Rapid adaptation of endocytosis, exocytosis, and eisosomes after an acute increase in membrane tension in yeast cells

  1. Joël Lemière
  2. Yuan Ren
  3. Julien Berro  Is a corresponding author
  1. Department of Molecular Biophysics and Biochemistry, Nanobiology Institute, Yale University, United States
  2. Nanobiology Institute, Yale University, United States
  3. Department of Cell Biology, Yale University School of Medicine, United States

Abstract

During clathrin-mediated endocytosis (CME) in eukaryotes, actin assembly is required to overcome large membrane tension and turgor pressure. However, the molecular mechanisms by which the actin machinery adapts to varying membrane tension remain unknown. In addition, how cells reduce their membrane tension when they are challenged by hypotonic shocks remains unclear. We used quantitative microscopy to demonstrate that cells rapidly reduce their membrane tension using three parallel mechanisms. In addition to using their cell wall for mechanical protection, yeast cells disassemble eisosomes to buffer moderate changes in membrane tension on a minute time scale. Meanwhile, a temporary reduction in the rate of endocytosis for 2–6 min and an increase in the rate of exocytosis for at least 5 min allow cells to add large pools of membrane to the plasma membrane. We built on these results to submit the cells to abrupt increases in membrane tension and determine that the endocytic actin machinery of fission yeast cells rapidly adapts to perform CME. Our study sheds light on the tight connection between membrane tension regulation, endocytosis, and exocytosis.

Introduction

During clathrin-mediated endocytosis (CME), the cell plasma membrane undergoes a dramatic change in topology to form an invagination that is subsequently pinched off into a vesicle. During this process, the endocytic machinery has to overcome the forces produced by membrane tension and the osmotic pressure that opposes membrane deformation and engulfment. In yeast cells, these resisting forces are particularly large because their internal turgor pressure is high, ranging from ~0.6 MPa for Saccharomyces cerevisiae to more than 1 MPa for Schizosaccharomyces pombe (Davì et al., 2018; Minc et al., 2009; Schaber et al., 2010). Consequently, the formation of a vesicle requires several thousands of piconewtons (pN) (Dmitrieff and Nédélec, 2015; Ma and Berro, 2021).

Previous studies have shown that actin dynamics is required for productive endocytosis in yeast (Aghamohammadzadeh et al., 2014; Basu et al., 2014; Carlsson and Bayly, 2014; Palmer et al., 2015) and in mammalian cells when membrane tension is high (Aghamohammadzadeh and Ayscough, 2009; Boulant et al., 2011; Hassinger et al., 2017), or when membrane scission proteins are absent (Ferguson et al., 2009). Actin assembly at the endocytic site is believed to provide the forces that overcome turgor pressure and membrane tension to deform the plasma membrane, but the precise mechanisms of force production remain unknown (reviewed in Berro and Lacy, 2018; Goode et al., 2015; Lacy et al., 2018). We also lack a quantitative understanding of the regulation of actin dynamics in response to membrane tension and turgor pressure changes. We expect that a better quantitative characterization of this response will allow us to infer the molecular mechanisms of force production and force sensing during CME.

The mechanisms by which membrane tension is regulated are not fully understood. The yeast cell wall is believed to buffer abrupt changes in turgor pressure, thanks to its high stiffness of ~50 MPa (Atilgan et al., 2015). In addition, similarly to mammalian cells’ caveolae, which change shape or disassemble in response to increased membrane tension, yeast eisosomes can also disassemble when cells without a cell wall, called protoplasts, are placed in low osmolarity media (Kabeche et al., 2015; Parton et al., 2020; Sinha et al., 2011). However, it remains unknown how eisosomes may regulate plasma membrane tension in intact cells and whether eisosome disassembly directly influences cellular processes such as CME. In addition, it remains unclear to which extent endocytosis and exocytosis may contribute to the regulation of membrane tension when the cells are challenged with an abrupt increase in membrane tension.

Fission yeast is an ideal model system to quantitatively study the regulation mechanisms of membrane tension and its influence on the endocytic machinery. First, because yeast turgor pressure is high, actin is required for CME. Second, contrary to mammalian cells, yeast cells are devoid of any adhesion machinery or actin cortex, which usually complicates membrane tension manipulation and result interpretation. Last, quantitative microscopy methods developed in fission yeast are able to uncover fine regulations of the endocytic machinery and precisely measure the local and global numbers of endocytic events at a given time (Arasada and Pollard, 2011; Berro et al., 2010; Berro and Lacy, 2018; Berro and Pollard, 2014a; Berro and Pollard, 2014b; Chen and Pollard, 2013; Lacy et al., 2019; Sirotkin et al., 2010).

To probe the contributions of each possible mechanism of membrane tension regulation and their influence on CME, we submitted yeast cells with or without a cell wall to different hypotonic shocks. Using quantitative fluorescence microscopy, we showed that, on one hand, yeast cells rapidly reduce their membrane tension by (a) disassembling eisosomes, (b) reducing their rate of endocytosis, and (c) increasing their rate of exocytosis and, on the other hand, actin assembly adapts to increased membrane tension to allow endocytosis to proceed.

Results

Eisosomes participate in the regulation of protoplasts’ membrane tension

Previous studies have proposed that eisosomes, furrows at the inner surface of the plasma membrane, have a mechanoprotective role under increased membrane tension in fungi by acting as a reservoir of membrane, similar to the protective role of caveolae in endothelial cells (Cheng et al., 2015; Kabeche et al., 2015; Lo et al., 2016; Sens and Turner, 2006; Sinha et al., 2011). Since the yeast cell wall plays a major role in the maintenance of cell integrity under extreme osmotic conditions, thanks to its high stiffness of ~50 MPa (Atilgan et al., 2015), it prevents large variations in membrane tension under hypotonic shocks. Hence, to exclude the effect of the cell wall and amplify membrane tension changes, we performed our experiments using cells devoid of a cell wall, hereafter referred to as ‘protoplasts’, instead of intact cells, hereafter referred to as ‘walled cells’.

First, we characterized how the removal of the cell wall affects eisosomes’ reorganization. We used a protocol that allowed us to manipulate protoplasts for up to ~1 hr after their formation, since they remain void of cell wall for about 3 hr (Flor-Parra et al., 2014). Because protoplasts are more fragile than walled cells, they were prepared in Edinburgh Minimum Media (EMM5S) containing 0.25–1.2 M sorbitol to balance turgor pressure and prevent cells from bursting, while keeping nutrient concentration constant (Basu et al., 2014; Kabeche et al., 2015; Stachowiak et al., 2014), and were imaged ~15 min later, once they reached steady state. In the rest of the paper, we will refer to this experimental condition as ‘steady state in X M’ or ‘chronic exposure to X M sorbitol’, where X is the sorbitol concentration.

Our data show that eisosomes in protoplasts at steady state in 1.2 M sorbitol are qualitatively similar to those in walled cells (Figure 1A), and the cellular concentration of Pi1lp is the same in both conditions (Figure 1E). However, the surface area of the protoplasts’ plasma membrane covered by eisosomes decreased with decreasing media osmolarity at steady state (Figure 1B and C) and correlated with increasing cell volume (Figure 1D). This result confirms previous results (Kabeche et al., 2015) showing that eisosomes are disassembled in media with low osmolarity and the disassembly of eisosomes may reduce membrane tension.

Eisosomes disassemble to buffer increases in membrane tension.

(A) Representative walled yeast cells (left column) and protoplasts (right column) at steady state in 1.2 M sorbitol expressing eisosome core protein Pil1-mEGFP (inverted contrast). Note that the cellular concentration of Pil1p-mEGFP is the same in walled cells and protoplasts (panel E). (B) Eisosomes labeled with Pil1p-mEGFP (inverted contrast) in wild-type protoplasts at steady state in different sorbitol concentrations. From left to right: 0.25, 0.4, 0.8, and 1.2 M sorbitol. (C and D) Density of eisosomes at the plasma membrane (C), measured as the ratio between the intensity of Pil1p-mEGFP on the plasma membrane and the surface area of the protoplast and volume (D), at steady state in 0.25 M (N = 26), 0.4 M (N = 34), and 1.2 M (N = 39) sorbitol. Error bars: standard deviations. (E) The total amount of Pil1-mEGFP in walled cells (N = 32) and protoplasts in 0.4 M sorbitol (N = 19) are not significantly different (Mann-Whitney test, p=0.65). Each point represents one measurement; bars are the mean and SEM. (F) Micropipette aspiration was used to measure membrane tension. Rc: cell radius; Rp: micropipette radius; l: length of the tongue inside the micropipette. (G) Membrane tension of protoplasts at steady state in 0.8 M sorbitol and ~5 min after a hypotonic shock (ΔC = −0.2 M) for wild-type (blue bars, N = 28 for steady state and N = 5 for the shock) and pil1Δ protoplasts (red bars, N = 42 for steady state and N = 7 for the shock). Error bars: standard deviation. p-values: non-significant (ns), p>0.05; **p≤0.01; ***p≤0.001. (H and I) Eisosomes of wild-type protoplasts disassemble rapidly after a hypotonic shock. (H) Time course of a representative protoplast expressing Pil1p-mEGFP over 10 min after a hypotonic shock (ΔC = −0.2 M) and initially at steady state in 0.4 M sorbitol (just before time 0 min). (I) Evolution of the surface area covered by eisosomes over time, as a fraction of the surface area covered at time 0 min (normalized to 1). Data are from three independent experiments (N = 15) and presented as mean ± 95% confidence interval. Scale bars in (A), (B), (F), and (H): 5 µm.

We then performed hypotonic shocks to abruptly increase the membrane tension of protoplasts and we imaged them before their long-term adaptation to changes in media osmolarity. Prior to the shocks, we let cells reach steady state by exposing them to media with a given sorbitol concentration for more than 15 min. We chose 0.4 M sorbitol as the steady-state concentration for this experiment as it corresponds to the estimated turgor pressure of walled cells (~1 MPa), and at this concentration, the dynamics of the actin endocytic machinery was virtually identical to the one in walled cells (Figure 7—figure supplements 1B). We performed acute hypotonic shocks by using a microfluidic system to rapidly exchange the steady state media with media containing a lower sorbitol concentration, hereafter noted as ΔC=–Y M, where Y is the difference in media osmolarity (note that the change in pressure ΔP in Pascal is related to the change in osmolite concentration ΔC in Molar as ΔP=ΔCRT~2.45106ΔC, where R is the gas constant and T, the absolute temperature, when the solute concentration is sufficiently low).

To quantitatively characterize eisosome disassembly in protoplasts after a hypotonic treatment, we measured the temporal evolution of the decrease in surface area covered by eisosomes after an acute hypotonic shock of ΔC = −0.2 M starting with protoplasts at steady state in 0.4 M sorbitol (Figure 1H and I). Eisosomes disassembled rapidly after the hypotonic shock, dropping to ~50% of the surface area covered by eisosomes before the shock within 5 min, indicating a fast response to counteract the hypotonic shock and an eventual change in membrane tension.

To test whether membrane tension is buffered by eisosomes, we measured membrane tension using a micropipette aspiration assay (Figure 1F). At steady state in 0.8 M sorbitol, the membrane tension was 0.45 ± 0.14 mN·m−1 for wild-type (WT) protoplasts and 0.39 ± 0.13 mN·m−1 for pil1Δ protoplasts (Figure 1G). We then repeated these measurements within 5 min after inducing a hypotonic shock of ΔC = −0.2 M. We observed a 1.6-fold increase in membrane tension for WT protoplasts (0.73 ± 0.21 mN·m−1) and a 4.5-fold increase for protoplasts lacking eisosomes (1.74 ± 0.61 mN·m−1). This result demonstrates that eisosomes participate in the adjustment of plasma membrane tension.

In protoplasts, eisosomes buffer moderate hypotonic shocks

Since WT protoplasts were able to withstand osmotic shocks by disassembling their eisosomes, we hypothesized that protoplasts lacking eisosomes – either because they lack the core eisosome protein Pil1p or because eisosomes are mechanically removed – are more sensitive to hypotonic shocks.

WT and pil1Δ protoplasts initially at steady state in 0.4 M sorbitol survived small hypotonic shocks (ΔC = 0.05 M) equally well (Figure 2A). However, pil1Δ protoplasts were more sensitive to moderate hypotonic shocks (ΔC = 0.1 M) since most of them were unable to survive 2 min after the moderate shock while virtually all the WT protoplasts were able to survive (Figure 2B, Figure 2—figure supplement 1).

Figure 2 with 1 supplement see all
Eisosomes protect protoplasts and walled cells from osmotic shocks.

(A–C) Percentage of wild-type (blue dots) and pil1Δ (red triangle) protoplasts that were alive at steady state in 0.4 M sorbitol, and after a ΔC = −0.05 M (A), ΔC = −0.1 M (B), and ΔC = −0.2 M (C) single hypotonic shock. Representative fields of view used to determine these percentages are shown in Figure 2—figure supplement 1. (D) Percentage of wild-type (blue dots) and pil1Δ (red triangles) protoplasts that were alive at steady state in 0.25 M sorbitol, and after a ΔC = −0.1 M hypotonic shock. In these conditions, before the shock, eisosomes in wild-type protoplasts were almost completely disassembled (Figure 1B). After the shock, wild-type cell survival was comparable to cells void of eisosomes because they lack Pil1p. (E) Timeline of repeated ΔC = 1.2 M osmotic shocks for walled cells. Each osmotic shock was performed by exchanging sorbitol concentration from 1.2 M (5 min) to 0 M (1 min). (F) Percentage of wild-type (blue dots, N = 273) and pil1Δ (red triangle, N = 197) walled cells that were alive after each osmotic shock. Note the progressive cell death induced by repeated osmotic shocks for pil1Δ cells. Combined data are from three independent experiments and plotted as mean ± standard deviation. (G) Representative images of wild-type (upper panel) and pil1Δ (lower panel) walled cells before shock and after the fourth shock. Dead cells were strongly stained by FM4-64 due to membrane damage. Scale bar: 10 µm.

Moreover, eisosomes were unable to protect protoplasts from larger hypotonic shocks (ΔC = 0.2 M), where most eisosomes are disassembled (Figure 1H and I) since most wild-type protoplasts were unable to survive longer than 4 min after these high hypotonic shocks, similarly to pil1Δ protoplasts (Figure 2C). To further confirm that increased death rate was due to the lack of eisosomes, we performed a moderate shock (ΔC = 0.1 M) on protoplasts originally at steady state in 0.25 M sorbitol, where eisosomes were mostly disassembled already. Under these conditions, WT protoplasts survival was comparable to the survival of pil1Δ (Figure 2D). This result further demonstrates that the presence of assembled eisosomes at the plasma membrane is indeed responsible for the adaptation of cells to acute hypotonic shocks, and the presence of Pil1p in the cytoplasm is not sufficient for this response.

Altogether, these experiments demonstrate that (a) eisosomes protect protoplasts from changes in their membrane tension, but only to a small extent, and (b) without eisosomes, protoplasts can withstand only minor increase in their membrane tension.

Eisosomes protect the integrity of walled cells during consecutive osmotic shocks

We observed that a significant number of both wild-type and pil1Δ protoplasts died after osmotic shocks, and the percentage of pil1Δ protoplasts that remained alive was significantly smaller than for wild-type protoplasts even under moderate shocks of ΔC=−0.05, −0.1, and −0.2 M (Figure 2A–C). In contrast, we found that both wild-type and pil1Δ walled cells can survive a single osmotic shock of ΔC = −1.2 M, which initially led us to think that eisosomes only have a minor protective role in walled cells (Figure 2F). However, we noticed that subsequent osmotic shocks led to a higher mortality of pil1Δ compared to wild-type walled cells. While almost all the wild-type walled cells remained alive after several shocks, around 10% of pil1Δ walled cells died after each subsequent shock (Figure 2E, F and G; supplementary video 1 and 2). These results demonstrate that, even in walled cells, eisosomes exert a protective role, likely by buffering sudden changes in membrane tension.

Membrane tension and eisosomes modulate the rate of endocytosis in cells

Within a few minutes of a hypotonic shock, the volume of WT protoplasts increased up to 50% and the volume of pil1Δ protoplasts increased up to 20% (Figure 3A and B, insets). However, the corresponding increase in surface area cannot be explained by eisosome disassembly alone – total eisosome disassembly could release about 5% of the total surface area of the plasma membrane, assuming eisosomes are hemi-cylinders with a diameter of ~50 nm and cells contain 1.6 μm of eisosomes per μm2 of plasma membrane on average (Kabeche et al., 2015). Therefore, another mechanism for protoplasts to gain plasma membrane occurs in the first few minutes after hypotonic shocks. We hypothesized that a decrease in the number of endocytic events happening in the cell after a hypotonic shock would gradually increase the surface area of the plasma membrane and reduce membrane tension.

Figure 3 with 1 supplement see all
The density of endocytic events rapidly adapts after acute osmotic shocks.

(A) Temporal evolution of density of endocytic events (average number of endocytic events at a given time in a cell divided by the cell length) in wild-type protoplasts initially at steady state in 0.4 M sorbitol and after an acute hypotonic shock of ΔC = −0.05 M (dark blue, Ncell ≥ 102), ΔC = −0.1 M (blue, Ncell ≥ 54), and ΔC = −0.2 M (light blue, Ncell ≥ 83). For ΔC = −0.1 and −0.2 M, the difference in the density of clathrin-mediated endocytosis (CME) events between steady state and 0 or 2 min after the shock is statistically significant (one-way ANOVA, p<10−4). In all conditions, the difference after 6 min is not significant (one-way ANOVA, p>0.12; details in Figure 3—source data 1). (B) Same as (A) but with pil1Δ protoplasts and hypotonic shocks of ΔC = −0.025 M (dark red, Ncell ≥ 70), ΔC = −0.05 M (red, Ncell ≥ 103), and ΔC = −0.1 M (light red, Ncell ≥ 78). In all conditions, the difference in the density of CME events between steady- state and any time after the shock is statistically significant (one-way ANOVA, p<10−3). For ΔC = −0.025 and −0.05 M, the differences between time points after 6 min are not significant (one-way ANOVA, p>0.09; details in Figure 3—source data 2). (A and B) Insets: relative volume increase after the hypotonic shocks (the volume at steady state is used as a reference). The numbers of cells used for each condition and each time point are given in Supplementary file 1a. The number of cells measured in the insets is the same as that in the main figures. *: the large majority of pil1Δ protoplasts were too damaged or dead 4 min after the hypotonic shocks at ΔC = −0.1 M (Figure 2B), which prevented us to measure the density of endocytic events and the volume after this time point. (C) Density of endocytic events in intact cells at steady state in different osmolarities, Ncell ≥ 80. In pil1Δ walled cells, the difference in the density of CME events between all pairs of conditions is statistically significant (one-way ANOVA, p<10−4). In wild-type walled cells, the difference is small but statistically significant (details in Figure 3—source data 3). The numbers of cells used for each condition and each time point are given in Supplementary file 1b. (D) Density of endocytic events in wild-type (blue circle) and pil1Δ (red triangle) walled cells initially at steady state in 1.2 M sorbitol and after an acute hypotonic shock of ΔC = −1.2 M, Ncell ≥ 44. The numbers of cells used for each condition and each time point are given in Supplementary file 1c. For wild-type and pil1Δ walled cells, the differences in the density of CME events after 2 min are not statistically significant (p>0.08; details in Figure 3—source data 4). (A), (B), (C), and (D): error bars are standard errors of the mean.

We measured the endocytic density, that is, the number of endocytic events in a cell normalized by the cell length, in wild-type and pil1Δ cells after a hypotonic shock using a ratiometric method (Berro and Pollard, 2014a). In brief, this method consists of imaging cells expressing a fluorescently tagged endocytic protein, here the actin filament crosslinking protein fimbrin (Fim1p) tagged with a monomeric enhanced green fluorescent protein (mEGFP), hereafter called Fim1p-mEGFP. First, we measure the temporal average intensity of the fluorescent protein at endocytic sites. Second, we measure the whole intensity of each cell from which the corresponding cytoplasmic intensity is subtracted – this number represents the sum of the intensities for all the fluorescently tagged proteins present at endocytic sites. The number of endocytic sites in each cell is then calculated as the ratio between those two numbers. The endocytic density is calculated by dividing this ratio with the cell length.

For all shocks tested in wild-type (ΔC = −0.05, −0.1, and −0.2 M) and pil1Δ protoplasts (ΔC = −0.025, −0.05, and −0.1 M) initially at steady state in 0.4 M sorbitol, the endocytic density in protoplasts significantly decreased immediately after the hypotonic shock (Figure 3A). The difference increased with increasing hypotonic shocks, up to 36% for wild-type protoplasts after a ΔC = −0.2 M shock, and 79% for pil1Δ protoplasts after a ΔC = −0.1 M shock (Figure 3B). These abrupt changes in the endocytic density were followed by a 2–6 min recovery back to the steady-state endocytic density, and recovery time depended on the magnitude of the hypotonic shock. Note that the change in cell volume (Figure 3A and B, insets) could not exclusively account for the observed decrease in the endocytic density in wild-type cells as the relative increase in cell volumes was larger than the relative decrease in endocytic density 2 min after the shocks and remained large 4 min after, while the endocytic densities recovered their pre-shock values.

Building on these results in protoplasts, we wondered whether the endocytic density in walled cells also adapts to hypotonic shocks. Indeed, immediately after the largest shock tested (ΔC = −1.2 M), we observed a similar decrease in the endocytic density for both wild-type and pil1Δ walled cells, 36% and 46%, respectively (Figure 3D). Recovery to steady-state endocytic densities occurred in less than 2 min in both wild-type and pil1Δ walled cells, faster than in protoplasts (Figure 3A, B and D). Our data show that the cell wall limits but does not completely cancel the effect of hypotonic shocks on endocytic rates. They also suggest that the regulation of the endocytic density supplements the regulation performed by the eisosomes to reduce membrane tension.

Wild-type and pil1Δ walled cells had a very similar adaptation after hypotonic shocks. However, we noticed a difference in the endocytic density at steady state in different sorbitol concentrations. For all concentrations tested (0–2 M), wild-type cells maintained roughly the same endocytic density. In contrast, the steady-state endocytic density in pil1Δ cells increased with increasing media osmolarity, up to 56% in 2 M sorbitol (Figure 3C). Our results suggest that eisosomes participate in maintaining a constant density of endocytosis independently of the media osmolarity, not only after an abrupt change in membrane tension, but also when they are at steady state in different osmolarities.

The exocytosis rate increases after a hypotonic shock in protoplasts but not in walled cells

Reciprocal to the decrease in the number of endocytic events observed after a hypotonic shock, we wondered whether the rate of exocytosis increases in the meantime to provide more surface area to the plasma membrane, as has been observed in mammalian cells (Gauthier et al., 2009).

To measure the rate of exocytosis in different conditions, we used the cell impermeable styryl dye FM4-64, whose fluorescence dramatically increases when it binds to membranes (Cochilla et al., 1999; Gachet and Hyams, 2005; Richards et al., 2000). After FM4-64 is introduced to the media, the plasma membrane is rapidly stained (Figure 4A). Fusion of unstained intracellular vesicles to the plasma membrane results in an increase in total cell fluorescence, because after each fusion event, a new unstained membrane from the interior of the cell is exposed to the dye. At this stage, if one wants to measure endocytosis rates, one would typically remove the FM4-64 dye from the media to destain the plasma membrane and quantify the internal fluorescence, which is proportional to the amount of membrane internalized by endocytosis. Here, we kept the FM4-64 dye in the media, and monitored the fluorescence of the plasma membrane and the interior of the cell. In this case, endocytic events do not increase the total cell fluorescence because they transfer patches of the plasma membrane that are already stained into the interior of the cell (Figure 4A, red arrow). Therefore, the increase in fluorescence we measured is due to the addition of new unstained lipids to the plasma membrane by exocytosis (Figure 4A, gray arrow). Note that the increase in total cell fluorescence could also be due to putative transfer of lipids by non-exocytic mechanisms (Reinisch and Prinz, 2021), but for simplicity and by lack of further evidence, hereafter, we will interpret the increase in fluorescence to an increase in exocytosis rate.

Figure 4 with 1 supplement see all
Exocytosis rate increases after an acute change in membrane tension in protoplasts but not in walled cells.

(A) Rationale of measurement of whole cell exocytosis rate through FM4-64 staining. After FM4-64 is flown in the imaging chamber, the dye rapidly binds to the cell surface in less than a minute. After this initial phase, the whole cell fluorescence increases every time the new (unlabeled) internal membrane is exposed to the cell surface by exocytosis. Note that endocytic events do not change the total fluorescence measured. (B) Measurement of yeast cell exocytosis rate at steady state in 0 M sorbitol. Cells were stained with 20 µM FM4-64 in Edinburgh Minimum Media (EMM5S) for 20 min before washing with EMM5S. During FM4-64 staining, the fluorescence intensity increases rapidly for 1 min before entering a slow linear phase over at least 20 min for wild-type cells. The fluorescence intensity at the end of the initial rapid increase phase corresponds to the complete staining of cell surface. It was normalized to 1, so that the subsequent increase in fluorescence intensity corresponds to a percentage of the plasma membrane surface area. After the dye was removed 20 min later, the decrease in fluorescence intensity suggests that the incorporation of FM4-64 didn’t interfere with the vesicle trafficking pathway of the cell. The rate of exocytosis (measured as a percentage of the plasma membrane surface area per minute) is the slope of a linear fit of the measured signal over the first 5 min (k0-5), 10 min (k0-10), or 15 min (k0-15). Example images of stained cells at different time points are shown in the middle panel (inverted contrast). (C–H) Rates of exocytosis at steady state and after hypotonic shocks. (C and D) The exocytic rate of wild-type walled cells is not changed after a ΔC = −1.2 M acute hypotonic shock (black, before shock, Ncells = 79; blue, after shock, Ncells = 68; three replicates each). The exocytic rate of pil1Δ walled cells does not change significantly in the same conditions (black, before shock, Ncells = 60; blue, after shock, Ncells = 96; three replicates each). All walled cells were at steady state in 1.2 M sorbitol before time 0 min. Curves for individual conditions in panels (C) and (D) are plotted in Figure 4—figure supplements 1A,B, respectively. (E) Summary of exocytic rates for wild-type and pil1Δ walled cells before and after hypotonic shock. (F and G) The exocytic rate of wild-type and pil1Δ protoplasts increases after ΔC = −0.2 M (black, before shock, Ncells = 20; light blue, after shock, Ncells = 37; four replicates each) and ΔC = −0.05 M (black, before shock, Ncells = 44; red, after shock, Ncells = 60; four replicates each) acute hypotonic shocks, respectively. Before time 0 min, all protoplasts were at steady state in 0.4 M sorbitol. Curves for individual conditions in panels (F) and (G) are plotted in Figure 4—figure supplements 1C,D, respectively. (H) Summary of exocytic rates for wild-type and pil1Δ protoplasts before and after hypotonic shock. (CH) Data from at least three independent experiments were pooled together to produce each curve. p-values: non-significant (ns), p>0.05; *p≤0.05; ***p≤0.001.

Staining of wild-type fission yeast with 20 µM FM4-64 in EMM5S (Figure 4B) showed that after a brief phase of rapid staining of the cell surface, the total cell fluorescence intensity grows roughly linearly for at least 20 min, and the slope of the normalized intensity corresponds to the exocytosis rate as a percentage of the plasma membrane surface area per unit of time (see 'Materials and methods') (Gauthier et al., 2009; Smith and Betz, 1996; Vida and Emr, 1995). Using this method, we measured that wild-type walled cells at steady state in EMM5S exocytose 4.6% of their plasma membrane surface area per minute (Figure 4B). FM4-64 staining did not seem to affect the endocytic and exocytic membrane trafficking of yeast cells, since stained vesicles were successfully released after washing cells with fresh media (Figure 4B).

We measured the exocytosis rates in the conditions that had the largest effects on the rates of endocytosis while keeping most cells alive, i.e., we used protoplasts at steady state in 0.4 M and performed a ΔC = −0.2 M shock for wild-type and ΔC = −0.05 M shock for pil1Δ. At steady state in 0.4 M sorbitol (Figure 4F and H), wild-type protoplasts had an exocytosis rate similar to walled cells in EMM5S in 0 M sorbitol (k0-5=4.4 ± 0.2% min−1). After a ΔC = −0.2 M shock, the exocytosis rate increased by 41% (k0-5=6.2 ± 0.4% min−1). At steady state in 0.4 M sorbitol (Figure 4G and H), the exocytosis rate of pil1Δ protoplasts was higher than that for walled cells in 0 M sorbitol (k0-5=6.2 ± 0.4% min−1). After a ΔC = −0.05 M shock, the exocytosis rate increased modestly (k0-5=6.8 ± 0.5% min−1). Therefore, in both wild-type and pil1Δ protoplasts, an acute hypotonic shock leads to an increased exocytosis rate, which increases surface area and likely reduces membrane tension. The more modest change in exocytosis rate in pil1Δ protoplasts than in wild-type cells highlights the role of eisosomes in buffering the change in the exocytosis rate in response to changes in osmolarity and membrane tension.

We wondered whether these changes in exocytosis rate also happen in walled cells. First, we measured exocytosis rate at steady state in solutions with different molarities and found that the rates were smaller than in protoplasts (Figure 4C–E). The exocytosis rate of wild-type walled cells at steady state in 1.2 M sorbitol (k0-5=3.1 ± 0.1 % min−1, Figure 4C and E) was 35% smaller than in 0 M sorbitol (k0-5=4.8 ± 0.1 % min−1, Figure 4B). In addition, in pil1Δ walled cells, the exocytosis rate of walled cells lacking eisosomes in 1.2 M sorbitol was only slightly smaller than wild-type cells in the same conditions (k0-5=2.6 ± 0.1% min−1, Figure 4D and E). After hypotonic shocks, the change in exocytosis rate in walled cells was very limited (Figure 4C–E). In fact, our strongest hypotonic shock of ΔC = −1.2 M did not significantly increase the exocytosis rate of wild-type or pil1Δ walled cells (Figure 4E). These data corroborate our previous finding that the cell wall limits but does not completely cancel the effect of hypotonic shocks in intact cells. In addition, they also demonstrate that eisosomes are involved in the regulation of the exocytosis rate.

Inhibition of exocytosis decreased the survival rate of protoplasts under acute hypotonic shocks and inhibition of endocytosis increased their survival rate

To further test our hypothesis that reducing the endocytosis rate and increasing the exocytosis rate help regulate membrane tension after a hypotonic shock, we blocked endocytosis or exocytosis with drugs and measured the survival rates of cells. We hypothesized that inhibition of endocytosis or exocytosis would have opposite effects on the survival of protoplasts under acute hypotonic shocks. Specifically, inhibition of endocytosis would help retain membranes on the surface of protoplasts, thereby reducing the probability of membrane rupture. Conversely, inhibition of exocytosis would reduce the transfer of membranes from intracellular vesicles to the surface of protoplasts, exacerbating the lack of plasma membrane in the face of imminent protoplast expansion. To observe the largest effects, we used pil1Δ protoplasts under ΔC = −0.2 M shock and exposed the cells to either Latrunculin A (LatA) or Brefeldin A (BFA) for 30 min before the shocks.

Blocking exocytosis with BFA increased the death rate of protoplasts after hypotonic shocks, confirming our hypothesis (Figure 5). Blocking actin assembly, and therefore endocytosis, with LatA made the protoplasts more resistant starting 4 min after the hypotonic shock, also confirming our hypothesis. Note that LatA treatment made the protoplasts less resistant to shock in the initial 2 min after the hypotonic shock, which seems in contradiction with our hypothesis. However, it is possible that prolonged treatment with LatA had other unidentified effects on protoplast survival or may indirectly affect the exocytosis rate since LatA affects all actin structures in the cell, including actin cables, which are needed for the transport of exocytic vesicles (Lo Presti et al., 2012).

Inhibition of exocytosis but not endocytosis decreased the survival rate of protoplasts under acute hypotonic shocks.

(A) pil1Δ protoplasts initially at steady state in 0.4 M sorbitol and supplemented with 25 μM Latrunculin A (LatA) (middle column) or 2 mM Brefeldin A (BFA) (right column) or nothing (control, left column) were submitted to a ΔC = −0.2 M hypotonic shock (t = 0 min). Cells were considered dead if they contained large amounts of intracellular red fluorescence from the FM4-64 dye, which is the consequence of a rupture of the plasma membrane. Scale bar: 10 µm. (B) Survival rate for all conditions. Black: control (N = 114); blue: 2 mM BFA (N = 83); red: 25 μM Latrunculin A (N = 70). Data were pooled from two independent experiments and plotted as Kaplan-Meier survival curves. Error bars: standard error of the mean by the Greenwood formula. *p≤0.05, logrank test.

Actin dynamics during clathrin-mediated endocytosis in wild-type walled cells is robust over a wide range of chronic and acute changes in media osmolarity

Next, we wondered how the actin machinery adapts to different changes in osmotic pressure and membrane tension. To monitor actin dynamics during CME, we imaged fission yeast cells expressing Fim1p-mEGFP (Figure 6A and B). Fimbrin is a bona fide marker for endocytosis in yeast since it has spatial and temporal co-localization with the classical endocytic marker End4p (the fission yeast homolog of mammalian Hip1R and budding yeast Sla2) during endocytosis (Figure 6—figure supplements 1A,B). Fimbrin’s time of appearance, disappearance, peak number of molecules, and spatial localization follow those of actin in wild-type cells and all mutants tested so far (Arasada et al., 2018; Berro and Pollard, 2014b; Chen and Pollard, 2013; Sirotkin et al., 2010). Fimbrin is the most abundant endocytic protein that is fully functional when tagged with a fluorescent protein at either N- or C-terminal. Tagged fimbrin is a more robust marker for actin dynamics than tagged actin or actin-binding markers such as LifeAct or calponin-homology domains, because they require over-expression, which is difficult to control precisely in fission yeast and potentially creates artifacts (Courtemanche et al., 2016; Suarez et al., 2015). Fimbrin is also a central player in force production during CME in yeast (Ma and Berro, 2019; Ma and Berro, 2018; Picco et al., 2018; Planade et al., 2019). We optimized our imaging protocols, and improved tracking tools and temporal super-resolution alignment methods (Berro and Pollard, 2014a) to (a) easily collect hundreds of endocytic events in an unbiased manner and (b) achieve high reproducibility between different samples, fields of view, and days of experiment (Figure 6C, Figure 6—figure supplements 2A). These improvements in our quantitative microscopy protocol have allowed us to detect small differences between mutants or conditions that would have been missed with previous methods. We confirmed that Fim1p accumulates at endocytic sites for about 10 s and then disassembles while the vesicle diffuses away from the plasma membrane (Figure 6C, Figure 6—figure supplements 2A; Sirotkin et al., 2010; Skau et al., 2011). As a convention, the peak number of Fim1p molecules is set to time 0 s and corresponds to vesicle scission in intact wild-type cells (Berro and Pollard, 2014a; Berro and Pollard, 2014b; Sirotkin et al., 2010).

Figure 6 with 4 supplements see all
CME in walled cells is robust over a wide range of conditions.

(A) Schematic of the plasma membrane deformations and the main components of the actin machinery during clathrin-mediated endocytosis (CME). Fimbrin (Fim1p, red) crosslinks actin filaments (blue) present at endocytic sites and is used as a proxy to monitor the amount of actin assembled (Figure 6—figure supplement 1). (B) Wild-type walled fission yeast cell expressing Fim1p-mEGFP (inverted contrast). Top: cell outlined with a dashed line; scale bar: 5 µm. Bottom: montage of a representative CME event. The interval between each frame is 1 s. (C) The number of molecules of Fim1p-mEGFP detected, tracked, and aligned with temporal super-resolution (Berro and Pollard, 2014a) is highly reproducible between fields of view (one-way ANOVA on the number of molecules at time 0 s, p=0.74). Each curve with a dark color represents the average of several endocytic events from a different field of view of the same sample (N ≥ 64), and the light colors are the 95% confidence intervals. For each average curve, the peak value corresponds to time 0 s, when vesicle scission happens. (D) Number of molecules of Fim1p-mEGFP in wild-type walled cells at steady state in media supplemented with different sorbitol concentrations. There is no statistically significant difference in the number of molecules at time 0 s between the three conditions (one-way ANOVA, p=0.29). N ≥ 388. (E) Number of molecules of Fim1p-mEGFP in pil1Δ walled cells at steady state in media supplemented with different sorbitol concentrations (N ≥ 342). The difference in the number of molecules at time 0 s between all pairs of conditions is statistically significant (one-way ANOVA, p<10−5). (F) Timeline of the hypotonic shock experiments and notations. By convention, hypotonic shocks start at time 0 min and are defined by the difference in concentrations of sorbitol in the steady-state media before the shock (CSS) and after the hypotonic shock (Cfin), ΔC=Cfin-CSS. Data for a given time point correspond to endocytic events happening within 1 min after this time point (e.g., the data at t = 0 min correspond to endocytic events happening between 0 and 1 min after the shock). These time intervals are represented by gray bars on the time axis. (G) Number of molecules of Fim1p-mEGFP for wild-type walled cells initially at steady state in 1.2 M sorbitol and after an acute osmotic shock of ΔC = −1.2 M. There is no statistically significant difference in the number of molecules at time 0 s between the three conditions (one-way ANOVA, p=0.95). Black: steady state in 1.2 M sorbitol; light to dark blue in top panel: 0, 2, 4, and 6 min after the acute hypotonic shock (N ≥ 103). (H) Number of molecules of Fim1p-mEGFP in pil1Δ walled cells before and after an acute osmotic shock (ΔC = −1.2 M). The difference in the number of molecules at time 0 s between all pairs of conditions is statistically significant (one-way ANOVA, p<0.03) except between 0 and 2 min after the shock (one-way ANOVA, p=0.18). Black: steady state in 1.2 M sorbitol before the hypotonic shock (N = 583); light to dark red in top panel: 0, 2, and 4 min after the acute hypotonic shock (N ≥ 145). (C, D, E, G, and H) Dark colors: average; light colors: average ± 95% confidence interval. The number of molecules and speed versus time for each condition are plotted separately in Figure 6—figure supplements 2C, Figure 6—figure supplements 3D,G, and Figure 6—figure supplement 4E,H (E and H). The numbers of endocytic events used in each curve are given in Supplementary file 1d (C), Supplementary file 1e (D), Supplementary file 1f (E), Supplementary file 1g (G), and Supplementary file 1h (H).

For all tested osmolarities at steady state in walled wild-type cells, we observed no significant difference in the dynamics of fimbrin recruitment or disassembly, maximum molecule number, or endocytic patch movements (Figure 6D). Our results indicate that wild-type walled cells have adaptation mechanisms for chronic exposure to a wide range of osmolarities, which allow them to perform CME in a highly reproducible manner.

We then tested the robustness of the endocytic actin machinery when cells experienced a hypotonic shock, which aimed to abruptly increase the tension of their plasma membrane. To observe the highest possible effect, we imaged cells grown at steady state in 1.2 M sorbitol and rapidly exchanged the media with a buffer free of sorbitol (Figure 6F and G), therefore performing an acute hypotonic shock of ΔC = −1.2 M. Despite the high hypotonic shock, which represents a ~3 MPa drop in pressure, CME proceeded quite similarly to steady-state conditions (Figure 6G, Figure 6—figure supplements 3). The maximum number of fimbrin proteins was the same before and after the hypotonic shock, but fimbrin assembly and disassembly were ~15% faster after the shock.

Eisosomes mitigate the response of the endocytic machinery to acute and chronic changes in media osmolarity

The robustness of the endocytic process under a wide range of chronic and acute exposure to different media osmolarities is consistent with our previous results showing that fission yeast cells rapidly regulate plasma membrane tension. To better understand the role of eisosomes and amplify the change in membrane tension after hypotonic shocks, we repeated our experiments in pil1Δ cells that lack eisosomes (Figure 6E).

Dynamics of Fim1p during CME for wild-type and pil1Δ walled cells at steady state in media free of sorbitol was identical (Figure 6—figure supplement 1C). However, at steady state in media with high sorbitol concentrations, cells lacking eisosomes recruited slightly fewer fimbrin molecules to endocytic patches than wild-type cells (Figure 6E). The maximum number of Fim1p assembled at CME sites in pil1Δ cells in buffers containing 0.8 and 1.2 M sorbitol was 10 and 17% lower, respectively. Within the first 2 min of an acute hypotonic shock from 1.2 M sorbitol to 0 M (ΔC = −1.2 M), the maximum number of Fim1p increased by 30%, while its timing was shortened by ~30% compared to steady state (Figure 6H). Four minutes after the hypotonic shock, the dynamics of fimbrin stabilized at its steady-state dynamics in 0 M sorbitol (Figure 6E and H). Overall, our data show that the endocytic actin machinery in cells lacking eisosomes is more sensitive to acute and chronic changes in media osmolarity than in wild-type cells, consistent with a role for eisosome in regulating membrane tension at endocytic sites.

CME in protoplasts is sensitive to chronic changes in osmolarity

Endocytosis in wild-type protoplasts at steady state in a medium containing 0.4 or 0.8 M sorbitol was able to proceed normally by recruiting almost the same number of fimbrin molecules as in walled cells, but with a slightly longer timing (Figure 7—figure supplement 1B). In contrast, in a medium with 1.2 M sorbitol, the timing of fimbrin recruitment was dramatically longer, and endocytosis failed to proceed normally, as reported by the virtually null speed of patches during the entire time fimbrin was present at the endocytic site (Figure 7—figure supplement 1B). Cells lacking eisosomes had very similar phenotypes but endocytosis started failing at 0.8 M sorbitol (Figure 7—figure supplement 1C).

At 0.25 M sorbitol, both wild-type and pil1Δ protoplasts were able to perform endocytosis but required a larger amount of Fim1p (Figure 7—figure supplements 1B,C). In these conditions, the eisosomes covered only half of the plasma membrane surface area they cover at 0.4 M sorbitol (Figure 1B and C), and our data suggest that the plasma membrane was under high tension (Figure 1G). This result indicates that during CME, the actin machinery is able to adapt to mechanical cues by mechanisms that are independent of the cell wall.

For both wild-type and pil1Δ protoplasts in 0.4 M sorbitol, the temporal evolution of the number of fimbrin molecules and the speed of patches were close to the same metrics measured in walled cells in media without sorbitol (Figure 7—figure supplements 1B,C). These results suggest that the osmotic pressure at these concentrations, which is equivalent to a pressure of 1 MPa, is close to the naturally maintained turgor pressure of walled fission yeast cells, in good agreement with previous measurements (Minc et al., 2009). Therefore, to keep protoplasts in conditions close to walled cells, the steady-state media used in our following experiments on protoplasts contained 0.4 M sorbitol.

The endocytic actin machinery rapidly adapts to increases in membrane tension

To characterize the adaptation of the endocytic actin machinery to a rapid increase in turgor pressure and membrane tension, we repeated our acute hypotonic shocks (ΔC = −0.05, −0.1, or −0.2 M) on protoplasts initially at steady state in media containing 0.4 M sorbitol. After low (ΔC = −0.05 M) and medium (ΔC = −0.1 M) acute shocks in wild-type protoplasts, we did not observe any stalled endocytic events – when cells started the recruitment of the actin machinery, endocytosis proceeded to successful completion (Figure 7A–C and Figure 7—figure supplements 5). The recruitment of fimbrin did not significantly change over time. In contrast, 2 min after a ΔC = −0.2 M shock, endocytic sites recruited 20% more fimbrin, and it took ~25% longer to perform endocytosis (Figures 7C,D and Figure 7—figure supplements 5). The actin machinery restored its steady-state behavior less than 4 min after the shock (Figure 7D).

Figure 7 with 7 supplements see all
The actin endocytic machinery adapts to increases of membrane tension in protoplasts.

(A and B) Representative wild-type protoplasts expressing Fim1-mEGFP (inverted contrast) at steady state in 0.25 M sorbitol (A, left panel) and immediately after (0 min) an acute osmotic shock of ΔC = −0.1 M (B, left panel). Right panels: kymographs of the fluorescence under the yellow lines in the left panels. Black dashed lines: protoplast outline. Scale bars: 5 µm. (C and F) Number of Fim1p-mEGFP molecules in wild-type (C) and pil1Δ (F) protoplasts at steady state in 0.4 M sorbitol (purple), 0 min (brown) and 10 min (orange) after a hypotonic shock of ΔC = −0.05 M (left panels), ΔC = −0.1 M (middle panels), and ΔC = −0.2 M (right panels), N ≥ 95. Data for each condition are plotted separately in Figure 7—figure supplement 3 (wild type) and Figure 7—figure supplement 4 (pil1Δ). The speeds of Fim1p-mEGFP for each condition are shown in Figure 7—figure supplement 5 (wild type) and Figure 7—figure supplement 6 (pil1Δ). The numbers of endocytic events used in each curve are given in Supplementary file 1i. Note that the large majority of pil1Δ protoplasts were too damaged or dead 2 min after hypotonic shocks larger than or equal to ΔC = −0.1 M to allow us to track enough endocytic events and produce a curve (Figure 2B,C and Figure 7—figure supplement 1). In panel (C), the difference in the number of molecules at time 0 s at steady state and 0 min after the shock is statistically significant for all shocks (one-way ANOVA, p<0.03), and the difference between steady state and 10 min after the shock is not statistically significant (one-way ANOVA, p>0.2; details in the data file for C). In panel (F), the difference at steady state and 0 min after the shock is statistically significant for all shocks (one-way ANOVA, p<10−5; details in the data file for Figure 5F). (D) Temporal adaptation of the peak number of Fim1p-mEGFP in wild-type protoplasts initially at steady state in 0.4 M sorbitol and 0–10 min after a ΔC = −0.2 M osmotic shock. The condition for this figure is the same as the condition with the blue star in (C). The difference between steady state and 0 or 2 min after shock is statistically significant (one-way ANOVA, p<10−3; details in the data file for D). The difference between steady state and 4, 6, 8, and 10 min after shock is not statistically significant (one-way ANOVA, p>0.2; details in the data file for D). (E) Montage of representative endocytic events (Fim1-mEGFP, inverted contrast) in pil1Δ protoplasts (one frame per second) at steady state in 0.4 M sorbitol (first row) and immediately after (0 min) hypotonic shocks of ΔC = −0.05 M (second row), ΔC = −0.10 M (third row), and ΔC = −0.20 M (fourth row). (G) Temporal adaptation of the peak number of Fim1p-mEGFP in pil1Δ protoplasts initially at steady state in 0.4 M sorbitol and 0–10 min after a ΔC = −0.05 M shock. The condition in this figure is the same as the condition with the red star in (F). The difference between steady state and 0, 2, 4, 6, or 8 min after shock is statistically significant (one-way ANOVA, p<0.01; details in the data file for F). The difference between steady state and 10 min after shock is not statistically significant (one-way ANOVA, p>0.3; details in the data file for F). (D and G) Error bars are 95% confidence intervals. The numbers of endocytic events at each time point are given in Supplementary file 1j. (H and I) Number of molecules of Fim1p-mEGFP for wild-type (H) and pil1Δ (I) protoplasts at steady state in 0.25 M sorbitol (purple dashed) and immediately after (0 min) a hypotonic shock of ΔC = −0.1 M (brown), N ≥ 67. The difference in the number of molecules at time 0 s at steady state and 0 min after the shock is statistically significant for all conditions (one-way ANOVA, p<10−16). The speed data for each condition are plotted in Figure 7—figure supplement 7. The numbers of endocytic events used in each curve are given in Supplementary file 1k. The survival rates for the wild-type and pil1Δ protoplasts in these conditions are plotted in Figure 2D.

We repeated these experiments with pil1Δ protoplasts to eliminate the role of eisosomes in the reduction of membrane tension during hypotonic shocks. Immediately (0 min) after the lowest hypotonic shock tested (ΔC = −0.05 M), fimbrin recruitment took slightly longer and the number of proteins recruited was higher than at steady state (Figure 7E–G and Figure 7—figure supplements 6). While fimbrin restored its steady-state dynamics in less than 4 min after a high acute hypotonic shock (ΔC = −0.2 M) in wild-type protoplasts (Figure 7D), recovery of fimbrin dynamics to its steady-state behavior in pil1Δ protoplasts occurred over 10 min, even for the most modest hypotonic shock, ΔC = −0.05 M (Figure 7G). The changes in fimbrin dynamics in pil1Δ protoplasts became increasingly larger for ΔC = −0.1 and −0.2 M hypotonic shocks – endocytic sites assembled a peak number of fimbrin, respectively, ~25 and ~50% larger and took ~85 and ~50% longer.

Wild-type protoplasts at steady state in 0.25 M sorbitol contained significantly fewer assembled eisosomes despite expressing normal amounts of Pil1p (Figure 1B, C and E). We took advantage of this condition to test whether the absence of eisosome structures at the plasma membrane, and not the absence of the protein Pil1p, is responsible for changes in actin dynamics after an acute hypotonic shock. We subjected wild-type protoplasts at steady state in 0.25 M sorbitol to an acute hypotonic shock of ΔC = −0.1 M (Figure 7H and I). Two minutes after the shock, endocytic sites accumulated 73% more fimbrin and took ~60% longer (Figure 7H and Figure 7—figure supplement 7A). This behavior was nearly identical to fimbrin dynamics in pil1Δ protoplasts under the same conditions (Figure 7I and Figure 7—figure supplement 7B).

Altogether, these experiments demonstrate that (a) the endocytic actin machinery adapts to compensate the increase in membrane tension and (b) actin dynamics restores its steady-state behavior within a few minutes, providing the protoplasts survived the hypotonic shock.

Discussion

Mechanisms of tension regulation and homeostasis of the plasma membrane

Our results demonstrate that the regulation of membrane tension in hypotonic environment is performed via a combination of at least three mechanisms: the mechanical protection by the cell wall, the disassembly of eisosomes, and the temporary shift in the balance between endocytosis and exocytosis (Figure 8). Our data indicate that all three mechanisms are used in parallel since wild-type walled cells are less sensitive to acute hypotonic shocks than wild-type protoplasts and pil1Δ walled cells, and they experience a temporary decrease in their endocytic density for about 2 min after the shock. In addition, our data allow us to estimate the relative contribution of each mechanism in the regulation of membrane tension.

Schematic of the adaptation of fission yeast endocytosis, exocytosis, and eisosome after acute hypotonic shock-induced increase in membrane tension.

(A) In an isotonic solution, endocytosis and exocytosis rates are largely balanced, and proteins including Pil1p are assembled at the plasma membrane to form eisosomes. Actin is recruited to endocytic sites to provide the forces needed to reshape the membrane under normal membrane tension. When present, cell wall makes fission yeast cell resistant to significant changes in the osmolarity of extracellular solution. (B) Acute hypotonic shock results in an increase in membrane tension, which leads to a decrease in endocytosis rate, an increase in exocytosis rate, and a rapid disassembly of eisosomes, within ~2 min. The proteins of the actin machinery are recruited in larger amounts to endocytic sites to provide larger forces for successful endocytosis under increased membrane tension. Failure of adaptation to the increase in membrane tension leads to membrane rupture and cell death in both protoplasts and walled cells.

The cell wall provides the largest protection during chronic and acute hypotonic shocks. Wild-type walled cells are virtually insensitive to osmotic downshifts, and pil1Δ walled cells are much less sensitive than pil1Δ protoplasts. Removal of the cell wall dramatically affects actin dynamics at endocytic sites and eisosome assembly at the plasma membrane (Figures 1B, C and 7 – Supplement 1B and 7 – Supplement 1C), and greatly increased the effect of hypotonic shocks on exocytosis (Figure 4). It is surprising that endocytosis in protoplasts still proceeds in media with osmolarity as low as 0.25 M, where a large fraction of eisosomes is disassembled. In fact, the actin endocytic machinery can overcome membrane tensions high enough to rupture the plasma membrane since we did not see stalled actin patches, or actin comet tails, in any of our experiments. Our results contrast with recent data in S. cerevisiae (Riggi et al., 2019) where endocytosis is blocked and actin comet tails are formed within 2 min of a hypotonic shock. These differences may highlight species specificities.

Our results add to a growing body of evidence that eisosomes play a critical role in the regulation of membrane tension and membrane integrity through dynamic remodeling and scaffolding of the plasma membrane (Kabeche et al., 2015; Moseley, 2018). Endocytosis in wild-type walled cells is not sensitive to chronic or acute hypotonic changes, whereas pil1Δ walled cells’ endocytosis is (Figure 6). Conversely, exocytosis seems to respond more strongly to acute hypotonic shocks in wild-type walled cells than in pil1Δ walled cells (Figure 4). The protective role of eisosomes is even more striking in protoplasts under acute hypotonic shocks. Wild-type protoplasts whose plasma membrane is covered with eisosomes are largely insensitive to increases in membrane tension, whereas protoplasts with little to no eisosomes are extremely sensitive to increases in membrane tension and their plasma membrane is easily damaged (Figure 2A–C). Eisosomes retain this protective function even in walled cells, which becomes evident when cells are put under repeated osmolarity shocks (Figure 2E–G). Our micropipette aspiration experiments also demonstrate that eisosomes are critical to keep membrane tension low during an acute hypotonic shock. Therefore, our data indicate that membrane tension is decreased via the disassembly of eisosomes, through release of excess membrane surface area. Assuming eisosomes are hemi-cylinders with a diameter of ~50 nm and cells contain 1.6 μm of eisosomes per μm2 of plasma membrane on average, total eisosome disassembly could release about 5% of the total surface area of the plasma membrane over ~3 min after a hypotonic shock (Kabeche et al., 2015), although a mild shock of ΔC = −0.2 M disassembled close to ~50% eisosomes over 5 min, or about 2.5% of the surface area of the plasma membrane (Figure 1H and I). Single-molecule imaging in our lab demonstrated that at steady state, Pil1p undergoes rapid exchange at the eisosome ends (Lacy et al., 2017), potentially providing a convenient route for rapid eisosome disassembly, analogous to filament depolymerization, in combination with eisosome breaking. Disassembled eisosome components have altered phosphorylation level or sub-cellular localization, which potentially relays the signaling from eisosome integrity to endocytosis and/or exocytosis (Riggi et al., 2018; Walther et al., 2007), possibly via TORC2 (Riggi et al., 2019).

Our study highlights a third mechanism to reduce membrane tension by increasing the surface area of the plasma membrane via a temporary reduction in the endocytosis rate and an increase in the exocytosis rate. Using our data, we estimate that cells endocytose about 2% of their surface area per minute through CME, confirming our previous measurements (Berro and Pollard, 2014a; Berro and Pollard, 2014b). During acute hypotonic shocks, a reduction of the endocytosis rate plus an increase in the exocytosis rate for a few minutes would allow for a net addition of surface area to the plasma membrane. For example, in pil1Δ protoplasts, initially at steady state in 0.4 M sorbitol, the endocytosis rate is reduced by ~25% for ~10 min after an acute hypotonic shock of ΔC = −0.05 M, while the exocytosis rate increased by ~10%. The net surface area added over that period by reduction in endocytosis and increase in exocytosis corresponds to a 5% + 6% = 11% increase in the protoplast surface area, close to the ~12% surface area increase we measured. These results confirm and quantify previous reports of control of surface tension by increasing the surface area via a modulation of endocytosis and exocytosis rates in other eukaryotes (Apodaca, 2002; Homann, 1998; Morris and Homann, 2001). These estimates demonstrate that modulating the endocytosis and exocytosis rates is an efficient way to increase the surface area of the plasma membrane by large amounts, but this process is relatively slow compared to eisosome disassembly. The slowness of this process might explain why pil1Δ and pre-stretched wild-type protoplasts that have about half the normal amount of eisosomes on their surface do not survive even relatively small hypotonic shocks, being unable to provide enough membrane in a short amount of time to reduce the tension of their plasma membrane.

For the calculations presented above, we have assumed that the size of endocytic and exocytic vesicles remain constant in all osmotic conditions. However, further experiments will be necessary to validate or invalidate this assumption. In addition, our FM4-64 data do not allow us to distinguish between lipid addition to the plasma membrane via exocytosis or via a putative transfer of lipids by non-exocytic mechanisms (Reinisch and Prinz, 2021), and for simplicity and by lack of further evidence, we have discussed our data as an increase in the exocytosis rate. Not much is known about lipid transfer proteins at the plasma membrane, and further experiments will be necessary to determine whether the activity of these proteins is enhanced by abrupt increases in membrane tension or hypotonic shocks.

Molecular mechanisms driving the adaptation of the actin endocytic machinery and the rate of endocytosis under various membrane tensions

Our data demonstrate that fission yeast CME is very robust and can proceed in a wide range of osmolarities and membrane tensions. Even cells devoid of a cell wall and eisosomes were able to perform endocytosis after an acute change in membrane tension, as long as their plasma membrane was not damaged and cells remained alive. Even in the most extreme conditions tested, i.e., cells devoid of a cell wall and lacking the majority of their eisosomes, the peak number of molecules and timing of fimbrin at endocytic sites were only two times larger than those observed in wild-type walled cells.

Under conditions where membrane tension and turgor pressure were significantly increased, we observed that the endocytic actin machinery took longer and assembled a larger number of fimbrin molecules to successfully produce endocytic vesicles. This effect increased with increasing membrane tension, up to tensions high enough to rupture the cell plasma membrane. This result strongly supports the idea that the actin machinery provides the force that counteracts membrane tension and turgor pressure and deforms the plasma membrane into an endocytic pit.

The precise molecular mechanism that regulates this enhanced assembly remains to be uncovered. Our data suggest that actin dynamics is controlled via a mechanical or geometrical regulation, where actin assembles until the plasma membrane is deformed and pinched off. An alternative, and non-mutually exclusive, hypothesis is that the activity and/or recruitment of proteins upstream of the actin nucleators may be enhanced by increased membrane tension. A third hypothesis is that the decrease in the number of endocytic events after an increase in membrane tension leads to an increase in the concentration of endocytic proteins in the cytoplasm, which can then enhance the reactions performed at the endocytic sites. Sirotkin et al., 2010 measured that 65–85% of the total cellular content of key proteins involved in the endocytic actin machinery is localized to endocytic sites at any time. A 20% decrease in the number of endocytic sites would increase their cytoplasmic abundance by roughly 40–80%. This percentage is larger than the volume changes we measured, resulting in a net increase in the cytoplasmic concentration of these proteins, which would allow larger amounts of protein to assemble at the endocytic sites.

Conversely, the decreased endocytosis rate could be attributed to the larger number of endocytic proteins assembled at each endocytic site, which would decrease their cytoplasmic concentration. Indeed, Burke et al., 2014 showed that modulating actin concentration modulates the number of endocytic sites in the same direction. However, it is more likely that one or several early endocytic proteins are sensitive to membrane tension, and they either fail to bind the plasma membrane or prevent the triggering of actin assembly when membrane tension is high. This idea would be consistent with results from mammalian cells demonstrating that the proportion of stalled clathrin-coated pits increases when membrane tension increases (Ferguson et al., 2017). In addition, several endocytic proteins that arrive before or concomitantly with the activators of the actin machinery contain BAR domains (such as Syp1p, Bzz1p, and Cdc15p), and other members of this domain family (which also includes Pil1p) have been shown to bind membranes in a tension-sensitive manner. Further quantitative studies of early endocytic proteins will help uncover the validity and relative contributions of each one of these hypotheses.

We expect our results to be relevant to the study of CME and membrane tension regulation in other eukaryotes since the molecular machineries for endocytosis, exocytosis, and osmotic response are highly conserved. In addition, regulation of membrane tension and CME are particularly critical during cell polarization (Mostov et al., 2000), during neuron development and shape changes (Urbina et al., 2018), and at synapses where large pools of membranes are added and retrieved on a very fast time scale (Nicholson-Fish et al., 2016; Watanabe and Boucrot, 2017).

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Schizosacchoromyces pombe)Fim1p-mEGFPBerro and Pollard, 2014aSpJB57fim1-mEGFP-NatMX6 ade6-M216 his3-Δ1 leu1-32 ura4-Δ18 h+
Strain, strain background (Schizosacchoromyces pombe)Pil1p-mEGFPLacy et al., 2017SpJB204pil1-mEGFP-kanMX6 ade6-M216 his3-Δ1 leu1-32 ura4-Δ18 h-
Strain, strain background (Schizosacchoromyces pombe)pil1Δ Fim1p-mEGFPThis studySpJB234pil1Δ fim1-mEGFP-NatMX6 ade6-M216 his3-Δ1 leu1-32 ura4-Δ18 h-
Strain, strain background (Schizosacchoromyces pombe)mScarlet-I-End4p mEGFP-Fim1pThis studySpJB566mScarlet-I-end4 mEGFP-fim1 fex1Δ fex2Δ ade6-M216 his3-Δ1 leu1-32 ura4-Δ18 h-
Chemical compound, drugFM4-64Biotium, Fremont, CA
Chemical compound, drugSulforhodamine BMP Biomedicals LLC, Santa Ana
Chemical compound, drugLatrunculin AMillipore, MA
Chemical compound, drugBrefeldin ASanta Cruz Biotechnology Inc, TX
Software, algorithmMatlabMathworksR2016a, R2018b
Software, algorithmFIJI ImageJSchindelin et al., 2012; Schneider et al., 2012
Software, algorithmTrackmateTinevez et al., 2017
Software, algorithmPatchTrackingToolsThis study, Berro, 2018Version 2015.12.16https://bitbucket.org/jberro/patchtrackingtools/src/stable/, copy archived at swh:1:rev:226505c08c2197584c1299462b5de85433d1fcf1.
Software, algorithmPatchTrackingDataPostprocessing.DeterministicThis study, Berro, 2021#1B100-4https://bitbucket.org/jberro/patchtrackingdatapostprocessing.deterministic/src/master/, copy archived at swh:1:rev:3d96cd606f47a692dd6db55ba2aab0a7f6e0c4f2.
OtherGlass micropipetteWorld Precision Instruments, SarasotaVersion 160801
Chemical compound, drugβ-caseinMillipore-SigmaSaint-Louis
Software, algorithmGraphPad PrismSoftware, La Jolla, CAversion 8.4.2

Yeast strains and media

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The S. pombe strains used in this study are listed in the Key Resources Table. Yeast cells were grown in YE5S (yeast extract supplemented with 0.225 g/l of uracil, lysine, histidine, adenine, and leucine), which was supplemented with 0–1.2 M D-sorbitol, at 32°C in the exponential phase for about 18 hr. Cells were washed twice and resuspended in filtered EMM5S (Edinburgh Minimum Media supplemented with 0.225 g/l of uracil, lysine, histidine, adenine, and leucine), which was supplemented with the same concentration of D-sorbitol, at least 10 min before imaging so they can adapt and reach steady state.

Protoplast preparation

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S. pombe cells were grown in YE5S at 32°C in exponential phase for about 18 hr. 10 ml of cells were harvested and washed two times with SCS buffer (20 mM citrate buffer, 1 M D-sorbitol, pH = 5.8), and resuspended in SCS supplemented with 0.1 g/ml Lallzyme (Lallemand, Montreal, Canada) (Flor-Parra et al., 2014). Cells were incubated with gentle shaking for 10 min at 37°C in the dark except for experiments in Figure 5, where cells were digested at room temperature with gentle shaking for 30 min in the presence of inhibitors. The resulting protoplasts were gently washed twice in EMM5S with 0.25–1.2 M D-sorbitol, spun down for 3 min at 960 rcf between washes, and resuspended in EMM5S buffer supplemented with 0.25–1.2 M D-sorbitol at least 10 min before imaging so that they can adapt and reach steady state.

Microscopy

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Microscopy was performed using a spinning disk confocal microscope, built on a TiE inverted microscope (Nikon, Tokyo, Japan), equipped with a CSU-W1 spinning head (Yokogawa Electric Corporation, Tokyo, Japan), a x100/1.45NA phase objective, an iXon Ultra888 EMCCD camera (Andor, Belfast, UK), and the NIS-Elements software v. 4.30.02 (Nikon, Tokyo, Japan). The full system was switched on at least 45 min prior to any experiment to stabilize the laser power and the room temperature. Cells were loaded into commercially available microfluidic chambers for haploid yeast cells (Y04C-02-5PK; Millipore-Sigma, Saint-Louis) for the CellASIC ONIX2 microfluidics system (Millipore-Sigma). Each field of view was imaged for 60 s, and each second, a stack of six z-slices separated by 0.5 µm was imaged. The microscope was focused such that the part of the cell closest to the coverslip was captured.

Acute hypotonic shocks

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Walled cells or protoplasts were first imaged in their steady-state media (EMM5S supplemented with 0–1.2 M D-sorbitol). The steady-state media were exchanged with media supplemented with a lower D-sorbitol concentration (the concentration difference is noted, ΔC), with an inlet pressure of 5 psi. These hypotonic shock media were labeled with 6.7 µg/ml of sulforhodamine B (MP Biomedicals LLC, Santa Ana), a red cell-impermeable dye that allowed us to (a) monitor the full exchange of the solution in the microfluidic chamber prior to image acquisition and (b) monitor the plasma membrane integrity of the cells after the shock. In each condition, the first movie was started when the sulforhodamine B dye was visible in the field of view. For clarity, this time point is labeled t = 0 min in all our figures, but note that we estimate it may vary by up to ~30 s between movies and conditions. We imaged cells by taking one stack of six z-slices per second for 60 s. After the end of each movie, we rapidly changed field of view and restarted acquisition 1 min after the end of the previous movie, so that movies started every 2 min after the acute hypotonic shock. Tracks from cells that contained red fluorescence from the sulforhodamine B dye were excluded from the analysis, because this indicated that cell membrane had been damaged.

Inhibition of endocytosis and exocytosis during acute hypotonic shocks

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Endocytosis or exocytosis was inhibited by including, respectively, 25 µM Latrunculin A (Millipore, MA) or 2 mM Brefeldin A (Santa Cruz Biotechnology Inc, TX) in the solution used to prepare the protoplasts and to perform the hypotonic shocks. Hypotonic shock solution also included 20 µM FM4-64 (Biotium, Fremont, CA) to stain dead protoplasts (Vida and Gerhardt, 1999; Figure 5A), and inlet pressure was set at 4 psi.

Measurement of the temporal evolution of the number of proteins and speed

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Movies were processed and analyzed using an updated version of the PatchTrackingTools toolset for the Fiji (Schindelin et al., 2012) distribution of ImageJ (Berro and Pollard, 2014a; Schneider et al., 2012). This new version includes automatic patch-tracking capabilities based on the Trackmate library (Tinevez et al., 2017) and is available on the Berro lab website http://campuspress.yale.edu/berrolab/publications/software/ and the lab bitbucket https://bitbucket.org/jberro/. Prior to any quantitative measurement, we corrected our movies for uneven illumination and camera noise. The uneven illumination was measured by imaging a solution of Alexa 488 dye, and the camera noise was measured by imaging a field of view with 0% laser power. We tracked Fim1-mEGFP spots with a circular 7-pixel diameter region of interest (ROI), and measured the temporal evolution of the fluorescence intensities and the position of the centers of mass. The spot intensity was corrected for cytoplasmic background using a 9-pixel median filter and was then corrected for photobleaching. The photobleaching rate was estimated by fitting a single exponential to the temporal evolution of the intensity of cytoplasmic ROIs void of any identifiable spots of fluorescence (Berro and Pollard, 2014a). Only tracks longer than 5 s and displaying an increase followed by a decrease in intensity were kept for the analysis. Individual tracks were aligned and averaged with the temporal super-resolution algorithm from Berro and Pollard, 2014a and post-processed to generate figures using the Matlab R2016a (Mathworks) scripts PatchTrackingDataPostprocessing.Deterministic, available on the Berro lab website http://campuspress.yale.edu/berrolab/publications/software/ and the lab bitbucket https://bitbucket.org/jberro/. In brief, this method realigns temporal signals that have low temporal resolution and where no absolute time reference is available to align them relatively to each other. It iteratively finds the temporal offset, which has a higher precision than the measured signal and minimizes the mean square difference between each measured signal and a reference signal. For the first round of alignments, the reference signal was one of the measurements. After each realignment round, a new reference was calculated as the mean of all the realigned signals, which is an estimator of the true underlying signal.

To control and calibrate the intensity of our measurements, we imaged wild-type walled cells expressing Fim1p-mEGP each imaging day. Intensities were converted into number of molecules with a calibration factor such that the peak intensity of our control strain corresponded to 830 molecules (Berro and Pollard, 2014a).

In all figures presenting the temporal evolution of the number of molecules or the speed, time 0 s corresponds to the time point when the number of molecules is maximum (also called the peak number). The ‘speed vs time’ plots help determine whether endocytosis completes normally in different conditions. At a given time point, the speed corresponds to the average movement of the endocytic structure in the plane of the membrane between two consecutive images. For an endocytic event that completes normally, the speed is close to 0 while the endocytic pit elongates before the vesicle is pinched off. Note that the speed is not exactly 0 because of localization errors and putative small movements of the endocytic structure in the plane of the membrane. The speed increases after the vesicle is pinched off and it diffuses freely in the cytoplasm. Since this movement is mostly diffusive, the standard deviations of the speeds are large.

Statistical tests between conditions were performed at time 0 s with a one-way ANOVA test using the number of tracks collected to build the figure. To avoid extrapolating the data, we compared the relative duration of assembly and disassembly between conditions using the time at which the average number of molecules reaches half the peak number.

Measurement of the density of CME events

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We used the S. pombe profiling tools for ImageJ (Berro and Pollard, 2014a) to measure the number of endocytic events at a given time in each cell. In brief, on a sum-projected z-stack, we manually outlined individual cells, and, for each position along the long axis of a cell, we measured the sum of fluorescence orthogonal to the long axis. We corrected the intensity profile in each cell for its cytoplasmic intensity and media fluorescence outside the cell. We estimated the number of patches in each cell by dividing the corrected fluorescence signal with the temporal average of the fluorescence intensity of one endocytic event. We calculated the linear density of endocytic events as the ratio between the number of endocytic events in a cell and its length.

We estimated the percentage of plasma membrane internalized by endocytosis per minute as follows. We measured ~120 endocytic sites per cell at a given time on average. Since the actin meshwork assembles and disassembles in about 20 s, we estimated that 360 endocytic vesicles are formed per minute. Assuming endocytic vesicles have a diameter of 50 nm, this corresponds to an ~2.8 μm2 area endocytosed per minute or ~2% of the total surface area, considering the average cell is around 12 μm long and has a 132 μm2 area of plasma membrane (assuming the cell shape is a cylinder capped by two hemispheres).

Measurement of the exocytosis rate with FM4-64 staining

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The exocytosis rate was measured by combining the acute hypotonic shock with FM4-64 staining, in a similar approach as has been reported (Gauthier et al., 2009; Smith and Betz, 1996; Vida and Emr, 1995). The cell impermeable dye FM4-64 (Biotium, Fremont, CA) was diluted to a final concentration of 20 µM in any of the media used. When cells are exposed to FM4-64, the dye rapidly stains the outer leaflet of the plasma membrane. Upon endocytosis, the dye is trafficked inside the cell without change in fluorescence. The total cell fluorescence intensity was measured after segmenting the cells by thresholding the fluorescence signal above background levels. The fluorescence intensity was normalized to the intensity reached at the end of the fast increase ~1 min after the dye was flowed in, which corresponds to the intensity of total surface area of the plasma membrane (Figure 4B). After this fast phase (<20 s), the fluorescence signal increased more slowly every time the unstained membrane was exposed to the cell surface by exocytosis. At a short time scale (~5–20 min depending on the exocytosis rate), recycling of stained membrane is negligible and one can assume that all exocytosed membrane is virtually unstained. Since the intensity at the beginning of the slow phase was normalized to 1, the slope of the linear increase of fluorescence is equal to the amount of membrane exocytosed per minute, expressed as a fraction of the surface area of the plasma membrane. For all measurements, images were taken at 5 s intervals at the middle plane of cells with the help of Perfect Focusing System (Nikon, Tokyo, Japan), with minimal laser excitation in order to reduce toxicity and photobleaching to negligible values. Curve fitting and slope calculation were performed in GraphPad Prism (GraphPad Software, La Jolla, CA).

Measurement of eisosomes’ density on the plasma membrane

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We imaged full cells expressing Pil1p-mEGFP by taking stacks of 0.5-µm-spaced z-slices. We corrected these z-stacks for uneven illumination and manually outlined individual cells to determine the surface area of each cell. To determine the total amount of eisosome-bound Pil1p-mEGFP, we subtracted the cytosolic intensity of Pil1-mEGFP using a pre-determined threshold and summed all the z-slices. We measured the mean membrane intensity of each cell on the thresholded sum-projection image. The eisosome density was determined by dividing this mean intensity by the surface area of each protoplast.

To quantify the relative changes in area fraction of eisosomes after acute hypotonic shocks, wild-type protoplasts expressing Pil1p-mEGFP were loaded into ONIX2 microfluidics system (Millipore-Sigma, Saint-Louis), and time-lapse fluorescent images were taken at a single z-slice at the top of protoplasts during media change. After background correction, the total area fraction of eisosomes at the beginning of the hypotonic shock was set to 1.0 for normalization, and the normalized values of area fraction were fit to a single exponential decay curve in GraphPad Prism (GraphPad Software; La Jolla, CA).

Measurement of membrane tension

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Protoplasts were loaded in a custom-built chamber, which was passivated with 0.2 mg/ml β-casein (Millipore-Sigma, Saint-Louis) for 30 min and pre-equilibrated with EMM5S supplemented with 0.8 M D-sorbitol. A glass micropipette (#1B100-4; World Precision Instruments, Sarasota) was forged to a diameter smaller than the average protoplast radius (~2.5 µm) and was connected to a water reservoir of adjustable height to apply a defined aspiration pressure. Before and after each experiment, the height of the water reservoir was adjusted to set the aspiration pressure to 0. Cells were imaged with a bright-field IX-71 inverted microscope (Olympus, Tokyo, Japan) equipped with a 60x/1.4NA objective, and images were recorded every second. Aspiration pressure was gradually increased every 30 s and the membrane tension σ was calculated as σ=P.Rp/[2(1-Rp/Rc)], where Rp and Rc are, respectively, the micropipette and the cell radius, P is the aspiration pressure for which the length of the tongue of the protoplast in the micropipette is equal to Rp (Evans and Yeung, 1989). To limit the effects of the adaptation of cells’ membrane tension, all measurements were performed within the first 5 min after the hypotonic shock, which greatly limited the throughput of our assay (one measurement per sample), compared to the measurements at steady state (around six measurements per sample).

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files.

References

Decision letter

  1. Vivek Malhotra
    Senior Editor; The Barcelona Institute of Science and Technology, Spain
  2. Christophe Lamaze
    Reviewing Editor; Institut Curie, France

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "Adaptation of actin dynamics and membrane tension control for yeast endocytosis" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Aurélien Roux (Reviewer #3).

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

The reviewers concurred on the quality and the overall significance of your data. However, the paper appears to be composed of two disjointed stories that are not mature yet. Considering the effort in terms of additional experiments required to present a conceptual advance, we regrettably cannot move forward with the paper at eLife. We encourage you to resubmit a new version that contains the data requested or you may choose to submit this work elsewhere for its timely publication.

Reviewer #1:

This study is of interest as the role of eisosomes in fission yeast remains poorly explored and the findings presented by Lemière and Berro are likely to be relevant for the study of CME and its relation with membrane tension in mammalian cells. Several controls need however to be performed to better support the conclusions of this otherwise timely contribution.

1. My main concern has to do with the readout used in this study. The authors assume that following the dynamics of fimbrin allow to monitor endocytosis and actin dynamics. What is the evidence that fimbrin molecules are recruited to endocytic patches? Several control experiments should be performed to validate this claim including clathrin colocalization to fimbrin patches. In addition, inhibiting CME (dyn mutant or else) should then result in decrease of fimbrin recruitment. Several of the effects of osmotic shocks described on fimbrin dynamics could occur independently from CME and the authors ought to correlate these effect with cargo uptake. What is the effect of fimbrin deletion on endocytosis? As for actin dynamics, why did not the authors follow actin directly? Following the dynamics of actin or another actin binding protein would reinforce this aspect of the work.

2. I am somehow concerned by the degree of novelty of this study as it has long been known in mammalian cells that membrane tension controls the rate of endocytosis and exocytosis (see Sheetz seminal papers and more recently papers such as Boulant et al.,). Moreover, eisosomes have been recently established as membrane reservoirs in a study with similar data in yeast (protective role of the wall and disappearance of eisosomes through flattening under increased osmolarity in protoplasts as decribed in Kabeche, JCS 2015). As a matter of facts, these published data should be better acknowledged in the results and discussion. For instance, on line 178, the authors write that their data "suggest that eisosomes might be disassembled to reduce membrane tension in media with low osmolarity…" this has been already demonstrated in Kabeche, JCS 2015. In this context, the new finding reported in the current study is mostly about the role of endocytosis on membrane tension, that is, that endocytosis is transiently reduced to decrease membrane tension. However, in none of the presented experiments have the authors addressed the selective contribution of endocytosis. The authors should therefore re-examine their findings when endocytosis is blocked by drugs or mutants. Thus, measuring membrane tension variations under osmotic shock when endocytosis is blocked would allow to estimate the relative contribution of endocytosis in membrane tension buffering.

3. The absence of eisosomes should be documented in Pil1∆ mutants. Line 244: what is the evidence that WT protoplasts express normal amounts of Pil1p when they assemble fewer eisosomes? Data presented in Figure 6 should be correlated with the visualization of eisosomes as in Figure 4B.

Reviewer #2:

Lemiere et al., use quantitative fluorescence microscopy in the model organism fission yeast to address the complex relationship between cellular processes such as endocytosis and cellular structures such as eisosomes to regulate and equilibrate plasma membrane tension. Dissecting individual processes that mechano and/or biochemically regulate plasma membrane tension remains to be challenging. Single cellular organisms such as fission yeast therefore provide a versatile and easy to manipulate model system to tackle these particular very important regulatory processes and can translate crucial findings to higher eukaryotic model systems.

The authors use their previously established quantitative fluorescence microscopy and analytical approach to measure the response of the actin crosslinker fimbrin to changes in the number of molecules, lifetimes and speed to osmolarity media changes that are meant to change plasma membrane tension. The authors describe in a combination of experiments conducted in walled cells and protoblasts, both also mutated for the Pil1 eisosome component, and changed dynamics of fimbrin. These findings provide very exciting new results about how the actin machinery adapts to osmolarity changes. These are even combined with membrane tension measurements. Together with determining the density of endocytic events after hypotonic shock treatment, the authors conclude that the cooperation of eisosome disassembly, decrease of endocytic events and ongoing exocytosis are responsible for equilibrating plasma membrane tension changes.

The findings in the manuscript are of descriptive nature and do not identify mechno-regulatory or biochemical mechanisms that control the membrane reservoir or the adaption of actin dynamics in response to changing media osmolarities. Our major concern is that the overall main point of the paper is not clear. It seems that the authors present two very interesting half stories that have not yet have been completed and that do not stand on their own in a complete story. Given the allowed time frame of revision, it is not clear that the authors could make adequate revisions. If they can, we would suggest that they concentrate on one of their respective stories and endeavor to provide more mechanistic insights into either "the adaption of actin dynamics in yeast endocytosis in response to membrane tension changes" or "cellular mechanisms that control the plasma membrane reservoir in response to osmolarity changes". Detailed comments can be found below.

1. As mentioned above, it seems that the authors pursued two different stories in this manuscript and tried to cover them under one main point. This creates confusion for the reader throughout the manuscript especially in Figure 4 and Figure 5.

Therefore, we suggest that the authors concentrate on one of the two take subjects that they address. The two different stories that could be expanded for a manuscript are:

a. One on the adaptive dynamics of the actin crosslinker fimbrin. The authors precisely report on the number of molecules, lifetimes and speed changes that happen under hypotonic shock treatments.

However, the authors do not pursue how actin dynamics mechanistically adapt to hypotonic shock treatments. Fimbrin is a crosslinker and there are many other interesting candidate proteins whose dynamic behavior under hypotonic shock treatments the authors could explore. Thus, it would be possible for the authors to obtain precise mechanistic details about how "actin dynamics adapt to membrane tension increase to control yeast endocytosis". Previously published work of the last author shows that these strains are available. We would like to see experiments equivalent to the one conducted with the fimbrin-FP expressing yeast strain with Sla1, Arp2/3 and an early endocytic marker. If the authors are willing to add these experiments, we would also like to see the measurements of membrane tension of the protoblasts treated in the conditions used in Figure 5. A direct relationship between actin cytoskeleton protein binding dynamics and quantified membrane tension is of high interest for the endocytic community and would make this manuscript very valuable and significantly new. Then the authors could explain and discuss the differences they see in fimbrin dynamics, which they do not include on in the main text. In Figure 2E and Figure3D, fimbrin lifetimes are faster under hypotonic treatment. However, in Figure 5, lifetimes decrease under hypotonic treatment. We do not understand what the authors are looking at in these analyses. Determining membrane tension for all experimental conditions is essential for the correct interpretation of these results (of course, this is only possible for protoplasts). In addition, the authors should think about showing experimentally how faster or slower fimbroin dynamics can be related to productive endocytic events.

b. The other main point the authors pursue is the regulatory mechanism for membrane tension equilibration in response to osmolarity changes. Here the authors suggest focus on these regulatory components: the endocytic machinery, eisosomes, the cell wall and the exocytic machinery. With their experiments the authors only cover the dynamic behavior of the actin crosslinker fimbrin as the representative marker for the dynamics of endocytic events in response to hypotonic treatment and the observation that PIL1 deleted cells are more prone to hypotonic shock.

What follows are conclusions made by the authors despite a lack of experimental evidence:

- 'eisosomes play a critical role in the regulation of membrane tension and membrane integrity through dynamic remodeling and scaffolding of the plasma membrane' p9, line 342.

- the 'protective role of eisosomes is even more striking in protoplasts under acute hypotonic shocks'.

- 'our data indicate that membrane tension is decreased via the disassembly of eisosomes, through release of excess membrane surface area.' p9, line 352.

- Lines 361 – 367: 'Our study identified a third, new mechanism to reduce membrane tension by temporarily reducing the endocytic density. Using our data and previous measurements (18,19), we estimate that cells endocytose about 2% of their surface area per minute. Assuming that at steady state the surface area added to the plasma membrane via exocytosis balances the surface area that is endocytosed, a reduction of the endocytic rate while keeping the exocytic rate constant for a few minutes would allow for a net addition of surface area to the plasma membrane'.

The authors only show images of eisosomes at steady state e.g. Figure 4a,b,c. To investigate the regulatory mechanism and provide convincing evidence for their claims, we suggest that the authors also track Pil1 dynamics under different hypotonic treatments and the dynamics of an exocytic marker. Here again, a direct relation to membrane tension values measured under the experimental conditions used would be crucial.

2. The title of the manuscript, "Adaption of actin dynamics and membrane tension control for yeast endocytosis," does not describe the work well since the authors only perform experiments on the actin crosslinker fimbrin and quantification of membrane tension was only performed for one experimental condition. We suggest that the authors choose a more specific title that states the main point of the revised manuscript.

3. "Temporal super-resolution" is not yet a common method that the reader should be expected to know about. Moreover, the reader might not be familiar with the improvements of this method performed by the authors (Line 98). We suggest that they schematically describe the method of temporal super-resolution in a supplementary Figure to Figure 1 and also explain the improvements. It is also not clear in which plane the cells are imaged and why only a subset of fimbrin tracks where kept for the further analysis (see comment 4.)

4. It would be helpful if the authors would provide a thorough analysis of how many fimbrin tracks there are in total, what classes of behavior can be seen and on which subset the authors are focusing and why.

The authors state in their methods, that all tracks were discarded that did not match the following criterion: 'longer than 5s and displaying an increase followed by a decrease in intensity' p12 line 497. This introduces a significant bias towards productive tracks. Tracks that are stalled or last longer than the observation window would not enter the analysis.

Why did the authors did not discard tracks that 'failed to proceed normally, as reported by the virtually null speed' p6 line 196? Or did they use a different criterion for these tracks?

5. Quality of Figures: Overall the plot quality could be improved by reporting the confidence interval differently. The plots become very busy and the lines overlap strongly, so that the reader has to look at the plots with a higher zoom to differentiate between the actual data lines and the lines of the confidence interval. We suggest that they plot the lifetimes of endocytic events separately and report statistical tests on the lifetime distributions. The way the Figure legends are placed in Figure 4 is very confusing and can be improved in the other figures for clarity, as well.

6. A model Figure summarizing their main conclusions at the end of the manuscript would clarify the take home message for the reader. The summary could make points about: What factors influence membrane tension control? Or, they could provide a schematic representation for how the actin machinery adapts to increases in membrane tension.

7. The plots in Figure 5H and I and the conclusion in the text about the Pil1 cytoplasmatic pool are very confusing and would be better placed in a supplementary Figure.

Reviewer #3:

The paper by Lemiere and Berro focuses on the dynamics of endocytic events and eisosomes in walled cells and protoplasts, to investigate the role of membrane tension in both endocytic bud formation and eisosomes. It builds on previous work that has shown that clathrin endocytosis dnamics is affected by changes in membrane tension, and that eisosomes participate into the regulation of plasma membrane tension. The paper is clearly written, and has interesting findings, among which are:

1. That membrane tension changes do not strongly affect CME in walled cells

2. Embrane tension strongly affect CME in protoplasts

3. Eisosomes, while having a limited role in walled cells, have a strong role in regulating membrane tension in protoplasts.

I have a few concerns about the paper in the current state. The authors want to compare WT (walled) cells and protoplasts, but the comparison is not done thoroughly, which makes the paper a bit unbalanced. Also, the fact that the paper is written in two parts, first the walled cell results, and then the protoplasts results, the comparison is not very easy to follow. The authors may reconsider the text organization by clustering similar experiments done in wall cells and protoplasts. I understand that some experiments cannot be performed in protoplasts or in walled cells, but the following comments are here to extend as much as possible the comparison.

– The density of eisosomes should be better characterized in walled cells and in protoplasts, before and after the shock (see Figure 4B and 4C, if I understood well, this is only for cells at steady state, and different osmotic pressure). The authors model proposes that eisosomes regulate tension through disassembly. The authors have to show this. I would recommend to also perform hypertonic shocks that should increase the number of eisosomes.

– Generally speaking, it sounds strange that eisosomes would have an impact only on protoplast. The observation that when grown at higher osmotic pressure, cells have more eisosomes suggest that cells prepare to stringer hypo-osmotic shocks in this condition. How can the author rationalize these results?

– I was wondering how sure the Fim1p reporter reports actin density? I guess this has been published elsewhere as the authors rely on published work on this point, but it would still be nice to have a figure panel that shows a co-labelling with actin.

– I find surprising that the dynamics of actin is not different upon osmotic shocks in WT walled cells. I was wondering if the actin treadmilling was modified upon osmotic shocks. This should be easy to test through FRAP experiment of Fim1p at the endocytic sites.

– Reported tension values are extremely small (10-4 nM/um, which is 10-7 N/m). Fibroblastic cells have tensions of typically 10-4 – 10-5 N/m. I expect yeast cells to have even higher tension become they live in higher turgor pressure. Also at 10-7 N/m, membrane usually have large fluctuations which are visible by any type of microscopy, which are not seen in the authors’ experiments. Either the calculation has a problem, or the unit is wrong. Please use standard units for membrane tension in N/m or in mN/m (or m-1).

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Rapid adaptation of endocytosis, exocytosis and eisosomes after an acute increase in membrane tension in yeast cells" for further consideration by eLife. Your revised article has been evaluated by Vivek Malhotra (Senior Editor) and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed before acceptance, as outlined below:

As you will see in the verbatim comments attached, the first reviewer agrees that the revised manuscript contains a substantial amount of work and that it has increased in length. While Reviewer 1 acknowledges that some aspects of the manuscript have greatly improved, the reviewer is still concerned by the readability of the manuscript, a concern raised already for the initial submission. This is exacerbated by the addition of a third part on exocytosis. In particular, the reviewer finds it difficult to follow the comparison between walled cells and protoplasts. The reviewer expresses some concerns about the readability of the plots expressed as "speed vs time". The reviewer also disagrees with the use of deltaP for accounting for osmolarity changes. Finally, related to the new part on exocytosis, the reviewer is asking for a better characterization of its regulation by osmotic shock.

Reviewer 2 is somehow less critical and agrees that the manuscript strength resides in the highly quantitative characterization of endocytosis dynamics and in the study of the 3 co-operating mechanisms at play during the response to hypo-osmotic shock.

Considering the positive appreciation of your work by the two reviewers, I am willing to leave the door open for a revised version. Since this work is a resubmission of a manuscript already reviewed in 2018, we will be very strict on the revised manuscript. It should comply with all of the 2 reviewers concerns with a special attention to the readability of the manuscript. I would like to insist that another round of reviewing will not be granted at this stage.

Reviewer #1:

This submission is a revised version of a previous submission. The authors have done a substantial amount of work, and the paper has increased in length. Some aspects have greatly improved, such as the characterization of the roles of the cell wall, tension and actin (using drugs and mutants to change actin organization) in controlling endocytosis. The role of eisosomes in buffering membrane tension has been characterized in further details, providing the first evidence that eisosomes could play a similar role in yeast than caveolae in cells in buffering membrane tension. Finally, the authors added a study on the evolution of exocytosis during hypoosmotic shocks, which reacts in opposite direction than endocytosis.

While I value the efforts and work provided by the authors in their revised version, they in fact took an opposite direction in their revised manuscript than what was initially suggested by the reviewers: – split the manuscript in two, and strengthen the characterization of tension/endocytosis on the one hand, and eisosomes/tension on the other. Instead, the authors kept the two parts in the same manuscript, and added a third part on exocytosis. They also extended their comparison of walled cells and protoplasts, which is indeed interesting, but makes the flow hard to follow because the reader needs to go back-and-forth between different figures. This choice is at the expense of the manuscript's readability, which became lengthy and difficult to follow.

While I value the efforts made by the authors, and the improvement of the characterization of endocytosis, tension and eisosomes, I am embarrassed to say that the main criticism remains. I understand that the authors wanted to broaden their study, and get more general conclusions, but it does not go through well. I unfortunately cannot recommend publication in the present form, and would certainly suggest again to split the manuscript in two (which could be back-to-back in ELife for example), one on endocytosis/exocytosis and tension (where pil1delta and protoplasts would be used as a tool to dysregulate tension) and one on eisosomes/tension buffering.

– While the mechanism at play to regulate dynamics of endocytosis and eisosomes during osmotic shock became well characterized, the new part on exocytosis is very short and does not provide any further understanding of the regulation at play here. Is it tethering, or SNARE function that is regulated by tension? Is that higher tension strengthens the actin dome around exocytic vesicles to trigger fusion? A better characterization, at least at the same level of other parts would be needed here.

– Some plots are difficult to read. The plot "Speed" vs "time", which zero the invagination time, is not easy to read. The total duration of endocytic events is changing with conditions, and if I understood well, that is the main effect that the author look at, but it took me quite some time to figure out how to read these. I think a classic plot with average duration would be easier to follow for most readers. I understand that in some cases, the authors see no invagination, which is also reported in this graph, but I think 2 plots would do a better job than a single one in this case. Or perhaps, explain better how to read those plots, as they are uncommon, and at the basis of the authors' argumentation.

– I disagree with the use of deltaP to account for a change of osmolarity. Pressure should be in Pa (or any corresponding unit), not in Moles per Liter. Either the authors use deltaC, or they measure the osmotic pressure of their solution and can use Osm in this case for pressure. Also, at concentrations about 0.5M, there is usually a substantial discrepancy between Osmolarity and Molarity, and as the authors are using Molarities above 1M, I think it would be safer to estimate the osmotic pressure of those solutions.

Reviewer #2:

Lemiere and colleagues show in this manuscript that endocytosis is remarkably robust against osmotic changes in normal walled yeast cells. However, in cells without cell walls endocytosis is much more sensitive to osmotic changes, presumably due to changing membrane tension. Eisosomes are shown to be important for regulating membrane tension after osmotic shocks.

Changes in turgor pressure are also shown to transiently affect the frequency of endocytic events. Eisosomes are shown to be important for restoring the normal endocytic frequency after shocks. Finally, exocytosis is suggested to be a mechanism that restores normal membrane tension after hypotonic shocks.

The strength of the manuscript is the highly quantitative and careful experimentation and the fact that three different plasma membrane processes are considered together (endocytosis, exocytosis, eisosomes), which is surprisingly rarely done, even though they must be functionally connected.

I have some suggestions for the authors to improve their manuscript:

Lines 217-223: Does the plasma membrane area remain constant after hypotonic treatment? I.e. are the eisosomes really disassembled and not just diluted if the plasma membrane area increases?

Figure 4H: Looking at the time series images it seems like the plane of focus shifts during the experiment. At time zero, and at 1 minute, the focus is at the surface of the cell, but later it appears that the focus shifts toward the center of the cell. During the second half of the time series the remaining eisosomes are seen at the periphery of the cell, which could be explained by a focus shift. Is this a representative example?

Lines 325 : The authors could clarify a bit more clearly whether they quantify the density of endocytic events, or the density of Fim1 labeled endocytic sites, which would depend on the lifetime of the Fim1 patches. In other words, is the reported value the number of endocytic vesicles formed per area?

Line 340: I don't understand why the authors say that "volume increased faster than the change in endocytic density". Aren't they both significantly changed already at time 0, and it takes longer for the volume to reach maximum change?

Lines 379-382: This is the only place where potential non-vesicular mechanisms are brought up. Lipid transfer, for example, via membrane contact sites is a possible mechanism for expanding the plasma membrane area. The method used cannot distinguish between such non-vesicular transport and exocytosis. Therefore, the authors should bring this caveat up also in the discussion.

Lines 370-382: I think that the exocytosis assay is not totally intuitive. It might be useful for the readers if the authors tried to clarify the explanation a bit. In other words: how is it measuring exocytosis and not endocytosis, which FM4-64 is usually used to measure?

Line 520: How do the authors arrive at the value of 2%. Please elaborate.

Line 583: What is "higher order eukaryote"? (I would also advise against using "higher eukaryote", which, although commonly used, is not a clearly defined term, nor really biologically meaningful.)

https://doi.org/10.7554/eLife.62084.sa1

Author response

[Editors’ note: the authors resubmitted a revised version of the paper for consideration. What follows is the authors’ response to the first round of review.]

Reviewer #1:

This study is of interest as the role of eisosomes in fission yeast remains poorly explored and the findings presented by Lemière and Berro are likely to be relevant for the study of CME and its relation with membrane tension in mammalian cells. Several controls need however to be performed to better support the conclusions of this otherwise timely contribution.

1. My main concern has to do with the readout used in this study. The authors assume that following the dynamics of fimbrin allow to monitor endocytosis and actin dynamics. What is the evidence that fimbrin molecules are recruited to endocytic patches? Several control experiments should be performed to validate this claim including clathrin colocalization to fimbrin patches. In addition, inhibiting CME (dyn mutant or else) should then result in decrease of fimbrin recruitment. Several of the effects of osmotic shocks described on fimbrin dynamics could occur independently from CME and the authors ought to correlate these effect with cargo uptake. What is the effect of fimbrin deletion on endocytosis? As for actin dynamics, why did not the authors follow actin directly? Following the dynamics of actin or another actin binding protein would reinforce this aspect of the work.

We thank Reviewer 1 for their interest in our study. We thank the reviewer for pointing out the lack of clarity about the marker we used to monitor actin dynamics. We have clarified our rationale in the main text. In summary, fimbrin is an actin filament crosslinker whose recruitment to CME structures has been well documented (Adams et al., 1991; Berro and Pollard, 2014; Drubin et al., 1988; Sirotkin et al., 2010; Sun et al., 2019). Fimbrin has been routinely used as an actin marker during CME because it is abundant and follows exactly actin dynamics during CME (it is recruited, peaks and disappears at the same time as actin). In addition, GFP-tagged fimbrin is fully functional. Fimbrin deletion in S. pombe has a very mild phenotype, probably because of compensatory effects from other crosslinkers or other proteins such as tropomyosin (Skau and Kovar, 2010). We did not use actin as a marker because GFP-actin is not fully functional, and one needs to overexpress GFP-actin on top of endogenous actin. Overexpression promoters in fission yeast lack precision and have significant cell-to-cell variability, which makes the data more noisy and difficult to quantitatively analyze and interpret. Colocalization of fimbrin with clathrin and adaptor proteins has been shown before (Kaksonen et al., 2005; Sirotkin et al., 2010). To make our argument more convincing, we have added in the paper a panel showing colocalization of fimbrin with the adaptor protein End4p, which is recruited before fimbrin and is a marker of endocytic structures and vesicles (Figure 1D and 1E). Note that we did not choose to use clathrin as a second marker because clathrin also localizes to endosomes in addition to the plasma membrane, which makes the signal more dispersed. It has been shown before that if early stages of endocytosis are inhibited, actin does not assemble at CME sites, neither does fimbrin (Sun et al., 2015). If actin dynamics is inhibited with drugs, endocytosis is blocked before any membrane ingression (Ayscough et al., 1997; Kukulski et al., 2012).

The successful formation of a vesicle can be assessed by the movements of the markers after they peak. Our new data using the FM4-64 dye (Figure 8) demonstrates that the dye is internalized even in the worst hypotonic conditions tested, which further demonstrates that vesicles are indeed formed. Figure 8B shows example images of internalized vesicles stained with the FM4-64 dye in wild-type walled cells. Author response image 1 is representative wild-type protoplasts stained with FM4-64 for 10 minutes after a hypotonic shock of -0.2 M (the movie from which this snapshot was extracted was used to produce Figure 8C).

Author response image 1

2. I am somehow concerned by the degree of novelty of this study as it has long been known in mammalian cells that membrane tension controls the rate of endocytosis and exocytosis (see Sheetz seminal papers and more recently papers such as Boulant et al.,). Moreover, eisosomes have been recently established as membrane reservoirs in a study with similar data in yeast (protective role of the wall and disappearance of eisosomes through flattening under increased osmolarity in protoplasts as described in Kabeche, JCS 2015). As a matter of facts, these published data should be better acknowledged in the results and discussion. For instance, on line 178, the authors write that their data "suggest that eisosomes might be disassembled to reduce membrane tension in media with low osmolarity…" this has been already demonstrated in Kabeche, JCS 2015. In this context, the new finding reported in the current study is mostly about the role of endocytosis on membrane tension, that is, that endocytosis is transiently reduced to decrease membrane tension. However, in none of the presented experiments have the authors addressed the selective contribution of endocytosis. The authors should therefore re-examine their findings when endocytosis is blocked by drugs or mutants. Thus, measuring membrane tension variations under osmotic shock when endocytosis is blocked would allow to estimate the relative contribution of endocytosis in membrane tension buffering.

We thank Reviewer 1 for pointing out the lack of clarity in what was previously known and what is new in our study. We have edited the text to further describe what was already known before we performed our study and provided better reference to previous papers. We have also edited the text to better describe what are the novelties of our paper.

Some links between membrane tension, endocytosis and exocytosis have been described before, but our study brings a new quantitative characterization of these processes to determine (1) how much more actin is needed when membrane tension is increased, (2) how eisosomes influence this actin regulation, and (3) how much membrane is added via regulation of the rates of endocytosis and, in this revised version of the paper, via regulation of the rate of exocytosis. Kabeche et al., showed eisosomes disassemble under hypotonic shocks but did not prove that hypotonic shocks increased membrane tension or that membrane tension is reduced by eisosome disassembly. Our micropipette aspiration data, and our data on actin requirement during CME demonstrate that membrane tension is indeed reduced.

We thank the reviewers for suggesting us to determine the effects of blocking endocytosis on membrane tension. We have now included this experiment and an experiment where we blocked exocytosis (new Figure 9 in the revised paper). However, since direct membrane tension measurements with micropipette aspiration is difficult, low throughput and slow (it requires several minutes, which is the timeline over which the systems goes back to equilibrium), we do not think that the temporal resolution of membrane tension measurement using micropipette aspiration is high enough to monitor the effects of osmotic shocks or drug treatments. Instead, we used the fact that the plasma membrane ruptures and cells are more likely to die when membrane tension is high to determine the effects of blocking endocytosis or exocytosis (Figure 9 in the revised paper).

3. The absence of eisosomes should be documented in Pil1∆ mutants. Line 244: what is the evidence that WT protoplasts express normal amounts of Pil1p when they assemble fewer eisosomes? Data presented in Figure 6 should be correlated with the visualization of eisosomes as in Figure 4B.

We have clarified the text to better explain that previous studies have shown that the deletion of the gene coding for the major scaffolding eisosome protein Pil1p is sufficient to remove all eisosomes from the cell (Kabeche et al., 2011; Olivera-Couto et al., 2011).

Since wild-type protoplasts are made and imaged within 50 min, we did not expect any significant change in the expression level of Pil1p between intact cells and protoplasts. Indeed, we have now checked the concentrations and found no difference (Figure 4 – Supplement 1A). We have also added new panels H and I in Figure 4 to quantify the amount of eisosome that disassembles during a shock and the kinetics of the disassembly.

Reviewer #2:

Lemiere et al., use quantitative fluorescence microscopy in the model organism fission yeast to address the complex relationship between cellular processes such as endocytosis and cellular structures such as eisosomes to regulate and equilibrate plasma membrane tension. Dissecting individual processes that mechano and/or biochemically regulate plasma membrane tension remains to be challenging. Single cellular organisms such as fission yeast therefore provide a versatile and easy to manipulate model system to tackle these particular very important regulatory processes and can translate crucial findings to higher eukaryotic model systems.

The authors use their previously established quantitative fluorescence microscopy and analytical approach to measure the response of the actin crosslinker fimbrin to changes in the number of molecules, lifetimes and speed to osmolarity media changes that are meant to change plasma membrane tension. The authors describe in a combination of experiments conducted in walled cells and protoblasts, both also mutated for the Pil1 eisosome component, and changed dynamics of fimbrin. These findings provide very exciting new results about how the actin machinery adapts to osmolarity changes. These are even combined with membrane tension measurements. Together with determining the density of endocytic events after hypotonic shock treatment, the authors conclude that the cooperation of eisosome disassembly, decrease of endocytic events and ongoing exocytosis are responsible for equilibrating plasma membrane tension changes.

The findings in the manuscript are of descriptive nature and do not identify mechno-regulatory or biochemical mechanisms that control the membrane reservoir or the adaption of actin dynamics in response to changing media osmolarities. Our major concern is that the overall main point of the paper is not clear. It seems that the authors present two very interesting half stories that have not yet have been completed and that do not stand on their own in a complete story. Given the allowed time frame of revision, it is not clear that the authors could make adequate revisions. If they can, we would suggest that they concentrate on one of their respective stories and endeavour to provide more mechanistic insights into either "the adaption of actin dynamics in yeast endocytosis in response to membrane tension changes" or "cellular mechanisms that control the plasma membrane reservoir in response to osmolarity changes". Detailed comments can be found below.

1. As mentioned above, it seems that the authors pursued two different stories in this manuscript and tried to cover them under one main point. This creates confusion for the reader throughout the manuscript especially in Figure 4 and Figure 5.

Therefore, we suggest that the authors concentrate on one of the two take subjects that they address. The two different stories that could be expanded for a manuscript are:

a. One on the adaptive dynamics of the actin crosslinker fimbrin. The authors precisely report on the number of molecules, lifetimes and speed changes that happen under hypotonic shock treatments.

However, the authors do not pursue how actin dynamics mechanistically adapt to hypotonic shock treatments. Fimbrin is a crosslinker and there are many other interesting candidate proteins whose dynamic behavior under hypotonic shock treatments the authors could explore. Thus, it would be possible for the authors to obtain precise mechanistic details about how "actin dynamics adapt to membrane tension increase to control yeast endocytosis". Previously published work of the last author shows that these strains are available. We would like to see experiments equivalent to the one conducted with the fimbrin-FP expressing yeast strain with Sla1, Arp2/3 and an early endocytic marker. If the authors are willing to add these experiments, we would also like to see the measurements of membrane tension of the protoblasts treated in the conditions used in Figure 5. A direct relationship between actin cytoskeleton protein binding dynamics and quantified membrane tension is of high interest for the endocytic community and would make this manuscript very valuable and significantly new. Then the authors could explain and discuss the differences they see in fimbrin dynamics, which they do not include on in the main text. In Figure 2E and Figure3D, fimbrin lifetimes are faster under hypotonic treatment. However, in Figure 5, lifetimes decrease under hypotonic treatment. We do not understand what the authors are looking at in these analyses. Determining membrane tension for all experimental conditions is essential for the correct interpretation of these results (of course, this is only possible for protoplasts). In addition, the authors should think about showing experimentally how faster or slower fimbroin dynamics can be related to productive endocytic events.

We thank the reviewers for finding our results exciting. The goal of our paper is to describe how the actin machinery is regulated after a sudden increase in membrane tension and how membrane tension rapidly adapts to change. We believe that the description of these phenomena and the relative contributions of the cell wall, eisosomes, endocytosis and exocytosis in this process will be useful for the community. We acknowledge that our paper does not identify all the underlying molecular mechanisms involved but lay the foundations to do so. It is in our plans to follow up on both parts of our study to determine the molecular mechanisms at the origin of the effects we observed. We believe that identifying molecular mechanisms for each part will each lead to a full paper and are therefore beyond the scope of our current study.

In our paper, we present two related effects of osmotic shocks because the data from the same experiments have been used to determine at the same time a) the amount of fimbrin assembled and b) the rate of endocytosis. Because these complementary results are extracted from the same datasets, we believe it is legitimate to present them in the same paper. In addition, the results of both parts are consistent with each other and presenting each part independently would make the results more puzzling and difficult to interpret.

To complete the story about membrane homeostasis, and to substantiate our speculations on the rates of exocytosis, we have adapted a method to determine the rates of exocytosis using the FM4-64 dye (new Figure 8 in the revised paper). We now show that (a) the rate of exocytosis is increased after a hypotonic shock in protoplasts but not in walled cells, and (b) change in endocytosis and exocytosis rates almost exactly account for the change of membrane surface area. We believe that these new data significantly complement and clarifies the second part of our paper.

Measuring membrane tension in different osmotic shocks so as to correlate the change in actin dynamics to membrane tension (instead of osmolarity) is an excellent suggestion. However, measuring membrane tension using micropipette aspiration is slow, slower than the membrane tension adaptation. Therefore, it would be challenging to accurately correlate the dynamics of the actin machinery with membrane tension. We and our collaborators recently obtained funding to build a new setup which includes optical tweezers and will allow us to make more precise measurements and also allow us to follow the temporal evolution of the adaptation of membrane tension. We believe data obtained with this setup.

b. The other main point the authors pursue is the regulatory mechanism for membrane tension equilibration in response to osmolarity changes. Here the authors suggest focus on these regulatory components: the endocytic machinery, eisosomes, the cell wall and the exocytic machinery. With their experiments the authors only cover the dynamic behavior of the actin crosslinker fimbrin as the representative marker for the dynamics of endocytic events in response to hypotonic treatment and the observation that PIL1 deleted cells are more prone to hypotonic shock.

What follows are conclusions made by the authors despite a lack of experimental evidence:

- 'eisosomes play a critical role in the regulation of membrane tension and membrane integrity through dynamic remodeling and scaffolding of the plasma membrane' p9, line 342.

- the 'protective role of eisosomes is even more striking in protoplasts under acute hypotonic shocks'.

- 'our data indicate that membrane tension is decreased via the disassembly of eisosomes, through release of excess membrane surface area.' p9, line 352.

- Lines 361 – 367: 'Our study identified a third, new mechanism to reduce membrane tension by temporarily reducing the endocytic density. Using our data and previous measurements (18,19), we estimate that cells endocytose about 2% of their surface area per minute. Assuming that at steady state the surface area added to the plasma membrane via exocytosis balances the surface area that is endocytosed, a reduction of the endocytic rate while keeping the exocytic rate constant for a few minutes would allow for a net addition of surface area to the plasma membrane'.

The authors only show images of eisosomes at steady state e.g. Figure 4a,b,c.

To investigate the regulatory mechanism and provide convincing evidence for their claims, we suggest that the authors also track Pil1 dynamics under different hypotonic treatments and the dynamics of an exocytic marker. Here again, a direct relation to membrane tension values measured under the experimental conditions used would be crucial.

We thank the reviewer for pointing out the ambiguity of these statements or their overinterpretation in the initial version of our paper. We have modified them to make sure credit is given to the previous studies that support these statements and how our data support these claims (originally l. 342 and 352). In addition, the new version of the paper contains data about the rates of exocytosis which support the last statement mentioned above (originally l. 361-367). We have also added new data showing the temporal evolution of eisosomes at the plasma membrane after a hypotonic shock (Figure 4H and 4I).

2. The title of the manuscript, "Adaption of actin dynamics and membrane tension control for yeast endocytosis," does not describe the work well since the authors only perform experiments on the actin crosslinker fimbrin and quantification of membrane tension was only performed for one experimental condition. We suggest that the authors choose a more specific title that states the main point of the revised manuscript.

We have modified the title to better reflect the main message of the revised paper.

3. "Temporal super-resolution" is not yet a common method that the reader should be expected to know about. Moreover, the reader might not be familiar with the improvements of this method performed by the authors (Line 98). We suggest that they schematically describe the method of temporal super-resolution in a supplementary Figure to Figure 1 and also explain the improvements. It is also not clear in which plane the cells are imaged and why only a subset of fimbrin tracks where kept for the further analysis (see comment 4.)

We have extended the description of the temporal super-resolution method in the

“Measurement of the temporal evolution of the number of proteins and speed” section of the Materials and methods. We have also made sure that we clearly explain that we imaged the “bottom” surface of the cell, i.e. the planes closest to the objective and the coverslip.

4. It would be helpful if the authors would provide a thorough analysis of how many fimbrin tracks there are in total, what classes of behavior can be seen and on which subset the authors are focusing and why.

The authors state in their methods, that all tracks were discarded that did not match the following criterion: 'longer than 5s and displaying an increase followed by a decrease in intensity' p12 line 497. This introduces a significant bias towards productive tracks. Tracks that are stalled or last longer than the observation window would not enter the analysis.

Why did the authors did not discard tracks that 'failed to proceed normally, as reported by the virtually null speed' p6 line 196? Or did they use a different criterion for these tracks?

We thank the reviewers for pointing out the ambiguities in our description of the methods. While developing our analysis pipeline, we were careful not to introduce bias when we chose the methods and parameters for tracking and selecting the tracks to keep. Sites of endocytosis are quite dense in most areas of the cell, which means that a very large majority of the fluorescence signal from endocytic events overlap with each other and cannot be used for quantitative analysis. It is difficult for tracking algorithm to identify overlapping spots, independent spots that appear consecutively in a diffraction limited region, or spots that go out of focus, etc. Our methods aimed at rejecting those spots automatically when possible, or manually after visual inspection. In fact, after visual inspection, long tracks always corresponded to multiple endocytic events that overlapped (which can usually be determined by the shape of the fluorescence spot or a sequence of multiple fluorescence increase and decrease), and tracks shorter than 5 s never covered the full time course of an endocytic event.

Speed was not one of our criteria because overlapping patches cannot be distinguished by speed alone. Therefore, we did not reject tracks that did not move much and noticed that some tracks had very low mobility in some conditions (e.g. l. 196 of the original paper as Reviewer 2 noted).

We do not believe there are several classes of endocytic events, because even when tracked manually or automatically but with very lenient parameters, we identified only a single class of behavior for single endocytic sites.

5. Quality of Figures: Overall the plot quality could be improved by reporting the confidence interval differently. The plots become very busy and the lines overlap strongly, so that the reader has to look at the plots with a higher zoom to differentiate between the actual data lines and the lines of the confidence interval. We suggest that they plot the lifetimes of endocytic events separately and report statistical tests on the lifetime distributions. The way the Figure legends are placed in Figure 4 is very confusing and can be improved in the other figures for clarity, as well.

We thank Reviewer 2 for these suggestions. To increase clarity, we have added new supplemental figures where we have plotted the curves individually for different conditions whenever they significantly overlapped in the main figures. We have also performed statistical tests when it was relevant.

Patch lifetimes are virtually impossible to determine accurately since our tracking method detects spots only when the signal to noise ratio is high enough. Therefore, we likely miss at least one or two time points at the beginning and the end of each track. Determining lifetimes with high precision would require us to extrapolate the data, which we prefer not to do. In the paper, we discuss relative differences in lifetimes by comparing the times when the number of molecules is half the maximum value (which is reached at time 0 s). Since this number is evaluated from the mean number of molecules we have only one data point per condition and we cannot perform statistical analysis between conditions. To highlight the lack of precision in these numbers, we only discuss the difference between conditions as a percentage, round these numbers to the closest multiple of 5 and added an approximate sign (~) before them.

6. A model Figure summarizing their main conclusions at the end of the manuscript would clarify the take home message for the reader. The summary could make points about: What factors influence membrane tension control? Or, they could provide a schematic representation for how the actin machinery adapts to increases in membrane tension.

We thank Reviewer 2 for this suggestion. We have added a new figure at the end of the paper with a schematic summarizing our results (Figure 10).

7. The plots in Figure 5H and I and the conclusion in the text about the Pil1 cytoplasmatic pool are very confusing and would be better placed in a supplementary Figure.

The rationale for showing these figures in the main text is to demonstrate that once we mechanically get rid of most of the eisosomes (by putting protoplasts in a solution with a very low osmolarity), endocytosis behaves the same way as in cells where the eisosome is genetically removed. We believe this further validates our conclusion that eisosomes are the first line of defense from a mechanical shock. We think these figures are an important piece of data to make our point so we have left them in the main figures. However, we have edited the text to make these ideas clearer.

Reviewer #3:

The paper by Lemiere and Berro focuses on the dynamics of endocytic events and eisosomes in walled cells and protoplasts, to investigate the role of membrane tension in both endocytic bud formation and eisosomes. It builds on previous work that has shown that clathrin endocytosis dynamics is affected by changes in membrane tension, and that eisosomes participate into the regulation of plasma membrane tension. The paper is clearly written, and has interesting findings, among which are:

1. That membrane tension changes do not strongly affect CME in walled cells

2. Membrane tension strongly affect CME in protoplasts

3. Eisosomes, while having a limited role in walled cells, have a strong role in regulating membrane tension in protoplasts.

I have a few concerns about the paper in the current state. The authors want to compare WT (walled) cells and protoplasts, but the comparison is not done thoroughly, which makes the paper a bit unbalanced. Also, the fact that the paper is written in two parts, first the walled cell results, and then the protoplasts results, the comparison is not very easy to follow. The authors may reconsider the text organization by clustering similar experiments done in wall cells and protoplasts. I understand that some experiments cannot be performed in protoplasts or in walled cells, but the following comments are here to extend as much as possible the comparison.

We thank Reviewer 3 for finding our findings interesting. As we explained in our response to other reviewers, we have kept both parts together because most of the results from both parts come from the same experimental datasets, and the results of both parts support each other. We have extended the membrane tension and homeostasis regulation parts with new data on the rates of exocytosis. We have considered reorganizing the text around mutants but we think our message is still clearer when the results on the dynamics of the actin machinery and on membrane tension regulation are separated. We believe this organization is also better justified and clearer now that we have added new data about the regulation of the rates of exocytosis (Figure 8).

– The density of eisosomes should be better characterized in walled cells and in protoplasts, before and after the shock (see Figure 4B and 4C, if I understood well, this is only for cells at steady state, and different osmotic pressure). The authors model proposes that eisosomes regulate tension through disassembly. The authors have to show this. I would recommend to also perform hypertonic shocks that should increase the number of eisosomes.

We thank Reviewer 3 for this suggestion. We have added new live imaging data of Pil1p that allowed us to quantify the timing and extent of eisosome disassembly after an acute hypotonic shock (Figure 4H and 4I in the resubmitted manuscript). These data strengthen our argument that eisosome disassembly regulate membrane tension.

We agree with Reviewer 3 that understanding the dynamics of eisosomes under hypertonic shocks is an interesting and important question. However, we believe answering this question is beyond the scope of our current paper, which focuses only on increases in membrane tension (i.e. under hypotonic shocks). There are several pieces of evidence in the literature that argue that the response to hypertonic shocks triggers different pathways than the response to hypotonic shocks. To keep our paper clearer we have decided to leave the study of hypertonic shocks on eisosomes for a future study.

– Generally speaking, it sounds strange that eisosomes would have an impact only on protoplast. The observation that when grown at higher osmotic pressure, cells have more eisosomes suggest that cells prepare to stringer hypo-osmotic shocks in this condition. How the author can rationalize these results?

Our initial data showed that eisosomes did not seem to have a significant effect on walled cells after a single hypotonic shock while had a protective effect in protoplasts. However, while performing experiments for the revisions of this paper, we serendipitously discovered that eisosomes do protect walled cells from dying when we performed consecutive osmotic shocks (Figure 6D and 6E). We have now revised our conclusions to reflect the protective role of eisosome more accurately.

– I was wondering how sure the Fim1p reporter reports actin density? I guess this has been published elsewhere as the authors rely on published work on this point, but it would still be nice to have a figure panel that shows a co-labelling with actin.

As we explained in our answers to other reviewers, fimbrin is a marker for actin dynamics that we and other have used extensively in endocytosis because it is very abundant and exactly follows the dynamics of actin in wild-type and all mutants tested. To clarify this point we have added new panels showing colocalization of fimbrin and the clathrin-coated pit adaptor End4p (homolog of Sla2 in S. cerevisiae and Hip1R in mammals) (Figure 1D and 1E).

– I find surprising that the dynamics of actin is not different upon osmotic shocks in WT walled cells. I was wondering if the actin treadmilling was modified upon osmotic shocks. This should be easy to test through FRAP experiment of Fim1p at the endocytic sites.

We say in the text that the overall dynamics of actin does not change after an osmotic shock. However, the microscopy method we used cannot tell whether the turnover of actin dynamics is different in these conditions. This would be an interesting property to measure. However, performing and analyzing FRAP data on such transient system is very difficult. We recently collected some FRAP data for another paper (Lacy et al., 2019) but we realized that even at steady-state (i.e. not in shock conditions) the resolution of the data would not be high enough for us to make definitive conclusions.

– Reported tension values are extremely small (10-4 nM/um, which is 10-7 N/m). Fibroblastic cells have tensions of typically 10-4 – 10-5 N/m. I expect yeast cells to have even higher tension become they live in higher turgor pressure. Also at 10-7 N/m, membrane usually have large fluctuations which are visible by any type of microscopy, which are not seen in the authors's experiments. Either the calculation has a problem, or the unit is wrong. Please use standard units for membrane tension in N/m or in mN/m (or m-1).

We thank Reviewer 3 for picking up this typo. Indeed, the units we meant to report are actually 103 larger than initially reported on the figure. Our numbers are indeed consistent with the values mentioned in the reviewer’s comment and the values previously reported in the literature.

[Editors’ note: what follows is the authors’ response to the second round of review.]

Reviewer #1:

This submission is a revised version of a previous submission. The authors have done a substantial amount of work, and the paper has increased in length. Some aspects have greatly improved, such as the characterization of the roles of the cell wall, tension and actin (using drugs and mutants to change actin organization) in controlling endocytosis. The role of eisosomes in buffering membrane tension has been characterized in further details, providing the first evidence that eisosomes could play a similar role in yeast than caveolae in cells in buffering membrane tension. Finally, the authors added a study on the evolution of exocytosis during hypoosmotic shocks, which reacts in opposite direction than endocytosis.

While I value the efforts and work provided by the authors in their revised version, they in fact took an opposite direction in their revised manuscript than what was initially suggested by the reviewers: – split the manuscript in two, and strengthen the characterization of tension/endocytosis on the one hand, and eisosomes/tension on the other. Instead, the authors kept the two parts in the same manuscript, and added a third part on exocytosis. They also extended their comparison of walled cells and protoplasts, which is indeed interesting, but makes the flow hard to follow because the reader needs to go back-and-forth between different figures. This choice is at the expense of the manuscript's readability, which became lengthy and difficult to follow.

While I value the efforts made by the authors, and the improvement of the characterization of endocytosis, tension and eisosomes, I am embarrassed to say that the main criticism remains. I understand that the authors wanted to broaden their study, and get more general conclusions, but it does not go through well. I unfortunately cannot recommend publication in the present form, and would certainly suggest again to split the manuscript in two (which could be back-to-back in eLife for example), one on endocytosis/exocytosis and tension (where pil1delta and protoplasts would be used as a tool to dysregulate tension) and one on eisosomes/tension buffering.

We thank Reviewer #1 for her/his positive comments about the new work presented in this revised version. We thank her/him for pointing out to us that the paper is still difficult to read, even more so after the revisions. On their suggestion, we have thoroughly reorganized the paper into two parts, the first one addressing the regulations of membrane tension by eisosomes, endocytosis and exocytosis (Figures 1 to 5) and a second part on the effects of increased membrane tension on the dynamics of actin at sites of endocytosis, focusing on wild-type and pil1Δ protoplasts as tools to amplify the increase in membrane tension (Figure 6 and 7). We thank the reviewer for this suggestion because we think this new organization indeed makes the flow of the paper better and its messages clearer. We seriously considered splitting our study into two papers, but we eventually decided to keep everything in a single paper because we believe the data we present are quite interconnected and splitting the paper into two would create other readability issues and may force readers to go back and forth between papers. That said, if Reviewer #1 still finds our new version difficult to read we should be able to split it in two papers.

– While the mechanism at play to regulate dynamics of endocytosis and eisosomes during osmotic shock became well characterized, the new part on exocytosis is very short and does not provide any further understanding of the regulation at play here. Is it tethering, or SNARE function that is regulated by tension? Is that higher tension strengthens the actin dome around exocytic vesicles to trigger fusion? A better characterization, at least at the same level of other parts would be needed here.

Our new data quantitatively describe how much more membrane is added by exocytosis to regulate membrane tension. Understanding the precise molecular mechanisms at play during the regulations of exocytosis when membrane tension increases is indeed an important and interesting question. However, it is not an easy question to address and there have been debates about the effect of high membrane tension on exocytosis (Kozlov and Chernomordik, 2015). In particular, two contradictory effects happen at the same time: a) the initiation of the fusion of a lipid vesicle to a lipid bilayer is less favorable under high tension, but b) once fusion starts, high membrane tension favors the completion of the fusion. In addition, tethering complexes (like the exocyst) and the SNARE complex may also be influenced differentially by membrane tension. In other words, the question is complex and it would not be straightforward to answer it in vivo, especially over the two months allocated for the revisions. Note we can already exclude that actin plays a direct role in favoring vesicle fusion in yeast cells because, unlike mammalian cells, they do not have an acto-myosin cortex, and the only actin at the plasma membrane is at site of endocytosis. The only indirect role of actin in exocytosis may be in vesicle delivery to the plasma membrane via actin cables.

– Some plots are difficult to read. The plot "Speed" vs "time", which zero the invagination time, is not easy to read. The total duration of endocytic events is changing with conditions, and if I understood well, that is the main effect that the author look at, but it took me quite some time to figure out how to read these. I think a classic plot with average duration would be easier to follow for most readers. I understand that in some cases, the authors see no invagination, which is also reported in this graph, but I think 2 plots would do a better job than a single one in this case. Or perhaps, explain better how to read those plots, as they are uncommon, and at the basis of the authors' argumentation.

The “speed vs time” plots allow us to determine whether endocytosis completes normally in different conditions. At a given time, the speed corresponds to the average movement of the endocytic structure in the plane of the membrane (XY plane) between two consecutive images. For an endocytic event that completes normally, the speed is close to 0 in the XY plane while the endocytic pit elongates along the Z axis before the vesicle is pinched off. Note that the speed is not exactly 0 because of localization errors and putative small movements of the endocytic structure in the plane of the membrane. The speed increases after the vesicle is pinched off and it diffuses freely in the cytoplasm. Since this movement is mostly diffusive, the standard deviations of the speeds are large. To clarify the text, we have added an explanation on how to read these figures in the Materials and methods.

Note that the durations of endocytic events in different conditions can be compared directly from the “number of molecule vs time” plots, but they cannot be precisely determined because fluorescence intensities are too low at the very beginning and very end of each endocytic event to be detected unambiguously, which prevents us to determine the exact time for appearance and disappearance.

– I disagree with the use of deltaP to account for a change of osmolarity. Pressure should be in Pa (or any corresponding unit), not in Moles per Liter. Either the authors use deltaC, or they measure the osmotic pressure of their solution and can use Osm in this case for pressure. Also, at concentrations about 0.5M, there is usually a substantial discrepancy between Osmolarity and Molarity, and as the authors are using Molarities above 1M, I think it would be safer to estimate the osmotic pressure of those solutions.

We have changed our nomenclature to express molarity changes as ΔC instead of ΔP.

Reviewer #2:

Lemiere and colleagues show in this manuscript that endocytosis is remarkably robust against osmotic changes in normal walled yeast cells. However, in cells without cell walls endocytosis is much more sensitive to osmotic changes, presumably due to changing membrane tension. Eisosomes are shown to be important for regulating membrane tension after osmotic shocks.

Changes in turgor pressure are also shown to transiently affect the frequency of endocytic events. Eisosomes are shown to be important for restoring the normal endocytic frequency after shocks. Finally, exocytosis is suggested to be a mechanism that restores normal membrane tension after hypotonic shocks.

The strength of the manuscript is the highly quantitative and careful experimentation and the fact that three different plasma membrane processes are considered together (endocytosis, exocytosis, eisosomes), which is surprisingly rarely done, even though they must be functionally connected.

We thank Reviewer #2 for their positive remarks about our revised manuscript.

I have some suggestions for the authors to improve their manuscript:

Lines 217-223: Does the plasma membrane area remain constant after hypotonic treatment? I.e. are the eisosomes really disassembled and not just diluted if the plasma membrane area increases?

The plasma membrane area increased after hypotonic shock, but the normalized area fraction of eisosomes (total area of eisosomes divided by total area of protoplast, in units of um2) decreased as quantified in Figure 1I. Therefore, our conclusion is that eisosomes disassembled rather than being just diluted. The disassembly of eisosomes can also be observed in the representative images in Figure 1H (compare the second and the third frame), where the drop in eisosome coverage in the middle of the protoplast is stronger than the increase in protoplast surface area.

Figure 4H: Looking at the time series images it seems like the plane of focus shifts during the experiment. At time zero, and at 1 minute, the focus is at the surface of the cell, but later it appears that the focus shifts toward the center of the cell. During the second half of the time series the remaining eisosomes are seen at the periphery of the cell, which could be explained by a focus shift. Is this a representative example?

Because Figure 1H (originally Figure 4H) is a projected image, eisosomes at the periphery appear darker because fluorescence intensities are summed over several Z-slices whereas only one slice contributes to the fluorescence in the center of the projected image. From our experience with this setup, the focus is usually quite stable for at least 30 minutes.

Lines 325: The authors could clarify a bit more clearly whether they quantify the density of endocytic events, or the density of Fim1 labeled endocytic sites, which would depend on the lifetime of the Fim1 patches. In other words, is the reported value the number of endocytic vesicles formed per area?

The reported values are indeed the number of endocytic vesicles normalized to the cell length (we normalize to the cell length, not the cell area because cell area is more difficult to measure accurately due to the cell shapes). The fact that the lifetime of Fim1p may be different in different condition is accounted for by the method (we recalculate the temporal average of Fim1p at one endocytic site for each condition). To clarify this point, and because the counting method is not commonly used, we added a short description of the method in the main text.

Line 340: I don't understand why the authors say that "volume increased faster than the change in endocytic density". Aren't they both significantly changed already at time 0, and it takes longer for the volume to reach maximum change?

We clarified this sentence as follows:

“Note that the change in cell volume (Figures 3A and 3B, insets) could not exclusively account for the observed decrease in the endocytic density in wild-type cells as the relative increase in cell volumes were larger than the relative decrease in endocytic density 2 minutes after the shocks and remained large 4 minutes after while the endocytic densities recovered their pre-shock values.”

Lines 379-382: This is the only place where potential non-vesicular mechanisms are brought up. Lipid transfer, for example, via membrane contact sites is a possible mechanism for expanding the plasma membrane area. The method used cannot distinguish between such non-vesicular transport and exocytosis. Therefore, the authors should bring this caveat up also in the discussion.

We now discuss these alternative non-vesicular mechanisms also in the discussion and added the following paragraph:

“For the calculations presented above, we have assumed that the size of endocytic and exocytic vesicles remain constant in all osmotic conditions. However, further experiments will be necessary to validate or invalidate this assumption. In addition, our FM4-64 data does not allow us to distinguish between lipid addition to the plasma membrane via exocytosis or via a putative transfer of lipids by non-exocytic mechanisms (Reinisch and Prinz, 2021) and for simplicity and by lack of further evidence, we have discussed our data as an increase in the exocytosis rate. Not much is known about lipid transfer proteins at the plasma membrane and further experiments will be necessary to determine whether the activity of these proteins is enhanced by abrupt increases in membrane tension or hypotonic shocks.”

Lines 370-382:

I think that the exocytosis assay is not totally intuitive. It might be useful for the readers if the authors tried clarify the explanation a bit. In other words: how is it measuring exocytosis and not endocytosis, which FM4-64 is usually used to measure?

We have modified the main text and provided references where similar approaches were used to better explain our assay. The paragraph now reads:

“To measure the rate of exocytosis in different conditions, we used the cell impermeable styryl dye FM4-64, whose fluorescence dramatically increases when it binds to membranes (Cochilla et al., 1999; Gachet and Hyams, 2005; Richards et al., 2000). After FM4-64 is introduced to the media, the plasma membrane is rapidly stained (Figure 4A). Fusion of unstained intracellular vesicles to the plasma membrane results in an increase of total cell fluorescence, because after each fusion event new unstained membrane from the interior of the cell is exposed to the dye. At this stage, if one wanted to measure endocytosis rates, one would typically remove the FM4-64 dye from the media to destain the plasma membrane, and quantify the internal fluorescence which is proportional to the amount of membrane internalized by endocytosis. Here, we kept the FM4-64 dye in the media, and monitored the fluorescence of the plasma membrane and the interior of the cell. In this case, endocytic events do not increase the total cell fluorescence because they transfer patches of the plasma membrane that are already stained into the interior of the cell (Figure 4A). Therefore, the increase in fluorescence we measured is due to the addition of new unstained lipids to the plasma membrane by exocytosis.”

Line 520: How do the authors arrive at the value of 2%. Please elaborate.

To clarify our calculation, we added the following paragraph in the Materials and methods:

“We measured ~120 endocytic sites per cell at a given time on average. Since the actin meshwork assembles and disassembles in about 20 seconds, we estimate that 360 endocytic vesicles are formed per minute. Assuming endocytic vesicles have a 50-nm diameter, this corresponds to a ~2.8 μm2 endocytosed per minute, or ~2% of the total surface area, considering the average cell is around 12 μm long and has a 132 μm2 of plasma membrane (assuming the cell shape is a cylinder capped by two hemispheres).”

Line 583: What is "higher order eukaryote"? (I would also advise against using "higher eukaryote", which, although commonly used, is not a clearly defined term, nor really biologically meaningful.)

We have corrected this typo to “other eukaryotes”.

https://doi.org/10.7554/eLife.62084.sa2

Article and author information

Author details

  1. Joël Lemière

    1. Department of Molecular Biophysics and Biochemistry, Nanobiology Institute, Yale University, New Haven, United States
    2. Nanobiology Institute, Yale University, West Haven, United States
    Present address
    UCSF, Department of Cell and Tissue Biology, San Francisco, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Yuan Ren
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9017-1959
  2. Yuan Ren

    1. Department of Molecular Biophysics and Biochemistry, Nanobiology Institute, Yale University, New Haven, United States
    2. Nanobiology Institute, Yale University, West Haven, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Joël Lemière
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7155-7664
  3. Julien Berro

    1. Department of Molecular Biophysics and Biochemistry, Nanobiology Institute, Yale University, New Haven, United States
    2. Nanobiology Institute, Yale University, West Haven, United States
    3. Department of Cell Biology, Yale University School of Medicine, New Haven, United States
    Contribution
    Conceptualization, Data curation, Software, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing - original draft, Writing - review and editing
    For correspondence
    julien.berro@yale.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9560-8646

Funding

National Institute of General Medical Sciences (R01GM115636)

  • Julien Berro

American Cancer Society (IRG 58-012-58)

  • Julien Berro

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Yale West Campus Imaging core for providing access to the spinning disc confocal microscope, members of the Berro lab for insightful discussions, and R Fernandez for providing mutant yeast strains. We thank Ramesh Ramji and Kathryn Miller-Jensen for initial help with the microfluidics part of this project. We gratefully thank Millipore Sigma for lending us a CellASIC unit and Erdem Karatekin for allowing access to his micropipette aspiration setup. We also thank Samantha Dundon, Mike Lacy, and Matt Akamatsu for their comments on our manuscript. This research was supported in part by National Institutes of Health/National Institute of General Medical Sciences Grant R01GM115636 and by seed funding from the American Cancer Society Institutional Research Grant #IRG 58-012-58.

Senior Editor

  1. Vivek Malhotra, The Barcelona Institute of Science and Technology, Spain

Reviewing Editor

  1. Christophe Lamaze, Institut Curie, France

Publication history

  1. Received: August 13, 2020
  2. Accepted: May 12, 2021
  3. Accepted Manuscript published: May 13, 2021 (version 1)
  4. Version of Record published: April 27, 2022 (version 2)

Copyright

© 2021, Lemière et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Joël Lemière
  2. Yuan Ren
  3. Julien Berro
(2021)
Rapid adaptation of endocytosis, exocytosis, and eisosomes after an acute increase in membrane tension in yeast cells
eLife 10:e62084.
https://doi.org/10.7554/eLife.62084
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