et al., 2012; Stewart et al., 2011). These cellular forces together lead to mitotic cell rounding that provides an ideal cell geometry to undergo mitosis and cytokinesis (Lancaster et al., 2013). Establishing and maintaining cell surface tension to sustain a minimum cell height for a functional cell geometry during mitosis is particularly important when cells are dividing in a physically confined tissue environment (Cattin et al., 2015; Matthews et al., 2020). Within a tissue environment, cells constantly experience mechanical stresses such as compressive stress from surrounding cells and the extracellular matrix. This is particularly significant in an overgrowth tumor environment where the excessive cell proliferation, cell infiltration and matrix deposition results in an accumulation of solid stress or compressive stress inside the tumors (Nia et al., 2016; Nia et al., 2020). Cells that are not able to withstand the compressive stress during mitosis have defects in their mitotic spindle stability and form multipolar spindles that lead to chromosome mis-segregation and aneuploidy (Lancaster et al., 2013; Matthews et al., 2020). Thus, the ability to generate and sustain cell surface tension in mitotic cells in a crowded environment is important in maintaining their genome integrity (Cadart et al., 2014; Taubenberger et al., 2020). How mechanistically cells maintain cell surface tension during mitosis in a confined environment remains unclear.
Cells progressively increase their surface tension from prophase and maintain a constant high level of the surface tension throughout mitosis with the actin cortex homeostatically maintained (Nishimura et al., 2019; Stewart et al., 2011). Cell surface tension is generated by contractile forces derived from an actomyosin-based cell cortex. In the cell cortex, actin filaments turn over to maintain a homeostatic network for the molecular motor non-muscle myosin II (NMII) to slide and generate forces (Chew et al., 2017; Chugh et al., 2017). As cells enter mitosis, a Rho guanine nucleotide exchange factor (RhoGEF) Ect2 activates RhoA, which in turn binds to the cell membrane and recruits its effectors such as anillin, formin and Rho-associated kinase (ROCK) to activate actomyosin contractility at the cell cortex (Maddox and Burridge, 2003; Matthews et al., 2012; Ramanathan et al., 2015). Interestingly, NMIIs display mechanoresponse by accumulating at the cell cortex in response to mechanical stress (Effler et al., 2006; Schiffhauer et al., 2016; Schiffhauer et al., 2019). In addition, a tension-dependent activation of RhoA signaling has been detected at the cell-cell adherens junctions and the cell matrix focal adhesion to reinforce NMII contraction (Acharya et al., 2018; Guilluy et al., 2011). A similar mechanosensitive RhoA activation has not been demonstrated in mitosis to regulate NMII’s mechanoresponse despite ROCK is required for NMII’s mechanoresponse at the cell cortex (Schiffhauer et al., 2016).
Results and discussion
To study how cell surface tension is maintained during mitosis, we made a human MCF10A mammary epithelial cell stably expressing fluorescently tagged non-muscle myosin IIB (mEmerald-NMIIB) and active RhoA biosensor consisting of the RhoA-binding domain of anillin (mRuby2-AHPH) (Budnar et al., 2019; Piekny and Glotzer, 2008). NMIIB localizes to the cell cortex in the early mitosis and functions downstream of active RhoA to generate cortical tension (Taneja et al., 2020). Live-cell microscopy imaging revealed that NMIIB and active RhoA localized to the cell cortex starting from early mitosis and enriched at the division site during mitosis and cytokinesis, consistent with previous studies and their roles in tension generation at the cell cortex during cell division (Fig. S1 A) (Piekny and Glotzer, 2008; Taneja et al., 2020). When mitotic cells were treated with a RhoA inhibitor C3 exoenzyme that prevents activation of RhoA, both active RhoA and NMIIB disappeared from the mitotic cell cortex (Fig. S1 B). This indicates that mRuby2-AHPH reliably reports the active state of RhoA, and NMIIB and anillin AHPH localizations report the status of cellular contractility during mitosis and cytokinesis.
We first addressed whether the maintenance of cortical NMIIB and cell surface tension during mitosis required an active NMIIB turnover. To this end, we treated mitotic cells with low, medium and high concentrations of the low cytotoxic and photostable NMII inhibitor para-nitro-blebbistatin (p-nitroblebb), which lowers the actin affinity of NMII heads (Kepiro et al., 2014; Rauscher et al., 2018). Mitotic cells treated with a low concentration of p-nitroblebb showed punctate cortical NMIIB patterns (Fig. 1 A, insets). Higher concentrations of p-nitroblebb resulted in a decreasing NMIIB intensity at the mitotic cell cortex (Figs. 1 A and B). Analyses of the NMIIB turnover at the mitotic cell cortex with the fluorescence recovery after photobleaching technique (FRAP) showed an increase of the immobile fractions of NMIIB upon increased concentrations of p-nitroblebb (Figs.1 C and D; DMSO: 0.27 ± 0.08; [Low]: 0.45 ± 0.01; [Medium]: 0.50 ± 0.01; [High]: 0.59 ± 0.03). The recovery half time of cortical NMIIB decreased when increasing concentrations of p-nitroblebb were used (Fig. 1 D; DMSO: 20.33 ± 2.09 s; [Low]: 12.61 ± 1.46 s; [Medium]: 11.30 ± 0.59 s; [High]: 9.49 ± 0.05 s). The FRAP analysis suggests that a decrease of NMIIB recovery half time with higher immobile fractions characterizes inhibition of NMIIB function at the cell cortex.
Next, we measured cell surface tension in mitotic cells treated with p-nitroblebb using the micropipette aspiration technique. The mitotic cell surface tension decreased to 56%, 44%, 31% of the control group after treatments with low, medium and high concentrations of p-nitroblebb, respectively (Fig. 1 E; DMSO: 894.90 ± 186.74 pN/μm; [Low]: 500.42 ± 90.64 pN/μm; [Medium]: 393.87 ± 121.08 pN/μm; [High]: 281.58 ± 60.84 pN/μm). Our data suggests that a low concentration of p-nitroblebb reduced NMIIB capacity in tension generation despite having punctate localizations at the cell cortex after treatment. This is similar to other contractile actomyosin machinery in which the absence of NMII turnover results in NMII aggregation and low tension generation (Thiyagarajan et al., 2021). Perturbation of NMII turnover by p-nitroblebb did not affect the actin cortex significantly as its functional thickness remained largely similar after treatment (Fig. 1 F). Taken together, we showed that an active NMII turnover at the cell cortex is required to maintain a contractile actomyosin architecture and cell surface tension during mitosis.
Interestingly, we found that there was a decrease of the cortical localization of active RhoA when cell surface tension has decreased for 3-fold in mitotic cells treated with a high concentration of p-nitroblebb, suggesting that RhoA could respond to the cortical tension generated by NMII during mitosis (Fig. 1 G). This contrasts with the upstream function of active RhoA in regulating NMII and implies a mechanosensitive activation of RhoA during mitosis. To test whether RhoA and NMIIB could be mechanically regulated, we compressed the mitotic cells with a soft elastic gel and a weight (Fig. 2 A). When compressive forces were exerted on the mitotic cells, the cell has enlarged its diameter, and the cell height has decreased from 22.89 ± 1.88 μm to 15.19 ± 1.19 μm (Figs. 2 B and C). Within 2 minutes after compression, we observed increased intensity of active RhoA at the mitotic cell cortex. In average, there was 1.54 ± 0.31-fold increase of cortical active RhoA intensity in individual cells (Fig. 2 D). Concomitantly, NMIIB increased its intensity at the mitotic cell cortex by 1.45 ± 0.29-fold compared to that of before compression (Figs. 2 E and F). Approximately 76.5% individual cells showed an increase of both active RhoA and NMIIB at the mitotic cell cortex after compression (Fig. S2), suggesting that compressive forces exerted on the mitotic cell cortex activate RhoA, which in turn activates NMIIB to generate higher cell surface tension to withstand the mechanical stress. Consistently, we found that compression in mitotic cells has slowed down the turnover of the cortical NMIIB, with an increase of the recovery half time of 27.75 ± 0.86 s as compared to that of in the uncompressed mitotic cells that was 18.85 ± 1.79 s (Figs. 2 G and H). Thus, compressive forces stabilize NMIIB at the mitotic cell cortex leading to a mechanosensitive accumulation of NMIIB. Taken together, our results showed that active RhoA and NMIIB demonstrate a mechanosensitive accumulation at the cell cortex to generate cell surface tension to withstand compressive forces during mitosis.
We next studied how NMII’s functions in cell surface tension generation and mechanoresponse are regulated and interrelated during mitosis. NMII is regulated by the phosphorylation of its regulatory light chain (RLC) that associates with the head domain and by the phosphorylation of the tail domain. Phosphorylation of RLC at the NMII head domain increases NMII actin-activated ATPase activity and promotes the formation of an unfolded active NMII to slide on actin filaments. Phosphorylation of the tail domain is involved in regulating myosin Il filament assembly (Heissler and Sellers, 2016). We examined the contribution of NMII head and tail domain phosphoregulations on the tension generation and the mechanoresponse of NMII during mitosis. Phosphorylation of RLC at the NMII head domain is driven by a RhoA / ROCK signaling pathway (Fig. 3 A) (Amano et al., 1996). When mitotic cells were chemically inhibited by the ROCK inhibitor Y-27632, NMIIB decreased for about 2-fold at the cell cortex while active RhoA did not show significant changes (Figs. 3 A and B). Consistently, there was about 20% decrease of cell surface tension from 965.34 ± 133.34 pN/μm in the control group to 770.04 ± 160.08 pN/μm in the Y-27632 treated group (Fig. 3 C). Of note, a decrease of cell surface tension by the concentration of Y-27632 used in this study was not sufficient to reduce active RhoA from the cell cortex.
When mitotic cells were compressed for about 8 minutes in the presence of Y-27632, there was a decrease of cortical NMIIB as compared to the control group that showed an increase of cortical NMIIB after compression (Figs. 3 D, E and F). Active RhoA showed a mechanosensitive accumulation at the mitotic cell cortex after compression in the presence and absence of Y-27632 (Figs. 3 D and E and F). Importantly, the lack of NMIIB mechanosensitive accumulation at the mitotic cell cortex after compression has further decreased the cell height after compression (Fig. 3 G; 13.62 ± 0.83 μm in DMSO treated compressed group and 11.08 ± 1.22 μm in Y-27632 treated compressed group), which is a readout of how much the cells could resist the compressive stress. Thus, mitotic cells were not able to generate sufficient cell surface tension to withstand the compressive force when RhoA / ROCK pathway was defective. Collectively, our results showed that inhibition of the RLC phosphorylation by ROCK has reduced NMII capacity in tension generation and in mechanoresponse. Thus, NMII head domain phosphoregulation by RhoA / ROCK signaling is required for both tension generation and mechanoresponse of NMII.
We next tested the role of NMIIB tail domain phosphorylation in regulating cell surface tension and NMIIB mechanoresponse during mitosis. Phosphorylation of NMIIB S1935 lowers the ability of NMIIB to assemble into myofilaments (Fig. 4 A) (Juanes-Garcia et al., 2015; Schiffhauer et al., 2019). We first generated three cell lines stably expressing wild type NMIIB, NMIIB containing a non-phosphorylatable alanine residue at S1935 (S1935A), and NMIIB containing a phosphomimetic aspartate residue at S1935 (S1935D), respectively. The endogenous NMIIB in these cell lines was depleted using CRISPR interference (CRISPRi) that switches off gene expression from its promoter and does not interfere with the transgene expression (Fig. S3 A and B). Cells expressing NMIIB S1935D showed a decrease in the cytoskeletal fraction compared to cells expressing wild type NMIIB or NMIIB S1935A, indicating less NMIIB myofilaments were assembled (Fig. 4 B and C).
Next, we examined the mitotic cell surface tension and the NMIIB mechanoresponse in cells expressing NMIIB tail phosphomutants. Interestingly, mitotic cells expressing NMIIB S1935D generated lower cell surface tension (1007.94 ± 130.21 pN/μm in wild type NMIIB and 677.67 ± 111.03 pN/μm in NMIIB S1935D) but were able to demonstrate mechanosensitive accumulation of NMIIB S1935D after compression (Figs. 4 D and E). Mitotic cells expressing NMIIB S1935A exhibited a slightly higher level of cell surface tension (1069.76 ± 158.13 pN/μm in NMIIB S1935A) and displayed a similar mechanoresponse as in cells expressing wild type NMIIB (Figs. 4 D and E). Thus, phosphorylation of NMIIB S1935 at the tail domain affected NMIIB’s function in cell surface tension generation but less on the mechanoresponse of NMIIB, in line with the previous study showing that the NMIIB tail domain fine-tuned the NMIIB mechanosensitive dynamics and the phosphomutants could accumulate with varying degree at the cell cortex in epithelial cells in response to mechanical stress (Schiffhauer et al., 2019). Importantly, the NMIIB S1935D generated sufficient mitotic cell surface tension to withstand the compressive force exerted on the cells as evidenced by the similar cell heights as in the cells expressing wild type NMIIB after compression (Fig. 4 F). Thus, the mechanosensitive accumulation of NMIIB compensates for the decreased ability to generate tension of the NMIIB phosphomimetic tail mutant. By accumulating more NMIIB at the cell cortex has allowed cells to generate sufficient cell surface tension during mitosis under compression.
Our study revealed that during mitosis, cell surface tension is maintained by a mechanosensitive RhoA / ROCK pathway that leads to the NMIIB accumulation at the cell cortex in a mechanically constrained environment. Mechanosensitive accumulation of NMIIB is a result of a slowdown of NMIIB turnover at the cell cortex, which we showed is important in maintaining a contractile actomyosin architecture that sustains the mitotic cell surface tension. We further show that tension generation and mechanoresponse are two mechanoproperties of NMIIB that can be uncoupled. The phosphoregulation by RhoA / ROCK pathway at the NMII head domain is involved in both mechanoproperties, whereas the phosphoregulation at the NMII tail domain is required mainly for the tension generation and for fine-tuning the mechanoresponse of NMIIB.
Mechanical activation of RhoA happens at the cell-cell adherens junctions and the cell-matrix focal adhesion to reinforce actomyosin contractility in response to mechanical stress (Acharya et al., 2018; Guilluy et al., 2011). This study, to our knowledge, is first to show mechanoactivation of RhoA at the cell cortex to sustain cell surface tension during mitosis. Mechanosensitve accumulation of active RhoA at the cell cortex during mitosis was not previously identified although inhibition of its downstream ROCK led to a decreased mechanoresponse of NMII (Schiffhauer et al., 2016). In this study, we used a widely used live-cell active RhoA biosensor (Koh et al., 2021) and focused only on mitotic cells using a microscopy with confocality and high temporal resolution. This has facilitated the detection of a tension-dependent activation of RhoA at the cell cortex when mitosis is mechanically challenged.
We showed that NMIIB’s function in tension generation and its mechanoresponse can be uncoupled by a phosphomimetic point mutation at S1935 of the tail domain, which perturbs assembly of NMIIB myofilaments. This perturbation lowers NMIIB’s ability to generate cell surface tension but does not decrease NMIIB’s ability to accumulate at the cell cortex in response to mechanical stress. The functional uncoupling enables cells to withstand the mechanical stress by compensating with more NMIIs at the cell cortex even the NMII is weaker in tension generation, enabling robust maintenance of cell surface tension during mitosis.
NMII shows a decrease of ADP-release kinetics when experiencing opposing loads (Kovacs et al., 2007). This increases the lifetime of actin-bound state of NMII and hence its duty ratio, which could lead to the mechanoaccumulation of NMII upon compression (Luo et al., 2013). This is in line with our FRAP analysis that NMIIB stabilizes at the cell cortex when cells experience compressive forces. We propose that mechanosensitive activation of RhoA / ROCK signaling pathway leads to RLC phosphorylation, which associates with an increase of the ATPase kinetics of myosin head domain to increase the flux of ATP hydrolysis into ADP (Amano et al., 1996). This together with the load-dependent decrease of ADP-release kinetics of NMII collectively increase the ADP-bound NMII that has a higher actin-binding lifetime, leading to its accumulation at the cell cortex in response to mechanical stress to maintain cell surface tension during mitosis.
Materials and methods
Cell lines and culture
MCF10A cell lines were cultured in DMEM F-12 (Shanghai BasalMedia) with Glutamax (Gibco), 5% horse serum (Biological Industries), 20 ng/ml EGF (Gibco), 0.5 mg/ml Hydrocortisone (MedChemExpress), 100 ng/ml Cholera toxin (Sigma), 10 μg/ml Insulin (Biological Industries), 1% Pen/Strep (Biological Industries) at 37°C with 5% CO2. The MCF10A cell line was a gift from Dr. Kuan Yoow Chan from Zhejiang University. The cells were authenticated by Sangon Biotech and were routinely checked for mycoplasma contamination in the lab using Myco-Blue Mycoplasma Detector (Vazyme). MCF10A cells expressing mRuby2-AHPH and mEmerald-NMIIB were prepared by lentivirus transduction and were enriched using the fluorescence-activated cell sorting (FACS). MCF10A cells expressing mEmerald-NMIIB WT, mEmerald-NMIIB S1935A, and mEmerald-NMIIB S1935D, respectively, were prepared by lentivirus transduction and selected with 2 μg/ml puromycin (Sangon Biotech) and were enriched using the FACS. To knockdown the endogenous NMIIB, the lentivirus expressing pCRISPRi0003 containing the guiding RNA (gRNA) targeting the NMIIB’s 5’ UTR region was transduced into the cells expressing NMIIB phosphomutants. Cells were selected with 6 μg/ml blasticidin (Solarbio) for 6 days and were continued to grow without the antibiotic for another 1 to 3 days before the experiments.
Drug treatments
The following drugs were used in the study: NMII inhibitor: p-nitro-blebbistatin (Cayman Chemical, 24171); Rho inhibitor: C3 exoenzyme (Cytoskeleton, CT04); ROCK inhibitor: Y-27632 (MedChemExpress, HY-10583); CDK1 inhibitor: RO-3306 (Selleckchem, S7747). DMSO was used to dissolve p-nitro-blebbistatin, Y-27632, RO-3306. C3 exoenzyme was dissolved in water. Details of concentrations used in the study are described in figure legends.
Lentiviral expression constructs
The following lentiviral transfer plasmids were used in the study: pTGL0427 containing mRuby2-AHPH in which AHPH was subcloned from an Addgene plasmid #68026 (a gift from Michael Glozter) and fused with mRuby2; pTGL0193 containing mEmerald-NMIIB, which was subcloned from an Addgene plasmid #54192 (a gift from Michael Davidson); pTGL0373 containing mEmerald-NMIIB S1935A; pTGL0374 containing mEmerald-NMIIB S1935D; pTGL0386 containing KRAB and dCas9 for CRISPRi (a gift from Jorge Ferrer, Addgene plasmid #118154); pCRISPRi0001 containing gRNA 5’ GTGCTAAAGGAGCCCGGCGG 3’ cloned into pTGL0386; pCRISPRi0002 containing gRNA 5’ GCTGGATCTGTGGTCGCGGC 3’ cloned into pTGL0386; pCRISPRi0003 containing gRNA 5’ GGACTGAGGCGCTGGATCTG 3’ cloned into pTGL0386. The 3rd generation lentiviral packaging system was used to prepare lentiviral particles. In brief, packaging plasmids pRSV-Rev, pMDLg/pRRE, pMD2.G (gifts from Didier Trono) were chemically transfected with the transfer plasmid into the 293Ta packaging cell line (Genecopoeia, LT008) using GeneTwin transfection reagent (Biomed, TG101). After 3 days, the culture medium was collected, filtered through a 0.45 μm filter, and then concentrated using the lentivirus concentration solution (Genomeditech, GM-040801) before adding to MCF10A cells.
Mitotic cell preparation
To increase the mitotic cell number, cells cultured on the imaging chambered coverglass were treated with 7.5 μM RO-3306 (Selleckchem) for 16 to 18 hours at 37 °C and were washed with the pre-warmed culture medium for 3 times to release cells into mitosis and added with 1 ml of fresh medium. Experiments were performed 40 minutes after the drug wash-off.
Elastic polyacrylamide gel preparation
The polyacrylamide gel used for cell compression was prepared as described in Matthews et al. and Le Berre et al. with modifications (Le Berre et al., 2014; Matthews et al., 2020). Briefly, 18 mm glass coverslips (Sangon, F518211) were treated with 10 μl Binding-silane (Sangon, C500226) for 10 minutes and then were rinsed with 100% ethanol and air-dried. To prepare soft elastic gels (2 kPa), 1 ml polyacrylamide gel solution was prepared by mixing 125 μl 40% w/v acrylamide (Sangon), 35 μl 2% bis-arcylamide (Sangon), 10 μl APS (10% in water, Sangon), and 830 μl water. After adding and mixing 1 μl TEMED (Sangon) into the gel solution, about 350 μl of the final gel solution was immediately transferred onto a flat glass slide and covered by the coverslip pre-treated with the Binding-silane. After polymerization for 20 minutes, the gel and the attached coverslip were gently removed from the glass slide using a surgical blade and were soaked in PBS for at least 2 hours and followed by the incubation in cell culture media for overnight. In experiments that involved drug treatment, the gels were incubated in media containing the drugs for overnight.
Micropipette aspiration (MPA)
The micropipette was prepared by first pulling a borosilicate glass capillary (Sutter Instrument, B100-58-10) using the P-97 Micropipette puller (Sutter Instrument). Then, the thin end of a borosilicate glass capillary was cut by a Microforge (Narishige) to a diameter of about 5.5 μm. The micropipette was bent to a desired degree by heating it over an alcohol lamp and was filled with phosphate-buffered saline (PBS) using the MicroFil (World Precision Instruments, MF34G-5). The micropipette was installed to a micropipette holder, which was connected to a syringe pump (Harvard apparatus) and a 25 ml serological plastic pipettes via a three-way valve. The micropipette was positioned under an inverted microscope (Olympus, IX73) equipped with Olympus 60x objective lens (N.A. 1.35, U Plan super apochromat), a micromanipulator system (Eppendorf, TransferMan 4r), and a CMOS high speed camera (Vision Research, Phantom 410L, 333.33 nm / pixel).
To perform micropipette aspiration, cells were cultured on a one-well chambered coverglass (Cellvis, C1-1.5H-N) for 24 hours and then were treated with 7.5 μM RO-3306 for 18 hours. Cells were washed 3 times with fresh media and were added with fresh media containing 20 mM HEPES. Cells were incubated for another 40 minutes to reach mitosis. When a mitotic cell was aspirated by the micropipette, the microscopy image was captured and analyzed using Fiji to obtain the pipette radius (Rp) and the cell radius (Rc). The cell length of about half a diameter of the pipette opening was aspirated in. The water pressure changes (ΔP) as displayed on the 25 ml serological plastic pipette was also recorded. Cell surface tension (T) was then calculated based on the Laplace equation, T = ΔP / 2 (1/Rp – 1/Rc).
Spinning-disk confocal microscopy for live-cell imaging
Spinning-disk confocal microscopy was equipped with a Nikon Eclipse Ti2-E inverted microscope, Nikon 60x oil-immersion objective lens (N.A. 1.40, Plan Apochromat Lambda), a spinning-disk system (Yokogawa Electric Corporation, CSU-W1), a Photometrics Prime 95B sCMOS camera, a Piezo Z stage (Physik Instrumente), and a live-cell stage top chamber with humidified CO2 (Okolab). Images were acquired using the Metamorph with a z-step size of 0.5 μm and a x-y plane resolution of 183.33 nm / pixel. The fluorophores were excited by laser lines at wavelengths of 488, 561, or 640 nm. For experiments involving cell compression, a z-stack of 30 μm was acquired before cells were compressed and a stack of 22 μm was acquired after cells were compressed. This ensured that the entire cell volume was covered during imaging. To quantitate the actin cortex thickness using the super-resolution microscopy in Figure 1 F, images were acquired using the LiveSR super-resolution module (Gataca) installed on the spinning-disk confocal microscope. For microscopy imaging not involving cell compression, the μ-Slide 8-well chambered coverglass (ibidi, 80826) was used for cell culture and imaging. Mitotic chromosomes were labeled with 0.2 μM SiR-DNA in live cells during imaging (Cytoskeleton, CY-SC007).
Mitotic cell compression and live-cell imaging
To compress cells, 8 × 104 cells were first seeded on a 20 mm two-well chambered cover glass (Cellvis, C2-1.5H-N). Forty-eight hours later, cells were treated with 7.5 μM RO-3306 to synchronize cells at the G2/M boundary for 16 to 18 hours. Prior to live-cell imaging with compression, synchronized cells were washed 3 times with the pre-warmed medium and added with 1 ml of fresh medium and incubated in the stage-top chamber attached to the spinning-disk microscopy for 40 minutes. When compressing the mitotic cells, the coverslip coated with the elastic gel was put over the cell layer and followed by a 5 g weight. Elastic gels and weights were pre-heated to 37 °C in the chamber. In experiments where cells were treated with DMSO or 20 μM Y-27632, 1 ml medium containing 40 μM Y-27632 or containing equivalent volume of DMSO was added to cells grown in 1 ml medium on the chambered coverglass to achieve a final concentration of 20 μM. The elastic gel that was pre-incubated with 20 μM Y-27632 for overnight was then put on top of the cell layer and followed by a 5 g weight. Cells at the same position were imaged before and after compression. The cell compression efficiency was validated by their decrease of cell heights.
Image analysis and processing
Microscopy images were analyzed using Fiji. To quantitate fluorescence intensity of mEmerald-NMIIB and mRuby2-AHPH in mitotic cells (Figs. 1 B, 1 G, S1 B, 3 A) and in mitotic cells for compression experiments (Figs. 2 D, 2 F, 3 E, 3 F, 4 E), image stacks containing 5 slices around the mid-plane were projected along the z-axis using the sum-intensity projection (Fiji/image/stacks/Z project). Background subtraction (Fiji/process/subtract background) was performed on projected images using the rolling ball radius of 20 pixels. Then, a segmented line with a line width of 4 (for mitotic cells) or 5 (for mitotic cells in compression experiments) was used to select the cortical fluorescent signals over the entire cell circumference. The mean fluorescence intensity of the selected line region from the same cells before and after treatment and/or compression was measured. The fold change was then calculated by dividing the intensity after treatment and/or compression to the intensity before treatment and/or compression. For time series in Fig. 1 B, the mean intensity of the cortical fluorescence was measured every 45 s. To quantitate the cell height in cell compression experiments, the number of slices from top to bottom of a cell were multiplied by 0.5 μm. To quantitate the actin cortex thickness, a line was drawn across the cortical actin to obtain a point spread function. The full-width-half-maximum of the point spread function was used to indicate the thickness.
FRAP
FRAP was performed using the above-mentioned spinning-disk confocal microscopy equipped with Nikon 100x oil-immersion objective lens (N.A. 1.45, Plan Apochromat Lambda) and iLAS2 FRAP system (Gataca). Images were acquired using the Metamorph software at 110 nm / pixel. The mEmerald-NMIIB fluorescence signal was photobleached by using a short pulse of high power 488 nm laser lines and imaged by using a low power 488 nm laser lines. A small bleaching rectangular region-of-interest (ROI) of about 2 × 10-μm was drawn on the cortical mEmerald-NMIIB in mitotic cells. Three frames of pre-bleached images were acquired with 1 second interval before photobleaching, followed by time-lapse imaging of the same cells with 2 seconds interval for 120 seconds. The bleaching ROIs were recorded to locate the regions of bleaching during analysis.
Fiji was used to quantitate the fluorescence intensity of FRAP images. Background subtraction (Fiji/process/subtract background) was performed on the images with the rolling ball radius of 20 pixels. A segmented line ROI (line width 5) was used to select the FRAP regions for quantification. The mean intensity of ROIs for pre-bleached images and after bleached images (Ibleached) was calculated for each time point. A mean pre-bleached intensity (Iprebleach) was derived from the average of three pre-bleached images. The ratio of Ibleached / Iprebleach at each time point was calculated and was subsequently deducted by the ratio of the first image after photobleaching to obtain ratiot. The ratiot of each time point was plotted as the recovery curve and fitted using a non-linear fitting function (one-phase association function constrained by Y0 = 0 and plateau < 1) in the Prism 9 software (GraphPad). The recovery time was derived from the half-time parameter calculated from the fitting. The mobile fraction was derived from the span parameter calculated from the fitting. The immobile fraction was calculated as a difference between 1 and the mobile fraction.
Cytoskeletal fractionation and immunoblotting
Cells were washed once with ice-cold PBS. To a single well of a 6-well cell culture plate, 200 μl of lysis buffer (50 mM PIPES, pH 6.8, 46 mM NaCl, 2.5 mM EGTA, 1 mM MgCl2, 0.5% Triton-X 100, protease inhibitor cocktail) was added. Lysates were harvested and centrifuged at 13000 g for 20 minutes at 4°C. The supernatant (soluble fraction) was separated from the pellet (cytoskeletal fraction). The pellet was then resuspended in a same volume of lysis buffer as the supernatant. Both fractions were added with 5 x Laemli sample buffer and boiled for 5 minutes. The pellet suspension was sonicated in a water bath sonicator for 1 minute with 20% power to improve solubility. Protein samples were separated in a pre-cast SDS-PAGE gel (GenScript, M00657) and blotted to a PVDF membrane (EMD Millipore, immobilon-P, IPVH00010). Blots were blocked with 5% skim milk for 1 hour at room temperature and were incubated at 4°C overnight with the NMIIB antibody (Proteintech, 19673-1-AP) or the β-actin antibody (HuaBio, ET1701-80). The secondary antibody used was Alexa Fluor Plus 800-conjugated goat anti-rabbit IgG secondary antibody (ThermoFisher, A32735). Blots were imaged using the LI-COR Odyssey CLx imaging system. The signal intensity of protein bands was calculated using the LI-COR Image Studio software. The cytoskeletal fraction was calculated by dividing the cytoskeletal fraction intensity by the sum intensity of cytoskeletal and soluble fractions. Measurements in Fig. 4 C were obtained from four proteins blots derived from four independent experiments. For the blot in Fig. S 3 A, primary antibodies used were NMIIB antibody (Proteintech, 19673-1-AP) and GAPDH antibody (Cell Signaling Technology, #5174). The secondary antibody used was Alexa Fluor Plus 800-conjugated goat anti-rabbit IgG secondary antibody (ThermoFisher, A32735).
Quantitative real-time PCR (qPCR) for checking endogenous NMIIB
To validate the knockdown of endogenous NMIIB in cells expressing the phosphomutants used in Figure 4 E and 4 F, total RNAs were prepared from cells using the FastPure Cell/Tissue Total RNA Isolation Kit (Vazyme, RC101-01) and were reverse transcribed into cDNA using the HiScript II Q RT SuperMix for qPCR (Vazyme, R223-01). The expression level of cDNAs was determined using the real-time PCR with the ChamQ Universal SYBR qPCR Master Mix (Vazyme, Q711-02). The reaction mix was assembled on the hard-shell PCR plate (Bio-Rad, HSP9655), sealed with the Microseal ‘B’ seal (Bio-Rad, MSB1001), and was performed in the CFX96 Touch Real-Time PCR Detection System (Bio-RAD, C1000). Primers used in the qPCR: GAPDH, 5’ CAGGAGGCATTGCTGATGAT 3’ and 5’ GAAGGCTGGGGCTCATTT 3’; NMIIB, 5’ CCTCATGCTGACCTTGCAAA 3’ and 5’ GGACACAAAACCAATATTCCCATT 3’. The primer pair for NMIIB targets the 3’ UTR of NMIIB gene, allowing checking of endogenous NMIIB expression. The CT value for GAPDH was used for normalization to obtain the relative expression level.
Statistical analysis
Statistical analysis was performed using Prism 9 (GraphPad). Datasets were analyzed by Student t-test. If datasets were not normally distributed, they were analyzed using the Mann-Whitney test. All graphs were plotted using Prism 9 (GraphPad).
Author contributions
J.D. and C.W. designed, performed experiments, and analyzed data. S.D. prepared elastic gels used in the study. Q.W and X.G provided the devices and technical expertise for micropipette aspiration. T.G.C conceived the study, designed experiments, supervised the study, and prepared the manuscript with inputs from all authors.
Competing interests
The authors declare no competing interests.
Acknowledgement
This study was funded by the Zhejiang University International Campus start-up grant (130000-541902/016) given to T.G.C. We thank Dr. Di Chen for sharing the CRISPRi vector with us. We thank Dr. Mikael Bjorklund for sharing the 293Ta packaging cell line with us. We thank Dr. Wanzhong Ge and Dr. Junqi Huang for their comments on the manuscript. Special thanks to Dr. Fang Kong from the Nanyang Technological University, Singapore for designing the imaging chamber adaptor used in the microscopy imaging for cell compression. Q.W and X.G would like to acknowledge the support from the National Natural Science Foundation of China (12072198).
Footnotes
↵* Co-first authors