Monitoring Candida albicans filamentous growth in micro-fabricated chambers
To investigate C. albicans hyphal growth, we took advantage of micro-fabrication approaches using the elastomer polydimethylsiloxane (PDMS) that, in particular, have been reported as single-cell force sensors for fission yeast cells [21]. We generated PDMS arrays with approximately 105 microchambers, which were cylindrical in shape with a diameter of 10 μm, a depth of 5 μm, and 15 μm spacing between adjacent chambers (Fig. 1a). C. albicans cells in micro-fabricated PDMS chamber arrays were visualized with inverted microscopes; imaging was carried out through an upright array of 150–200-μm-thick PDMS. Figure 1b shows an XZ confocal reflectance scan through the PDMS microarray with the chambers and media at the top (highest position) and the coverslip below for support (a zoom of chambers is shown in Fig. 1c). C. albicans cells were mixed with fetal calf serum, added to the PDMS array, incubated for ~ 1 h, and subsequently, filamentous growth was followed over time (Fig. 1d). With low-stiffness PDMS (a high polymer to cross-linker ratio of 40:1) we observed two predominant filamentous growth modes: non-invasive growth on the PDMS surface and invasive growth within PDMS (Fig. 1e, f). By examination of the focal plane of the PDMS surface and the fungal filaments, using DIC optics, we were able to distinguish between non-invasive (surface) and invasive growth, referring to whether the filament tip is on or within the PDMS, respectively. Invasive growth was also confirmed by labeling the PDMS surface and filamentous cells (see below). Furthermore, we observed that the blastospore (round cell) portion of the filamentous cells, which grew in the microchambers, pushed back against the chamber wall upon PDMS filament penetration and the filament frequently buckled within PDMS, presumably due to the resistive force during growth within the elastomer (Figs. 1f and 2a). These results indicate that, in addition to having ideal optical properties, PDMS is compatible with C. albicans filamentous growth.
Growth modes depend on substrate stiffness
We followed C. albicans filamentous growth in PDMS of different stiffness, i.e. the extent to which an object resists deformation in response to an applied force, by varying the ratio of polymer to cross-linker. We observed two main growth modes from cells initially in chambers depending on PDMS stiffness: invasive growth, which predominated with less stiff PDMS (40:1) (Fig. 2a) and dramatic bending in the stiffer PDMS (30:1) chambers, which was predominantly subapical (Fig. 2b, top panel). In contrast to Schizosaccharomyces pombe [21], extensive deformation of the chambers was not observed with C. albicans, which we attribute to the different sizes, geometries, and growth modes of these fungi; fission yeast has a radius of ~ 2 μm, compared to C. albicans hyphal filaments with a radius of ~ 1 μm [22] (and our observations), resulting in a greater than 4-fold difference in the cross-sectional area. At most, a slight chamber deformation was observed with C. albicans, as cells frequently popped out of the chambers comprised of stiff PDMS or penetrated this material, when less stiff. Occasionally, at intermediate PDMS stiffness, we observed filamentous cells growing on the PDMS surface that appeared to be probing the surface with a “nose down” growth (Fig. 2b, bottom panel), as similarly observed [19]. This type of growth was suggestive of the filaments attempting to penetrate into PDMS and, consistent with this, we observed these filaments buckling and/or bending subapically at each attempt, prior to the tip popping out and forward. Buckling is defined as a sudden change in the shape of a component under load, i.e. change in the shape of the filament due to the physical forces it experiences. Subapical bending is, additionally, defined as a change in the direction of growth that results in curved filaments (Fig. 2c). In buckling, it is expected that the shape changes are largely reversed upon removal of the external forces, whereas in bending, the shape changes are not a result of the mechanical forces directly. Buckling can occur with a filament initially straight or bent/curved.
We next examined whether cells, initially in chambers, which were unable to invade, underwent bending. Figure 2d shows that as the stiffness of PDMS increased (from 50:1 to 30:1 PDMS to cross-linker), there was an increase in the percentage of cells undergoing bending, concomitant with a decrease in those invading PDMS. During invasive growth, we also frequently observed buckling of the filament (Fig. 2a red arrowheads, c), i.e. a growth-dependent curvature that typically occurred at the portion of the filament within PDMS. Figure 2e shows that such buckling was dependent on the PDMS stiffness, with over half of invasive filaments buckling in the two stiffest PDMS (40:1 and 35:1).
To determine the mechanical properties of the different PDMS preparations, we used dynamic mechanical analysis for which measurements were reproducible over a range of PDMS stiffness. Oscillating strain at a frequency of 10 Hz was applied to PDMS samples, and stress (σ)-strain (ε) curves were obtained (Additional file 1: Figure S1), from which Young’s modulus was determined (initial dσ/dε). Young’s modulus is the quantitation of the stiffness, i.e. the ratio stress/strain for a uniaxial load, where stress is the force per unit area and strain is the proportional deformation (change in length divided by original length) and is dimensionless. Figure 2f shows the Young’s modulus of different ratios of PDMS to cross-linker, which are in good agreement with published values [23,24,25,26]. The stiffness of lower ratio samples (10:1 and 20:1), intermediate ratios (30:1 to 40:1), and higher ratios (45:1 and 50:1) was similar to that of medical silicone implants [27], with a Young’s modulus of ~ 1 MPa; stiff tissues such as the myocardium [7], with a Young’s modulus of ~ 0.1–0.2 MPa; and less stiff tissues such as the epithelia [5, 6], with a Young’s modulus of ~ 40–70 kPa, respectively.
Penetration into and escape from PDMS
Given that active penetration is critical during the process of C. albicans epithelium invasion [10,11,12, 14], we examined in further detail this process in PDMS. Figure 3a shows a filamentous cell that penetrates PDMS after 4 min (II; note that I, not shown, is prior to the filament contacting the chamber wall); subsequently grows invasively within PDMS (III); deforms the adjacent chamber (IV), resulting in a dramatic invagination; and exits PDMS into the adjacent well at 2:04 (V), followed by penetration into the opposing chamber at 2:08 (VI) and subsequent invasive growth (2:12; VII). The resistive force revealed by buckling of the filament, as well as deformation of the initial chamber during invasive growth (III), likely increases upon deformation and subsequent piercing into the adjacent well (IV), as the portion of the filament within PDMS buckled during this time (1:22–2:02), resulting in an S-shaped filament (Fig. 3a). The tension on the filament was released upon exiting PDMS into the adjacent well (V), as the tip of the filament appears to jump forward (2:04). The resistive force from the final step of growth (VII) also resulted in buckling of the filament (portion in the well) leading to an M shape (2:42–3:00). This escape from PDMS is analogous, in some respects, to filaments bursting out of a macrophage [28,29,30,31]. Here, the filament pushes into a circle resulting in a deformation that does not require expansion of the surface area but rather local invagination of the chamber, which is easier to detect (Fig. 3a). Indeed, such a bursting out of PDMS was observed a number of times, and Additional file 1: Figure S2 shows such examples in different PDMS stiffness (40:1, 110 kPa; 35:1, 150 kPa; and 30:1, 250 kPa).
In order to better visualize the invasive growth within PDMS during these different steps, we followed cells in which GFP was targeted to the plasma membrane [32], by confocal spinning disk microscopy acquisition over a range of z-positions. Figure 3b and c show a typical time-lapse acquisition in which the analysis of the cell outline did not reveal a substantial change in the shape of the filament tip during invasive growth and bursting into the next well (Fig. 3c, d; Additional file 1: Figures S3A and S3B). Indeed, the radius of the curvature of the cell tip was identical to that of surface-growing cells, and there were no changes upon burst out of PDMS. Buckling of the filament was evident upon invasive growth and occurred over 35–45 min, prior to the appearance of a septum (Fig. 3b (red arrowheads) and Fig. 4a, two examples). Analyses of the angle of the filament at which the septum formed ultimately, indicate that during invasive growth, cytokinesis occurs the majority of the time after the filament buckles (Fig. 4b).
To analyze the physical constraints during penetration, invasive growth, and tip escape from the PDMS matrix, we established a physical experimental model, which consisted of a steel probe that mimics the filament shape, continuously advancing up to and into a cylinder of PDMS of different stiffness (Fig. 5a, b). The steel probe tip approximated the shape of the filament tip (Fig. 5c) with a radius of curvature (± 45°) of 1.1 μm when normalized to the hyphal filament, compared to that of 1.0 ± 0.1 μm for the surface and invasively growing hyphal filaments. Figure 5d shows the probe prior to PDMS rupture and subsequent to exiting from the PDMS. Experiments were carried out with a range of probe displacement rates encompassing that of the filament extension rates (~ 0.3 μm/min [33]), when scaled down to the filament diameter. Figure 6a shows an example of such a force versus displacement curve. The initial phase of increasing force corresponds to the elastic compression of PDMS (Felast compr; analogous to growth stage II, Fig. 3a), culminating in the force required to break the PDMS surface, Fcrit. The next phase corresponds to the extension within PDMS, analogous to invasive growth, Finvas (analogous to growth stage III, Fig. 3a) culminating in PDMS exit (Fexiting; analogous to the end of growth stage IV, Fig. 3a). The final phase corresponds to when the end of the probe has emerged from the PDMS (Fout; analogous to growth stage V, Fig. 3a). Figure 6b shows the Fcrit from the physical probe experiments as a function of PDMS stiffness. Filaments are able to penetrate PDMS of ratio 35:1, for which we measured Fcrit of ~ 7 N with a 1-mm-diameter probe. Scaled to the diameter of a hyphal filament, this would correspond to a force of ~ 31 μN, indicating that hyphal filaments generate forces larger than 31 μN to penetrate PDMS. Scaling to the diameter of a hyphal filament was done assuming a constant critical stress, given the filament diameter is several orders of magnitude greater than the PDMS mesh size, and was calculated by taking the ratio of the metal probe/filament radius squared. Furthermore, our results indicate that at a stiffness of 200 kPa, for which little PDMS penetration is observed (Fig. 2d), the Fcrit is ~ 8 N with a 1-mm-diameter probe, i.e. ~ 35 μN when scaled to a hyphal filament, which would correspond to the growth stalling force.
Resistive force affects hyphal extension and morphology
The buckling of the filaments, as well as the deformation of the PDMS wells during invasive growth, indicated that these filaments were responding to resistive force, whose magnitude we have measured in the physical model. The percentage of cells that penetrate PDMS is dependent on Young’s modulus (Fig. 2d, f), and analyses of percent of PDMS invasion at two stiffness values indicate that the threshold for invasion is between 120 and 200 kPa (Fig. 7a). Hence, to investigate the effects of resistive force on filamentous growth, we determined the length of the filaments over time from cells growing on and within PDMS in this range of stiffness. Figure 7b shows that cells extend at a constant rate, which is reduced by ~ 30% within PDMS from an average of 0.28 ± 0.07 μm/min (n = 29) for surface growth to 0.19 ± 0.05 μm/min (n = 32) when filaments were growing within PDMS with a stiffness of ~ 100 kPa (Fig. 7c). This filament extension rate was further reduced to 0.15 ± 0.08 μm/min (n = 23) upon growth in PDMS with a stiffness of ~ 150 kPa (Fig. 7c). To confirm that the reductions in extension rate during invasive growth were not due to the substantial changes in the z-position of the growing filament apex, cells were grown in PDMS chambers that had been stained with fluorescent ConA and z-section images were projected onto the XZ plane (Fig. 7d). These projections show that the filaments grow slightly downward in PDMS, below the bottom of the chamber, with maximally 5 μm displacement in the z-axis for a 25-μm filament, resulting in at most a 2% reduction in extension rate upon projection in the XY plane. In contrast, Fig. 8a shows that the mean filament extension rate of cells grown on the surface is not dependent on the substrate’s stiffness, with indistinguishable rates on PDMS with Young’s modulus from 100 to 200 kPa. Furthermore, the extension rate of invasive growth normalized for that of surface growth from each experiment correlates with substrate stiffness (Fig. 8b). Extrapolation to the y-intercept, where the invasive extension rate, equals the surface rate indicates a substrate stiffness of ~ 20 kPa, suggesting that during filamentous growth on PDMS, the cells experience this resistive force from adhesion. Consistently, the surface extension rate was slightly reduced compared to that in liquid media (0.26 ± 0.09 μm/min compared to 0.32 ± 0.01 μm/min; p = 0.001) (Fig. 8a). Of note, very few cells invaded PDMS at 200 kPa, on average ~ 5% (Fig. 7a), but, strikingly, the filament extension rate for these “escapers” was similar to that of less stiff PDMS, raising the attractive possibility that these cells may play a critical role in tissue invasion.
The difference in filament extension rate could either be due to an overall reduction in cell growth or a reduction in polarized, apical growth. To differentiate between these possibilities, we determined the length and diameter of compartments (between 2 septa) for cells growing on the surface and within PDMS (100 kPa). Figure 9a and b show that the compartment length decreased ~ 30%, from 24.6 ± 3.0 μm (n = 100) for surface growing cells to 16.6 ± 1.8 μm (n = 120) during invasive growth, and the filament diameter increased concomitantly from 2.1 ± 0.2 μm for surface growing cells to 2.5 ± 0.2 μm during invasive growth. As a result, the compartment volume remained constant (83 ± 17 μm3 for surface growing cells compared to 80 ± 18 μm3 for invasively growing cells), indicative of altered polarized growth. Consistently, analyses of filamentous cells grown within stiffer PDMS (150 kPa) revealed a further decrease in compartment length (14.6 ± 2.5 μm) and an increased diameter (2.8 ± 0.3 μm). This altered morphology is dependent on growth against a resistive force in PDMS as the diameter in the part of the filament outside PDMS was similar to that of surface growing cells (Fig. 10a). In addition to the comparison of cells growing on the surface and within PDMS, we examined the relatively rare occurrence of cells transitioning between these growth modes. Figure 10b and c show an example of such a transition, in which the extension rate is reduced during invasive growth in PDMS and increases exiting PDMS. Measurements of the filament diameter just before and after the filament exited PDMS revealed an increased filament diameter during invasive growth that was significantly reduced upon exiting PDMS (2.7 ± 0.2 μm compared to 2.3 ± 0.3 μm, Fig. 10d).
Reduced filament extension rate in response to a resistive force suggested that similar effects could be observed in cells undergoing non-invasive, dramatic subapical bending in chambers of stiff PDMS (Additional file 1: Figure S4A). Additional file 1: Figure S4B shows that, in such conditions, there was indeed a dramatic reduction in filament extension rate with an average (over 100–150 min) of 0.10 ± 0.01 μm/min. Surface growth rates were constant over time, whereas extension rates of cells undergoing dramatic subapical bending decreased concomitantly with the cell filling the well (Additional file 1: Figure S4C); initial rates of extension were 3-fold reduced from surface growth, and these were further reduced 3-fold after 2 h of growth. Due to the complex geometries during such a growth mode, we were unable to determine the resistive force that the filament experiences while it fills up the chamber; however, the initial extension rate is similar to that of filaments growing invasively in 150 kPa PDMS.
Determination of the effective turgor pressure
From the comparison of extension rates within PDMS of different stiffness (Fig. 7c), we determined the effective turgor pressure in C. albicans hyphae, using the viscoplastic growth model [21]. This determination makes use of Finvas values, measured from the physical model (Fig. 6), after scaling these forces from a cylinder with a radius of 0.5 mm to that of 1.04 μm. In order to correctly extrapolate the macroscopic measurements to the microscopic scale of filamentous cells, we analyzed the physical forces at play. The mode of extension during hyphal growth and in this physical experimental model is different, as new material is incorporated into the hyphal tip, i.e. growth occurs via apical extension, whereas in the physical experimental model, the probe is pushed into the PDMS from the back. Given that only a small portion of the filamentous cell apex extends in the PDMS, we removed the contribution from friction/adhesion due to the displacement of a 1-mm-diameter probe within PDMS by subtracting the Fout value from the Finvas (Fig. 6c). These corrected Finvas values, i.e. Fin, were largely independent of probe displacement rates over a range equivalent to cell filament extension rates when scaled down (0.2–0.4 μm/min). We used the equation that was established for S. pombe by Minc and colleagues;
$$ \frac{V_{(F)}}{V_o}=\left(1-\frac{F_{\left(\mathrm{PDMS}\right)}}{\pi {R}^2\Delta P}\right) $$
(1)
V(F) and Vo are the filament extension rates within PDMS and on the surface, respectively; F(PDMS) is the resistive force of PDMS during filament displacement within this material (Fin); R is the filament radius; and ∆P is the effective turgor pressure. The F(PDMS) was 1.5 ± 0.7 N and 0.7 ± 0.3 N at PDMS to cross-linker 35:1 (Young’s modulus of 150 kPa) and 40:1 (Young’s modulus of 100 kPa), respectively; scaling to the size of the hyphal filament (radius 1.04 μm for surface growth) yielded 6 ± 3 μN and 3.2 ± 1.4 μN, respectively. From these values, we determined the effective turgor pressure, ∆P, to be 6 ± 3 MPa and 2 ± 1 MPa in these two conditions, respectively. Given that the hyphal filament diameter increases during invasion (radius 1.42 μm at Young’s modulus of 150 kPa and 1.24 μm at Young’s modulus of 100 kPa), the calculated ∆P are 3.1 ± 1.5 MPa and 1.5 ± 0.7 MPa, suggesting that the hyphal turgor pressure is 1–3 MPa. This value for turgor pressure is within the range reported both for planktonic and biofilm C. albicans cells, ~ 1.2 MPa [19] and ~ 2 MPa [34], respectively, as well as S. pombe, 0.85–1.5 MPa [21, 35]. Nonetheless, it must be noted that these values are effective turgor pressure. In other words, ∆P is the turgor pressure exceeding the critical stress needed to deform the cell wall [21]. Hence, a combination of local compartment turgor pressure alteration, difference in cell wall deformability, or potentially finer adjustments in tip geometry may play important roles in penetration and invasion.
Resistive force affects cell polarity
The change in morphology during invasion, resulting in shorter and wider cells, could be explained by tip growth becoming more isotropic in response to a resistive force, raising the possibility that cell polarity is adversely affected. In S. pombe, it was observed that reducing the growth rate chemically, genetically, or mechanically destabilized active Cdc42 polarization, a cell polarity master regulator [36]. To investigate whether cell polarity was altered in hyphal filaments growing invasively in PDMS, we examined the distribution of active Cdc42 (Cdc42•GTP), using a CRIB-GFP reporter [37]. Surprisingly, we observed a striking increase in polarized active Cdc42 at the filament tip throughout invasive growth, compared to surface growth (Fig. 11a, b). We determined, using a tailor-made MATLAB program, that this results from an increase in the concentration of Cdc42•GTP at the tip, rather than an alteration in the position of the maximum signal or spread of active Cdc42 further down the filament (Additional file 1: Figure S5A-D). These results suggest that, in response to a resistive force, there is an increase in cell polarization, perhaps reflecting a direct response to such external forces. We speculate that this higher level of active Cdc42 during invasive growth is due to the increased recruitment of the Cdc42 activator, Cdc24 [38]. We next examined active Rho1, as cell wall stress mediated by the cell surface mechanosensors Wsc1/Mid2 results in Rho1 depolarization in S. cerevisiae [39, 40]. Figure 11c and d show that, in contrast to the increase in tip localized active Cdc42, active Rho1 is depolarized during invasive growth. We attribute this depolarization of active Rho1 to the mechanical properties of PDMS, which are likely to impose a uniform force over the hyphal filament surface, in addition to the resistive force in response to the tip extension.
The increase in tip-localized active Cdc42 during invasive growth suggests that the increase in filament diameter does not result from growth becoming more isotropic in response to resistive force. To examine whether this morphological change results from mechanical forces, we compared cell morphology over time during growth on the surface and within PDMS. Figure 12a and b show that the relative filament diameter (D1) was not altered during surface growth (mean diameter 2.26 ± 0.15 μm initially compared to 2.44 ± 0.06 μm after 2 h growth), in contrast to the invasive growth where there was a striking increase (2.38 ± 0.22 μm initially compared to 2.98 ± 0.15 μm, p < 0.0001). Specifically, the diameter of the filament compartment increased even > 10 μm back from the tip (Fig. 12b). This ~ 25% increase in diameter during invasive growth could either occur upon tip growth or subsequent to tip growth. The hyphal tip diameter was constant over 2 h of invasive growth and only slightly wider than that of cells growing on the surface (2.53 ± 0.11 μm compared to 2.24 ± 0.09 μm; p = 0.0003) (Fig. 12c, d). In contrast, Fig. 12d shows that there was a significant difference between the mean diameter at the tip of the apical cell and that of the cell proximal to the apical cell during invasive growth (mean proximal cell diameter 2.99 ± 0.17 μm; p < 0.0001). Together, these results indicate that relatively small changes in the tip morphology are not sufficient to explain the altered morphology of the filament, back from the tip, which are due to external mechanical forces.