Abstract

Macropinocytosis is a fundamental mechanism that allows cells to take up extracellular liquid into large vesicles. It critically depends on the formation of a ring of protrusive actin beneath the plasma membrane, which develops into the macropinocytic cup. We show that macropinocytic cups in Dictyostelium are organised around coincident intense patches of PIP3, active Ras and active Rac. These signalling patches are invariably associated with a ring of active SCAR/WAVE at their periphery, as are all examined structures based on PIP3 patches, including phagocytic cups and basal waves. Patch formation does not depend on the enclosing F-actin ring, and patches become enlarged when the RasGAP NF1 is mutated, showing that Ras plays an instructive role. New macropinocytic cups predominantly form by splitting from existing ones. We propose that cup-shaped plasma membrane structures form from self-organizing patches of active Ras/PIP3, which recruit a ring of actin nucleators to their periphery.

DOI: http://dx.doi.org/10.7554/eLife.20085.001

eLife digest

Cells can use a process known as macropinocytosis to take up fluid from their surroundings. This process plays an important role in many situations. For example, it allows human immune cells to sample their environment to search for harmful microbes and viruses and helps cancer cells to collect more nutrients so that they can grow more rapidly. During macropinocytosis, a protein called actin – which provides structural support to cells – drives the formation of cup-shaped structures from the membrane that surrounds the cell. Several signaling molecules control when and where the “cups” form, but it was not known exactly how the different types of molecules work together.

Here Veltman et al. used a technique called lattice light sheet microscopy to investigate how the macropinocytic cups form in a single-celled amoeba known as Dictyostelium. The experiments revealed that to make a cup, the actin first arranges to form a ring. The ring copies a template in the membrane, which consists of high concentrations of signaling molecules, and then extends outward to form a hollow cup by which fluid is taken up. The most important signaling molecule identified in these patches of membrane is a protein called Ras, which is mutated and hyperactive in many different types of cancer. In Dictyostelium cells that have a genetic mutation that makes Ras more active, the patches of signaling molecules and macropinocytic cups were larger than in normal cells.

The findings of Veltman et al. provide new details about how cells engulf fluids from their surroundings. The next steps will be to investigate how the signaling molecules form patches in the first place, and how they attract actin molecules. Also, more research is necessary to find out whether all cells take up fluid in a similar way or if other methods have evolved in mammalian cells.

DOI: http://dx.doi.org/10.7554/eLife.20085.002

Main text

Introduction

Macropinocytosis provides cells with an efficient way of taking up large volumes of medium into intracellular vesicles, from which they can extract nutrients, antigens and other useful molecules (Bloomfield and Kay, 2016; Egami et al., 2014; Maniak, 2001; Swanson, 2008; Swanson and Watts, 1995). It is an ancient process, used for feeding by amoebae (Hacker et al., 1997; Thilo and Vogel, 1980), but one that is important for a wide spectrum of human biology, including uptake of drugs, and large-scale sampling of extracellular medium for antigens by immune cells. It has also been hijacked by pathogens as a major route of entry (Mercer and Helenius, 2012). Recent data suggest that macropinocytosis is a principal and widely used method for sustaining the excessive metabolic demands of cancer cells (Commisso et al., 2013; Kamphorst et al., 2015) and may be implicated in the spread of neurodegenerative disease within the brain (Münch et al., 2011).

Considering its biological importance, macropinocytosis is not well understood. Macropinosomes form from cup-shaped extensions of the plasma membrane, often known as circular ruffles, which are extended by actin polymerisation. The leading rims of these ruffles must be driven outwards to enclose liquid – often for a very significant fraction of the cell’s diameter – but the base must be held static. The resulting cups can be several microns in diameter, and eventually close by constriction of their rim, with membrane fusion producing an endocytic vesicle. Here we address a critical and mysterious question about this process - how do cells organize actin to polymerize in a ring and so form the walls of the cup?

In the closely related process of phagocytosis, in which solid particles are taken up, it is proposed that cup formation is guided by engaging receptors with the particle to be engulfed in a zippering process (Freeman and Grinstein, 2014; Griffin et al., 1975). However, macropinosomes take up fluid and so cannot use a particle as a template in this way. Nor is there any known equivalent in macropinocytosis of the coat that organises clathrin-mediated endocytosis. Thus it appears that macropinocytic cups must form by a self-organizing process within the actin polymerization machinery and its regulators.

The dynamic actin that polymerises around macropinocytic cups is probably initiated by a number of nucleators, including both formins, such as ForG, which is needed for the basal part of phagocytic cups in Dictyostelium (Junemann et al., 2016), and the Arp2/3 complex (Insall et al., 2001), which produces dendritic structures (Pollard and Borisy, 2003), like the actin that drives pseudopods. Assembly of Arp2/3 based actin is controlled by the WASP family of nucleation promoting factors; the two family members that act at the plasma membrane are WASP and SCAR/WAVE (hereafter called SCAR). WASP is important for actin polymerisation during clathrin-mediated endocytosis (Taylor et al., 2011), and SCAR, acting in a five-membered complex (Eden et al., 2002), for the formation of pseudopods (Seastone et al., 2001; Veltman et al., 2012). It is not known which is responsible for macropinocytosis.

Ras and phosphoinositide signalling help organize the cytoskeleton for macropinocytosis and phagocytosis (Bar-Sagi and Feramisco, 1986; Bloomfield and Kay, 2016; Bohdanowicz and Grinstein, 2013; Rodriguez-Viciana et al., 1997; Swanson, 2014). There is evidence that Ras activity stimulates macropinocytosis in both mammalian and Dictyostelium cells, and macropinocytic cups are associated with an intense domain, or ‘patch’, of PIP3 (Araki et al., 2007; Parent et al., 1998; Yoshida et al., 2009), which is essential for their function (Araki et al., 1996; Buczynski et al., 1997; Hoeller et al., 2013; Zhou et al., 1998).

In macrophages, which often evolve macropinocytic cups from linear ruffles, it has been suggested that ruffle circularisation creates a diffusion barrier in the membrane leading to intensified PIP3 signalling and a domain of PIP3 in the centre of the circular ruffle (Welliver et al., 2011). This domain then drives the further progression of the macropinocytic cup.

In axenic strains of Dictyostelium, which grow efficiently in liquid medium, macropinocytosis is massively up-regulated due to mutation of the RasGAP neurofibromatosis-1 (NF1) (Bloomfield et al., 2015; Hacker et al., 1997; Kayman and Clarke, 1983). These strains are thus an excellent starting point for research into the organising principles behind macropinocytic cup formation. Here we examine macropinocytosis with unprecedented 3D detail using lattice light sheet microscopy, and map the spatial and temporal control of actin regulators such as SCAR and WASP with respect to signalling molecules including PIP3 and active Ras. This leads us to propose a new and general hypothesis for the formation of cups from the plasma membrane.

Results

The origins of macropinosomes in axenic strains of Dictyostelium

To determine whether macropinosomes form in Dictyostelium by circularization of linear ruffles, as reported for macrophages (Welliver and Swanson, 2012), we used lattice light sheet microscopy (Chen et al., 2014), which allows unparalleled high-resolution imaging of light-sensitive and dynamic cells over prolonged periods. In axenic cells expressing an F-actin reporter, three types of large F-actin structure are routinely detected: macropinocytic cups, which predominate, pseudopods and basal waves.

3D movies show that the majority of macropinocytic cups initiate by splitting of existing ones (62%; n = 152, Figure 1E). Splitting occurs by a variety of routes, including: simple division in the middle; detachment of a small ruffle that grows into a new macropinocytic cup (Figure 1A, Video 1); and abortive fragmentation of a parental macropinocytic cup into multiple daughter cups. We examined a reporter for active Rac in some of these movies and found that it is spatiotemporally associated tightly with F-actin in this morphological process (Figure 1B, Video 2).

Figure 1.
Download figureOpen in new tabFigure 1. Life histories of macropinocytosis, pseudopods and basal waves.

Vegetative axenic cells of strain Ax2 were followed in 3D time-lapse movies made by lattice light sheet microscopy. Images show maximum intensity projections. Cells express a marker for F-actin (LimEΔcoil-RFP) unless otherwise indicated. Numbers indicate time in seconds. (A) A new macropinocytic cup formed by splitting (arrow marks nascent daughter macropinocytic cup). (B) Close-up of a large macropinocytic cup, viewed en-face, that fragments and forms multiple smaller macropinocytic cups (arrowed). (C) Origin of a cluster of macropinocytic cups from a small basal F-actin structure (arrow). (D) Detail of a cell initiating a de novo dorsal macropinocytic cup (ie not in contact with substratum). (E) Table of macropinocytic cup origins. (F) Close-up of macropinocytic cup closure, viewed en face. (G) Growth of a pseudopod from a small F-actin structure close to the substratum (arrow). (H) Growth of a basal F-actin wave from a small F-actin punctum (arrow), to eventually encompass the entire basal surface of the cell. See also supplementary movie 1-7 for full movies of panel A, B, C, D, F, G and H respectively.

DOI: http://dx.doi.org/10.7554/eLife.20085.003

Video 1. Macropinosomes are generated by splitting.

A vegetative Ax2 cell expressing an F-actin marker imaged using lattice light sheet microscopy and viewed from three perpendicular angles.

DOI: http://dx.doi.org/10.7554/eLife.20085.004

Video 2. Macropinocytic cups are generated by splitting.

Detail of a vegetative Ax2 cell expressing markers for active Rac and F-actin, imaged using lattice light sheet microscopy and viewed from three perpendicular angles.

DOI: http://dx.doi.org/10.7554/eLife.20085.005

The remaining macropinocytic cups form de novo, expanding from places where no previous F-actin activity was detected. In more than 90% of cases the initiation is close to the base of the cell, even though most mature macropinocytic cups are present on the top, and are commonly described as 'crowns'. In the example illustrated in Figure 1C (Video 3), the parental ruffle first emerges close to the substratum (t = 0, white arrow) and cannot unequivocally be classified as either pseudopod or circular ruffle. The ruffle then quickly sweeps to the top of the cell, during which time it grows in size and splits several times, to produce multiple full-grown macropinocytic cups. As with splitting macropinocytic cups, the F-actin in de novo macropinosome cups is closely associated with signalling molecules, as illustrated by active Ras in Figure 1D (Video 4) and discussed later. Circular ruffles can persist on the cell surface for prolonged periods before either closing successfully or regressing back into the cell body. Closure of the cup can be quite abrupt and often appears to involve the inward collapse of the rim (Figure 1F, Video 5) (Swanson et al., 1999).

Video 3. Macropinocytic ruffles often initiate on the substratum.

A vegetative Ax2 cell expressing an F-actin marker imaged using lattice light sheet microscopy and viewed from three perpendicular angles.

DOI: http://dx.doi.org/10.7554/eLife.20085.006

Video 4. A small fraction of de novo macropinocytic cups are initiated off- substratum.

Detail of a vegetative Ax2 cell expressing markers for active Ras and F-actin, imaged using lattice light sheet microscopy and viewed from three perpendicular angles.

DOI: http://dx.doi.org/10.7554/eLife.20085.007

Video 5. Closure of a macropinocytic cup.

A vegetative Ax2 cell expressing an F-actin marker imaged using lattice light sheet microscopy and viewed from two perpendicular angles.

DOI: http://dx.doi.org/10.7554/eLife.20085.008

The other large F-actin projections in growing Dictyostelium cells are pseudopods. These are distinguished from macropinocytic circular ruffles by their shape, which is convex instead of concave. De novo pseudopods also initiate close to the substratum (Figure 1G, white arrow and Video 6) and expand steadily to their full size. Pseudopods are surprisingly rare in growing axenic cells, accounting for less than 5% of all large F-actin structures.

Video 6. Pseudopods are distinct from macropinocytic ruffles.

A vegetative Ax2 cell expressing an F-actin marker imaged using lattice light sheet microscopy and viewed from three perpendicular angles. A pseudopod is initiated on the right hand side of the cell at t = 1 min.

DOI: http://dx.doi.org/10.7554/eLife.20085.009

Finally, we could follow the enigmatic actin waves that move across the basal surface of vegetative cells (Bretschneider et al., 2004, 2009; Gerisch, 2010). These waves also generally originate from existing ruffles by splitting. When the splitting ruffle in Figure 1H (Video 7) contacts the substratum, it initiates an actin wave that spreads across the entire footprint of the cell, becoming so dominant that other large F-actin structures are suppressed and flattening the cell into a smooth bell-shape.

Video 7. Basal F-actin wave that originates by splitting from a nascent macropinosome.

A vegetative Ax2 cell expressing an F-actin marker imaged using lattice light sheet microscopy and viewed from three perpendicular angles. Image jitter in this movie was due to technical issues with the microscope's Z-drive).

DOI: http://dx.doi.org/10.7554/eLife.20085.010

These observations show that the large macropinocytic cups of axenic Dictyostelium cells generally form by splitting or by expanding de novo from a small focus, as in fibroblasts (Bernitt et al., 2015), rather than by circularization of linear ruffles (Welliver and Swanson, 2012). The smaller macropinocytic cups of wild-type cells (see later) also more normally form de novo or by splitting, rather than by circularization.

PIP3, Ras and SCAR are required for normal fluid phase uptake

We tested the involvement of PIP3 and Ras signalling in macropinocytosis using an isogenic set of mutants in which we measured both fluid uptake and growth in liquid medium (Supplementary material, Table 1). Either increased or decreased PIP3 levels (PTEN and PI3-kinase mutants) are deleterious to fluid uptake and growth in liquid medium, as expected from earlier work (Clark et al., 2014; Hoeller and Kay, 2007). Two independent RasG- mutants are substantially impaired in growth in liquid medium, as previously described, but contrary to the earlier report (Khosla et al., 2000), both are also defective in fluid uptake. Compensation by other Ras proteins and genetic background differences may account for the discrepancy (Bloomfield et al., 2008; Bolourani et al., 2010). RasC null cells have no growth defect and a lesser defect in fluid uptake, while RasS (not tested here) may also contribute to macropinocytosis (Chubb et al., 2000). Notably, we confirm that the Arp2/3 activator, SCAR, is required for efficient fluid uptake (Seastone et al., 2001).

Table 1.

Growth and fluid uptake by mutants.

DOI: http://dx.doi.org/10.7554/eLife.20085.011

Strain

Mutated protein

Genotype

MGT±SEM (hr)

Fluid uptake ± SEM (nl/10^six cells/h)

Ax2

parental

9.26 ± 0.26 (26)

114.7 ± 10.8 (9)

HM1505

RasC

rasC-

9.27 ± 0.26 (7)

84.3 ± 6.3 (3)

HM1497

RasG

rasG-

27.6 ± 2.9 (7)

55.3 ± 8.4 (3)

HM1514

RasG

rasG-

19.1 ± 0.24 (3)

67.6 ± 1.9 (3)

HM1200

PI3K1-5

pikA-, pikB-, pikC-, pikF-, pikG-

103 ± 1.1 (3)*

9.0 ± 0.6 (3)

HM1289

PTEN

ptenA-

19.0 ± 1.4 (9)

12.4 ± 0.6 (3)

HM1809

SCAR

scrA-

27.8 ± 2.4 (6)

56.8 ± 9.7 (3)

HM1818

SCAR

scrA-

38.4 ± 4.6 (10)

19.7 ± 5.0 (3)

  • * taken from Hoeller et al. (2013).

A ring of active SCAR forms around PIP3 domains at the rim of macropinocytic cups

The SCAR complex is mostly cytosolic and basally inactive, but when recruited to the plasma membrane it causes actin polymerization through the Arp2/3 complex (Steffen et al., 2004; Ura et al., 2012). A GFP reporter tagged at the HSPC300 subunit accumulates at sites of actin polymerization (Veltman et al., 2012). To confirm that this accumulation signifies the presence of activated SCAR complex, we correlated the signal with the expansion of pseudopods, using this as a proxy for actin polymerization. The results clearly show that the reporter is recruited during expansion phases but lost in stalls (Figure 2A,B). This correlation holds true globally: SCAR reporter intensity along the membrane correlates well with the local membrane expansion speed (Figure 2C) as all pixels with high SCAR reporter are associated with positive instantaneous membrane speed. Note that the small set of pixels with very high membrane speeds and no SCAR (Figure 2C, red arrow) are due to blebs (Zatulovskiy et al., 2014).

Figure 2.
Download figureOpen in new tabFigure 2. Macropinocytic cups contain a central domain of PIP3 surrounded by a ring of SCAR.

(A, B, C) Evidence that fluorescently tagged SCAR complex faithfully marks regions of active actin polymerisation: (A) An aggregation-competent cell moving under an agarose overlay (optimal conditions for visualising pseudopods) showing HSPC300-GFP recruited to the two pseudopods; (B) Kymograph of expansion of the pseudopod arrowed in (A) showing that the SCAR reporter is present during periods of expansion, but absent in the plateaus when the pseudopod is not expanding; (C) Membrane speed and SCAR complex accumulation are positively correlated. The HSPC300-GFP signal and local membrane speed was measured at 100 points along the membrane of a motile cell. Data of eight independent cells was combined and plotted as a 2D histogram. Green arrow indicates data points with high levels of SCAR and positive displacement. Red arrow indicates data points due to blebs, which are actin-free and expand much faster than pseudopods (Zatulovskiy et al., 2014). (D) SCAR is recruited as a ring to the lip of macropinocytic cups. The upper panels show top and side views of a surface rendering of a cell with three macropinocytic cups and the lower shows the same cell with a SCAR reporter. (E) SCAR is recruited to pseudopods in distinct blocks, not as a ring. Pitted appearance of the 3D surface is a rendering artefact caused by small vesicles that reside just underneath the cell membrane. (F) SCAR is recruited to the edge of an intense PIP3 patch in the macropinocytic cup. The white dotted line in the left panel corresponds to the position of the vertical plane in the right panel. (G) 3D reconstruction of the cell in the previous panel. (H) F-actin is nearly uniformly distributed in the macropinocytic cup and does not predict the localization of SCAR. Ax2 cells were used in all panels. HSPC300 was used as a marker for the SCAR complex, PH-CRAC as a reporter for PI(3,4,5)P3 and Lifeact as a reporter for F-actin. 3D images were reconstructed from Z-stacks taken on a spinning disk microscope.

DOI: http://dx.doi.org/10.7554/eLife.20085.012

In images recorded in 3D, the reporter reveals a thin, sometimes broken ring of active SCAR around the lip of macropinocytic cups (Figure 2D and Video 8). The presence of a ring could not be predicted by imaging the actin cytoskeleton itself, as actin filaments are distributed rather uniformly throughout the cup (Figure 2H). These circular SCAR structures are not seen in pseudopods, where 3D images show the same discrete blocks of SCAR as in 2D images (Figure 2E).

Video 8. The rim of macropinocytic cups is traced by a thin line of SCAR.

Shown is a vegetative Ax2 cell expressing the SCAR marker HSPC300. A Z-stack was collected using a spinning disk microscope and deconvolved using a calculated point spread function. Left panel shows a surface render of the cell outline and the right panel shows a 3D reconstruction of the fluorescence signal using maximum intensity projection.

DOI: http://dx.doi.org/10.7554/eLife.20085.013

The discovery of these remarkable rings immediately raises the question of how individual SCAR molecules are coordinated to maintain the ring shape. We visualised PIP3 using a double reporter that co-expresses the PH-domain of CRAC fused to RFP (Insall et al., 1994). This revealed a second remarkable feature of SCAR rings: they follow the edges of intensely stained domains, or ‘patches’ (Postma et al., 2004) of PIP3. In all macropinocytic cups examined, of whatever size, the concave cup contains a patch of PIP3 and SCAR is present as a ring around this patch, without detectable recruitment to its centre (Figure 2F and G and Video 9).

Video 9. Macropinocytic cups are defined by a patch of PIP3 that is circumscribed by a thin line of SCAR.

Shown is a vegetative Ax2 cell expressing the SCAR marker HSPC300-GFP and the PIP3 marker PH-CRAC-RFP. A Z-stack was collected using a spinning disk microscope and deconvolved using a calculated point spread function. Image shown is a 3D reconstruction of the fluorescence signal using maximum intensity projection.

DOI: http://dx.doi.org/10.7554/eLife.20085.014

This was confirmed in a larger sample by measuring fluorescence intensity at the centre and rim of 17 macropinocytic cups from nine cells rendered in 3D (Figure 3A–B). All cup centres contained high levels of PIP3, but SCAR consistently followed the rim of the cup with the mean fluorescence of the SCAR reporter significantly higher than cytosolic background (p<0.01), while signal at the centre of the PIP3 patch was not statistically different from the cytosolic background. This is also clear in 2D images, but is easily overlooked as the narrow SCAR ring appears only as tiny puncta in the cross sections obtained from confocal microscopy.

Figure 3.
Download figureOpen in new tabFigure 3. SCAR is present at the edge but not the centre of the macropinocytic cup.

(A, B) Quantification of PIP3 and SCAR in macropinocytic cups identified visually. (A) Method of analysis: Macropinocytic cups were identified morphologically from 3D rendered images and the position of their center (blue arrow) and edge (orange arrows) was noted. Fluorescence intensity along the cell membrane of the boxed macropinocytic cup was plotted and the intensity was measured at the marked center and edge. (B) PIP3 and SCAR fluorescence intensity at center and edge of 17 visually identified macropinocytic cups. Bar indicates the mean, box indicates the second and third quartile. Whiskers indicate the range of the data. (C, D) Analysis of SCAR in PIP3 patches. (C) Method of PIP3 patch analysis: Membrane-bound fluorescence intensity of the respective markers was measured for individual cells. PIP3 patches were defined as those membrane regions where the fluorescence intensity is greater than mean cytosol plus one standard deviation (dotted line) and these regions are marked in grey. (D) Quantification of fluorescence intensity at the centers and edges of 31 identified PIP3 patches. Bar indicates the mean, box indicates the second and third quartile. Whiskers indicate the range of the data. (E) Anti-correlation between SCAR and PIP3 reporter intensity. In this analysis the intensity of fluorescence in all membrane pixels was compared, irrespective of the morphological structure in which they lay. The plot shows the combined data from 16 cells. Vegetative Ax2 cells expressing the SCAR complex reporter HSPC300-GFP and the PI(3,4,5)P3 reporter PH-CRAC-mRFP were used in all experiments. The asterisk marks significant differences (p<0.01).

DOI: http://dx.doi.org/10.7554/eLife.20085.015

We further tested the spatial relation between PIP3 and SCAR in two ways not requiring visual recognition of macropinocytic cups. In the first, a number of growing axenic cells was analysed as follows. Membrane areas with fluorescence intensity of the PIP3 reporter greater than cytosolic background plus one standard deviation were defined as PIP3 patches, and the associated SCAR signal was measured. In all cases SCAR is consistently and significantly enriched at patch edges (p<0.01), and never at their centres (Figure 3C–D). In the second test, the fluorescence intensity of the SCAR and PIP3 reporters was extracted from the circumference of a number of growing cells and the results plotted as a 2D histogram (Figure 3E). In pixels with high PIP3 signal, the SCAR signal is low, and conversely in pixels with high SCAR, PIP3 is low. This method cannot show whether high SCAR and PIP3 pixels are adjacent, but it does confirm that SCAR and PIP3 do not co-localise but instead are anti-correlated.

SCAR is associated to the periphery of PIP3 patches throughout macropinocytic cup lifetime

The complete lifetime of a de novo macropinocytic cup is shown in Figure 4A (Video 10 shows another example). The PIP3 patch first becomes visible at t = 1 and this sub-micron sized patch is already flanked by puncta of SCAR. As the patch of PIP3 grows the SCAR puncta remain dynamically associated with its edge right up to closure of the macropinocytic cup, after which the SCAR signal quickly disappears and the vesicle is internalised. The SCAR is not detected at the centres of the patches above background.

Figure 4.
Download figureOpen in new tabFigure 4. A SCAR ring encircles PIP3 patches throughout their lifetime.

(A) Detail of a cell initiating a de novo macropinocytic cup. Puncta of SCAR flank the PIP3 patch at all times. Time is indicated in seconds. (B) Kymograph of the fluorescence intensity along the membrane of a vegetative cell completing several macropinocytosis events. SCAR is associated with the edge of PIP3 patches during their entire lifetime and never with their centers. (C) Quantification of peripheral SCAR signal during macropinocytosis. A line was drawn through the edge of the macropinocytic cup and extending into the cytosol for each frame of a time lapse of the lifetime of a complete macropinocytosis event. Line plots were concatenated resulting in kymographs as shown in panel (D). Orange dotted line indicates the closure event. Numbers indicate independent macropinocytosis events. (E) The SCAR signal at the edge of macropinocytic cups is present from start to finish. The fluorescence intensity at the macropinocytic cup edge was quantified and averaged for all six analysed macropinocytic cups. Vegetative Ax2 cells expressing the SCAR complex reporter HSPC300-GFP and the PI(3,4,5)P3 reporter PH-CRAC-mRFP were used in all experiments. Images are representative of typical macropinocytosis events.

DOI: http://dx.doi.org/10.7554/eLife.20085.016

Video 10. SCAR remains dynamically associated to the edge of PIP3 patches throughout macropinocytosis.

Vegetative Ax2 cell expressing the SCAR marker HSPC300-GFP and the PIP3 marker PH-CRAC-RFP. Images were taken on a spinning disk microscope.

DOI: http://dx.doi.org/10.7554/eLife.20085.017

This is also shown in a kymograph of the membrane pixels of a single cell as it makes several macropinosomes (Figure 4B). The SCAR signal, though sometimes weak, can be traced along the edge of the PIP3 patches from the start of a patch to its abrupt loss at invagination. Combining the data from several macropinocytosis events confirms this continuous association (Figure 4C–E). Thus despite changes in size and shape of PIP3 patches, SCAR remains associated with their edges, and only their edges, throughout the macropinocytic cup lifetime.

All PIP3 patches, whatever their origin, recruit SCAR to their periphery

It seemed possible that as a rule of cytoskeletal organization in Dictyostelium, PIP3 patches always recruit SCAR to their edges. Four other examples of PIP3 patches support this: two from growing cells, and two from starved cells, which are highly migratory, chemotactically sensitive, and morphologically very different from growing cells:

During phagocytosis Dictyostelium cells make a PIP3 patch where they contact the particle to be ingested (Clarke et al., 2010; Marshall et al., 2001). In the yeast case shown in Figure 5A, SCAR is recruited to the edges of the PIP3 patch, and not the centre, while a 3D view reveals a full ring of SCAR around the rim of the phagocytic cup. Indeed, a clear ring of SCAR could be detected in all such cases, provided expression of the SCAR reporter was low enough to avoid excessive background fluorescence.

Basal waves have a core of PIP3 surrounded by a ring of F-actin (Bretschneider et al., 2004, 2009; Gerhardt et al., 2014; Gerisch, 2010), and again, the basal PIP3 patches are invariably surrounded by a ring of SCAR (Figure 5B). Basal waves are favourable for microscopy, and we found that WASP is also excluded from PIP3 patches and forms a ring around them, though weaker and less coherently than SCAR (Figure 5C). Similarly, WASP forms rings at the edges of PIP3 patches of normal macropinocytic cups, again more weakly than SCAR (Figure 5D). The remaining Arp2/3 activator in Dictyostelium, WASH, does not associate with PIP3 patches (Figure 5—figure supplement 1).

During chemotactic aggregation, developing cells form small chains and streams with strong head-to-tail adhesions between them and PIP3 patches in their front (Dormann et al., 2002). These patches are invariably surrounded by a ring of SCAR. In the example shown in Figure 5E–G, a cell strongly expressing reporters is situated between two poorly expressing cells. The strongly expressing cell forms a PIP3 contact patch, with SCAR present as a clear ring and excluded from the centre.

Cells respond to cyclic-AMP by making PIP3, initially homogenously and then, after about a minute, in patches at the membrane (Postma et al., 2004). In the low light conditions required for time-lapse imaging, the SCAR signal is weak, but where detected, it is clearly at the edges of the PIP3 patches (Figure 5H–J and Video 11). These patches have sometimes been regarded as new pseudopods (for example [Chen et al., 2003]), but many become concave and close to engulf a drop of medium, indicating that the cell is performing macropinocytosis, not a chemotactic response.

Video 11. Chemoattractant-stimulated PIP3 patches recruit SCAR to their edges, not to their centres.

A developed Ax2 cell expressing the PIP3 marker PH-CRAC-RFP and the SCAR marker HSPC300-GFP was stimulated at t = 0 with 1 μm cyclic AMP. Images were collected using a spinning disk microscope.

DOI: http://dx.doi.org/10.7554/eLife.20085.020

PIP3 patches are based on active Ras but do not require F-actin ruffles

PIP3 is largely made by Ras-activated PI3-kinases (Clark et al., 2014; Funamoto et al., 2002; Hoeller and Kay, 2007). We confirmed that a patch of activated Ras exactly coincides with each PIP3 patch (Figure 6A) (Sasaki et al., 2004; 2007). Similarly, plots of intensity, pixel-by-pixel, show exceptional correlation between the Ras and PIP3 signals (Figure 6B). Thus PIP3 patches have a matching patch of activated Ras, which could sustain them by activating PI3-kinase.

Figure 6.
Download figureOpen in new tabFigure 6. PIP3 patches are supported by coincident patches of activated Ras, which can recruit weaker SCAR rings.

(A, B) PIP3 and activated Ras domains are essentially coincident in vegetative Ax2 cells: (A) dorsal patches (macropinocytic cups) and basal patches. Left and right panels show different cells; (B) 2-D histogram showing strong correlation between PIP3 and activated Ras reporters. Fluorescence intensity values of the active Ras reporter and PIP3 reporter along the perimeter of 35 cells are plotted against each other. (C) Macropinocytic signal patches additionally coincide with patches of active Rac. Shown is a representative image of a macropinocytic cup in a vegetative Ax2 cell co-expressing a marker for active Ras (Raf1-RBD) and active Rac (pakB-CRIB). (D, E) mutant lacking all Ras-activated PI3-kinases (strain HM1200) still forms Ras patches, both off the substratum (D) and basally- see (E). (E, F) Basal patches of the PI3-kinase mutant recruit SCAR to their periphery, though less strongly than the wild-type, Ax2 (TIRF images). Error bars indicate the standard deviation. Thus, loss of PI3K signalling does not allow SCAR to trespass on the Ras patch. (G, H) Ras patches (indicated by white arrow) remain discrete, despite globally high levels of PIP3 in PTEN-null cells (strain HM1289) and these domains still invariably recruit a complete SCAR ring to their edges.

DOI: http://dx.doi.org/10.7554/eLife.20085.021

Similarly, the Ras/PIP3 patch overlaps a patch of active Rac1, as detected by the CRIB domain (Figure 6C) (Manser et al., 1994). Rac1 is an upstream regulator of SCAR, and has been implicated in macropinocytosis (Dumontier et al., 2000; Palmieri et al., 2000). However, its broad distribution cannot simply account for the much narrower SCAR ring. Alternatively, Rac1 may define a permissive area where SCAR can be activated or other Rac isoforms may be involved, such as RacB, RacC or RacG (Lee et al., 2003; Seastone et al., 1998). No specific markers exist for their activated state, but the RacG molecule itself is modestly enriched at the rim of phagocytic cups (Somesh et al., 2006).

It has been proposed that PIP3 patch formation requires a positive feedback loop where PIP3 activates Ras (Sasaki et al., 2007). We tested this by genetically manipulating PIP3 levels (Clark et al., 2014; Hoeller and Kay, 2007). A mutant without Ras-activated PI3-kinases and producing only 10% of wild-type PIP3 levels still forms patches of activated Ras at a similar frequency to parental cells (Figure 6D). The SCAR signal in confocal cross sections of macropinocytic cups is too small for an accurate comparison of SCAR ring formation between mutants and therefore we used basal waves as a proxy for macropinocytic cups. Ras patches on the basal surface of PI3-kinase null cells still exclude SCAR from their centre and recruit a peripheral ring of SCAR as normal, albeit more weakly than in parental cells (Figure 6E–F). Conversely, when PIP3 levels are increased 10-fold by eliminating the PTEN phosphatase, the activated Ras domains do not expand correspondingly (Figure 6G) and remain associated with rings of SCAR (Figure 6H). Thus Ras, rather than PIP3, is the primary determinant of patches and SCAR rings.

It has also been proposed that PIP3 patch formation requires an enclosing circular ruffle to act as a diffusion trap (Welliver and Swanson, 2012). We tested this by controlled use of the actin inhibitor latrunculin-A to inhibit ruffle formation. Latrunculin-A at 1 µM leaves some actin polymerisation intact, and at 5 µM abolishes all visible actin filaments, resulting in spherical cells (Figure 7A). Neither treatment abolishes the patches of PIP3, which become larger but less numerous, with a fluorescence intensity not significantly different from control cells (Figure 7B–E). We tested whether the sharp boundaries of patches are affected by latrunculin-A by measuring the intensity across the edges of more than 30 patches for each condition, (Figure 7F–H). It is clear that latrunculin-A has little effect on the sharpness of the patch, suggesting that a diffusion barrier is not required to maintain its strong spatial coherence.

In summary, a circular ruffle is not essential to create signalling patches, which appear to largely depend on Ras, with PIP3 playing a secondary though still important role.

The intensity of Ras signalling controls patch and macropinocytic cup size

To test whether Ras plays an instructive role in macropinocytic cup morphogenesis, directly regulating their formation and size rather than acting as a remote trigger or passive participant, we examined the effect of genetically increasing Ras activity. The RasGAP NF1, encoded by the Dictyostelium axeB gene, is present in the wild-isolate NC4 but inactivated in its axenic derivatives, including the standard Ax2 used here. We found that macropinocytic cups in NC4 maintain exactly the same organization as in Ax2, with a central patch of PIP3 surround by a ring of SCAR, but are much smaller and shorter-lived and often arise de novo (Figure 7I,J and Videos 12, 13 and 14). To confirm that macropinocytic cup size is controlled by NF1 we compared an isogenic NF1 knock-out with its parent (DdB; also derived from NC4; [Bloomfield et al., 2008]). Cells from each strain were cultivated for 48 hr in axenic medium to maximally induce the rate of macropinocytosis. Under these conditions the axeB-null cells that have lost NF1 make significantly larger macropinocytic patches compared to cells from the parental strain (p<0.01, Figure 7K–L).

Video 12. Large circular ruffles are absent in vegetative wild-type NC4 cells.

Shown is a maximum intensity projection of the fluorescence intensity of the F-actin marker LimEΔcoil. Images were taken on a lattice light sheet microscope.

DOI: http://dx.doi.org/10.7554/eLife.20085.024

Video 13. The morphology of vegetative cells from axenic strains is dominated by large circular ruffles.

Shown is a maximum intensity projection of the fluorescence intensity of the F-actin marker LimEΔcoil (Jitter in this movie was due to technical issues with the microscope's Z-drive). Images were taken on a lattice light sheet microscope.

DOI: http://dx.doi.org/10.7554/eLife.20085.025

Video 14. SCAR follows PIP3 patches in macropinocytic cups of wild-type NC4 cells.

Detail of a vegetative cell from the wild strain NC4 expressing a marker for PIP3 and a marker for the SCAR complex. Macropinocytosis is rapid and small but the cups still show SCAR puncta on their edges. Images were taken on a spinning disk microscope. It should be noted that due to the small size of the macropinocytic cups, the top and bottom of the cup are frequently in the focal plane, resulting in an overlap in both signals.

DOI: http://dx.doi.org/10.7554/eLife.20085.026

The effect of the loss of NF1 on basal PIP3 patches (basal actin waves) is equally striking. Basal PIP3 patches are prevalent in axenic laboratory strains, especially during early starvation, but absent from all wild-type strains tested (Figure 7—figure supplement 1 and compare Videos 15 and 16). axeB knockout cells that have lost NF1 form abundant basal PIP3 patches, but their wild-type parent does not (Figure 7—figure supplement 1D). Thus the intensity of Ras signalling governs the size and frequency of SCAR rings in macropinocytic cups and basal waves, showing that Ras must play an instructive role.

Video 15. Basal patches are dominant in axenic cells during early starvation.

Cells from the axenic strain Ax2 were washed free of nutrients and left to develop autonomously under non-nutrient buffer. Time-lapse images were taken using a confocal microscope. Z-plane was set so that the basal membrane of the cell was in focus. Shown is an overlay of the fluorescence signal of a PIP3 marker with the trans-illuminated image.

DOI: http://dx.doi.org/10.7554/eLife.20085.027

Video 16. Basal patches are absent from non-axenic wild type cells.

Cells of the wild-type strain NC4 were washed free of nutrients and left to develop autonomously under non-nutrient buffer. Time-lapse images were taken using a confocal microscope. Z-plane was set so that the basal membrane of the cell was in focus. Shown is an overlay of the fluorescence signal of a PIP3 marker with the trans-illuminated image.

DOI: http://dx.doi.org/10.7554/eLife.20085.028

We therefore propose that Ras patches, assisted by PI3-kinase and Rac, cause macropinocytic cup formation by recruiting rings of SCAR/WAVE complex to their edge.

Discussion

Macropinosomes develop from cup-shaped projections of the plasma membrane, whose walls are driven outwards by actin polymerization. They contain a central patch of activated Ras and PIP3 throughout their life and we find that in Dictyostelium, this patch is invariably associated with a ring of active SCAR at its edge. We propose that this ring of active SCAR is recruited by the signalling patch and drives a hollow ring of F-actin to extend the walls of the macropinocytic cup.

A possible alternative mechanism comes from immune cells, which make abundant linear ruffles. These occasionally fold back to form circular ruffles, which have been described as diffusion traps that can intensify signalling within them, leading to the formation of a patch of active Ras and PIP3, (Welliver et al., 2011). In this model, PIP3 patches form as a consequence of circular ruffle formation, rather than as a cause of it. Despite the evidence that sharply curved membrane areas such as those present at the leading edge of lamellipods can act as a diffusion barrier (Weisswange et al., 2005), this idea does not easily extend to Dictyostelium, where linear ruffles are much less common, and the central PIP3 patch of macropinocytic cups can still form when ruffle formation is inhibited. However it remains possible that a diffusion barrier forms by a ruffle-independent mechanism, for example by septin-like molecules (Golebiewska et al., 2011), or perhaps by cross-linking components within the patch. Further, Dictyostelium patches become larger when Ras signalling is increased by NF1 inactivation, showing that Ras plays an instructive part in their formation.

PIP3 patches are coincident with patches of activated Ras, which presumably support them by activating PI3-kinase, and also of activated Rac. Previous work suggest that patches are self-organising structures, which can form independently of input from G-protein coupled receptors (Sasaki et al., 2007) and are likely dependent on positive feedback loops between their components (Postma et al., 2003, 2004). Our results argue against an essential role for feedback from PIP3 to Ras, because activated Ras patches can form independently of type-1 PI3-kinases and are still able to recruit SCAR to their edges, albeit less efficiently than when PI3-kinases are present. Thus it appears that the kinetics that lead to patch formation must lie largely within the compass of the GEFs and GAPs activating and inactivating Ras.

We can only speculate on how SCAR is recruited to the periphery of Ras/PIP3 patches. One possibility is that SCAR and Arp2/3 are preferentially recruited by newly synthesised F-actin (Ichetovkin et al., 2002) produced by formins (Jasnin et al., 2016), which might therefore be the initial actin nucleator to be recruited. However, this seems unlikely in the light of recent work showing that ForG contributes to the base of the macropinocytic cup, but seemingly not to the extending lip (Junemann et al., 2016). Alternatively, SCAR might be moved to the periphery of Ras/PIP3 patches, perhaps by myosin-1 motors. Early work showed that myosin-1 is genetically important for macropinocytosis in Dictyostelium (Novak et al., 1995; Titus, 2000). In support of the genetic evidence, myosin-1 isoforms are recruited to macropinocytic cups in both Dictyostelium and Acanthamoeba (Brzeska et al., 2012; Ostap et al., 2003), most likely due to their affinity for PIP3 (Chen et al., 2012). The PIP3-binding MyoE and MyoF are recruited in the centre and MyoB at the periphery of macropinocytic cups, forming a striking ‘bull’s eye’ pattern (Brzeska et al., 2016; Dieckmann et al., 2010). In such a scenario, CARMIL may provide the link between myosin-1 and SCAR (Jung et al., 2001).

Our work also has implications more specific to Dictyostelium biology. First, the basal actin waves, which give a valuable window into actin dynamics (Bretschneider et al., 2004, 2009; Gerisch, 2010), appear to be formed as a consequence of the loss of NF1 in standard laboratory axenic strains. Knowing this should allow for better manipulation of these waves and for modelling to take account of their underlying need for activated Ras (Arai et al., 2010; Khamviwath et al., 2013; Sasaki et al., 2007; Taniguchi et al., 2013). Second, we consider that all patches of PIP3 and activated Ras are related by their common recruitment of SCAR to their periphery and are therefore likely to organise circular rings of actin polymerization, rather than the solid blocks characteristic of pseudopods. Therefore, the proposed role of these patches in chemotaxis, where they have been mistaken for pseudopods, needs to be re-evaluated.

In summary, our work suggests a general hypothesis for the formation of cupped actin structures: that these structures arise from a ring of actin polymerization formed by recruiting actin nucleators to the periphery, but not the centre, of self-organizing patches of intense Ras and PIP3 signalling. This hypothesis suggests many new lines of experimentation.

Materials and methods

Cell strains, cultivation and fluid uptake assay

The following Dictyostelium discoideum strains were used: Ax2 (R. Kay lab strain), NC4 (from K. Raper, obtained via P. Schaap), DdB (from M. Sussman, obtained via D. Welker), NC66.2 (from D. Francis), Ax3 (R. Chisholm laboratory strain, obtained via Stock Center) and Ax4 (W. Loomis laboratory strain, obtained via Stock Center). Axenic strains were cultured in Petri dishes under HL5 medium (Formedium, Hunstanton, UK) using standard methods. Non-axenic strains were cultivated on SM agar plates with a lawn of live Klebsiella pneumoniae and where necessary washed free of bacteria by repeated low-speed centrifugation from KK2 (20 mM KH2PO4/K2HPO4, 2 mM MgSO4, 0.1 mM CaCl2, pH 6.2) (for detailed protocols of these standard techniques, see (Kay, 1987) and dictybase.org/techniques/). Mutant strains, all in the parental Ax2 (Kay) background, are listed in Supplementary Material, Table 1.

Fluid uptake was measured using TRITC-dextran and flow cytometry. Cells were grown on bacterial lawns, washed free of bacteria, resuspended in HL5 with antibiotics, 50 µl aliquots were distributed into 96 plates and allowed to adapt for about 18 hr, until macropinocytosis was maximally up-regulated. TRITC-dextran was added to 0.5 mg/ml in HL5 to the wells and the cells incubated for various times, after which the TRITC dextran was removed, the cells washed once and uptake terminated with ice-cold, 5 mM NaN3, which also detaches the cells. Fluorescence in individual cells was then measured by flow cytometry, and the rate determined while uptake was linear with time (first 45–60 min).

DNA constructs and transfection

Single and dual expression vectors were used for all experiments (Veltman et al., 2009). Specifically, the following vectors were used: plasmid pDM1219 - expression of mCherry-LimEΔcoil (residue 1–145 of Dd LimE), pDM767 - dual expression of HSPC300-GFP and PH-CRAC-mRFPmars (residue 1–126 of Dd DagA), pDM1492 - dual expression of mCherry-RBD-Raf1 (residue 1–134 of Hs Raf1) and PH-PkgE-mCherry (residue 1–100 of Dd PkgE), pDM1383 - dual expression of HSPC300-GFP and mCherry-RBD-Raf1 and pDM1424 - dual expression of HSPC300-GFP and PH-PkgE-mCherry.

The act6 promoter that drives the resistance marker on the expression vectors is not active when cells are cultivated using bacteria as a food source. Therefore, this promoter was replaced by the coaA promoter (bp −293 to bp −1 relative to the start codon of coaA) for those vectors that were used to transfect non-axenic, wild-type cells.

Transfection of non-axenic cells was performed as follows: 5 × 106 cells were harvested from the feeding front of an SM agar plate, washed once in H40 buffer (40 mM HEPES/KOH pH 7.0, 1 mM MgCl2), and resuspended in 100 μl H40 buffer. Cells were mixed with 5 μl miniprep DNA (~0.5–1 μg total) and put on ice. Cells were then electroporated with two square waves of 350 V, 8 ms, 1 s apart using a Gene Pulser Xcell (Biorad) and immediately transferred to a Petri dish with SorMC buffer (15 mM KH2PO4, 2 mM Na2HPO4, 50 μM MgCl2, 50 μM CaCl2, pH 6.0) supplemented with live Klebsiella pneumoniae at an OD600 of 2. Selection marker was added after 5 hr (10 μg/ml G418 or 100 μg/ml hygromycin).

Image acquisition

Lattice light sheet microscopy 3D images were acquired as described (Chen et al., 2014), using a massively parallel array of coherently interfering beams comprising a non-diffracting 2D optical lattice as light sheet illumination focused by 0.65 NA objective for excitation (Special Optics). This creates a coherent structured light sheet that can be dithered to create uniform excitation in a 400 nm thick plane across the entire field of view determined by the length of the light sheet. In order to obtain the array of lattice light sheet, a binary spatial light modulator (SXGA-3DM, Forth Dimension Displays) is placed conjugate to the sample plane, and a binarized version of the desired structured pattern at the sample is projected on the display. In the time-lapse dithered mode, 3D stacks were acquired either by moving the detection objective (Nikon, CFI Apo LWD 25XW, 1.1 NA, 2 mm WD), which is synchronized with the scanning galvo mirror, or moving the sample by fast piezoelectric flexure stage (Physik Instrumente, P-621.1CD) with 100 ~150 z planes, to have about 20 µm in z axis with respect to the detection objective. Exposure time was 5 or 10 msec per plane, for a total exposure time of ~1 s for one 3D stack and a 1 s pause was added between each time point to have time series data. Raw data was deconvolved via a 3D iterative Lucy-Richardson algorithm in Matlab (The Mathworks, Natick, MA) utilizing an experimentally measured point spread function.

Spinning disk microscopy was performed on an Andor Revolution system with a Yokogawa CSU spinning disk confocal unit. The microscope was fitted with a 1.49 Plan Apo 100x oil immersion objective and an additional 1.2x magnification lens. GFP and mCherry signals were separated by a Tucam beam splitter and detected using two Andor iXon Ultra backlit EMCCD cameras with 16 µm pixel size. Z-scans were performed with the 1.5x optovar in place using 70 ms exposure per frame and a Z-spacing of 0.19 µm. Typically, 80 frames were collected from each camera in a total of 8 s.

TIRF microscopy was performed using a Nikon N-STORM system fitted with a 1.49 Plan Apo 100x oil immersion objective and the 1.5x optovar in place. GFP and mCherry fluorescence signals were recorded sequentially on an Andor iXon Ultra backlit EMCCD camera.

Confocal microscopy was performed on a Leica SP8 system using a 1.4 NA plan apo oil immersion objective and GFP/mCherry fluorescence was detected using two HyD detectors. All microscopy was performed at room temperature.

Image analysis

General image handling, such as brightness/contrast adjustments and generation of kymographs was done using ImageJ (NIH). 3D cellular fluorescence images were generated as follows. A Z-stack was recorded on a spinning disk microscope using previously indicated settings. The dataset was deconvolved with Huygens Professional software (Scientific Volume Imaging) using a calculated point spread function. The images presented are maximum intensity projections of the deconvolved dataset.

Correlation between speed and membrane fluorescence intensity was analysed using Quimp11 (www.warwick.ac.uk/quimp). Identification of membrane pixels and measuring their fluorescence intensity was done using a custom-written MATLAB (The MathWorks) script (Supplementary file 1 and Source code 1).

Image sets that were used for quantification were taken from at least two independent transfections. Only those cells with very low HSPC300-GFP expression were included for analysis, as overexpression dramatically reduces image contrast. For the quantification of SCAR fluorescence on the edge and centre of macropinocytic cups a paired 2-tailed T-test was used in Figure 3B and D and a 2-tailed T-test was used in Figure 6F.

All lattice light sheet microscopy movies (1–7 and 12–13) show a maximum intensity projection of the fluorescence intensity. The F-actin marker LimEΔcoil is used in all images unless otherwise specified. Images were deconvolved using a custom-written Richardson-Lucy algorithm. The maximum intensity projection was generated using Huygens software. Indicated time is in the min:sec format.

References

Acknowledgements

We wish to thank Sean Munro for comments on the manuscript and the Biotechnology and Biological Sciences Research Council (Grant number BB/K009699/1 to RRK and DV), Medical Research Council (reference number U105115237 to RRK) for support.

Decision letter

Joel Swanson, Reviewing editor, University of Michigan Medical School

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

[Editors’ note: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration. The first decision letter after peer review is shown below.]

Thank you for submitting your work entitled "A plasma membrane template for macropinocytic cups" for consideration by eLife. Your article has been evaluated by Vivek Malhotra (Senior Editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors. Our decision has been reached after consultation amongst the reviewers. Based on these discussions and the individual reviews pasted below, we regret to inform you that your work will not be considered further for publication in eLife.

Dr. Joel Swanson has agreed to reveal his identity as one of the peer reviewers of your paper.

All in all, the reviewers were satisfied with the images, the high quality of data and your interesting hypothesis. However, they were concerned that your conclusions were not sufficiently justified by the data especially with respect to the mechanism of ring formation and the actual function of the ring in cup formation. We feel their major concerns cannot be addressed within the usually permitted period of 2-3 months. But, given the potential importance of the findings, we would be willing to consider a new manuscript for review by the same reviewers, provided you can address the reviewers’ concerns. Otherwise, we strongly urge you to submit this work elsewhere.

Reviewer #1:

This paper investigates the mechanisms driving the formation of macropinocytic cups in Dictyostelium. Using high end imaging (lattice sheet microscopy), the authors uncover that the actin nucleation promoting factor SCAR accumulates in a thin ring at the edge of PIP3 patches. They then explore PIP3 patches in a variety of other cellular processes, such as phagocytosis and basal actin waves, and demonstrate that similar SCAR rings are present. Finally, the authors investigate the role of Ras in the formation of PIP3 patches.

The imaging is beautiful and the results do suggest a morphogenetic mechanism. However, the work remains at a rather preliminary level and does not go much beyond the observation of the SCAR rings. Many open questions remain: What is the mechanism of SCAR ring formation? The authors discuss the possible involvement of myosin 1, could they explore this experimentally?

If the SCAR ring is indeed responsible for the curving of the membrane and the formation of a circular actin ruffle, why do all the structures with such a ring not adopt a similar geometry? How does the morphogenetic mechanism suggested by the authors actually work? It would have been useful to follow the dynamics of SCAR during macropinosome formation. Is the PIP3 patch initially flat and then the membrane curves out? And if so, does this event correspond with SCAR ring formation? Or does the ring initially promote patch expansion, like in an actin wave, and if so, how does the transition to outwards polymerisation occur? Addressing any aspect of the mechanism of ring formation or of its morphogenetic effect would bring the paper beyond the level of an interesting observation.

1) The authors use the correlation between HSPC300-GFP recruitment and forward extension of pseudopods as a proof that this construct localizes active SCAR. This is not entirely clear. Is SCAR itself (in an inactive form) still present at the edge of stalled pseudopods? And if not, could HSPC300 localize together with SCAR independently of its activity?

2) If CRAC PH-domain binds both PIP3 and PIP2 as stated by the authors (subsection “SCAR is recruited to the periphery of PIP3 domains in macropinocytic cups”, third paragraph), why do they consider it a readout of PIP3 localization?

3) Some of the figures (e.g. most of the data displayed on Figure 4; Figure 5F-H) lack quantification, and while they display beautiful images, it would be helpful to quantify the readouts, or at least mention in the figure legend of how many cells and experiments the presented image is representative.

4) Figure 4 is interesting but raises several questions: why has WASP been localized only for basal actin waves? Does WASP also follow the same trend as SCAR in the other cases of PIP3 patches? In panel H, SCAR appears to form larger domains that overlap with PIP3 to some extent. How does this fit with the other observations?

5) It is not clear how the measurements displayed in panel 5E were actually done: is the background measured outside the cell? And if so, is this the right normalization? And if not, where would the background region be in pictures like those displayed in panel D? Also, the authors claim that SCAR recruitment is less strong in PI3K-null cells than in wt, however, the fluorescence at center also seems lower in this case. It would be better to plot the ratio of center to edges (both measured in the same patch), which would also avoid the use of the background signal.

6) The authors discuss the possible presence of a diffusion barrier at the edge of Ras/PIP3 domains, could they discuss what would and actin based barrier (as suggested) prevent from diffusing? How would such a barrier work?

Reviewer #2:

The manuscript by Veltman, et al., examines the dynamics of F-actin, PIP3 Ras-binding domains (RBD) and SCAR/WAVE during macropinosome formation and related movements in Dictyostelium. Using state-of-the-art fluorescence microscopy, the authors demonstrate a distinct spatial arrangement of PIP3 RBD and SCAR in forming macropinocytic cups, with PIP3 and RBD localizing to patches of plasma membrane within cups and SCAR (inferred by imaging the SCAR component HSPC300-GFP) localizing to cup rims, at the edges of ruffles that define PIP3 patches. The images provide a striking correlation between the localization of PIP3 and RBD patches and of SCAR at the edges of ruffles that delineate those patches. They show further that similar patterns can be detected in wild-type amoebas which make macropinosomes more infrequently than those which are adapted to broth culture.

My major concern is with the statements, in the Abstract and elsewhere, that "patches of intense PIP3 signalling […] recruit SCAR/WAVE complex to their edges, […] driving a ring of actin polymerization that forms the walls of the cups" (Abstract). This implies that PIP3 patches precede the SCAR localization to patch edges, which is not demonstrated by the experiments. The figures provided, especially Figure 4H, indicate that PIP3 patches form coincident with SCAR concentration at leading edges. Instead, I would interpret the images as supporting a model in which SCAR defines a boundary, perhaps a diffusion barrier, that permits amplification of PI 3-kinase activity within a circular domain in the inner leaflet of plasma membrane; that is, SCAR either prescribes the PIP3 patch or assembles coincidently into the concentric arrangement. This essential claim of the manuscript about which comes first could be addressed by experiments which measure the timing of PIP3 patch formation relative to that of SCAR localization and ring formation.

Secondly, mechanistic interpretations (i.e., PIP3 patches recruit SCAR) would require more than the correlative evidence provided here, such as localization after experimental manipulations of the system components. Some mechanism is suggested by the experiments of Figure 5, which show RBD patches forming without corresponding patches of PIP3, and which suggest that PIP3 patches do not prescribe SCAR rings. However, the mechanism implied by this result remains speculative.

Reviewer #3:

In this manuscript the authors study the organization of macropinocytic cups in Dictyostelium cells. They first characterize the dynamic structure of the macropinocytic cups in 3D using the new lattice light sheet microscopy technology. They also compare and contrast the macropinocytic structures to basal actin waves and to pseudopodia. The authors show that active SCAR protein localizes to the rim of the macropinocytic cups. The SCAR protein, and to a lesser extent WASP, follows the edge of a membrane patch enriched in PIP3 and Ras activity. The localization of SCAR to the edge of PIP3 patches is shown to be very robust as it is seen under various growth conditions and genetic backgrounds. The authors suggest that the SCAR pattern explains the formation of the cup shaped actin structures driving the formation of the macropinosomes. Finally, the authors show using mutant cells that PIP3, may contribute, but is not required for the formation of the SCAR pattern.

The finding that SCAR localizes to the rim of PIP3 patches is interesting and likely to be important for mechanistic understanding of the actin driven formation of the macropinosomes. However, the molecular mechanism that forms the SCAR pattern around the PIP3 patches remains unknown. Several hypotheses are discussed, but not tested in the manuscript. Testing these models may be beyond the scope of this manuscript.

The manuscript is clearly written, and the presented data is very beautiful, quantitative and of high technical quality. The key conclusions of the manuscript are well supported by the data.

I would, however, suggest toning down some overstated conclusions that might be misleading for some readers.

1) At the end of the subsection “All PIP3 patches, whatever their origin, recruit SCAR to their periphery” the authors state that: "[…] any process that causes patches of PIP3 will cause recruitment of SCAR at the edges […]". This is a rather exaggerated conclusion based on colocalization data, especially when the authors later show that PIP3 is not needed for SCAR localization.

2) At the end of the subsection “Macropinosomes in wild-type cells show the same as organisation as in axenic cells, but basal PIP3 waves do not form” they also conclude "[…] that PIP3 patches cause peripheral SCAR and actin rings to form […]". Again this is too strong a conclusion. No evidence is presented that PIP3 causes the SCAR localization.

3) In the first paragraph of the Discussion it says: "This work offers an explanation how the architecture of these rings is controlled." The manuscript describes the SCAR ring architecture, but doesn't really explain the mechanisms that control it.

[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]

Thank you for resubmitting your work entitled "A plasma membrane template for macropinocytic cups" for further consideration at eLife. Your revised article has been favorably evaluated by Randy Schekman (Senior editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors.

The manuscript has been improved but there are some remaining issues that need to be addressed before acceptance. No additional experimental work is needed, but the Discussion section should be revised to clarify points identified by reviewers 2 and 3.

1) "The authors should address what is known about the role of specific Racs that have been described as having a role in macropinocytosis, such as Rac1A and Rac1C and perhaps RacB, so as to remove some of the ambiguity from the presentation of the data and synthesis of the results. "

2) Known roles for RasC or RasG in macropinocytosis, if any, should be mentioned.

3) Known roles for myosin 1 isoforms in macropinocytosis should be mentioned.

4) Citations of Hoeller et al. and Seastone et al. should be included with the reference to Clark et al. 2014.

5) Additional thoughts about the nature of the diffusion barrier that remains after latrunculin treatment are optional but encouraged.

Reviewer #1:

This manuscript analyzes the spatial and temporal organization of actin PIP3, Ras, Rac and SCAR during the formation of macropinosomes in Dictyostelium. It demonstrates that SCAR localizes to the edges of plasma membrane patches of PIP3 coincident with patch formation. The PIP3 patches, which also contain active Ras and Rac, prescribe the localization of SCAR and the accompanying actin polymerization that forms the cup. Additional experiments support a model in which the size of the PIP3 patch is prescribed by active Ras, and not vice versa; this is an important mechanistic insight. An alternative hypothesis, that the PIP3 patch is prescribed by the forming actin-rich cup, is excluded by experiments showing that PIP3 patches can form in cells whose actin has been depolymerized by latrunculin A.

The revised manuscript adequately addresses the critiques from the first reviews. The additional experiments provide mechanistic analyses that were missing from the first manuscript. Revised text addresses earlier concerns about interpretation of the results.

The title of the manuscript is acceptable. Supplementary videos provide important supporting data for the claims of the paper, especially regarding the relative distributions of PIP3 and SCAR.

Reviewer #2:

Cells can internalize large volumes of liquid via macropinocytosis, a dynamic process driven by actin polymerization. Veltman et al. beautifully describe macropinocytosis in the Dictyostelium amoebae. They show that macropinocytic cups do not arise from flat membrane protrusions or pseudopods as had been proposed but rather arise de novo from the membrane of a Dictyostelium cell. The Arp2/3 regulator SCAR is tightly localized to the rim of the extending cup, surrounding a membrane patch enriched for both active Ras and PIP3, where it is maintained all throughout its formation, expansion and closure. Cells with significantly reduced levels of PIP3 continue to make macropinocytic cups, with a core patch of activated Ras bordered by SCAR, establishing that activation of small G proteins is the initial driver of cup formation. Support for this comes from the finding that true wild type strains extend significantly fewer and smaller macropinocytic cups, however upon disruption of the NF1 RasGAP (which is mutated in axenic laboratory strains) macropinocytosis is dramatically upregulated. Together with previous work in the field, the results establish that Ras/Rac activation is critical for macropinosome formation that promotes the localized production of PIP3 on the membrane and selectively recruits and/or activates SCAR at the border that, in turn activates Arp2/3 mediated actin polymerization to drive cup extension.

Activated Ras, PIP3, and SCAR are known to have significant roles in macropinocytosis, but it has not been clear which is the primarily responsible for initiating cup formation. Here the authors provide new insights into this problem by showing that Ras/Rac activation at the membrane corresponds with the recruitment of SCAR to the periphery, likely playing critical role in initiating a ring of actin polymerization surrounding the patch at the membrane. PIP3 appears to be dispensable for this initial step but must play a role in subsequent steps. The results also demonstrate that membrane patches of PIP3 are not associated with pseudopods but are rather mainly involved in the formation of macropinocytic structures, an important clarification for the field that has focused a good deal of attention to the role of PIP3 in directed migration. Naturally, these findings raise the critical question of how specific small GTPases with roles in macropinocytosis act together with PI3K to orchestrate subsequent events. Activated Ras/Rac is permissive for SCAR activation at the edges of the forming and mature cup but how the relatively large region of active Ras on the membrane causes such restrictive localization of SCAR remains an intriguing mystery.

Specific comment:

Reporters for activated Ras and Rac are localized to the macropinocytic cup and a number of small GTPases, such as RasC and RasG, are implicated in macropinocytosis, as previously shown by others and also described here. The authors should address what is known about the role of specific Racs that have been described as having a role in macropinocytosis, such as Rac1A and Rac1C and perhaps RacB, so as to remove some of the ambiguity from the presentation of the data and synthesis of the results.

Have the authors examined either the RasC or RasG null mutant to see what step in macropinocytosis either of these small GTPases affects – macropinosome formation or cup size/extension? The addition of such data could be an important first step towards clarifying the role of specific small GTPases in initiating macropinocytosis.

Reviewer #3:

This study dissects the spatial and temporal patterns of PIP3, SCAR and actin in the genesis of macropinocytic lamellae and cups with unprecedented precision. The study integrates, extends and explains previous studies on the generation, propagation and function of actin waves at the ventral surface of the cell, by showing their relationship with ruffles and cups that give rise to macropinosomes on the dorsal surface.

One of the major claims is that PI3K does not play an instructive role in the generation of cups, but instead demonstrates such a role for Ras and not for Rac.

Another extremely important discovery is that the RasGAP ortholog of NF1, which absence or lack of functionality is determinant for the capacity of Dictyostelium cells to live from fluid phase uptake, dictates the size of the cups.

Overall, this revised version of the manuscript presents very strong experimental evidence for the claims, is richly documented both by time-lapse microscopy with unprecedented 3D and temporal resolution, and exquisite reconstructions and analyses. All important results are quantified and analysed with appropriate statistical tools.

It is also worth noting that I was not among the reviewers of the original manuscript, but am fully satisfied by the answers given to all criticisms, mainly in the rich and complete additional experimental evidence as well as in the reformulation of the claims and the overall more objective tone of the text.

Therefore, I do not have strong criticisms, but only a few points for which I would encourage the authors to bring some additional clarification and discussion.

1) Every PIP3 patch has a rim of SCAR, but it is not really clear to me whether every SCAR-positive structure (dot/patch/ring) has a core of underlying PIP3. The same can potentially be asked for combinations of activated Ras or Rac.

2) The claim that the LatA experiments rule out that a diffusion barrier is required to restrict the PIP3 patch is too general. It shows that an actin-dependent diffusion barrier is not required. But one might envisage other membrane heterogeneities that could serve as diffusion barrier (Lo versus Ld phases, rafts etc.).

3) Grinstein and colleagues had shown that the phagocytic cup lip serves as diffusion barrier, possibly implying that the geometry - the curvature - was the mechanism. The findings here with LatA-treated spherical cells refute this hypothesis. This should be mentioned.

4) Maybe a slightly expanded discussion on the possible roles of class I myosins would be cool. They were implicated in the ventral actin waves; they are known to participate in the recruitment and activation of the Arp2/3 complex; the absence of MyoK induces morphological similarities with the "smooth bell-shaped cell" morphology induced by a wave spreading through the cell footprint; absence of many class I myosins result in macropinocytic but also phagocytic and trafficking phenotypes.

DOI: http://dx.doi.org/10.7554/eLife.20085.031

Author response