# Cell cycle phases, their effect on cell mechanical properties and the impact on Candida-host cell interaction

Easter Ndlovu, Zinnat Shahina, Tanya E S Dahms, Etienne Dague

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Easter Ndlovu, Zinnat Shahina, Tanya E S Dahms, Etienne Dague. Cell cycle phases, their effect on cell mechanical properties and the impact on Candida-host cell interaction. 2025. hal-05381178

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1 Cell cycle phases, their effect on cell mechanical properties
2 and the impact on Candida-host cell interaction

3

4 Easter Ndlovu¹, Zinnat Shahina¹, Tanya E. S. Dahms*¹, Etienne Dague*²

5 ¹Department of Chemistry and Biochemistry University of Regina, 3737 Wascana

6 Parkway, Regina, SK, S4S 0A2, Canada

7 ²National Centre for Scientific Research, Laboratory for Analysis and Architecture of

8 Systems (LAAS). 7 avenue du Colonel Roche, BP 54200, 31031, Toulouse, cedex 4,

9 France

10 *Co-corresponding authors: edague@laas.fr (ED), tanya.dahms@uregina.ca (TESD)

11 Abstract

12 Cell mechanics is essential in many biological phenomena such as cell division, and
13 migration. Further, cell mechanobiological measurements can distinguish between
14 healthy and diseases cells, thus, investigations of cell mechanics have led to the
15 development of tools to study the elasticity and the viscosity of cells. Cell mechanics can
16 be affected by factors such as cell morphology, and cytoskeletal remodelling and cell
17 intrinsic factors and these are important in understanding disease progression. Here we
18 use an atomic force microscopy (AFM)- microrheology with a colloidal probe for a
19 dynamic mechanical analysis of host cell elasticity and viscosity at 6 frequencies ranging
20 from 1 Hz to 200 Hz. Epithelial cells exhibit a more “liquid-like” behavior as the frequency
21 increases, whereas cancerous cells transition into this viscous, fluid-like state at lower
22 frequencies (48 and 63 Hz) compared to normal cells (92 Hz).

23 Cell mechanical measurements inherently exhibit heterogeneity due to physical factors
24 such as cell shape and the position of the probe on the cell surface. In addition to this
25 physical variability, biological parameters — notably the cell cycle phases — also
26 contribute to mechanical heterogeneity. In this study, we specifically investigated the
27 influence of cell cycle phases on cell mechanical properties. Using chemically
synchronized normal and cancerous cells in different phases of the cell cycle, shows that
the actin cytoskeleton undergoes rapid reorganization as the cell cycle progresses.
Results show that as actin becomes disorganized the elastic moduli decreases, the loss
tangent is larger coupled with a lower phase shift frequency. Cells in the G₁ and S phase
had the lowest elastic moduli (G') meaning they were softer than cells in G₂/M phase.
During disease onset the pathogen adheres and invades the host, a process which leads
to cytoskeleton arrangement and thus, changes in cell elasticity and thus, to understand
the impact of the cell cycle on host invasion we probed the interaction of C. albicans with
HeLa, HCT 116 and HaCaT cells using AFM in single cell force spectroscopy mode. There
was a significant increase in force of interaction during the S phase which could be
attributed to the disorganized cytoskeleton. This shows the importance of cytoskeletal
organization and cell cycle phase in cell mechanical properties and pathogen-host
interaction.

Keywords

Microrheology, Atomic force microscopy, elasticity, viscosity, cell cycle phases, adhesion,
cytoskeleton
## Introduction

There is a resurged interest in studying cell and tissue mechanics, as they are now known to govern cell behaviour and biological function.¹⁻³ Most studies have examined cell elasticity or stiffness, representing the ability of cells to return to their original state after deformation in response to an external stress.² Elasticity impacts cellular events such as adhesion and differentiation, and thus, aberrations in the elastic moduli resulting from various physiological and pathological processes can be associated with disease pathogenesis and progression.⁴

Cell mechanics are closely linked to the cytoskeletal structure, which enables the cell to maintain its viscoelasticity and fluidity.⁴,⁵ The cytoskeleton is a three-dimensional network of protein filaments, comprised mainly of three polymers namely, actin filaments (F-actin), intermediate filaments (IF) and microtubules (MT). These protein filaments serve as mechanical elements, differing in their magnitude of stiffness, with microtubules stiffer than the other two polymers.⁵⁻⁷ For instance a complex of F-actin with myosin (stress fibers) is stiffer than the surrounding cytoplasm, and therefore, dictate cell shape by resisting stress, thereby controlling cellular processes.⁶

Living cells dynamically change their internal arrangement during the progression of the cell cycle, affecting the intrinsic ability of cells to uptake drugs for disease treatment.⁸⁻¹⁰ Progression of the cell cycle is a tightly regulated process, with cells induced to divide as part of the physical or homeostatic process.¹¹⁻¹³ The cell cycle is depicted by phases associated with DNA synthesis (S phase) and mitosis (M phase) which are separated by two growth phases, G₁ and G₂.¹⁴,¹⁵ Following the mitotic phase, during the G₁ phase, cells begin growing, which helps maintain a stable size for the entire duration of the cycle.¹³
During the cell cycle the cytoskeleton is reorganized as the cells change shape, thus, affecting cellular mechanics.<sup>16,17</sup>

To access mechanical forces that regulate the various functional processes and interaction between single cells during the cell cycle, a wide variety of techniques have been developed which include traction force microscopy (TFM) and atomic force microscopy (AFM) and FluidFM<sup>18-21</sup>. TFM measures endogenous forces by monitoring, using fluorescent beads, the surface tractions imparted by cells on a flexible substrate. Fluorescent beads are mixed into the substrate, and their displacement can be tracked using an optical microscope<sup>18,22,23</sup>. Although traction force microscopy does spatial measurements over a wide range of force scales, it does not allow for the intracellular measurement of cell viscosity and stiffness<sup>24</sup>. Over the past years, atomic force microscopy (AFM) has advanced from just being used for surface topography imaging of cells<sup>25</sup> to one capable of characterizing a range of biochemical, properties at the subnanometer resolution.<sup>26-28</sup>

FluidFM, connected to AFM allows for the measurement of forces of interaction without the need of chemically attaching cells onto an AFM cantilever but uses suction to attach the cell to the aperture of the cantilever connected to a fluidic channel<sup>29</sup>. The development of robotized FluidFM with an automatic controlled XY-stage allows for high throughput single cell force spectroscopy (SCFS) measurements<sup>30</sup>. Although, these techniques has been used to measure the effect of the cell cycle on cell adhesion and evaluating mechanical properties of cells, TFM does not allow for the intracellular measurement of cell viscosity and elasticity while studies using robotic FluidFM and AFM measured only the Young's modulus (elasticity)<sup>8,21,31</sup>. Furthermore, the associated elastic properties are
evaluated by applying the Hertz model to the approach portion of the force-distance curve, which assumes a purely elastic sample. In reality, most biological samples are viscoelastic, and therefore, microrheology has recently been used to probe both the viscous and elastic properties of cells over a broad range of frequencies.³²,³³

Since cell mechanical properties are associated with biological function, and defects in stiffness and viscosity has been related to several malfunctions and diseases, an in-depth understanding of these parameters is essential.³⁴,³⁵ In our previous work we reported that disruption of the actin cytoskeleton, both directly or through altered cell shape, led to a change in Candida adhesion to host cells.³⁶ Further, the cytoskeletal organization differs between normal and aberrant cells, the latter having an irregular network with reduces cell stiffness.³⁷ Considering that cell shape and cytoskeletal structure changes during cell cycle progression, we hypothesized that the biomechanical properties would change over the different phases of the cell cycle. Microrheology allows a systematic characterization of phase shifts between solid- and liquid-like states to better understand their contribution to disease etiology. Such mechanical properties can serve as disease biomarkers, for example cancerous versus non-cancerous human cells to shed light on their susceptibility to pathogens. We demonstrate that cell mechanics differ over the cell cycle, with phase shifts at lower dynamic frequencies in cancerous cells which are more elastic cells as compared to normal cells which are stiffer cells.

124
125
126
127
128 Results

129 Microrheology measurements reveal differential cellular mechanics

130 Microrheology measurements examined the elastic and viscous modulus of two
131 cancerous (HeLa and HCT 116) and one normal cell line (HaCaT) over a range of six
132 different frequencies (Figure 1a-d, S1). Figure 1 a-d histograms show a random
133 percentage frequency distribution of G' (elastic modulus) and G" (viscous modulus) at 1
134 Hz and 200 Hz. The histograms and whisker plots (Figure 1a-d inserts) show distributions,
135 and the latter also the means, highlighting the results from the frequency distribution plots
136 (Figure 1c-d). G' at 1 Hz was the lowest for HeLa cells at 128 ± 56 Pa, compared to 354
137 ± 191 Pa for HCT 116 cells and 374 ± 240 Pa for HaCaT cells. On the contrary, the G"
138 was 41 ± 18, 119 ± 58, 112 ± 66 Pa for HeLa, HCT 116 and HaCaT cells, respectively.
139 The same trend was observed at 200 Hz (Figure 1c,d) and other frequencies (Figure
140 S1a). For all cell lines, at 1 Hz G" ranged between 1 and 400 Pa, while G' were as high
141 as 1400 Pa, whereas at 200 Hz we see the opposite trend (Figure 1a-d).
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170 Individual cells, randomly selected and further analyzed, show heterogeneity within a cell line, for which transition frequencies range from 20Hz to above 100 Hz (S1b). These transition frequencies were grouped as ≤ 50, 51 – 100 and > 100 Hz, showing HeLa cells having more cells transitioning at frequencies below 50 Hz and HaCaT having the most above 100 Hz (Figure 1g).

175 Cytoskeletal arrangement, cell topography and properties vary with cell cycle phase.

176 Abidine et al 2018 report that cell rheology measurements depend on the tip location on the cell (nucleus, perinuclear region, and edge)³³ and other factors such as cell shape may contribute to heterogeneity, thus leading to the analysis of cells in different cell cycle phases. HeLa, HCT 116 and HaCaT cells chemically synchronized at the G₁, S and G₂/M phase had different cytoskeletal arrangements (Figure 2a) and cell surface topography (Figure 2b), height, surface area and roughness (Figure 2c). Cells in G₁ were the smallest having multiple radiating branches indicative of cellular growth following mitosis, those in S-phase were enlarged with actin spread throughout the cytoplasm as expected for cells that have grown in preparation for division, and those in G₂/M phase were spherical with actin condensed around the nucleus as expected for cells preparing or mitosis (Figure 2a-b).
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189

**Figure 2.** Analysis of parameters for chemically synchronized epithelial cells. HaCaT, HeLa and HCT 116 cells were chemically arrested at G₁, S and G₂/M phase. Synchronized cells were either (a) stained with phalloidin and imaged by epifluorescence (λ<sub>ex</sub> = 495 nm, λ<sub>em</sub> = 518 nm) for (b) or fixed and imaged by AFM in QI mode. (c-e) Average height, roughness and estimated cell surface area of cells at different phases of the cell cycle was calculated from AFM-QI topography images. Column graphs show data and error bars represented as the ±SEM. Statistical significance was tested by one-way Anova and Bonferroni post-test, where ns indicates no statistical significance and asterisks represent statistical significance: ****(p< 0.0001), ***(p < 0.001), **(p < 0.01), *(p < 0.05), ns, not significant (p > 0.05). Scale bars are a) 50 µm and b) 10 µm.
200 HeLa cells in G₂/M were the tallest (8.1 µm), but with no statistical difference observed
201 for the height of G₁ and S phase cells (Figure 2c). HCT 116 cells were also largest in G₂/M
202 (height = 8.6 µm), and smallest in G₁ (7.2 µm). However, there was no significant
difference in height for HaCaT as a function of cell cycle phase (Figure 2c). For all three
203 cell lines (Figure 2d), G₁ cells had the lowest surface area (~300 µm²), while those in S
204 phase had the largest surface area (~1250 µm² for HaCaT, 1132 µm² for HeLa and 558
205 µm² for HCT cells. Lastly, the cytoplasmic area was more granulated and rougher
206 compared to the nuclear area for all cell lines at all phases. S phase cells had the
207 smoothest surfaces, while G₂/M phase cell surfaces were the roughest (Figure 2e). There
208 was an inverse relationship between cell surface area and cell roughness (Figure 2d,e).
209 Taken together, the results indicate that cell morphology and ultrastructure vary with cell
210 cycle phase.
211

212 Elasticity and viscosity vary with cell cycle phase

213 Based on the transition pattern heterogeneity we observed for randomly selected cells in
214 a given population and the well-established idea that cell geometry determines
215 mechanical properties, we expected that the cell cycle phase would play a role in cellular
216 rheological properties. We therefore, compared the elastic moduli and the viscous moduli
217 of chemically arrested cells using a spherical bead. HeLa cells had the lowest elasticity in
218 G₁ at frequencies below 50 Hz (Figure 3a,S2) while G₂/M cells had the greatest elasticity
219 at all frequencies (Figure 3a,S2). At all frequencies, the HCT 116 cells in G₁ were more
220 elastic than those at G₂/M and S, with those at G₂/M being stiffer, especially at 200 Hz
221 (Figure 3a,S2). On the other hand, HaCaT cells in S phase were more elastic than those
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G' G"

G₁

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S

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G₂/M

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**Figure 3.** Elasticity and viscosity assessment of synchronized cells. (a,b) Microrheology measurements at 1Hz showing the elastic and viscous moduli at G₁, S and G₂/M phases. (c-e) Relationship between G' and G" reveals the transition frequency from an elastic state to a more viscous state as a function of oscillation frequency at G₁, S and G₂/M cell cycle phases. (f) The ratio of G" and G', shows loss tangent as a function of frequency at G₁, S and G₂/M. Plots are the mean and the error bars show the ±SEM of at least 10 cells.

in G₁ (Figure 3a,S2). Viscous moduli were higher for HeLa (46 ± 9 to 850 ± 271) and HaCaT (76 ± 26 to 1375 ± 402) in G₁ from 1 to 200 Hz, while HCT 116 in S phase had the highest viscosity (57 ± 23 to 988 ± 298). Furthermore, cells in G₂/M had the lowest viscosity for all 3 cells lines at every oscillation frequency (Figure 3b, S2). Further, there was an inverse relationship between elasticity and viscosity, with cells in G₁ having the lowest G' value and those in G₂/M the highest, while G" was highest for cells in G₁ and lowest for cells in G₂/M (Figure 3a,b,S2).

To further understand the behaviour of these cells, we examined their fluidity by comparing G' and G" as for non-synchronized cells (Figure 1e), revealing behaviour at the different cell cycle phases. Transition from a more solid phase to a more liquid one was observed at lower frequencies for cells in G₁ (Figure 3c-e), corroborating their lowest elasticity (Figure 3a,S2). As expected, the transition period for cells in S phase was observed at frequencies below 50 Hz (Figure 3c-e), as cells in S phase closely resemble those in G₁ (Figure 3a,S2). Lastly, cells in G₂/M transitioned at the highest frequencies, indicating the greatest stiffness (Figure 3c-e). Loss tangent showed that indeed, for HeLa, HCT 116 and HaCaT, G₂/M had the most solid like nature for the longest period following applied strain, becoming greater than 1 at higher frequencies compared to cells in S or G₁ (Figure 3f). Interestingly, phase shift frequencies were almost the same within one cell line (Figure 3f).
248 Adhesion force of *Candida* to host cells depends on cell cycle phase
249 In our previous work, we reported that cell mechanical properties associated with the
cytoskeleton impact how the fungi *C. albicans* adheres to host cells, showing that *Candida*
250 adhere better to malignant cells or those having disrupted actin filaments, as compared
251 to normal cells.36 Given that cells at different cell cycle phases have different cytoskeletal
252 arrangements, sizes, textures and mechanical properties, we hypothesized that *Candida*
253 adhesion would vary with cell cycle phase.
254

255 Firstly, using the colony counting adhesion assay, the percentage of adhered *C. albicans*
256 as a function of colony forming units were statistically identical at all stages of the cell
257 cycle, for all 3 cell lines after 1h of co-incubation to synchronized HaCaT, HeLa and HCT
258 116 cells (Figure 4a). SCFS measurements were used to define the interaction between
259 the host and the pathogen as a function of maximal adhesion force by attaching *C.
260 albicans* to a tipless cantilever using to probe HeLa, HCT 116 and HaCaT at various
261 phases of the cell cycle. We observe a significant difference in maximal adhesion force
262 to cells in the S phase versus those in the G₂/M phase (Figure 4b), and in some cases
263 (HaCaT) differences in adhesion to cells in G₁ versus those in G₂/M (Figure 4b).

264 These results not only provide evidence that human cell cycle phase impacts candidal
265 adhesion but also underscore the importance of directly quantifying adhesion by AFM-
266 SCFS. A closer look at the retraction curves (Figure 4c) shows the presence of tethers,
267 associated with stretching of a cell membrane tube prior to unbinding, as well as multiple
268 jumps resembling step-like unbinding events, with a rise in the adhesion force before the
269 unbinding of a single adhesive unit.38,39 Force curves of cells in S phase, followed by
270 those in G₁, had more jumps, indicating the presence of additional molecular interactions.
<figure><img src="image_27.png" /></figure>

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<figure><img src="image_36.png" /><figcaption>272 **Figure 4.** *C. albicans* binds with varied force to cells at different phases of the cell cycle. (a)</figcaption></figure>

273 Colony formation from yeast cells co-incubated with synchronized host cells to measure bulk

274 adherence for 1 h, expressed as percent adhesion from the total number of yeast cells. (b) AFM-

275 SCFS adhesion interaction measurements, using *C. albicans* as a probe attached to a tipless

276 AFM cantilever. (c) Typical force-distance curves obtained from the interaction between C.
albicans and HeLa, HCT 116 and HaCaT cells. (d) Histograms of the adhesion work for the interaction. Results are representative of at least 10 cells with 16 force curves recorded per cell and error bars show the ±SEM. Statistical significance was tested by one-way Anova and Bonferroni post-test, where ns indicates no statistical significance and * represents, p < 0.05).

between the host and pathogen surfaces (Figure 4c). To further probe these interactions, the work of adhesion was calculated as the area between the retract force curve and the baseline of the force curve, showing that for HeLa cells in S phase, values ranged up to 60 000 x 10⁻¹⁸ joule (J), with 34 000 x 10¹⁸ J and 50 000 x 10⁻¹⁸ J for G₁ and G₂/M, respectively (Figure 4d). The adhesion work was proportional to the maximal adhesion force, with lower energy required to break bonds for the lowest recorded adhesion.

## Discussion

During most biological processes, deformation of the cytoskeletal structure leads to changes in gene expression and cell function, and thus, is strongly linked with the onset and progression of human disease.³⁷,⁴⁰ Many studies report that cancerous cells are softer compared to their non-cancerous counterparts, with a few exceptions.³⁷,⁴¹⁻⁴⁵ In cancerous cells there is dysregulation of proteins, including cytoskeletal ones, causing a rearrangement of cell dynamic structures.⁴⁶

As expected, the cancerous HeLa and HCT 116 cells were softer (lower G') as compared to the non-cancerous (HaCaT) human skin cell line (Figure 1), consistent with previous studies showing that prostate cancer DU145 cells are softer (lower G') compared to the non-cancerous Vero cells.⁴⁷ Indeed, cancerous liver and connective tissue (HUH-7, HT 1080) cells have elastic moduli of 0.5 kPa compared to 2 kPa for human dermal fibroblasts.⁴⁸ The actin cytoskeleton in breast and ovarian cancer cells is less dense with
fewer stress fibres compared to the non-cancerous MCF-10A.⁴² Furthermore, the nuclei of cancer cells are larger in volume with altered expression of nuclear envelope proteins, such as the lamins which play a role in cytoskeletal structure reorganization, likely contributing to decreased stiffness.⁴⁹

Since the cytoskeleton consists of a network of filaments with high water content, cells can store (solid-like state) and dissipate (liquid-like) energy according to the speed at which the stimulus is applied (Figure 1e,f).³² In this study, G' dominates over G'' at lower frequencies in a more solid-like phase, whereas at higher frequencies cells have a more viscous-like behaviour (Figure 1e,S1,3c-d), consistent prior studies showing cells to be more solid-like when stimulated at lower frequencies.⁴³,⁵⁰⁻⁵² Rother et al (2014) characterized the mechanical behaviour of 9 different cell lines (normal and cancerous) from different organs, including the breast, kidney and lungs, showing that cancer cell lines MCF-7, CaKi-1 and MDA-MB-231 had the lowest G' values at all frequencies⁵⁰, consistent with our results. Furthermore, it has been reported that Hela cells become softer and more liquid-like with an increase in frequency, with perturbation of the cytoskeleton leading to a cross from elastic state to a viscous state at lower frequencies as compared to the unperturbed HeLa cells.⁵¹,⁵²

The loss tangent is larger in cells that are more easily deformed (fluid like) (Figure 1f,3f), which was expected as these cells lack an intensive F-actin network and stress fiber formation, which is directly related to cell metastatic potential.³²,⁵⁰ Intuitively, increased cell stiffness correlates with more solid-like behaviour and *vice versa*. Thus, high internal contractile stress generated during the interaction of actin filaments and myosin motors
is related to a more solid like behaviour and vice versa.⁴ It has been hypothesized that under stress conditions cellular resources are insufficient to maintain a normal mechanical state, which requires energy, and thus cells become softer.⁴,⁵³ This explains our results (Figure 1e,3c-e) in which cells transition from a solid to a more liquid-like state.

In cancer cells, which are disrupted, this transition happens more easily, and as the frequency increases, they are unable to maintain their mechanical state. Further, a comparison of the transition frequency for cells having different degrees of invasiveness show that more invasive J82 cancer cells have a lower transition frequency, around 40 Hz.³³ The rearrangement of cytoskeletal networks during cell cycle progression are proposed to impact the intrinsic cell mechanical properties.⁵⁴ Indeed, HeLa cells changed their size and elasticity as the cell cycle progressed, with S phase cells being the softest⁸, while M phase cells have been reported to have a larger spring coefficient with corresponds to greater stiffness as compared to cells in other phases of the cell cycle³¹ consistent with our results (Figure 3). A recent study reported G' and G'' values of cell in S/G₂ phase were 40% and 30% larger than those in G₁ phase, similar to this study (Figure S2). Cell spreading, important for metastasis, is associated with a complex cytoskeletal reorganization and a larger cell surface area⁵⁴, related to changes in cell stiffness and permeability.⁸,⁵⁵ The height and surface area of HeLa cells was similar to that reported by Fan et al. (2017), and as cells spread out in the G₁ and S phase they became more elastic (Figure 3a). Cells were smoother (Figure 2b) in nuclear versus cytoplasmic regions, and more granulated and rougher in G₂/M, likely attributed to a loosely packed cytoskeleton. Further, normal cells were smoother than cancerous ones in both G₁ and S phases (Figure 2e), consistent with that of normal breast tissue and increased cell
roughness with cancer stage associated with a decrease in the SMAR1 tumor suppressor protein, which arrests cells at G₂/M.⁵⁶ Indeed, cancer drugs like cisplatin and colchine that interfere with DNA and RNA synthesis, disrupt microtubule, microfilament and intermediate filament function during mitosis, accompanied by increased cell roughness as observed with nocodazole.⁵⁷

Mitosis is accompanied by changes in morphology and mechanical properties, as demonstrated by increased intracellular pressure, cell surface tension and cortical stiffness during mitosis.⁵⁸ A 4-fold increase in the elasticity of Drosophila cells in the early stage of mitosis, further increased during division⁵⁹ and elastic moduli of MCF10A normal breast epithelial cells changed between interphase and mitosis.⁶⁰ The ability of mitotic cells to resist deformation, remaining in a more solid phase at higher frequencies (Figure 3c-e), can be attributed to acto-myosin. Cortical architecture, contractility and protein composition, which controls the cell shape, most likely alter mechanical properties between phases. Cells in the mitotic phase require increased cell stiffness for cell rounding, which is necessary for proper chromosomal segregation.⁵⁸

The cytoskeletal filaments, together with cell surface receptors and extracellular matrix proteins, are not only important for regulating cell morphology, but contribute to cell elongation and adhesion.⁶¹ B16-F10 melanoma cells have lower stiffness, which is inversely proportional to adhesion force to a silicon nitride probe.⁵⁷ Differences in cell elasticity from cytoskeletal rearrangement alters the distribution of adhesins⁵⁴, with an increase in candidal adhesion to host cells upon disruption of the actin filaments.³⁶ The variation in actin arrangement with cell cycle phase, known to regulate macromolecules
involved in cell-cell adhesion⁶², likely explains differences in the adhesion between C. albicans and epithelial cells at various cell cycle phases (Figure 4b). In fact, actin rearrangement, including filament assembly and polymerization, may account for the incorporation of Candida into the host, leading to disease.⁶³

## Conclusion

The mechanical properties of cells are governed by many factors, as such techniques such as AFM-microrheology have been helpful in understanding cell rheological dynamics. Of note, changes in cell elasticity have been implicated in the pathogenesis of many human diseases including cancer. The correlation between cell elasticity and cytoskeletal dynamics is critical towards understanding how cell mechanics can be used as a predictor of disease state. We confirm that quantitative measurement of cell rheological properties indeed, shows that, cancerous cells are distinct from normal healthy cells, with the former having a larger loss tangent accompanied by a lower phase shift frequency meaning they are more elastic, thus, softer. Further, the cell cycle is implicated to cell mechanical properties, noting that the phase does contribute to variations in cell elasticity. We concluded that this variation is attributed by alterations in cytoskeletal organization, with G₁ and S phase cells having the lowest elastic moduli and most dispersed actin arrangement. For the first time we report the role of cell cycle phase on the adhesion strength of pathogens to host cells. The adhesion of the pathogen C. albicans is modified in different phases of the cell cycle, with stronger adhesion at phases with have a lower elastic modulus. From this we deduce that if the host cells are a phase where they are softer with a loosely arranged actin organisation, it increases the chances
391 of a stronger adhesion of the host. We further, show the importance of using the AFM to
392 measure the binding forces as we saw a difference in the behavior of the pathogen when
393 forces were measured. Our results, yield a foundation to link mechanical and functional
394 conditions, of particular showing the role cell mechanobiology plays in disease
395 progression.

396 **Materials and methods**

397 Fungal cell culture

398 C. albicans (ABC Platform® Bugs Bank, Nancy, France) and C. glabrata KUE10082 ((Prof
399 Miguel Teixeira) strains were stored at –80 °C in 50 % glycerol and revived by streaking
400 onto yeast extract-peptone-dextrose (YPD) agar plates containing 1 % Bacto™-yeast
401 extract, 2 % Bacto™ peptone, 2 % glucose and Bacto™ agar. Plates were then incubated
402 at 30 °C for at least 24 h to grow single colonies. For experiments, single colonies were
403 inoculated into YPD broth and incubated at 30 °C with shaking at 200 rpm for 18 h.

404

405 Human cell culture

406 The human skin keratinocytes, HaCaT (American Type Culture Collection (ATCC), CRL-
407 2404), human cervical cancer epithelial cells, HeLa (ATCC, CCL-2) and human colorectal
408 cancer epithelial cells, HCT 116 (ATCC, CCL-247), were maintained in Dulbecco's
409 Modified Eagle's Medium (DMEM) (Gibco, ThermoFisher Scientific) containing 4.5 g/ml
410 glucose, phenol-red, supplemented with 10 % (v/v) fetal calf serum (Avantorsciences), 1
411 % (v/v) of 100 IU/ml penicillin and 100 µg/ml streptomycin (penstrep) (Gibco, Life
412 Technologies). All cell lines were cultured in 25 or 75 cm² cell culture flasks (Sarstedt,
413 Germany) at 37 °C with a relative humidity of 90 % and 5 % CO₂. Cells were passaged
414 when they had reached 70 – 80 % confluency and detached from the flask by the addition
415 of TrypLE™ express enzyme (Gibco, ThermoFisher Scientific) for 10 min. The proteolytic
416 activity was inhibited by adding DMEM. Thereafter, cells were pelleted by centrifugation
417 at 200 ×g for 5 min, resuspended in media and then used in subsequent experiments.

418 Cell synchronization

419 Human cells, cultured as described above, were seeded onto clean coverslips (18 × 18
420 mm) and grown until they had reached 70 % confluency. For G1 phase synchronization,
421 cells were treated with either a final concentration of 40 µM (HeLa, HCT) or 60 µM (HaCat)
422 Iovastatin (Sigma-Aldrich, USA) and then incubated for 24 h.⁶⁴ Cells were synchronized
423 in the S-phase by blocking the G1/S phase transition using a final concentration of 2 mM
424 hydroxyurea (Sigma-Aldrich, USA) for 18 h.⁶⁵ For the mitotic phase, cells were
425 synchronized by arresting the cell cycle at the G2/M phase transition by adding
426 nocodazole (Sigma-Aldrich, USA) to final concentration of 40 ng/ml and then followed by
427 a 12 - 14 h incubation.⁶⁶

428 Immunofluorescence

429 Epifluorescence microscopy was used to assess synchronization efficiency. Following cell
430 cycle arrest at either the G1, S or G2/M phase, media was aspirated, and cells washed
431 thrice with phosphate buffered saline (PBS; 10 mM sodium phosphate, 1.8 mM potassium
432 phosphate, 138 mM sodium chloride, 2.7 mM potassium chloride, pH 7.4). Cells were
433 then permeabilized with 0.2 % Triton X-100 for 3 min, washed again with PBS, fixed with
434 4 % paraformaldehyde and then stained with Phalloidin-iFluor 488 (1:1000 dilution;
435 Abcam; $\lambda_{ex} = 495$, $\lambda_{em} = 518$) for 20 min, followed by 3 rinses with 100 µL PBS. Coverslips 436 were then mounted on clean microscope slides, sealed with nail polish, and imaged on a 437 Zeiss Axio Observer Z1 inverted epifluorescence microscope (Zeiss, Oberkochen, 438 Germany) at 20 $\times$. 439

440 Atomic force microscopy

441 To analyse cell morphology, samples seeded on coverslips in air at room temperature 442 were imaged by AFM (Bruker, USA) mounted on an Axio Observer inverted fluorescence 443 microscope (Carl Zeiss, Germany) using the dendritic silicon nitride, MLCT probes 444 (MLCT-A; Bruker, USA) in quantitative imaging (QI) mode. The spring constant of the 445 cantilevers was calibrated prior to data collection using the thermal noise method$^{67}$, 446 typically having a sensitivity of approximately 37 to 39 nm/V and a spring constant (0.051 447 to 0.058 N/m), consistent with the nominal value (0.07 N/m; Bruker, USA). The cantilever 448 was brought into contact with the HaCaT, HeLa and HCT human cells, using an applied 449 force of 1 nN, a constant approach-retract speed of 100 µm/s and at least 10 images were 450 obtained for each condition per biological replicate. Data were analyzed using the JPK 451 data processing software (Bruker, USA).

452 Atomic force-microrheology

453 AFM-based microrheology was used to analyse cell viscoelastic properties. HaCaT, HeLa 454 and HCT cells were seeded in 10 mm dishes and grown overnight in DMEM. AFM- 455 microrheology was performed using a colloidal probe which was made by attaching a 5 456 µm diameter silica bead (Bangs Laboratories) to NPO10 cantilevers coated with a thin 457 layer of UV-curable glue (NOA 63, Norland Edmund Optics).$^{68}$ To maintain human cell
growth conditions, the microscope was equipped with a heated petri stage (Bruker, JPK 458 BIOAFM) to keep the sample at 37 °C and was connected to a tube delivering CO₂. Cells 459 were indented at 1 nN and the cell microrheological features were determined by 460 measuring the complex shear modulus (G*) at 6 frequencies (1, 10, 20, 50, 100, 200 Hz), 461 at the center of the cell using a resolution of 4 × 4 pixels over a 10 µm × 10 µm area. 462

Data was analysed using the JPKSPM Data Processing software (JPK, Berlin, Germany), 463 which takes account of the hydrodynamic drag and processes the batches automatically. 464 The complex shear modulus is determined by the analysis of the force response of small 465 amplitude oscillations around a fixed indentation at different frequencies. 466

For a spherical indenter of radius R, the force F and the indentation δ can be obtained 468 based on the Sneddon's elasticity model: where, E is the Young's modulus, v the Poisson 469 coefficient (v = 0.5) and a is the contact radius. 470

$$F = \frac{E}{1 - v^2} \left[ (a^2 + R^2) \frac{\delta}{a} - aR \right] \quad 471$$

To link the force to the indentation, which is not accounted for in the Sneddon model, the 472 maximum amplitudes of the force (AF) and indentation (Aδ) as well as the phase shift φ 473 for each frequency are measured. The complex Young's modulus can then be written as: 474

$$E^* = \frac{1 - v^2}{2a_0} \frac{A_F}{A_\delta} e^{i\omega} \quad 475$$

For each frequency, the dynamic shear modulus is defined as $G^* \equiv \frac{E^*}{2(1+\vartheta)}$ 476

$$G^* = G' + iG'' = \frac{1 - v}{4a_0} \frac{A_F}{A_\delta} e^{i\omega} \quad 477$$
479 Single cell force spectroscopy

480 Cantilever functionalization and sample preparation

481 NPO10 (Bruker, USA) triangular shaped AFM tipless cantilevers (Bruker, USA) were
482 functionalized using the lectin Concanavalin A (conA; Sigma Aldrich). Cantilevers were
483 first cleaned for 2 min using oxygen plasma at a pressure of 0.4 mbar, then immersed in
484 100 µg/mL of conA solution overnight, and washed in acetate buffer (18 mM, pH 5.2)
485 before use. Human cells were plated into 10 mm dishes at 6 × 10⁴ cells per cm² and left
486 to grow overnight.

487 SCFS measurements

488 To prepare yeast-AFM probes, stationary-phase yeast cells were attached to the conA-coated cantilevers by approaching a single yeast cell with the AFM cantilever and allowing approximately 30 s contact, after which the cantilever was retracted along with the yeast as shown in Figure S2 (Nanowizard III AFM, Bruker, JPK). Single human-Candida cell interactions were measured using the AFM CellHesion® module under human cell growth conditions in a heated (Bruker, JPK BIOAFM) petri dish to which CO₂ was delivered. The spring constant of the cantilevers was calibrated using the thermal noise method, having a sensitivity of approximately 18 to 35 nm/V and a spring constant (0.044 N/m), consistent with the nominal value (0.06 N/m) provided by the manufacturer (Bruker, USA). The probes were then used to measure interaction forces within a 3 µm × 3 µm area using an applied force of 1 nN, with 20 µm/s approach and retract speeds and a 10 s contact time over 10 × 10 pixels. Each force curve was analyzed using the JPK Data Processing software (JPK, Berlin, Germany) from at least three biological replicates.

501
502 Candida to human cell adhesion assay.

503 To analyse the extent of adhesion between *Candida* and human cells arrested at various
504 stages of the cell cycle, cell-cell adhesion was assayed using the agar plate method.

505 Briefly, HaCaT, HeLa and HCT cells were seeded at 6 × 10⁵ cells/ml in 24 well plates and
506 left to grow overnight. Cells were then chemically synchronized for arrest at the G1, S
507 and G2/M phases. Upon synchronization, human cells were washed with PBS, co-
508 incubated with *Candida* (multiplicity of infection (MOI) 1:10) for 1 h and then washed 3 ×
509 with PBS to remove non-adherent yeast. Human cells were then detached from the plate,
510 serially diluted, a 100 µL aliquot spread onto a YPD agar plate incubated at 30 °C, and
511 the formation of colonies monitored and counted after 24 – 48 h. Each condition was
512 tested for 3 biological replicates with at least 3 technical replicates per experimental
513 triplicate.

514

515 Statistical analysis

516 GraphPad Prism® Software (version 10.2.3) generated graphical representations and
517 statistical analyses. One-way ANOVA analysis of variance with a Bonferroni's post-test
518 was used to compare all test data. Statistically significant differences of p < 0.05(*), p <
519 0.01 (**), p < 0.001 (***) or p < 0.0001 (****) are indicated in figures, and the absence of
520 an asterisk indicates no statistical difference, (p > 0.05, ns). Error bars represent the
521 standard error (SE) from the mean.

522 Supporting Material

523 Supporting information is available at

524
525 Author Information

526 **Corresponding Authors**

527 **Etienne Dague** – National Centre for Scientific Research, Laboratory for Analysis and
528 Architecture of Systems (LAAS), Toulouse, cedex 4, 31031, France; orcid.org/0000-0003-529 3290-9166; Email:edague@laas.fr

530 **Tanya Elizabeth Susan Dahms** - Department of Chemistry and Biochemistry, University
531 of Regina, Regina, S4S 0A2, Saskatchewan, Canada; orcid.org/0000-0002-8378-7480;
532 Email: tanya.dahms@uregina.ca

533
534 **Authors**

535 **Easter Ndlovu** - Department of Chemistry and Biochemistry, University of Regina,
536 Regina, SK, S4S 0A2, Saskatchewan, Canada

537 **Zinnat Shahina** - Department of Chemistry and Biochemistry, University of Regina,
538 Regina, SK, S4S 0A2, Saskatchewan, Canada

539
540 **Author Contributions**

541 E.N. performed all the experiments and analyzed the data and wrote the manuscript. Z.S
542 and E.N. collected the fluorescence images. E.N., T.E.S.D., and E.D. contributed to the
543 hypotheses, experimental design, helped guide data analysis, interpretation and helped
544 write and edit the manuscript.

545
546 Acknowledgments

547 This work was supported by the Discovery Grant from Natural Science and Engineering
548 Research Council (NSERC DG; 06649-2018) to T.E.S.D. Agence Nationale de la
549 Recherche (ANR) AutoBioTip – grant number ANR-20-CE42-0017 to E.D. E.N. was
550 supported by the Faculty of Graduate Studies and Research at the University of Regina
551 and NSERC.

552
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